ADV IN MICROBIAL PHYSIOLOGY VOL 24 APL, Volume 24 (Advances in Microbial Physiology)

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Advances in

MICROBIAL PHYSIOLOGY

This Page Intentionally Left Blank

Advances in MICROBIAL PHYSIOLOGY edited by

A. H. ROSE

J. GARETH MORRIS

School of Biological Sciences Bath University England

Department of Botany and Microbiology University College of Wales Aberystwyth, Wales

D. W. TEMPEST Laboratorium voor Microbiologie Universiteit van Amsterdam The Netherlands

Volume 24 1983

ACADEMIC PRESS, INC. (Harcourt Brace lovanovich, Publishers)

London Orlando San Diego New York Toronto Montreal Sydney Tokyo

ACADEMIC PRESS INC. (LONDON) LTD. 24/28 Oval Road London NW 1 7DX United States Edition published by ACADEMIC PRESS, INC. Orlando, Florida 32887

Copyright 0 1983 by ACADEMIC PRESS INC. (LONDON) LTD.

AN Rights Reserved

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

ISBN &12-027724-7 ISSSN 0065 291 1

PRINTED I N T H E UNITED STATES OFAMERICA

85 86 87 88

9 8 7 6 5 4 3 2

Contributors J. P. van DIJKEN Laboratory of Microbiology, Delft University of Technology, P 0 Box 5 , Delft, The Netherlands A. A. GUFFANTI Mount Sinai School of Medicine of the City University of New York, New York 10029, USA W . HARDER Department of Microbiology, University of Groningen, Broerstraat 5 , Groningen, The Netherlands A. L. KOCH Department of Biology, Indiana University, Bloomington, Indiana 47405, USA T . A. KRULWICH Mount Sinai School of Medicine of the City University of New York, New York 10029, USA I. S. KULAEV Institute of Biochemistry and Physiology of Micro-organisms, Academy of Sciences of the USSR, 142292 Pushchino, Moscow Region, USSR V. M . VAGABOV Institute of Biochemistry and Physiology of Microorganisms, Academy of Sciences of the USSR, 142292 Pushchino, Moscow Region, USSR M . VEENHUIS Laboratory of Electron Microscopy, University of Groningen, Broerstraat 5, Groningen, The Netherlands J. G. ZEIKUS Department of Bacteriology, University of Wisconsin, Madison, Wisconsin 53706, USA

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Contents The Significance of Peroxisomes in One-Carbon Compounds in Yeasts J . P. VAN DIJKEN and W . HARDER I . Introduction . . . . . . . I1. Role of peroxisomes in methanol metabolism

the Metabolism of by M. VEENHUIS.

. . . .

. . . .

. . . .

. . . .

. . . . . .

A . Dissimilation of methanol . . . B. Assimilation of methanol . . . . C . Structure and function of peroxisomes . . . . . D . The molecular substructure of crystalline peroxisomes . . I11. Role of peroxisomes in methylated amine metabolism . . . . A . Methylated amines as a nitrogen source . . . . . . B. Peroxisomes and the metabolism of methylated amines . . . C . Peroxisomes involved in the concurrent metabolism of Carbon- and Nitrogen-sources . . . . . . . . . . IV . Biogenesis of peroxisomes . . . . . . . . . A . Regulation of the synthesis of peroxisomal enzymes. . . . B. Development of peroxisomes during vegetative growth . . . C . Development and function of peroxisomes during spore formation D . Assemblage of peroxisomes . . . . . . . . V. Inactivation of peroxisomal enzymes and degradation of peroxisomes . A . Regulation of enzyme activity by inactivation . . . . . B. Modification inactivation of peroxisomal enzymes . . . . C . Degradative inactivation of peroxisomal enzymes . . . . D . Subcellular events in peroxisomal degradation . . . . . VI . Concluding remarks . . . . . . . . . . VII . Acknowledgement . . . . . . . . . . References . . . . . . . . . . . .

2 5 6 13 16 24 30 30 31 37

40 41 47 53 55 58 58

60 64 69 76 76 76

Polyphosphate Metabolism in Micro-Organisms by IGOR S. KULAEV and VLADlMlR M. VAGABOV I . Introduction . . . . . . . . . . . . 83 A . Inorganic polyphosphates . . . . . . . . 84 B. Distribution in micro-organisms . . . . . . . 85 C . Methods of detection, identification and fractionation of inorganic polyphosphates . . . . . . . . . . 86

viii

CONTENTS

11. High molecular-weight polyphosphates . . . . . . . A. Intracellular localization . . . . . . . . . B. Enzymes involved in biosynthesis and degradation of polyphosphates C. Metabolism of polyphosphates in eukaryotes . . . . . D. New data on polyphosphate metabolism in prokaryotes . . . E. Concluding remarks on the physiological role of high molecular. . . . weight polyphosphates in microbial metabolism 111. Inorganic pyrophosphate: new aspects of metabolism and physiological . . . . . . . . . . . . role. A. Utilization of pyrophosphate in phosphorylation reactions in bacteria B. Energy-dependent synthesis of pyrophosphate during photosynthetic and oxidative phosphorylation. . . . . . . C. Relationship between pyrophosphate and polyphosphate metabolism in micro-organisms . . . . . . . . . . IV. Modem concepts about the role of high molecular-weight polyphosphates . . and pyrophosphate in evolution of phosphorous metabolism. V. General conclusions . . . . . . . . . . VI. Acknowledgements . . . . . . . . . . References . . . . . . . . . . . .

89 89 103 114 132 141 142 142 145 150 153 157 158 158

Physiology of Acidophilic and Alkalophilic Bacteria by TERRY A. KRULWICH and ARTHUR A. GUFFANTI I. Introduction . . . . . . . . . . . . 173 11. Acidophilic bacteria . . . . . . . . . . 175 A. Special problems of life at low pH values. . . . . . 175 B. Organisms described . . . . . . . . . 176 C. Physiological adaptations that meet the problems . . . . 178 D. Why can’t obligate acidophiles grow at neutral pH values? . . 186 111. Alkalophilic bacteria . . . . . . . . . 187 A. Special problems of life at high pH values . . . . . 187 B. Organisms described . . . . . . . . . 188 C. Physiological adaptations to meet the problems . . . . 191 D. Why can’t obligate alkalophiles grow at neutral pH values? . . 206 . . . . . . . . . 207 IV. Concluding remarks . V. Acknowledgements . . . . . . . . . 208 References . . . . . . . . . . . . 208 Metabolism of One-Carbon Compounds Anaerobes by J. G. ZEIKUS I. Introduction. . . . . . . . . . . . . . A. Definitions . B. History and scope . . . . . .

by Chemotrophic . . .

11. Transformation of one-carbon metabolites by anaerobes A. Production of one-carbon compounds . . . B. Consumption of one-carboncompounds. . .

. . . . .

.

. . . . .

.

. . . .

.

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215 216 217 218 218 221

CONTENTS

ix

111. One-carbon transformations in methanogens .

.

.

.

.

A . General physiology and species properties . . . . B. Catabolism . . . . . . . . . C . Cell carbon synthesis . . . . . . . . . D . Unification and regulation of metabolism . . . . . IV. One-carbon transformations in homo-acetogens . . . . . A . General physiology and species properties . . . . . B. One-carbon metabolism . . . . . . . . . V . General metabolic perspectives on unicarbonotrophy . . . . A . Relation of substrate-product thermodynamics to growth efficiency . B. Relation of chemotrophic anaerobes to phototrophs and aerobes . VI . Trends in the significance of one-carbon transformations . . . A . Environmental . . . . . . . . . . B. Evolutionary . . . . . . . . . . . C. Biotechnological . . . . . . . . . . VII . Acknowledgements and dedication . . . . . . . References . . . . . . . . . . .

226 227 235 243 251 257 257 264 272 273 275 276 277 282 285 288 289

The Surface Stress Theory of Microbial Morphogenesis by ARTHUR L. KOCH I . Introduction . . . . . . . . . . . . A . Caveats . . . . . . . . . . . . I1 . Methods . . . . . . . . . . . . A . Soapbubbles . . . . . . . . . . . B. Themathematics ofnarrow zonal growth . . . . . C. Diffusegrowth . . . . . . . . . . D . Problems of electron microscopy . . . . . . . E . Analysis of autoradiograms . . . . . . . . 111. Results . . . . . . . . . . . . . A . Gram-positive rods . . . . . . . . . . B . Gram-negative rods . . . . . . . . . . IV . Discussion . . . . . . . . . . . . A . What shape ought a bacterium to have? . . . . . . B. Stressonpeptidoglycancovalent bonds . . . . . . C . Surface stress theory for cylindrical elongation . . . . . D . Pole formation . . . . . . . . . . E . Where do the conserved and non-conserved regions join in Gram. . . . . . . . . . positive rods? . F . Variable T mechanisms . . . . . . . . . V . Summary . . . . . . . . . . . . VI . Acknowledgements . . . . . . . . . . References . . . . . . . . . . . Author Index Subject Index

. .

. .

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. .

. .

. .

. .

301 306 307 308 310 311 312 322 325 326 333 340 340 343 346 349

354 359 362 362 364

. 367 . 387

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The Significance of Peroxisomes in the Metabolism of One-Carbon Compounds in Yeasts M. VEENHUIS". J . P. VAN DIJKEN"" and W. HARDER*** Laboratory of Electron Microscopy. University of Groningen

"" Laboratory of Microbiology. Delft University of Technology Department of Microbiology. University of Groningen. The Netherlands I. Introduction . . . . . . . . . . . . I1. Roleofperoxisomesinmethanolmetabolism . . . . . . A. Dissimilation of methanol . . . . . . . . . B. Assimilation of methanol . . . . . . . . . C. Structure and function of peroxisomes . . . . . . . D . The molecular substructure of crystalline peroxisomes . . . . 111. Role of peroxisomes in methylated amine metabolism . . . . . A . Methylated amines as a nitrogen source . . . . . . B. Peroxisomes and the metabolism of methylated amines C. Peroxisomes involved in the concurrent metabolism of C- and N-sources . IV. Biogenesis of peroxisomes . . . . . . . . . 1. Regulation of the synthesis of peroxisomal enzymes . . . . B. Development of peroxisomes during vegetative growth . . . . C. Development and function of peroxisomes during spore formation and germination . . . . . . . . . . . D . Assemblage of peroxisomes . . . . . . . . V . Inactivation of peroxisomal enzymes and degradation of peroxisomes . . A . Regulation of enzyme activity by inactivation . . . . . B. Modification inactivation of peroxisomal enzymes . . . . . C. Degradative inactivation of peroxisomal enzymes . . . . . D . Subcellular events in peroxisomal degradation . . . . . VI . Concluding remarks . . . . . . . . . . VII . Acknowledgement . . . . . . . . . . . References . . . . . . . . . . . .

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ADVANCES IN MICROBIAL PHYSIOLOGY. VOL 24 ISBN 0-12-027724-7

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2 5

6 13 16 24

30 30 31

31 40 41 41

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Copyright Q 1983 Arademir P r ~ r sLondon AN rights ofrcprodurlion in an) form r e s c u d

2

M. VEENHUIS,

J. P. VAN DIJKEN AND W. HARDER

I. Introduction

Micro-organisms that are capable of growth on compounds which possess a single carbon atom and a level of oxidation between methane and carbon dioxide (i.e. the so-called one-carbon compounds) are abundant in Nature. Although bacterial utilization of these one-carbon compounds has been known for a long time (see Quayle, 1972),it was not until 1969 that growth of eukaryotic organisms at the expense of such compounds was reported (Ogata et al., 1969). This first report on the isolation of a methanol-utilizing Kloeckera sp. 2201 (recently re-identified as a strain of Candida boidinii) was quickly followed by several others which established that a variety of yeasts (see Lee and Komagata, 1980) and some filamentous fungi (Goncharova et al., 1977)are able. to utilize methanol as a sole source of carbon and energy for growth. Consistently successful isolations of methanol-utilizing yeasts have been reported either from batch-type enrichment cultures containing antibacterial compounds such as penicillin or cycloserine at pH 4.5 (van Dijken and Harder, 1974) or from continuous-flow enrichments conducted at low pH values (Levine and Cooney, 1973; Pal and Hamdan, 1979). These and other studies have indicated that the ability of yeasts to grow on methanol is restricted to only a few species of a limited number of genera. This conclusion was also reached during an examination of methanol assimilation by the type strains maintained at the Culture Collection of the Centraalbureau voor Schimmelcultures at Delft, The Netherlands (Hazeu et ul., 1972). This study revealed that the type strains of only 15 species within the genera Candida, Hansenula, Pichia and Torulopsis were capable of growth on methanol. A similar screening of moulds suggested that the ability of these organisms to grow on methanol is also limited to a few species (Goncharova et al., 1977). The results obtained with yeasts have been confirmed and extended in a recent taxonomic investigation of a large number of fresh isolates and strains known to assimilate methanol (Lee and Komagata, 1980). These authors argued that methanol-utilizing strains of Hunsenulu and Pichia must be closely related, both on the basis of their cultural and physiological characteristics and also with respect to chemotaxonomic criteria. Furthermore, they suggested that the methanol-utilizing strains within the genera Cundidu and Torulopsis may be regarded as imperfect forms of Hansenula and Pichia. Lee and Komagata (1980) therefore concluded that the methanol-utilizing yeasts, although at present classified in four different genera by current taxonomic criteria, must be considered closely related. An interesting finding made by these workers was that almost all of the strains examined were able to assimilatepectin. Since this compound yields methanol on hydrolysis of its methyl esters, it was

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

3

suggested that methanol-utilizingyeasts may play a role in the metabolism of methanol that originates in Nature from decomposing plant material. It is questionable whether methanol utilization by fungi is restricted to the few yeasts and moulds capable of growth on methanol. Already before the discovery of yeasts capable of utilizing methanol as a source of carbon and energy, the key enzyme of methanol oxidation in these organisms (i.e. alcohol oxidase) had been detected in a variety of basidiomycetous moulds (Janssen and Ruelius, 1968; Kerwin and Ruelius, 1969) which are unable to grow on methanol. This suggests the possibility that methanol may be used as an energy source by these organisms. Recently, Sahm and coworkers reported further evidence for this possibility in that the brown rot fungus Poria contigua, capable of degrading lignin, can synthesize alcohol oxidase (Bringer et al., 1979) as well as formaldehyde and formate dehydrogenases (Bringer, 1980). Although it is not known at present how widespread is this capacity of methanol oxidation among non-methylotrophic fungi, it is not unreasonable to postulate that this process may be of importance in Nature during degradation of non-CI compounds that carry methyl groups. In addition to methanol, various methylated amines can be utilized by yeasts, albeit as a source of nitrogen only. This property is much more widespread among yeasts than is the capacity to assimilate methanol (van Dijken and Bos, 1981). A similar utilization of methylated amines as a nitrogen source has been reported for non-methylotrophic bacteria (Bicknell and Owens, 1980). Utilization of a number of carbon and nitrogen sources for growth by yeasts is characteristically associated with the development of unique subcellular compartments in the cells. These compartments are surrounded by a single membrane and collectively called microbodies (Fukui and Tanaka, 1979a; Veenhuis et al., 1979a;Zwart et al., 1980).Microbodies were first described by Rhodin (1954) in mouse kidney tissue and have since been observed in a large variety of eukaryotic cells. This has led to the view that these organelles represent ubiquitous subcellular compartments in eukaryotic cells (Masters and Holmes, 1977). Microbodies may have a large variety of enzyme repertoires and functions. This property, which is unknown for any other type of subcellular organelle, has 'given considerable impetus to the study of microbodies but has also produced disagreement on terminology and a unified conceptual framework of their function. The pioneering studies of de Duve and his colleagues (de Duve and Baudhuin, 1966; de Duve, 1969a,b)led to the first biochemical formulation of the activity and function of microbodies. He and his collaborators demonstrated that in these organelles catalase disposes of harmful hydrogen peroxide that is generated in a previous reaction catalysed by one or more oxidases and they proposed the term peroxisome for these structures (de

4

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

Duve, 1965). As the name suggests, the function of these organelles was envisaged as involving production and degradation of hydrogen peroxide. de Duve (1969a) proposed that the peroxisome provided the first primitive respiratory system in early aerobic organisms which was able to perform a variety of oxidative and other metabolic functions such as fatty acid B-oxidation, the glyoxylate cycle, photorespiration, amino-acid transamination and purine catabolism. Subsequent to the later appearance of the mitochondria as the predominant oxidative organellein aerobic eukaryotes, a variety of peroxisomal types may have evolved, possibly through loss of one or more functions. For instance, the enzymes necessary for the oxidation of fatty acids and the complete operation of the glyoxylate cycle are found in glyoxysomes (a special class of microbodies) of germinating oil seedlings (Breidenbach and Beevers, 1967), whereas peroxisomes of mammalian cells do not exhibit any activity for enzymes of the glyoxylate cycle (see Masters and Holmes, 1977). Alternatively, peroxisomes may be able to adapt their enzymic composition and function to the specialized requirements of particular metabolic conditions. This has, for instance, been suggested for microbodies in cucumber cotyledon leaf (Trelease et al., 1971) and in filamentous fungi (Maxwell et al., 1975). Striking examples of adaptations of peroxisome function to the physicochemical composition of the environment of cells have been encountered in studies on the metabolism of one-carbon compounds in yeasts. During exponential growth of methylotrophic yeasts on glucose, peroxisomes are generally very difficult to detect and their physiological function is uncertain (Avers, 1971; van Dijken et al., 1975b; Parish, 1975; Sahm, 1977). However, when these yeasts are grown in media containing methanol as the carbon source, a number of large peroxisomes are present in the cells. These organelles harbour mainly catalase and the hydrogen peroxide-producing alcohol oxidase which are involved in the initial oxidation of methanol (Roggenkamp et al., 1975; Sahm, 1977; Tani et al., 1978; Veenhuis et al., 1979a).A similar response is seen in yeast cells grown on n-alkanes (Osumi el al., 1975) or during growth in the presence of methylamine as a nitrogen source (Zwart et al., 1980). In these cases, the fatty acid p-oxidation system (Fukui and Tanaka, 1979b), or amine oxidase, the key enzyme of amine metabolism, is contained in the peroxisomes. The process of peroxisome proliferation can be readily reversed. When methanol-grown cells are transferred into glucose-containing media, the peroxisomes present in these cells quickly disappear (Bormann and Sahm, 1978;Veenhuis et al., 1978a)as a result of active degradation. This rapid adjustment of peroxisome numbers and functions to environmental conditions indicates that yeasts provide ideal model systems for the study of the physiological function, biogenesis and turnover of these organelles.

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

5

It is the purpose of this review to highlight the role of peroxisomes in the metabolism of methanol and methylated amines in yeasts and to discuss the available information regarding their biogenesis and turnover. For general information on methylotrophic yeasts the reader is referred to recent reviews by Sahm (1977) and Tani et al. (1978). The function and biogenesis of peroxisomes in a variety of eukaryotic cells has been considered by Masters and Holmes (1977,1979)and was the central topic of a recent symposium held at the New York Academy of Sciences.

II. Role of Peroxisomes in Methanol Metabolism During studies on the physiology and biochemistry of methanol oxidation by methylotrophic yeasts, several research groups almost simultaneously discovered that adaptation of these organisms to growth on methanol is associated with the proliferation of large microbodies in the cells (Romanenko and Pidhorskyi, 1974,1976; van Dijken et al., 1975b; Sahm et al., 1975; Fukui et al., 1975a).This very characteristic adaptation (Fig. 1) has been observed in

FIG. 1. Survey of cells of Hunsenulu polymorphu showing the overall cell morphology after growth on glucose (a) and methanol (b) as the sole source of carbon. In the glucose-grown cell (batch culture on 0.25% glucose, harvested at A63 = 1.O) one small peroxisomal profile is observed. In the methanol-grown cell (chemostat culture, D =0.10 h-I) approximately 20 peroxisomal profiles are visible. 1 (a) is from Veenhuis et ul. (1979a). In all the electron micrographs shown in this article, the cells were fixed and postfixed with potassium permanganate unless otherwise mentioned. Abbreviations used: I, lipid droplet; m, mitochondrion; N, nucleus; P, peroxisome; V, vacuole. The bar marker represents 0.5 pm unless stated otherwise.

6

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

all methanol-utilizing yeasts studied so far. To evaluate the unique function and structure of the microbodies during methylotrophic growth in yeasts it is first necessary to consider pertinent biochemical and physiological aspects of methanol metabolism in these organisms. These are discussed in the first part of this Section. The second part deals with the structure and function of the microbodies in methanol-grown yeasts.

A . DISSIMILATION OF METHANOL

1. Oxidation of Methanol to Formaldehyde The enzymology of methanol oxidation in yeasts is fundamentally different from that encountered in bacteria. In the prokaryotes studied so far a dehydrogenase catalyses the first step of methanol metabolism. This enzyme which belongs to a new class of enzymes collectively called quinoproteins (Duine and Frank, 1981) probably donates its electrons to the electron transport chain at the level of cytochrome c. Yeasts and moulds growing on methanol as the carbon and/or energy source, however, do not oxidize methanol via components of the electron transport chain. In these organisms an alcohol oxidase (EC 1.1.3.13) is present which catalyses the initial oxidation of methanol and is dependent on oxygen as an electron acceptor. This alcohol oxidase has been purified from various fungi (Table 1). It is a high molecular-weight protein consisting of eight identical subunits, each of which contains one non-covalently-bound flavin adenine dinucleotide moiety as a prosthetic group. Apart from methanol, lower primary aliphatic alcohols also may serve as substrates and are oxidized according to the following general equation:

R-C-OH

I

H2

+02+R-C=O +H202 I

(1)

H

Formaldehyde in its hydrated form (methylene glycol) is also oxidized by alcohol oxidase. Alcohol oxidase is present in high amounts during growth of yeasts on methanol. Reported data on enzyme activities (Table 1) are difficult to compare since the activity of the enzyme is markedly affected by the assay conditions (see below). Furthermore, it is not known whether the V,,, values of the various preparations of alcohol oxidase listed in Table 1 reflect a true difference in activity or a difference in stability during purification. In this respect it is of interest that specific activities of alcohol oxidase in cell-free extracts have been detected which are equal or exceed the values for some of

TABLE 1. Properties of alcohol oxidase from various fungi

K n (mM)

Source

V,,, methanol Formal- (pmol min-* Molecular Methanol Ethanol dehyde (mg protein)-') weight

MOULDS Polyporus sp.

1.52

Poria contigua

YEASTS Kloeckera sp. 2201 Kloeckera sp. 220 1 Hansenula polymorpha Hansenula polymorpha Hansenula polymorpha Candida boidinii Candida N 16 Candida N16 Candida 25A Pichia pastoris Pichia sp.

Reference

10.0

-

25.1

300,000

0.2

1.o

6.1

20.0

610,000

Janssen et al. (1965); Janssen and Ruelius ( 1968) Bringer et al. (1979); Bringer (1980)

1.25 0.44 0.08 0.23 1.3' 2.0 2.12

2.5 2.5

-

2.4

11.0 8.5

-

-

4.4 7.2 2.62

2.6 4.7 5.7

11.3 56.3 3.4 3.5

570,000" 673,OOob 617,000 669,000 6 16,000 600,000 2 10,000 600,000 520,000 675,000 300,000

Tani et al. (1972a,b); Ogata et al. (1975) Kato et al. (1976); N. Kato, personal communication Ogata et al. (1975) Kato et al. (1976); N. Kato, personal communication van Dijken (1976) Sahm and Wagner (1973b); Sahm (1975) Fuji and Tonomura (1972) Fujii and Tonomura (1975) Yamada et al. (1979) Couderc and Baratti (1980) Pate1 el al. (1981)

-

-

-

-

-

-

0.019 1.4c 0.5

0.13

-

2.8 11.9 6.6

3.5

Determined by sedimentation velocity centrifugation. Determined by sedimentation equilibrium centrifugation. Determined at air saturation.

8

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

the purified enzymes (Table 1). For instance, extracts prepared from cells of Hansenula polymorpha grown in methanol/sorbose-limited chemostat cultures contained activities of alcohol oxidase up to 19.6 pmol min-' (mg protein)-' (Eggeling and Sahm, 1980). It should be noted that different assay methods have yielded different units of activity. When the enzyme is assayed with the peroxidase/chromogen method, activities are usually expressed as pmol hydrogen peroxide produced min- I, whereas polarographic estimations are generally given as pmol oxygen consumed min-' (mg protein)-'. For comparison, the latter activity has to be multiplied by a factor of 2 because of the catalytic action of catalase in cell-free extracts (J. P. van Dijken, unpublished observations; Th. Egli, personal communication). Relatively little is known about the kinetics of alcohol oxidase with respect to its second substrate oxygen. In the case of H . polymorpha (van Dijken et al., 1975a)and Pichiapastoris (Couderc and Baratti, 1980) it was shown that the enzyme has a low affinity for oxygen, a property that is shared by other hydrogen peroxide-producing oxidases (Table 2). Since the oxidase reaction is essentially a two-substrate reaction, the oxygen concentration has a significant effect on the affinities of the enzyme for the alcohol substrate. As a consequence, the apparent K m values listed in Table 1 cannot be directly compared since most of the K m estimations were performed with colorimetric assays in which the oxygen concentration in the incubation mixture was not reported. The low affinity of alcohol oxidase for oxygen means that in uiuo the enzyme works outside the range of oxygen concentrations wherein it expresses its maximal activity. This occurs in aqueous environments saturated with pure oxygen. This makes the enzyme essentially a poor catalyst for the physiological function that it fulfils in the intact cell. The low affinity of alcohol oxidase for oxygen is indeed evident in uiuo. Oxidation of methanol by washed suspensions of yeast cells is linearly proportional to the dissolved oxygen concentration, whereas in bacteria, for example Hyphomicrobium X, methanol is oxidized to formaldehyde via cytochrome oxidase so that methanol oxidation becomes oxygen-limited only at very low concentrations of dissolved oxygen (Fig. 2). Since the oxidation of methanol by alcohol oxidase results in the formation of hydrogen peroxide, it is imperative that this is decomposed. In methylotrophic yeasts this is carried out by catalase and enhanced concentrations of this enzyme generally parallel high alcohol oxidase activities (see Sahm, 1977). It has been reported that catalase may be functioning in a peroxidative fashion in which methanol is oxidized to formaldehyde by the hydrogen peroxide formed in the alcohol oxidase reaction (Fujii and Tonomura, 1972; Roggenkamp et al., 1974; van Dijken et al., 1975a). In addition to methanol, formaldehyde and formate can also be oxidized in uitro by catalase purified from H . polymorpha (Table 3; van Dijken et al., 1975a). Whether the enzyme

TABLE 2. Apparent K, values for oxygen of various hydrogen peroxide-producing oxidases 4

Enzyme

EC number ~

_

_

_

_

Source

Km (mM)

Reference

0.20 0.20

Gibson et al. (1964) Isherwood et al. (1960) Couderc and Baratti (1980) van Dijken et al. (1976a) Ackerman and Brill (1965) Dixon and Kleppe (1965) Large et al. (1980) and personal communication Bongaerts (1978)

~

Glucose oxidase Gulunolactone oxidase Alcohol oxidase Alcohol oxidase Xanthine oxidase D-Amino acid oxidase M i n e oxidase

1.1.3.4 1.1.3.8 1.1.3.13 1.1.3.13 1.2.3.2 1.4.3.3 1.4.3.4

Aspergillus niger Rat liver Pichia pastoris Hamenula polymorpha Bovine milk Hog kidney Cadi& boidinii

Urate oxidase

1.7.3.3

Bacillusfmtidiosus

1.o

0.4 0.24 0.18 0.09

1.o

b

m

9

0

z

f?

0

%m

0

z

10

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

0

I 0

0.05

0.10

0.15

Oxygen concentration

0.20

(mM)

FIG. 2. Rate of (excess) methanol oxidation by washed suspensions of methanolgrown Hansenula polymorpha (*)and methanol-grown Hyphomicrobium X ).( at different dissolved oxygen tensions (van Dijken et al., 1976a; J . B. M. Meiberg, unpublished results). The organisms were grown in methanol-limited chemostat cultures at D=0.10 h-I.

acts peroxidatively or catalatically in vivo is not known. Kinetic studies on systems in uitro cannot solve this question since it is difficult to evaluate to what extent the conditions for either of the two mechanisms are met in the growing cell. Peroxidative versus catalatic action of catalase is dependent on the ratio of the rate of hydrogen peroxide production and the catalase concentration (Ohshino et al., 1973; van Dijken et al., 1975a). Various reports have appeared implicating nicotinamide adenine dinucleotide (NAD +)-dependent dehydrogenase in methanol oxidation in yeasts (Mehta, 1975a,b; Egorov et al., 1977; Dudina et al., 1977; Simisker et al., 1977). The available information suggests that this reaction (whose rates are generally low) may be due to an aspecific activity of primary or secondary alcohol dehydrogenases. Furthermore, in some cases activities detected in cell-free extracts may have been due to a concerted action of alcohol oxidase and NAD +-dependent formaldehyde and formate dehydrogenases. It has become clear, however, that growth of yeasts on methanol is strictly dependent on the activity of alcohol oxidase and catalase since mutants lacking either of the two enzymes have lost the ability to grow on methanol (Sahm and Wagner, 1973a; L. Eggeling, unpublished work).

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

11

TABLE 3. Possible reactions catalysed by catalase during methanol metabolism in yeasts

Mode of catalase action

Reaction catalysed

Catalatic Peroxidatic

H202 H202+2H20 0 2 H202 CH3OH-t HCHO + 2H20 H202 + HCHO-t HCOOH + H2O H202 HCOOH +COz + 2H20

+ + +

+

2. Oxidation of Formaldehyde to Carbon Dioxide Complete oxidation of formaldehyde by yeasts proceeds via two NAD+dependent dehydrogenases. The first enzyme in this sequence, formaldehyde dehydrogenase, is strictly dependent on glutathione for activity. This is due to the fact that it is not free formaldehyde but the hemi-mercaptal of formaldehyde and glutathione that is the substrate for this enzyme, which forms S-formylglutathione as a product (Uotila and Koivusalo, 1974;Schutte et al., 1976; van Dijken et al., 1976b; Kato et al., 1979a). The formate dehydrogenase of yeasts shows striking differences when compared to the enzyme commonly encountered in methylotrophic bacteria. The most peculiar property of the yeast formate dehydrogenase is a very low affinity for formate, ranging from 6 to 55 mM in various organisms (Kato el al., 1974; Schutte et al., 1976; van Dijken et al., 1976b; Volfova, 1975). On the basis of kinetic experiments with partially purified formate dehydrogenase from Hansenulapolymorpha, van Dijken et al. (1976b) suggested that the product of the formaldehyde dehydrogenase reaction, namely S-formylglutathione, rather than free formate is the actual substrate for formate dehydrogenase. This enzyme only hydrolysed S-formylglutathione in the presence of NAD+. The route of complete oxidation of formaldehyde in H. polymorpha may thus be represented by: /OH H2C\ SG

‘SG

+NAD+ +H-C

P \

SG

+NADH + H+

(2)

12

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

However, recent studies of Kato et al. (1980) and Neben et al. (1980) showed that in a variety of methanol-utilizing yeasts, including H. polymorpha, an S-formylglutathione hydrolase is present which is independent of NAD+ for activity. The enzyme is absent from glucose-grown cells but shows a high activity in methanol-grown cells (up to 53 pmol min-' (mg protein)-'; Neben et al., 1980). Furthermore, in contrast to the findings made with H. polymorpha, the formate dehydrogenases purified from Kloeckera sp. 220 1 and Candida boidinii did not exhibit hydrolase activity towards S-formylglutathione. It was therefore concluded that in these organisms oxidation of formaldehyde to carbon dioxide requires the action of S-formylglutathione hydrolase and proceeds according to the following reactions:

'SG

'SG

\

SG

HCOOH + NAD+ +CO2+ NADH

+H +

(6)

In view of the low substrate affinity of the formate dehydrogenase in these organisms it remains to be elucidated how formate is oxidized in vivo. When it is assumed that the intracellular concentration of formate does not build up to millimolar concentrations, the activity of formate dehydrogenase in cell-free extracts of methanol-grown yeasts is too low to explain the growth rate. 3. Generation of Energy During Formaldehyde Oxidation

As already outlined, during growth of yeasts on methanol NADH is generated via the action of formaldehyde and formate dehydrogenases which are thought to be soluble proteins. Under these conditions all the NADH is generated outside the mitochondria. This is a unique situation which only occurs during growth of yeasts on methanol. The mechanism whereby the mitochondria in methanol-utilizing yeasts oxidize cytoplasmic NADH is still an unsolved problem. In mammalian cells various shuttle mechanisms have been implicated in the oxidation of cytoplasmic NADH (Dawson, 1979). In fungi and plants, however, oxidation of cytoplasmicNADH may proceed in a

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

13

different way. Mitochondria isolated from these organisms oxidize exogenous NADH in the absence of shuttle components (von Jagow and Klingenberg, 1970;Ohnishi, 1973; Moore and Rich, 1980). In plants and yeasts two NADH dehydrogenases are present in the mitochondria; one of these enzymes is located at the outer layer of the inner membrane and oxidizes cytoplasmic NADH. The other NADH dehydrogenase, located at the inner layer of the inner mitochondria1 membrane, oxidizes NADH generated in the matrix of the organelle by the enzymes of the tricarboxylic acid (TCA) cycle. Unlike internal NADH dehydrogenase, oxidation of NADH by the externally located system is always insensitive to inhibitors of phosphorylation site I such as rotenone but is sensitive to antimycin A and cyanide, suggesting that reduction equivalents are channelled into the electron transport chain at the level of ubiquinone (Ohnishi, 1973; Moore and Rich, 1980). When a similar mechanism operates during growth of yeasts on methanol this would have important energetic consequences. Whereas during growth of yeasts a theoretical maximum of 3 mol of ATP may be formed for each mol of NADH oxidized (as a result of the fact that the bulk of the NADH is always generated intramitochondrially), during growth of yeasts on methanol the ATP yield from NADH is maximally only 2 mol mol-l.

B . ASSIMILATION OF METHANOL __

Early studies on the assimilation of methanol in yeasts indicated that methanol is incorporated into cell material via phosphorylated hexoses (Fujii and Tonomura, 1973; Fujii et al., 1974). This suggested that a pathway of formaldehyde fixation operated in yeasts similar to that encountered in certain bacteria, namely the ribulose monophosphate pathway of formaldehyde fixation. The key reactions of this pathway are a condensation of formaldehyde with ribulose monophosphate to form ~-arabino-3-hexulose 6-phosphate which is subsequently isomerized to fructose 6-phosphate (Kemp, 1974; Ferenci et al., 1974). The view that these reactions are also responsible for the incorporation of formaldehyde in yeasts was strengthened by several reports on the presence of enzymes catalysing these reactions in extracts of methanol-grown yeasts (see Sahm, 1977). However, conflicting results have also been reported. Whereas some workers, using the routine spectrophotometric and isotope assays, detected high activities of these enzymes (Sahm and Wagner, 1974; Die1 et al., 1974; Trotsenko, 1975)others found only trace activities which depended on the presence of ATP (Fujii et al., 1975), or no activity at all (J. P. van Dijken and W. Harder, unpublished observations). Later work clearly eliminated the presence of hexulose phosphate synthase and hexulose phosphate isomerase in methanol-grown

14

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

yeasts (Kato et al., 1977; van Dijken et al., 1978) and thereby the bacterial route of formaldehyde fixation. Instead, studies on the enzyme profiles of sugar phosphate metabolism in methanol-grown yeasts suggested that in contrast to the ribulose monophosphate pathway-in which triose phosphates are formed from hexose phosphates-in these organisms hexose phosphates are formed from triose phosphates. This was indicated by significantly elevated concentrations of fructose 1,6-bisphosphatase during growth on methanol (van Dijken et al., 1978; Babel and Lomagen, 1979). These and other findings have led to the formulation of a xylulose phosphate pathway of formaldehyde fixation as depicted in Fig. 3. The key reaction of this cycle is a transketolase-type condensation of xylulose 5-phosphate with formaldehyde resulting in the formation of glyceraldehyde phosphate (GAP) and dihydroxyacetone.The enzyme catalysing this reaction has been given the trivial name dihydroxyacetone synthase (O’Connor and Quayle, 1980). Dihydroxyacetone is phosphorylated by a triokinase and subsequently condenses with glyceraldehyde phosphate to form fructose 1,Ci-bisphosphate. Via the action of fructose 1,6-bisphosphate aldolase and fructose, 1,6-bis-

DHAP

GAP

DHAP

cel I constituents

F6P

I

F6P

reorrongement reactions

/

FIG. 3. The xylulose monophosphate pathway of formaldehyde fixation in yeasts. Abbreviations: XuSP-xylulose 5-phosphate; DHAP-dihydroxyacetone phosphate; GAP-glyceraldehyde phosphate; FBP-fructose 1,6-bisphosphate; F6P-fructose 6-phosphate. After van Dijken et al. (1978).

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

15

phosphatase, fructose 6-phosphate is then formed. Two molecules of fructose 6-phosphate and one molecule of dihydroxyacetone phosphate are then rearranged to give three molecules of xylulose phosphate by reactions catalysed by transaldolase, transketolase, pentose phosphate isomerase and epimerase. The net result of the cycle is the formation of one mole of triose phosphate from three moles of formaldehyde at the expense of three moles of ATP: 3 HCH0+3ATP+1 GAP+3 ADP+2 Pi

(7)

There is now convincing evidence that the above cycle of formaldehyde fixation, which at the time of its formulation was largely hypothetical, is indeed operating in methanol-utilizing yeasts. All key enzymes of this pathway, including the condensation reaction between xylulose 5-phosphate and formaldehyde, have been detected in extracts of methanol-grown yeasts and are present in elevated concentrations compared to extracts of glucosegrown cells (van Dijken et al., 1978; Babel and Lomagen, 1979; O'Connor and Quayle, 1980). Furthermore, mutants of Hansenula polymorpha and Candida boidinii which lack or contain decreased amounts of triokinase or fructose bisphosphatase are impaired in their ability to grow on methanol whereas revertants have regained this property. This indicated that these enzymes are indispensable for methanol metabolism (O'Connor and Quayle, 1979). Xylulose 5-phosphate-dependent fixation of [I4C]formaldehyde by cell-free extracts of yeasts results in the formation of labelled dihydroxyacetone (Kato et al., 1979b; Waites and Quayle, 1980). A reinvestigation of the early labelled products after pulse labelling of whole cells showed that, apart from sugar phosphates, dihydroxyacetone is also an early intermediate in the fixation of formaldehyde by whole cells (Lindley et al., 1980). A most elegant proof of the operation of this pathway in uivo has been obtained in studies on the intramolecular distribution of I4C in hexose phosphates after pulse labelling of whole cells of methanol-grown H.polymorpha with ['4C]methanol. These studies showed that carbon atoms, 1, 3 , 4 and 6 were heavily labelled with I4C (Waites et al., 1981) which is to be expected on the basis of the operation of the xylulose 5-phosphate pathway of formaldehyde fixation. When ['4C]formaldehydeis fixed via this pathway, then radioactivity is fixed into dihydroxyacetone. Since dihydroxyacetone is a symmetrical molecule, hexoses derived from it after phosphorylation and isomerization can be expected to have the observed labelling pattern. In contrast, fixation of formaldehyde via the ribulose monophosphate pathway results in the formation of hexose phosphates predominantly labelled at C-1 (Kemp and Quayle, 1967).Thus far, studies on the mechanism of methanol assimilation in yeasts have been performed mainly with H. polymorpha and C . boidinii. It is to be expected, however, that most if not all yeasts capable of growth on

16

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

methanol possess this pathway. In this respect, it is of interest that methanol-assimilatingrepresentatives of the genus Pichia were also found to contain' key enzymes of the xylulose phosphate pathway of formaldehyde fixation when grown on methanol (J. P. van Dijken, unpublished observations; W. Hazeu, personal communication). The pathway has so far not been encountered in methylotrophic bacteria.

C. STRUCTURE A N D FUNCTION OF PEROXISOMES

Microbodies of methanol-grown yeasts show a number of characteristic properties: they appear in clusters in the cell and exist in close association with strands of endoplasmic reticulum (Fig. 4a). The individual organelles which are surrounded by a single membrane of approximately 7 nm (70 A) in width contain, without exception, crystalline inclusions (Fig. 4b,d) (van Dijken et al., 1975b; Fukui et al., 1975a; Sahm et al., 1975; Hazeu et al., 1975). In freeze-etch replicas the peroxisomal membranes show characteristic smooth fracture faces (Fig. 4c). Electron microscopical studies on the yeast Hansenula polymorpha have shown that the size, number and substructure of the organelles are dependent on cultivation conditions (Veenhuis et al., 1978b, 1979a). Cells from the exponential growth phase of batch cultures contained rounded organelles with a partly crystalline matrix (Fig. 4b). During vegetative reproduction, the size and number of peroxisomes per cell gradually increased, along with an increase in size of the crystalloids in the peroxisomal matrix. In batch cultures entering the stationary phase of growth the organelles became more rectangular in shape and contained a largely, or completely, crystalline matrix (Veenhuis et al., 1978b). The peroxisomes present in cells of methanol-limited chemostat cultures invariably showed a completely crystalline substructure (Fig. 4d). The volume fraction of the organelles in the latter cells increased with decreasing growth rate whereas the total number per cell remained approximately constant, indicating an increase in size of the individual organelles with decreasing growth rate. This increase in size was associated with an overall change in morphology of the organelles. Mature peroxisomes in cells growing in chemostat culture at high dilution rates generally had a rounded form whereas in cells growing at low dilution rates they were almost cubic in shape (Fig. 4e). Old cells characterized for instance by the presence of many bud scars invariably contained large numbers of peroxisomes. Up to 20 peroxisomalprofiles have been observed in such cells in which these organelles occupied 80% of the total intracellular volume (compare Fig. lb). Evidence that the microbodies of methanol-grown yeasts contain alcohol oxidase and catalase has been obtained via cell fractionation and cytochemi-

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

17

FIG. 4. Thin sections of (a) a methanol-grown cell of Candida boidinii, showing a typical cluster of intracellular peroxisomes, and (b) a peroxisome, isolated from methanol-grown Hansenulapolymorpha, showing the characteristic crystalloid present in these organelles and the dimensions of the surrounding membrane. After freeze-etchingthis peroxisome membrane shows the typically smooth fracture faces PF and EF (c; nomenclature after Branton et al. (1975). The arrow indicates the direction of shadowing). (d) and (e) show details of cells of Hansenula polymorpha grown on methanol in chemostat culture, showing that these cells contain completely crystalline peroxisomes (d) of a cubic shape (e); the arrow indicates the direction of shadowing.

18

M. VEENHUIS,

J. P. VAN DIJKEN AND W. HARDER

cal studies. Sahm et al. (1975) reported that 80% of the alcohol oxidase sedimented after a low-speed spin of a sphaeroplast lysate of methanol-grown Cundidu boidinii. However, only 30% of the catalase was similarly pelleted. Roggenkamp et al. (1975) partially purified microbodies from methanolgrown C. boidinii on Ficoll gradients and showed that the particulate activities of alcohol oxidase and catalase in sphaeroplast lysates were associated with those fractions which, on electron microscopical examination, were found to be highly enriched in microbodies. Polyacrylamide-gel electrophoresis showed that alcohol oxidase and catalase were the main protein components of these microbody-enriched fractions. Qualitatively similar results were obtained by Fukui et al. (1975b); 50% of the alcohol oxidase and catalase activities were pelleted after a 3000g spin of a sphaeroplast lysate of Kloeckeru sp. 2201 and an additional 20% was sedimented after centrifugation at 20,000 g. They also found that enrichment of the microbodies on sucrose gradients coincided with an increase in specific activity of alcohol oxidase and catalase. The enzymic composition of the microbodies in methanol-grown yeasts has also been studied with cytochemical techniques. Using the diaminobenzidine method, which had been successfully used for the demonstration of catalase activity in Saccharomyces cerevisiae (Hoffman et al., 1970; Todd and Vigil, 1972), van Dijken et al. (1975~)studied the localization of catalase activity in methanol-grown H. polymorpha. After incubation of glutaraldehyde-fixed cells with diaminobenzidine and hydrogen peroxide the reaction products were present in the microbodies and the mitochondria (Fig. 5a). Staining of the microbodies was prevented when incubations were performed in the presence of 3-amino-1,2,4-triazole, a known catalase inhibitor. After such incubations the mitochondrial staining was unaffected (Fig. 5b). Since the mitochondrial staining did not depend on the presence of hydrogen peroxide and was prevented by the addition of potassium cyanide to the incubation mixtures, it was concluded that catalase activity in methanol-grown H. polymorpha is exclusively present in the microbodies. The staining of the mitochondria is probably due to activity of cytochrome c peroxidase (Hoffman et al., 1970; Todd and Vigil, 1972). Catalase activity was also demonstrated with a modified diaminobenzidine procedure. This procedure relied on endogenous hydrogen peroxide production by the alcohol oxidase during aerobic incubations with methanol (Veenhuis et ul., 1976). Thus incubations of both glutaraldehyde-fixed and unfixed cells with diaminobenzidine and methanol resulted in positively stained microbodies. However, the peroxisomes were frequently not uniformly stained by this method; depending on cultivation conditions, cells contained peroxisomes that were only partly stained or, occasionally, not stained at all (Fig. 5c). Staining of peroxisomes was not observed during anaerobic incubations or during incubations in the presence of aminotriazole. The above procedure, which principally locates

FIG. 5. Thin sections of methanol-limited chemostat grown cells of Hansenulu polymorpha. (a) Shows positively stained peroxisomes after incubation of glutaraldehyde-fixed cells with 3.3-diaminobenzidine and hydrogen peroxide; this staining -but not the mitochondria-is absent after similar incubation in the presence of aminotriazole as an inhibitor of catalase activity (b). After incubation of unfixed cells with diaminobenzidine and methanol as the endogenous source of hydrogen peroxide the peroxisomes are only partly stained (c). Time-course incubations with diaminobenzidine and methanol showed that the first reaction products are localized in the central region of the peroxisomes (d), indicating that staining of the organelles cannot be considered as an artefact of diffusion.

20

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

catalase activity, does not necessarily disclose the localization of a peroxisoma1 oxidase. The hydrogen peroxide required for this reaction could have been produced elsewhere in the cell and may have reacted with catalase after diffusion towards this enzyme. However, time-course incubations with diaminobenzidineand methanol showed that the initial reaction products are localized in the central part of the peroxisomes (Fig. 5d). These results exclude the possibility of staining of the organelles as a result of diffusion of hydrogen peroxide, generated by the oxidase, from the cytoplasm into the organelle since this would have resulted in staining at the periphery of the organelles. Additional incubation with diaminobenzidine and hydrogen peroxide of glutaraldehyde-fixed cells, previously incubated with diaminobenzidinel methanol, showed that the parts of the peroxisomes which were originally unstained now became stained (Veenhuis et al., 1976). This staining was not observed in control experiments (i.e. the same procedure applied to unjixed cells or in the presence of aminotriazole) demonstrating that catalase activity is really present in the originally unstained parts. Taken together these results suggest that after incubations with diaminobenzidine/methanolthe diaminobenzidine reaction product is only formed at the sites of alcohol oxidase activity. Therefore, this technique can be regarded as a reliable method for the cytochemical localization of alcohol oxidase activity-and probably other peroxisomal oxidase activities-in yeasts. Alcohol oxidase has also been located by a direct cytochemical technique based on the use of cerous ions. This so-called cerium technique, originally developed by Briggs et al. (1975) for the localization of NADH-oxidase activity, proved extremely useful for the detection of intracellular hydrogen peroxide-producing oxidases. The method is based on trapping of hydrogen peroxide by Ce3+ ions resulting in the formation of an electron-dense complex, probably cerium perhydroxide (Ce(0H)ZOOH). Aerobic incubations of glutaraldehyde-fixed sphaeroplasts of H. polymorpha with this reagent confirmed the results obtained after incubations of cells with diaminobenzidine and methanol in that substrate-dependent reaction products were exclusively present in the microbodies (Fig. 6a). Therefore, according to the definition of de Duve (1973) these organelles may be considered as peroxisomes. The apparent solubility of part of the alcohol oxidase and catalase in the fractionation experiments of Sahm et al. (1975) and of Fukui et al. (1975b) is most likely due to leakage of these enzymes from the peroxisomes which, especially in yeasts, are known to be very fragile (Avers, 1971). The other possibility, a truly cytoplasmic localization of a substantial part of the enzymes, can be ruled out on the basis of the results of the cytochemical experiments since such high concentrations of soluble enzyme would have been well within the limit of detection of the applied cytochemical procedures.

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

21

Apart from alcohol oxidase and catalase other hydrogen peroxide-producing oxidases such as urate oxidase, D-amino acid oxidase and L-aihydroxy acid oxidase (Fig. 6b) have been detected in peroxisomes of methanol-grown yeasts (Veenhuis et al., 1976; Fukui et al., 1975b). However, time-course cytochemical staining experiments showed that the minimum incubation times required for the detection of these enzymes were 50-100 times longer than those required for alcohol oxidase (M. Veenhuis, unpublished observations), which is in accordance with the activities of these enzymes found in vitro (Veenhuis et al., 1979a). Proteins involved in the metabolism of methanol, other than alcohol oxidase and catalase, have not yet been detected in peroxisomes. In fractionation experiments, the other enzymes involved, namely formaldehyde and formate dehydrogenases, appeared to be soluble. Although no systematic study has been made of the subcellular localization of the enzymes involved in the assimilationof methanol the availableevidence indicates that they are also soluble enzymes (J. R. Quayle, personal communication). The localization of fructose 1,dbisphosphatase studied with cytochemical methods confirmed this view since reaction products have been observed only in the cytosol (Veenhuis et al., 1980~;Fig. 6c). On the basis of the results discussed so far, we may conclude that peroxisomes present in methanol-grown yeasts constitute a clear-cut and comparatively simple example of the physiological role of this type of organelle in intermediary metabolism. The first reaction of methanol dissimilation (the oxidation of methanol to formaldehyde) is carried out by peroxisome-borne enzymes and during growth on methanol all the carbon flows through this compartment, thereby providing the cytoplasm with formaldehydefor assimilation and energy generation (Fig. 7). Other examples of peroxisomal functions, which includes a significant unidirectional flow of carbon through these organelles, include photorespiration and B-oxidation of fatty acids. In green leaves of plants phosphoglycollateis formed as a result of the oxygenase activity of ribulose bisphosphatase. After dephosphorylation, glycollate is oxidized to glyoxylate by a peroxisomal glycollate oxidase and is further metabolized to malate (Tolbert and Yamazaki, 1969). In fat-storing seeds the B-oxidation system of fatty acids is located in peroxisomes which harbour fatty acyl-CoA oxidase (Beevers, 1969). Similarly, during growth of yeasts on n-alkanes fatty acids are oxidized via peroxisomal enzymes (Fukui and Tanaka, 1979b). In animal cells, the role of peroxisomes is less well defined and probably more diverse. Various oxidations which yield hydrogen peroxide can be carried out simultaneously by these organelles such as oxidation of a-hydroxy acids, D-amino acids and uric acid (de Duve and Baudhuin, 1966) as well as 8-oxidation of higher fatty acids (Lazarow,1978; Osmundsen et ul., 1979). In

22

M. VEENHUIS, J.

P. VAN DIJKEN AND W. HARDER

FIG. 6. Thin sections of methanol-limited chemostat grown cells of Hunsenulu polymorphu. (a) Shows positively stained peroxisomes after aerobic incubations with cerium chloride and methanol, indicating the presence of alcohol oxidase activity in the organelles. The presence of L-a-hydroxy acid oxidase was demonstrated after similar incubations with cerium chloride and glycollate (b). (c) Shows the presence of fructose bisphosphatase activity in the cytosol, demonstrated after incubations with cerium chloride and fructose bisphosphate.

addition, the peroxidative action of catalase may add a number of substrates which can be oxidized by peroxisomes (de Duve and Baudhuin, 1966). Catalase may, for example, significantly contribute to the disposal of alcohol by liver cells (Thurman et a!., 1975).The mammalian peroxisome is therefore a multifunctional organelle as opposed to the organelles in yeasts utilizing methanol or n-alkane or the organelles of green leaves and fat-storing seeds. Consequently, a number of peroxisomal reactions thought to be of physiological importance in mammalian cells are probably of secondary importance

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

I

PEROXISOME

1

23

1

FIG. 7. Schematic representation of the role of peroxisomes in methanol metabolism in yeasts; (1) peroxisomal oxidation of methanol; (2) chemical formation of a hemimercaptal OF Formaldehyde and glutathione (GSH); (3) oxidation OF formaldehyde by formaldehyde dehydrogenase; (4) hydrolysis of S-formylglutathione and oxidation of Formate; ( 5 ) peroxisomal Formation of formate by alcohol oxidase and/or catalase.

for peroxisomes in methanol-utilizingyeasts. For example, the possibility of a peroxidative activity of catalase does not provide additional physiological functions during methanol metabolism. Whether or not the catalase is acting peroxidatively (for example, in the oxidation of methanol to formaldehyde)is of no significance for the functioning of the organelles which already catalyse the oxidation of methanol to formaldehyde without the conservation of energy. Also, the low affinity of the peroxisomal oxidase for oxygen (see preceeding paragraphs) which, it has been suggested, may help to protect the mammalian cell from oxygen toxicity (de Duve and Baudhuin, 1966), does not seem to have a physiological role during growth of yeasts on methanol. Indeed, this property of alcohol oxidase may even be regarded as disadvantageous since it drastically lowers the rate of methanol oxidation and probably necessitates the presence of a large amount of enzyme. Although the function of peroxisomes in methanol metabolism in methylotrophic yeasts is well defined, it is not at all clear why they are implicated in this process in these organisms. It is evident from the routes of methanol metabolism in bacteria that methanol can be efficiently oxidized by a dehydrogenase and there is so far no clue as to why yeasts have developed a separate peroxisomal reaction which, by analogy, could have been carried out by a dehydrogenase associated with the electron transport chain in the mitochondrion.

24

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

D . THE MOLECULAR SUBSTRUCTURE OF CRYSTALLINE PEROXISOMES

The molecular architecture of the crystalloids present in peroxisomes of methanol-grown yeast has been described by Osumi et al. (1979). On the basis of biochemical and cytochemical evidence (Fukui et al., 1975b; Osumi and Sato, 1978) it was suggested that both catalase and alcohol oxidase were structural elementsof the crystalloids, contained in peroxisomes of methanolgrown Kloeckera sp. 2201. Tilting experiments with cryosections of this organism showed that the different crystalline patterns observed in the peroxisomal matrix were in fact artificial images all of which could be accounted for by superposition of two different types of particles arranged alternately in a tetragonal lattice. On the basis of their shape and dimensions, it was suggested that these particles represented alcohol oxidase and catalase molecules, respectively, and therefore it was argued that the lattice structure, as observed in cryosections, was composed of alternating molecules of catalase and alcohol oxidase (Osumi et al., 1979). Thus, the model proposed for Kloeckera sp. 2201 specifies that the two proteins which are considered to represent the structural elements of the crystalloids, are present in a fixed 1:1 ratio. A number of observations indicate that such a two-component crystalloid is probably not present in methanol-grown Hansenulapolyrnorpha. In this organism, the development of a crystalloid in the peroxisomal matrix was shown to be exclusively dependent on the synthesis of alcohol oxidase protein in the cells (Veenhuiset al., 1979a).This is in agreement with an earlier observation of Sahm et al. (1975) who showed that crystalloids are invariably absent from alcohol oxidase-negative mutants of this yeast although catalase was present in similar activity as in wild-type cells (Eggelingetal., 1977). Also, when wild-type cells were grown under conditions in which the cells lacked alcohol oxidase but contained increased concentrations of other peroxisomal enzymes such as amine oxidase, uricase and D-amino acid oxidase,crystalloids were never observed (Zwart et al., 1980; Veenhuis et al., 1981a). Furthermore, when H. polymorpha was grown under methanol-limitation in chemostat culture, the ratio of alcohol oxidase and catalase varied considerably with the growth rate (van Dijken, 1976). Yet, under these conditions, the peroxisomes were completely crystalline, irrespective of the growth rate of these cells. Since the activities of both enzymes are confined to peroxisomes (Veenhuis et al., 1976,1979a)it is difficult to envisage that these organellescan be composed of the two enzymes in a fixed 1:l ratio. The distribution of alcohol oxidase and catalase activities in completely crystalline peroxisomes present in chemostat-grown cells of H. polymorpha was investigated using cytochemical techniques. Time-course cytochemical experiments, performed with glutaraldehyde-fixed cells of this organism,

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

25

FIG. 8. Details of methanol-limited chemostat grown cells of Hunsenulu polymorpha after time-dependent incubations with cerium chloride and methanol (a) and diaminobenzidine and hydrogen peroxide (b). Since early reaction products are distributed evenly over the peroxisomal matrix, staining of the organellesas a diffusion artefact is excluded indicating that alcohol oxidase and catalase activities are present throughout these organelles.

indicated that the activities of both enzymes were present throughout the peroxisomal matrix (Fig. 8a,b). Catalase could be completely removed from isolated crystalloids obtained by a short osmotic shock treatment of protoplasts of H. polymorpha. This procedure had virtually no effect on the integrity of the crystalloids. Since alcohol oxidase activity was still detected in such organelles, it was concluded that alcohol oxidase protein is the main structural element of the crystalloids contained in peroxisomes in methanolgrown cells of H . polymorpha (Veenhuis et al., 1979b, 1980a). The molecular architecture of the crystalloids was further investigated in ultrathin cryosections (Veenhuis et al., 1981b). As in Kloeckera sp. 2201 (Osumi et al., 1979) different regular substructures were observed in the peroxisomal matrix, depending on the plane of sectioning (Fig. 9a). These images were all caused by superposition of molecules arranged in a lattice. On the basis of morphological characteristics (Kato et al., 1976) these were identified as octameric alcohol oxidase molecules. The three-dimensional reconstruction of the crystalloids is based on the assumption that not all the molecules visualized in a cryosection (as shown in Fig. 9b,c) are located in the same plane. As can be deduced from the tilted part of this section, probably every second molecule is positioned in a plane below that of sectioning. The

26

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

molecules present in the plane of sectioning measure approximately 10 nm (100 A) in diameter and have a centre-to-centre distance of approximately 23.5 nm (235 A; Fig. 9c). The lattice structure represented by the molecules in the plane underneath that of sectioning of Fig. 9(b) is shown in Fig. 9(d). Compared to the arrangement of molecules in the plane of sectioning (Fig. 9c), in this layer the molecules are displaced at an angle of 45" along an axis perpendicular to the plane of sectioning. The centre-to-centre distance between the individual molecules present in this layer is approximately 14 nm (140 A). A schematic representation of the arrangement of molecules in the layers described above is shown in Fig. lO(a) and (b). In the vertical plane the crystalloid is built up of the two alternating layers of alcohol oxidase molecules. A three-dimensional reconstruction of this model is shown in Fig. lO(c). In a vertical section through this model the repeating unit of the crystalloid can be observed, which consists of four molecules of alcohol oxidase (Fig. 10d). The proposed model for crystalline alcohol oxidase in peroxisomes of methanol-grown H. polymorpha represents a very open structure. The molecular arrangement permits the presence of mobile catalase molecules. This was also indicated during reconstitution experiments, performed on isolated crystalloids obtained after short osmotic shock treatment of sphaeroplasts. The catalase-negative crystalloids could be impregnated with exogenously added catalase and with other proteins such as glucose oxidase and urate oxidase (Veenhuis et al., 1980~). These results may therefore explain the fast rate of leakage of catalase observed during fractionation experimentsand osmotic shock treatment. It may also add to a further understanding of other characteristic phenomena such as the fragility of the organelles during and after isolation or after cryosectioning of unfixed cells (M. Veenhuis, unpublished results) and the ease of deformation of the organelles, which is apparent in budding cells during the process of the separation of small peroxisomes (Veenhuis et al., 1978b). The main difference between the models proposed for the three-dimensional architecture of the crystalloids contained in peroxisomes of Kloeckera sp. 220 1 and H. polymorpha resides in the way in which catalase is thought to be incorporated into these structures. This interpretation hinges on the view taken of the nature of the particles observed in cryosections of both species which are located between every two adjacent alcohol oxidase molecules. In the Hansenula model this particle is thought to be an alcohol oxidase molecule, displaced over 45" along an axis perpendicular to the plane of sectioningand located in a plane underneath that of sectioning, whereas in the Kloeckera model it is suggested to represent a catalase molecule, present in the plane of sectioningbetween the alcohol oxidase molecules. In the latter model, the presence of one catalase molecule of approximately 7 nm (70 A) between

FIG. 9. Details of cryosections of glutaraldehyde-fixed cells of Hansenulapolymorpha, grown in a methanol-limited chemostat. (a) Shows part of a peroxisomal matrix, which is partly distorted during drying of the section. For this reason we can observe the main crystalline patterns in one section (arrows). These include the individual molecules, which are recognized in the central part, arranged in perpendicular lines in the horizontal and vertical directions. In addition, broader lines are observed running in two directions, perpendicular to each other (arrows). At a higher magnification of a similar section (b) it can be seen, in the tilted part of this section, that every second moleculein the micrograph isin a plane beneath the planeofsectioning (arrows). Adetail of (b) clearly demonstrating this phenomenon is shown in (c). When compared to the arrangement of molecules in the plane of sectioning (b), the molecules present in the plane underneath that of sectioning of (b) and (c) are displaced over an angle of 45" along an axis perpendicular to that of sectioning (c and d). From Veenhuis et al. (1981b).

28

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

FIG. 10. (a) and (b) are schematic drawings of the arrangement of alcohol oxidase molecules in the two alternating planes shown in Fig. 9(b) and 9(d). The bar represents 10 nm. From Veenhuis et a/. (1981b). (c) and (d) are models showing the three-dimensional arrangement of alcohol oxidase molecules in the crystalloid, composed of alternating layers of molecules as shown in (a) and (b). (d) Shows a vertical section through this model. In this section the repeating unit of the crystalloid, composed of four alcohol oxidase molecules (A-D) can be observed. The bar represents 10 nm. From Veenhuis et al. (1981b).

two adjacent alcohol oxidase molecules, whose centres are & 22 nm (220 A) apart (Osumi et al., 1979), implies a gap of 5 nm (50 A) in each direction of such an alcohol oxidase/catalase unit. Although this model, as also the model proposed for H. polymorpha, suffers from the imperfection that it is deduced from glutaraldehyde-fixed cells and therefore does not necessarily reflect the situation in uiuo, the data reported by Osumi et al. (1979) make it difficult to envisage how, in viuo, these structural alcohol oxidase/catalase units may interact t o stabilize the crystalloid. Recent investigations (M. Veenhuis, unpublished results) of methanol-grown Candida boidinii indicated that in this organism the molecular arrangement of the crystalloids is probably similar to that represented by the model proposed for H. polymorpha. As in H .

PEROXISOME METABOLISM

OF ONE-CARBON COMPOUNDS

29

FIG. 11. (a) Cryosection of a methanol-grown cell of Cundida boidinii, showing a detail of the peroxisomal matrix (cf. Fig. 9a-d). (b) Shows the detail of recrystallized alcohol oxidase, purified from methanol-grown Hunsenulu polymorphu, negatively stained with uranyl acetate.

polymorpha, liberation of peroxisomes from methanol-grown cells of this organism by means of osmotic shock treatment of sphaeroplasts, resulted in virtually intact crystalloids that were devoid of catalase activity. In ultrathin cryosections the substructure of these crystalloids, and also the substructure of peroxisomes in intact cells, were identical to the peroxisomal substructure observed in H . polymorpha (Fig. 1 la). Therefore in C . boidinii, and probably also in other methylotrophic yeasts as judged by virtually identical crystalline patterns of the peroxisomal matrices (Fukui et al., 1975a; Hazeu et al., 1975; Veenhuis et al., 1976), the crystalloids are composed of alcohol oxidase molecules, arranged in a manner similar to that described for H . polymorpha. Cytochemical staining experiments have shown that apart from alcohol oxidase and catalase other hydrogen peroxide-producing oxidases such as D-amino acid oxidase, uricase, L-u-hydroxy acid oxidase and amine oxidase are also present throughout the peroxisomal matrix (Fukui et al., 1975b; Veenhuis et al., 1976, 1981a; Zwart et al., 1980). However, their activities are very low compared to that of alcohol oxidase. Depending on their size and charge the molecules of these enzymes may be present in a mobile form, as suggested for catalase, or may be incorporated into the crystalloids. Recrystallization experiments in uitro, performed with alcohol oxidase purified from methanol-grown H . polymorpha, showed that the periodicity of these needle-shaped crystals, as observed in thin sections after fixation with glutaraldehyde/osmium tetroxide/potassium bischromate, was similar to that of the crystalline peroxisomes in intact cells of this organism. However, negative staining of the crystals revealed that their molecular organization greatly differed from that of the crystalloids of methanol-grown H . polymor-

30

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

pha (Fig. 11b). Therefore, molecular arrangements of alcohol oxidase other than that described for crystalline peroxisomes present in methanol-grown H. polymorpha and C. boidinii may also, permanently or temporarily, occur. What arrangement is adopted may, for example, depend on the environmental conditions.

111. Role of Peroxisomes in Metbylated Amine Metabolism

Soon after the discovery that peroxisomes may play a key role in the metabolism of certain carbon sources in yeasts (see Section 11) it was anticipated that these organelles may also be involved in the metabolism of various nitrogen compounds. In this section, evidence is presented that growth of yeasts on nitrogen sources which are metabolized via hydrogen peroxide-producing oxidases is indeed associated with the formation of peroxisomes in the cells. One particular example, namely the utilization of amines, will be considered in detail.

A . METHYLATED AMINES AS A NITROGEN SOURCE

Apart from the commonly used compounds such as ammonia, nitrate and urea, many yeasts are also capable of utilizing a variety of other compounds as nitrogen sources. Among these are D- and L-amino acids, purines and pyrimidines, monoamines such as methylamine, ethylamine and benzylamine, the diamines diaminoethane, diaminopropane, diaminopentane (cadaverine), polyamines such as spermine and spermidine, N-substituted amines, such as trimethylamine and dimethylamine, and various amides (i.e. acetamide; van der Walt, 1962; Brady, 1965; La Rue and Spencer, 1968; Yamada et al., 1965). Recently, van Dijken and Bos (1981) screened 461 strains of the yeast collection of the Centraalbureau voor Schimmelcultures (CBS) for their ability to use a number of different amines as sole carbon and energy source and/or nitrogen source for growth. None of the primary and methylated amines tested (methylamine, dimethylamine, trimethylamine, tetramethylammonium chloride, choline, ethylamine, propylamine, butylamine and benzylamine) could serve as a carbon and energy source. However, the majority of yeasts (86%) were able to use one or more of these compounds as a nitrogen source in the presence of glucose as the carbon source. Preliminary observations suggested that the ability to use each of these compounds as a nitrogen source was independent of the nature of the carbon source. When glucose was replaced by ethanol those strains capable of growth on ethanol

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

31

TABLE 4. Percentage of yeast strains showing special assimilation patterns of primary alkyl amines as a nitrogen source. Datacalculated from van Dijken and Bos (1981) Source Methylamine Ethylamine Propylamine Butylamine Percentage

-

-

+ -

other patterns Abbreviations:

+ +

-

+ +

-

+ +

14.5 52.7 18.8

14.0

+, growth; -, no growth. A total of 461 strains were tested.

showed the same utilization patterns for the amines tested as they displayed during growth on glucose. Yeast strains capable of using primary and N-substituted amines were to be found in almost all genera tested. Distinctive utilization patterns were observed. The results listed in Table 4 show that in yeasts the capacity to use one particular amine as a nitrogen source frequently coincided with the ability to use other amines as well. For instance, almost all strains capable of utilizing methylamine also utilized ethylamine but not vice Versa. This frequent capacity to utilize more than one primary amine is probably attributable to a broad substrate specificity of both the transport and oxidation systems for these structurally related compounds. B . PEROXISOMES A N D T H E METABOLISM OF M E T H Y L A T E D AMINES

The metabolism of methylated amines has mainly been studied in bacteria which are able to use these compounds as a carbon and energy source. They are metabolized by very diverse routes (Anthony, 1975; Colby et al., 1979; Large, 1981). The general pattern underlying the oxidation of methylated amines in micro-organisms involves the successive and eventually complete demethylation of quarternary, tertiary, secondary and primary amines, which at each stage yields formaldehyde from one of the methyl groups and produces a less substituted methylated amine. This process is represented by the following equation: (CHs),,N-4CH3)"- 1N +formaldehyde The formaldehyde formed from methylated amines is either oxidized to generate energy or assimilated to produce intermediates for growth. This further metabolism of formaldehyde is no different from that which occurs in organisms growing on methanol.

32

M. VEENHUIS, .I. P. VAN DIJKEN AND W. HARDER

In bacteria, several different enzymes have been found that catalyse the oxidation of methylated amines (Large, 1981). These include various mono-oxygenases, dehydrogenases and oxidases. The metabolism of methylamine in the fungus Trichosporon sp., an organism that can utilize this compound as a nitrogen source, has been investigated by Yamada et al. (1966). They showed that in this organism methylamine was metabolized by way of a primary amine oxidase which oxidized it to formaldehyde and ammonia. Methylamine was not a specific substrate for this enzyme which also catalysed the oxidation of various primary amines according to the equation:

Hz R-C-NH~

H

I

+H ~ +O02+R-C=O +H ~ +ON H~ ~

(9)

The activity of the enzyme towards alkylated amines decreased with increasing chain length of their molecules. A similar enzyme has recently been detected in a number of yeasts grown on trimethylamine, dimethylamine or methylamine as a nitrogen source. It has been purified from the methylotrophic yeast Candida boidinii and in many respects it resembles the enzyme present in Trichosporon sp. (Large et al., 1980). In addition, a second amine oxidase has been purified from C . boidinii. This enzyme, which is present in enhanced concentrations during growth of the organism on higher amines such as benzylamine, does not oxidize methylamine. Its activity towards alkylated amines increased with increasing chain length (Large, 1981) and in this respect resembled the amine oxidase which was purified from Aspergillus niger grown with n-butylamine as the nitrogen source (Yamada et al., 1965). A detailed study of methylamine metabolism in yeasts has recently been made with Hansenulapolymorpha and Candida utilis (Zwart et al., 1980).After transfer into glucose plus methylamine media of cells of C . utilis which had been grown on glucose plus ammonium sulphate, immediate synthesis of amine oxidase was observed in the lag phase preceeding growth (Fig. 12). The specific activity of this enzyme reached its maximum value in the mid exponential growth phase. Similar patterns of synthesis were observed for catalase and formaldehyde dehydrogenase (Fig. 12). The specific activity of formate dehydrogenase remained low during the first hours of cultivation after the transfer but increased during later stages of growth. The enzyme profiles obtained after transfer of cells into methylamine-containing media were largely similar in the two organisms studied. Transfer of H. polymorpha and C . utilis from glucose plus ammonium sulphate into glucose plus methylamine media was followed by ultrastructural changes in the cells (Zwart et a / . , 1980). In both organisms the synthesis of amine oxidase was accompanied by the development of large microbodies in

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

10 -

33

1

- 100

-

h

-

- -80

I

-I

.-c a

32-

.-c

-

+

E

9

P

e

-

I -_ -

-E0

-

E

-

a

0.5

-

-

E v

.'c

A -40 0 :: P

- 5

a

In

c -

0.1

-

I

-60

-

c

Q

v

IXI

.-c

e

-

Q

I

c .a c

?k

0)

L F

-7

-20

0 0

c 'E

-0

5

s

3

v

ea

I

n U

0"

z -0 0

4

12

8 Time ( h )

FIG. 12. Growth and enzyme profiles of Candida utilis in batch cultures containing glucose and methylamine. 0-0, Growth; A-A, amine oxidase; A-A, formaldehyde dehydrogenase; 0-0, formate dehydrogenase; 0-0, catalase. After Zwart et al. (1980).

TABLE 5. Number and volume fractions of peroxisomes present in glucose-, methanol- and methylamine-grown cells of Hansenulapolymorpha and Candida utilis Candida utilis

Growth conditions Glucose plus ammonium sulphate (A663 = 1 .O) Glucose plus methylamine ( A 6 6 3 = 1 .O) Glucose plus methylamine stationary growth phase Methanol plus ammonium sulphate stationary growth phase

Hansenula polymorpha

volume volume number fraction number fraction

0.1

0.3

0.15 3.0

0.04 0.4

0.09 2.3

2.7

9.3

0.9

5.2

2.2

29.8

-

-

The cells were grown on 0.5% methanol or 0.5% glucose, in the presence of 0.25% ammonium sulphate or 0.25% methylamine as the nitrogen source. The number of peroxisomes is expressed as the average per section, the volume fraction as percentage of the cytoplasmic volume.

34

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

FIG. 13. Ultrastructure of Candidu utilis cells, grown in batch cultures on media containing glucose plus methylamine. (a) A cell from the mid-exponential growth phase showing a number of cytoplasmic peroxisomes; (b) shows clusters of peroxisomes. In (c) elongated organelles are visible but after fixation with glutaraldehyde/ osmium tetroxide/potassium chromate the organelles contain no crystalline inclusions (d). The presence of amine oxidase activity in the peroxisomes is demonstrated after aerobic incubations with caesium chloride and methylamine (e) and catalase activity is demonstrated with diaminobenzidine and hydrogen peroxide (f).

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

35

the cells (Fig. 13a). The organelles varied in shape from spherical to highly elongated and in dimensions from 0.5 to 1.0 pm (Figs 13a,c). Generally, the organelles were scattered throughout the cytoplasm but occasionally small clusters of microbodies were also observed (Fig. 13b). Apart from the general absence of clusters of microbodies a number of other characteristic differences distinguished organisms grown on methylamine from those grown on methanol. Thus during exponential growth on methylamine the microbodies contained no crystalline inclusions (Fig. 13d). However, in cells of H . polymorpha harvested from stationary phase cultures, a small crystalloid had developed in the peroxisomal matrix due to the derepression of alcohol oxidase activity in these cells (Eggeling and Sahm, 1978; Egli et al., 1980). Again, apart from associations with the endoplasmic reticulum, the close association of microbodies with mitochondria in particular, as observed in serial sections (Zwart et al., 1980),was characteristic of organisms grown on methylamine. Finally, the total number and volume fraction of the microbodies were substantially less in cells grown on methylamine than in those grown on methanol (Table 5). The enzymic composition of the microbodies present in cells grown with methylamine as the nitrogen source has been studied by cytochemical methods (Zwart et al., 1980). Both cytochemical staining techniques for the demonstration of oxidase activities, namely the direct method involving incubation of glutaraldehyde-fixed sphaeroplasts with methylamine and cerium chloride under aerobic conditions, as well as the indirect technique involving incubation of whole cells with methylamine and diaminobenzidine, resulted in positively stained microbodies indicative of the presence of amine oxidase activity in these organelles (Fig. 13e). The presence of catalase activity in these organelles was also demonstrated after incubations with diaminobenzidine and exogenous hydrogen peroxide (Fig. 13f). These cytochemical experiments indicated that both amine oxidase and catalase were present in all of the microbodies contained in one cell. Therefore, these organelles can be considered as peroxisomes (de Duve, 1973). The above findings on the utilization of methylamine by H . polymorpha and C . utilis show that in these yeasts the metabolism of this compound is basically similar to that of methanol. The substrate is oxidized in peroxisomes and the first oxidation product-formaldehyde-is further metabolized in the cytosol by NAD dependent dehydrogenases (Fig. 14). The other product, ammonia, is used as a source of intracellular nitrogen. As already mentioned, primary amines cannot serve as a carbon and energy source for yeasts. It could be argued that this might be due to the fact that the aldehyde reaction product cannot be utilized as an (intracellular) carbon source. Although this may hold, for example, for the utilization of propylamine or butylamine, it cannot explain the inability of yeasts to utilize, for +-

rx

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

36 CHSNHZ

HzO

amine oxidose

NH,

0 2

H20

+

HCHO

n

formaldehyde dehydrogenose . , HO

NAD+

s mH

NADH

forrnote dehydrogenase

NAD+

COP

NADH

02

FIG. 14. Metabolism of methylamine by Hansenulapolymorpha and Candida boidinii.

example, ethylamine as carbon and energy source (van Dijken and Bos, 1981) since acetaldehyde is also an intermediate in the metabolism of ethanol which can be used by a variety of yeasts as a carbon source (Barnett et al., 1979). Thus transfer of an organism growing on ethanol plus ammonium sulphate to ethanol plus ethylamine would only require the additional synthesis of an amine transport carrier, amine oxidase and catalase. However, ethanollimited chemostat cultures of the yeast C. utilis growing with (excess) ethylamine metabolized the amine only to the extent that the nitrogen requirement of the culture, set by the reservoir concentration of ethanol, was met (K. B. Zwart, unpublished observations). Free ammonium ions were not detected in the culture supernatant indicating that ethylamine is primarily utilized for nitrogen-assimilatory purposes. Similar observations were made with C. utilis, grown on methylamine as the nitrogen source. The regulatory mechanism underlying this phenomenon has not yet been elucidated. It may be that the concentration of free ammonium ions which would be produced in excess when the amine is utilized as a carbon and/or energy source, is a regulating factor which controls their own production via inhibition and/or repression of the synthesis of the amine transport carrier and amine oxidase. The above examples clearly illustrate that peroxisomes may have an important function in the nitrogen metabolism of yeasts. Enhanced levels of specific oxidases and proliferation of peroxisomes during utilization of D-amino acids (D-amino acid oxidase) and uric acid (urate oxidase) have also been observed (K. B. Zwart, unpublished observations) supporting the hypothesis that peroxisomes may carry several specific oxidases involved in nitrogen metabolism. It is to be expected that in the near future the number of examples in which peroxisomes develop in fungi as a consequence of the need to utilize an unusual nitrogen source will be extended to utilization of diamines and polyamines which are generally metabolized via an oxidative attack by oxidases that produce hydrogen peroxide (Blaschko, 1963; Zeller, 1963).

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

37

C . PEROXISOMES INVOLVED IN THE CONCURRENT METABOLISM OF CARBON A N D NITROGEN SOURCES

In the previous paragraphs examples have been discussed in which peroxisomes serve only one function, namely either to furnish the cell with all of the carbon compounds required for growth or to provide ammonia from a single nitrogen source. However, conditions have recently been described (Veenhuis et al., 198la) in which cells of Hansenulapolymorpha depend on the activity of peroxisome-borne oxidases for both their carbon and nitrogen supply. The latter is the case during growth of this organism on methanol as the carbon source in the presence of methylamine, urate or D-alanine as the nitrogen source. After transfer of glucose-grown cells of H . polymorpha into media containing methanol and methylamine, large peroxisomes developed in the cells. Cytochemical staining experiments indicated that these organelles contained catalase but also alcohol oxidase and amine oxidase, the key enzymes in methanol and methylamine metabolism respectively (Fig. 15a,b). Furthermore, these enzymic activities were demonstrable in each individual peroxisome. Therefore, these organelles are involved in the concurrent oxidation of both the carbon and the nitrogen source during growth of H. polymorpha on methanol and methylamine. A schematic representation of the significance of these organelles in the initial oxidation of methanol and methylamine is given in Fig. 16. Peroxisomes similarly involved in the concurrent metabolism of methanol and urate have been described during

FIG. 15. Cells of Hansenula polymorpha, grown in batch culture in media containing methanol and methylamine. The peroxisomes present in these cells contain catalase activity (a; demonstrated with diaminobenzidine and hydrogen peroxide) and also amine oxidase activity (b; demonstrated with cerium chloride and methylamine).

38

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

TABLE 6. Specific activities of alcohol oxidase, amine oxidase and catalase, compared to the number of peroxisomes and their volume fractions during growth of Hansenulapolymorpha in a methanol-limited chemostat (D =0.088 h-l) with methylamine as the nitrogen source and after transfer to ammonium sulphate as the nitrogen source. After transfer to ammonium sulphate the cells were harvested after three volume changes Specific activities of

Growth conditions Methanol plus methylamine Methanol plus ammonium sulphate

Amine Volume fraction of: Number of Alcohol oxidase oxidase ( x lo3) Catalase peroxisomes Peroxisomes Vacuole 2.94

17.8

69.9

3.2

40.6

11.9

3.70

1.6

78.0

3.3

50.6

3.6

Oxidase activities are expressed as pnol 0 2 min-' (mg protein)-', catalase activity as AA2a min-' (mg protein)-'. The number of peroxisomes is given as the average number per section; volume fractions are expressed as percentage of the cytoplasmic volume.

sporulation experiments with cells of H. polymorphu (Veenhuis et ul., 1980b). Unexpectedly, as was shown in methanol-limited chemostat cultures, growth of H . polymorpha in the presence of methylamine as the nitrogen source resulted in a decrease in both alcohol oxidase and catalase activities in the cells, as compared to the specific activitiesof thosdenzymesin cells grown with

amino ocids nucleic ocids

FIG. 16. Schematic representation of the significance of peroxisomes in the initial oxidation of methanol and methylamine in Hansenula polymorpha. The products of these obligatory peroxisomal reactions are used for energy generation, carbon assimilation and nitrogen assimilation. From Veenhuis et al. (1981a).

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

39

ammonium sulphate. In spite of the presence of amine oxidase the volume fraction of the peroxisomes in methanol plus methylamine-grown cells was also significantly lower than in cells grown on methanol plus ammonium sulphate. This was also observed in transfer experiments performed in a methanol-limited chemostat culture (Table 6). Substitution of ammonium sulphate by methylamine resulted in a decrease of enzyme activities and was accompanied by a decrease of 20 to 30% in the peroxisomal volume fraction. After replacing methylamine again by ammonium sulphate the values originally found for alcohol oxidase and catalase activity and the volume fraction of the peroxisomes were re-established. The substructure of peroxisomes was identical in organisms grown in methanol plus ammonium sulphate and in methanol plus methylamine, namely partly crystalline in cells harvested from the exponential growth phase in batch cultures and completely crystalline in cells taken from chemostat culture (see also Section 1I.C). One major difference in cellular morphology was the presence of a large vacuole in methylamine-grown cells (Table 6). During growth on methanol plus methylamine, relatively high amounts of phosphate were present in this organelle. The physiological function of this has not yet been satisfactorily explained. It should again be stressed that although the oxidation of the carbon source and the nitrogen source proceeds via the same intermediate (formaldehyde) the amine is in this instance also only used as a nitrogen source (see also Section III.B, p. 36). The above example once more illustrates that the enzymic composition and function of the peroxisomes may be manipulated by changing the composition of the growth medium and it seems reasonable to postulate that a variety of oxidases acting on nitrogen compounds may be simultaneously active in peroxisomes along with alcohol oxidase. So far, yeasts capable of utilizing methanol as well as n-alkanes have not been found. Since these compounds are currently the only known carbon sources metabolized in yeasts by peroxisomal enzymes, it is at present not possible to define conditions that would lead to the formation of peroxisomes involved in the simultaneous metabolism of two carbon sources. As shown in Fig. 17, this restriction does not apply to the role of peroxisomes in nitrogen metabolism since various yeasts are able to utilize two or even three nitrogen compounds of different classes which are metabolized via oxidases that produce hydrogen peroxide. Here and in Section I1 the metabolic function of peroxisomes in yeasts has been considered in relation to the external environment. However, it should be recognized that yeast peroxisomes may also play a role in processes involving oxidases which yield hydrogen peroxide but which are not primarily influenced by the environment of the cell. For example, uric acid oxidation during purine turnover in the cell may occur independently of the nature of the carbon and nitrogen sources utilized for growth and it may well be that

40

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER SPECIES UTILIZING AMINES

f=i 7

\

SPECIES UTILIZING I D-AMINO ACIDS OR URIC ACID

N-ALKANE UTILIZING

EAST<

FIG. 17. Venn diagram illustrating the occurrence amongst yeasts of the capacity to utilize carbon and nitrogen sources which require the activity of peroxisomal oxidases. The figure is drawn on the basis of taxonomic studies of Lee and Komagata (1980) on methanol utilization, by van Dijken and Bos (1981) on methylamine utilization, by Bos (1975) on n-alkane utilization and by La Rue and Spencer (1968) on D-amino acid utilization.

peroxisomes play a key role in this process. In addition, a variety of polyamines such as putrescine, spermine and spermidine, have been found in all living cells investigated so far (see Gaugas, 1980) and the existence of a peroxisomal polyamine oxidase (Holltta, 1977) indicates that peroxisomes may be involved in regulating the concentrations of these biologically important amines. Finally, fatty acid oxidation, which can be provoked in certain yeasts by growing them on n-alkanes, may occur in peroxisomes of yeasts which cannot grow on n-alkanes under conditions that require lipid turnover in the cell. However, in contrast to the now well authenticated role of peroxisomes in enabling certain yeasts to utilize an enlarged range of exogenous sources of carbon and nitrogen, any role of these organelles in intracellular “turnover” has yet to be substantiated. IV. Biogewsis of Peroxisornes

In the previous sections it has been shown that in yeasts the occurrence of

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

41

microbodies and their enzymic composition may vary considerably during the different stages of culture growth and in response to the presence of specific carbon and/or nitrogen sources in the environment. This not only suggests possible control functions of peroxisomes in the metabolism of these cells, but also raises the question of how synthesis of peroxisomal enzymes is regulated and how biogenesis of these organelles is accomplished. These latter aspects are considered below in the case of the metabolism of one-carbon compounds in yeasts.

A. REGULATION OF THE SYNTHESIS OF PEROXISOMAL ENZYMES

The synthesis of enzymes involved in the oxidation of methanol and methylamine in yeasts in response to the presence of these compounds in the growth medium and the repression of their synthesis by glucose and ammonium sulphate, respectively, is well documented (Eggeling and Sahm, 1978, 1980; Egli, 1980; Zwart e t a / . , 1980). Detailed studies on the regulation of the synthesis of peroxisomal enzymes involved in the initial oxidation of methanol (alcohol oxidase and catalase) have mainly been conducted in Candida boidinii and Hansenula polymorpha. When glucose-grown cells of C . boidinii were transferred to a medium containing methanol as the sole carbon and energy source, the activities of alcohol oxidase and catalase increased significantly within five to eight hours (Kato eta/., 1974; see also Sahm, 1977; Fig. 18). These results have been interpreted to indicate that synthesis of these enzymes is induced by methanol and it has been reported that in glucosegrown C. boidinii cells approximately 100 mM methanol at a cell density of 3 mg dry weight ml-I is required for maximal induction of the two peroxisomal enzymes (Sahm, 1977). Induction of synthesis of alcohol oxidase and catalase in this organism was also observed in the presence of formaldehyde and formate, although the organism was unable to grow on these compounds. Furthermore, in this case, the extent of induction was dependent upon the initial concentration of the inducer compound. With formaldehyde, maximal activities were found at 1 to 2 mM whereas 20 to 30 mM formate was required for optimal induction. In addition, it was shown that glucose severely repressed the synthesis of these two enzymes. Thus, when C. boidinii was inoculated into a medium containing both glucose and methanol, methanol was not utilized until the glucose was exhausted and a diauxic growth curve was obtained (Sahm, 1979; Fig. 19). Work from Sahm’s laboratory (Eggeling and Sahm, 1978) with H. polymorpha showed that in this organism, in contrast to C. boidinii, alcohol oxidase and catalase may be formed in the absence of CI compounds. Significant amounts of these enzymes were detected in cultures growing

42

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

Time ( h )

FIG. 18. Changes in specific activities of the C1-oxidizingenzymes in Candidaboidinii after transfer of glucose-grown cells into a medium containing 300 mM methanol (W-W, catalase; 0-0, alcohol oxidase). Alcohol oxidase activity is expressed as pmol hydrogen peroxide produced min- (mg protein)-', catalase activity as pmol hydrogen peroxide decomposed min-' (mg protein)-'. From Sahm (1977).

exponentially on glycerol, ribose, xylose or sorbitol, but not on glucose. However, on exhaustion of glucose from the culture, synthesis of alcohol oxidase and catalase was observed. In contrast, during growth on ethanol, alcohol oxidase activity was not detected in the exponential or stationary growth phase. These results indicated that in this organism the synthesis of these enzymes may be subject to control by repression/derepression mechanisms rather than induction by C , compounds and catabolite repression. This possibility was investigated in detail by Egli (1980) and by Egli et al. (1980) in the yeasts H. polymorpha and Kloeckera sp. 2201 (=C.boidinii). When these organisms were grown in a glucose-limitedchemostat culture (i.e. in the absence of methanol!), derepression of synthesis of alcohol oxidase and

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

43

15

=

0.6

-

0.4

-

9

0.3

-

P

0

E L

2

10

20

30

40

50

60

70

Time ( h )

FIG. 19. Changes in specific activities of alcohol oxidase (m-m) and consumption of methanol (0-0) during growth (0-0) of Candid0 boidinii in media containing 0.5% glucose and 1% methanol. Enzyme units as given in Fig. 18. From Sahm (1977).

catalase was found at low dilution rates, although the extent to which derepression occurred was different for the different enzymes and organisms. In H. polymorpha alcohol oxidase activity in the cells increased with decreasing dilution rates (Fig. 20a). Full derepression was not observed; at low dilution rates cells of glucose-limited chemostat cultures contained only 10 to 20% of the alcohol oxidase found in methanol-limited cells grown at the same dilution rate (compare van Dijken, 1976). The activity of catalase also increased with decreasing dilution rates (Fig. 20b) up to values of 30 to 40% of that typical of cells grown under methanol-limitation. Similar results were obtained with glucose-limited cultures of Kloeckera sp. 2201 (Fig. 20c,d) with one major exception: the repression of alcohol oxidase in this organism was only relieved at dilution rates below 0.1 h-' and to a much smaller extent than in H. polymorpha. The extent of repression of synthesisof alcohol oxidase in both organisms is most likely a function of the residual glucose concentration in the culture. This concentration was estimated for both organisms at different dilution rates and was found to fit the equation:

S

= Ks. W m a x - D)

in which Ks and pmax were 15mg I-' and 0.51 h-' for H. polymorpha and 35 mg 1-' and 0.42 h-' for Kloeckera sp. 2201. These data show that at a certain

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

44

0

0.1

0.2

0.3

0.4

0.5

0

0.1

0.2

0.3

0.4

0.5

Dilution rate (h-')

FIG. 20. Alcohol oxidase (pmol 02 consumed min-' (mg protein)-') and catalase (A& min-' (mg protein)-') activities in Hansenula polymorpha and Kloeckera sp. 2201, grown in glucose-limited chemostats, as a function of the dilution rate; (a) and (c) alcohol oxidase and catalase in Hansenulapolyrnorpha; (b) and (d) alcohol oxidase and catalase in Kloeckera sp. 2201. From Egli et nf. (1980).

dilution rate the residual glucose concentration in the Kloeckera cultures was significantly higher than in the Hansenula culture. This observation may explain how it was that in the Kloeckera culture derepression of synthesis of alcohol oxidase was accomplished only at much lower dilution rates than in H. polymorpha. Synthesis of alcohol oxidase and catalase in the two organisms was also investigated during growth on mixtures of methanol and glucose in continuous culture. Essentially similar results were obtained (Fig. 2 1a,b,d,e), except that the rate of production of alcohol oxidase in H. polymorpha was significantly increased under these conditions (Fig. 2 lc,f). The above observations and those of Eggeling and Sahm (1980) have shown that in at least two methanol-assimilating yeasts the peroxisomal enzymes involved in methanol oxidation may be synthesized in the absence of methanol. This must be considered as a case of gratuitous enzyme synthesis because under the conditions of cultivation these enzymes have no apparent

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

45

2.5r

50

,c3

%

in P

0

0

s

;-.^i_ .*

30

0

.'

0

10

V

0 0

0.1

0.2

0.3

Dilution rate (h-')

0.4

0

0.1

0.2

0.3

0.4

Dilution rate (h-')

FIG. 21. Enzyme profiles and productivity of alcohol oxidase and catalase in Hansenula polymorpha and Kloeckera sp. 2201, grown in chemostat cultures on methanol and glucose (Cl :C6= 62.2%: 38.8%, w/w), as a function of the dilution rate. (a), (b) and (c) show alcohol oxidase, catalase and productivity of alcohol oxidase in Hamenula polymorpha. (d), (e) and (f) show alcohol oxidase, catalase and productivity of alcohol oxidase in Kloeckera sp. 2201. In (c) and (f) . . . . , . indicates productivity of alcohol oxidase during growth on methanol as the sole source of carbon. Productivity is expressed as units (pmol min-' mg-' h-l), alcohol oxidase as pmol oxygen consumed min-' (mg protein)-' and catalase activity as AA24 min-' (mg protein)-'. From Egli (1980).

physiological function (Eggeling and Sahm, 1980). The kinetics of the observed derepression of the synthesis of alcohol oxidase and catalase in the two organisms when grown at low growth rates in glucose-limited chemostat cultures indicate that the extent to which this derepression is expressed depends on the concentration of glucose in the culture (Egli et af., 1980). It may be envisaged that this exogenous glucose in turn is in equilibrium with intracellular metabolites derived fi;om glucose which initiate the onset of

46

M. VEENHUIS. J. P. VAN DIJKEN AND W. HARDER

catabolite repression of peroxisomal enzymes. The nature of these catabolites is as yet unknown but it seems likely that similar, if not identical, metabolites are involved which can be derived from a variety of substrates (i.e. glucose, ribitol, glycerol or ethanol). The intracellular concentrations of these metabolites are probably determined by the nature of the carbon source. Whereas during growth on glycerol, ribose, xylose and sorbitol alcohol oxidase is already synthesized during exponential growth (high substrate concentrations), derepression in glucose-containingmedia is only observed at low glucose concentrations (i.e. in stationary growth phase in batch cultures or in carbon-limited chemostat cultures). Ethanol exerts an even stronger repression and is also able to repress alcohol oxidase synthesis under carbon-limitingconditions. The intracellular concentration of the repressor of alcohol oxidase synthesis may additionally be determined by the availability of intracellular energy. Thus addition of methanol to a glucose- or glycerollimited culture of these yeasts may enhance the rate of synthesis of alcohol oxidase through a decrease in concentration of the intracellular repressor. This may be attributable to the fact that extra energy is generated from the methanol which could enhance the flow of intermediates into the direction of biosynthesis, thereby lowering the concentration of the intracellular pools. Lower pool concentrations of intracellular metabolites may also be present during growth of H. polymorpha on glycerol, ribose and xylose. An essentially similar model has recently been proposed for the regulation (by repression/ derepression) of the synthesis of Calvin cycle enzymes in a facultatively autotrophic bacterium (Dijkhuizen and Harder, 1979). Very little is known about the regulation of the synthesis of other peroxisomal enzymes in yeasts. Several hydrogen peroxide-producing enzymes involved in the metabolism of nitrogen sources such as amine oxidase, D-amino acid oxidase and urate oxidase are synthesized when methylamine, D-alanine or uric acid, respectively, is present as the sole source of nitrogen for growth (Zwart et al., 1980;Veenhuis et al., 1981a; K. B. Zwart, unpublished findings). Synthesis of these enzymes also occurs during growth of C. boidinii or H. polymorpha on these compounds in the presence of glucose as the carbon source. This indicates that their synthesis is not subject to catabolite repression by glucose, although it is severely repressed by ammonium ions. The regulation of the synthesis of catalase in yeasts is currently obscure. Although this enzyme is invariably present in yeast peroxisomes along with oxidases, its synthesis is probably controlled in a rather more complex way than that of the oxidases described above. There is clearly a need for further work on the regulation of the synthesis of this enzyme. It appears that a unifying model which explains the regulation of peroxisomal enzyme synthesis in yeasts is not available at present. Such a

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

47

model must also explain the commonly made observation that the synthesis of the different peroxisomal enzymes is not co-ordinately regulated. There is the general observation, however, that in yeasts the synthesis of hydrogen peroxide-producing oxidases involved in carbon and/or nitrogen metabolism is associated with an increase in the volume fraction of peroxisomes in these cells (Veenhuis et af., 1979a; Egli et af., 1980; Zwart et al., 1980). This is considered in more detail below.

B. DEVELOPMENT OF PEROXISOMES D U R I N G VEGETATIVE GROWTH

In Section 1II.B evidence has been presented that the volume fraction of peroxisomes in glucose-grown methylotrophic yeasts is very low. This poses the question of their development during adaptation of the cells to growth on methanol. In mammalian cells peroxisomes are believed to develop as outgrowths of strands of smooth endoplasmic reticulum (de Duve, 1973; Masters and Holmes, 1977; Goldman and Blobel, 1978). However, the recent models proposed for the biogenesis of mammalian peroxisomes cannot explain the observations made in the case of yeast peroxisomes. Most experimental evidence on biogenesis and multiplication of peroxisomes during methylotrophic growth of yeasts indicates that these organelles develop from pre-existing ones and multiply by separation of small peroxisomes from mature organelles (Veenhuis et al., 1978b, 1979a). Similar observations have been made with n-alkane grown yeasts (Osumi et af.,1974, 1975). In methylotrophic yeasts peroxisomes generally develop in response to changes in the environment of the cells. This is the case for peroxisomes which are metabolically active and contain enzymes which are involved in the oxidative metabolism of the carbon source (Roggenkamp et al., 1975; Fukui et al., 1975b; van Dijken, 1976), the nitrogen source (Zwart et al., 1980) or both the carbon and the nitrogen sources (Veenhuis et al., 1981a). In the yeast Hansenula polymorpha peroxisomes having no apparent physiological function also develop in cells grown under conditions in which the synthesis of alcohol oxidase is partly derepressed (see Section IV.A, p. 42). The biogenesis of peroxisomes in methylotrophic yeasts under the different conditions mentioned above has been studied in detail in H. polymorpha after transfer of glucose-grown cells of this organism into media containing different sources of carbon and nitrogen. For use as an inoculum in these experiments cells were taken from cultures growing exponentially on glucose. These cells contained low levels of catalase activity and no alcohol oxidase activity (Veenhuis et al., 1979a). Ultrastructural observations showed that besides the usual cell organelles, generally one small peroxisome, irregular in

48

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

shape with dimensions of 0.1-0.2 pm and located in close proximity to the cell wall, was also present in these cells. This organelle contained no crystalline inclusions and was associated with strands of endoplasmic reticulum (Fig. 22a). Transfer of such cells into media containing methanol as the sole source of carbon caused an increase in the size of the small peroxisome originally present in the cells (Fig. 22b). The increase in size of this organelle, which commenced in the lag occurring after transfer, was associated with the development of a crystalline nucleus in the peroxisomal matrix approximately two hours after the transfer (Fig. 22c). The development of a crystalloid was strictly associated with the synthesisof alcohol oxidase in the cells (Fig. 22c,d). De n o m synthesis of peroxisomes in cells of H. polymorpha, transferred from glucose to methanol-containing media, was not observed.De no00 synthesisof

FIG. 22. Details of a cell of Hansenulupolymorpha: (a) from the exponential phase of batch culture growth on glucose, demonstrating the peroxisome present in such cells (compare Fig. 1); (b) four hours after transfer of the cells from glucose into methanol-containing media, demonstrating the increase in size of the peroxisome, originally present in the glucose-grown cell; (c) two hours after transfer from glucose to methanol, showing a positively stained peroxisome after aerobic incubation with cerium chloride and methanol. Note the presence of a small crystalloid in the matrix of this organelle; (d) six hours after transfer from glucose to methanol, demonstrating that the increase in size of the organelles is associated with an increase in size of the peroxisomal crystalloid.

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

49

peroxisomes after transfer of a number of methylotrophic yeasts, including three strains of Candida boidinii, into media containing methanol as the carbon source has been suggested by Meisel et al. (I 978). The organelles were reported to derive from the endoplasmic reticulum or elements of the GoIgi apparatus. In addition, peroxisomes also were thought to arise fyom reorganized mitochondria. Tsubouchi et al. (1976) also suggested de nouo synthesis of peroxisomes in Candida sp. N 16. In the lag phase occurring after transfer of cells from glucose into methanol-containing media amorphous bodies were observed in the cells, which subsequently turned into peroxisomes after being surrounded by a membrane derived from the endoplasmic reticulum. Such observations have not been made in cells of H . polymorpha. In the case of this organism transfer of cells to methylotrophic growth conditions is generally not associated with the proliferation of a well-developed Golgi-complex; in C. boidinii, however, this has been infrequently observed (Fig. 23a), but associations of clusters of microbodies with the Golgi apparatus have not been found. Although the results obtained with H. polymorpha, transferred from glucose to methanol, do not exclude de nouo synthesis of peroxisomes, our findings strongly suggest that the development of these organelles is exclusively due to growth and development of the peroxisome originally present in the glucose-grown cells. This view is based on the following observations: (a) during the first hours after transfer to methanol generally only one large peroxisome developed in the cells, and this contained a crystalline nucleus. In any individual cell such an organelle was never observed together with the peroxisome typical of growth on glucose; (b) degradation of the “glucose”-peroxisome has not been observed. This is in agreement with biochemical findings, which indicated that specific activities of

FIG. 23. Details of methanol-grown cells of (a) Candida boidinii, showing a well-developed Golgi apparatus and (b) Hansenula polymorpha, showing positively stained peroxisomes after aerobic incubation with cerium chloride and D-alanine, indicating the presence of D-amino acid oxidase activity in these organelles.

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

50

different oxidases present in these organelles, namely urate oxidase, L-ahydroxy acid oxidase and D-amino acid oxidase, remained approximately constant during the first hours of cultivation (Fig. 24; Veenhuis et a/., 1979a). A similar observation has been made with cells of Kloeckera sp. 2201 after transfer from glucose to methanol (Tanaka et al., 1976); (c) cytochemical experiments revealed that the enzymes, mentioned above, were now exclusively present in the newly developed organelle (Fig. 23b), together with alcohol oxidase activity (Veenhuis et al., 1979a). Taken together these results clearly indicate that peroxisomes that develop in H. polymorpha after transfer of cells from glucose- into methanol-containing media originate from the organelles originally present in the glucose-grown cells.

t

n

T"

(b)

I 100

50

6

-d

0

0

2

4

Time ( h )

6

FIG. 24. Growth and enzyme profiles in Hansenula polymorpha, grown in batch cultures on 0.5% methanol. The cultures were inoculated with cells from the mid-exponential growth phase of a culture growing on 0.25%glucose (A663 = 1 .O). (a) 0-0, growth; (b) t - m , alcohol oxidase; 0-0, catalase; -0 L-ahydroxy acid oxidase; A-A, D-amino acid oxidase; 0-0, urate oxidase. From Veenhuis ef al. (1979a).

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

51

During prolonged cultivation on methanol the number of peroxisomes per cell gradually increased by the separation of small peroxisomes from mature organelles. Such a gradually increasing number of peroxisomes was not observed in Kloeckera sp. 2201 (Tanaka et al., 1976). During the first four hours of cultivation after transfer of this organism from glucose to methanol generally five to six small peroxisomes developed per cell. This number remained constant during prolonged cultivation whereas the volume fraction of the peroxisomes increased as a result of growth of individual organelles (Tanaka et al., 1976). In glucose-grown cells of Kloeckera sp. 2201 peroxisomes were rarely detected and, although the authors could not demonstrate the origin of the peroxisomes, it was suggested that as in H. polymorpha they developed from a pre-existing one (Tanaka et ul., 1976).

FIG. 25. (a) and (b) Cells of Hunsenula polymorpha, taken from cultures growing exponentially on methanol, demonstrating different stages of peroxisomal division in the neck between mother cell and bud. From Veenhuis et al. (1978b).

During vegetative reproduction of methanol-grown H. polymorpha, peroxisomes have frequently been observed in developing buds already in a very early stage of bud formation. Similar to the process for the multiplication of peroxisomes in vegetative cells, these organelles originate from peroxisomes present in the mother cell. Separation of these small peroxisomes may occur in the mother cell, followed by a subsequent migration of the organellesinto the bud, but it also occurred in the neck between mother cell and bud (Fig. 25a,b). The movement of small peroxisomes into developing buds seems to be a well-controlled process which, among other factors, depends on cultivation conditions and the methanol concentration in the culture. The following example in which methanol-grown cells of H. polymorpha from the stationary phase of batch culture were inoculated into media containing 0.4% methanol, may serve to illustrate this. In the cells introduced as the inoculum, five to ten large peroxisomes were present, together representing about 28% of the

52

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

cytoplasmicvolume. However, in the first developing buds generally only one or two small peroxisomes were found. The volume fraction of peroxisomes in this first generation of cells amounted to about 0.4%, and was therefore less than 2% of the total peroxisomal volume in the inoculum cells. The movement of peroxisomes into the newly developing buds after transfer of cells into a fresh methanol medium appeared to be independent of the conditions of growth of the inoculum cells and was therefore independent of the amount of alcohol oxidase and catalase present in these cells. For instance, when cells pre-grown on glucose, glycerol or dihydroxyacetone were transferred to a fresh methanol medium, the first developing buds generally contained only one or two small peroxisomes. In the late exponential growth phase on methanol the number of peroxisomes present in the buds had increased to two to four, whereas in cells from a chemostat culture, especially those obtained at low dilution rates, five to seven peroxisomes were generally observed in developing buds. The substructure, ultimate shape and number of the organelles in methanol-grown H. polymorpha were all dependent upon the growth conditions and were generally correlated with the concentrations of alcohol oxidase and catalase in the cells (Veenhuis et af.,1978b, 1979a;see also Section II.C, p. 16-18). Furthermore, differences in morphology of peroxisomes may reflect differences in the stage of development of the organelles; the mean dimensions of mature peroxisomes varied from 0.9 to 1.3 pm. Recent investigations have shown that the biogenesis and subsequent development in cells of H. polymorpha of peroxisomes involved in nitrogen metabolism (see Section III.B, p. 32) or in simultaneous carbon and nitrogen metabolism (see Section III.D, p. 37), also the development of peroxisomes whose physiological function is uncertain, were similar to that described above for the development of peroxisomes in cells transferred from glucose to methanol (Zwart et al., 1980; Egli et al., 1980; Veenhuis et al., 1981a). However, under the different conditions employed, the presence of crystalline inclusions in peroxisomes contained in cells of H. polymorpha was strictly correlated with the occurrence of alcohol oxidase in the cells. The development of peroxisomes from pre-existing organelles in response to changes in the environment of the organism was also nicely demonstrated in cells of H. polymorpha, grown in a chemostat on methanol plus methylamine and transferred into glucose plus methylamine-containingbatch cultures. Owing to the presence of excess glucose, the crystalline peroxisomes in the cells were subject to degradation (Veenhuis et al., 198la; see also Section V, p. 64), resulting in a decrease of both alcohol oxidase and amine oxidase activities in the culture. However, along with an increase of amine oxidase activity, which was observed after two hours of cultivation (see Fig. 34), new peroxisomes developed which clearly originated from crystalline organelles

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

53

that had escaped degradation (Veenhuis et af., 1981a). These newly formed organelles mainly contained amine oxidase and catalase. Thus, peroxisomes present in cells of the yeast H . polymorpha, and probably also in other methylotrophic yeasts, originate from pre-existing organelles and develop by means of growth and division. Their ultimate shape, number, substructure and physiological function depend on the conditions prevailing during growth.

C. DEVELOPMENT AND FUNCTION OF PEROXISOMES DURING SPORE FORMATION A N D GERMINATION

The development of peroxisomes during generative reproduction of H . polymorpha on different sources of carbon in the presence of urate as nitrogen source has also been studied (Veenhuis et al., 1980b). It was observed that the development of peroxisomes in ascospores of H . polymorpha proceeded along identical lines as the development of these organelles during vegetative reproduction of haploid cells (Veenhuis et al., 1978b), irrespective of the sporulation conditions employed. As in vegetative cells, peroxisomes in ascospores of H . polymorpha originated from existing organelles (Fig. 26a) in the developing ascus. Furthermore, their volume fraction, ultrastructure and enzymic composition depended on the cultivation conditions maintained during vegetative growth. In cells grown on glucose and urate, in which the synthesis of alcohol oxidase is repressed, one small peroxisome was usually observed in each mature spore. These organelles were spherical in shape with dimensions of 0.24.3 pm (Fig. 26b). They had no crystalline inclusions and besides catalase also contained D-amino acid oxidase and urate oxidase (Fig. 26c). However, when vegetative cells had been grown under conditions in which the synthesis of alcohol oxidase was derepressed (namely during growth on methanol, glycerol and dihydroxyacetone media and on malt agar) the ascospores generally contained one to three peroxisomes which were spherical or cuboid with dimensions of 0.4-0.5 pm (Fig. 26d). These organelles showed a crystalline substructure (Fig. 26e) and, besides the enzymes present in peroxisomes of repressed cells, also contained alcohol oxidase (Fig. 26f). As in vegetative cells, the peroxisomes present in ascospores of H . polymorpha occurred in close association with strands of endoplasmic reticulum under all sporulation conditions employed. The peroxisomes present in young asci of H . polymorpha had no apparent physiological function in the process of ascosporogenesis as has been described for microbodies present in both sexual and asexual spores of higher fungi (Hess and Weber, 1974; Maxwell et al., 1977).Germination experiments indicated that mature ascospores of H . polymorpha cannot use their

FIG. 26. Electron micrographs of asci and ascospores from Hansenula polymorpha. From Veenhuis et al. (1980b). (a) Young ascus of H. polymorpha, developed in a methanol plus urate medium showing spore initials with incomplete prospore walls. In one spore initial (left below, arrow) a nucleus and a peroxisome are already present. (b) and (c) Survqy of part of an ascus of H . polymorpha, developed in glucose plus urate medium, showing the typical hat-shaped ascospores of this organism. In each spore a small peroxisome is visible (b). In these organelles D-amino acid oxidase activity was demonstrated after aerobic incubation with diaminobenzidine and D-alanine (c). (d) Ascospores of H. polymorpha that developed in a methanol plus urate medium. These generally contained more than one large peroxisome. (e) Shows ascospores developed on malt agar. The peroxisome has a crystalline substructure and contains alcohol oxidase activity (f) as demonstrated with diaminobenzidine and methanol.

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

55

endogenous reserves as a source of carbon and energy or as a nitrogen source. During germination a glyoxysomal function of the organelles in the spores, as proposed in developing seeds (Gerhardt, 1973) or zoospores of various fungi (Chong and Barr, 1973; Held, 1975; Powell, 1976), was not observed. The germination experiments showed that the function of the peroxisomes present in ascospores of H . polymorpha was identical to that in vegetative cells. They were involved in the oxidative metabolism of different carbon and nitrogen sources. Their enzymic profile, which is identical to that of peroxisomes in the vegetative cells, is rapidly adjusted to changes in the environment during germination.

D . ASSEMBLAGE OF PEROXISOMES

Very little is known about the various processes involved in the biogenesisand development of peroxisomes in methylotrophic yeasts. Most of our knowledge has come from studies of mammalian and plant cells. It is generally accepted that the peroxisomes in these tissues do not contain DNA or ribosomes (Lord, 1980; but see also Osumi and Sato, 1978) and hence their development cannot be regarded as an autonomous or semi-autonomous process. In both mammalian and plant cells peroxisomes were believed to develop as outgrowths of smooth endoplasmic reticulum (de Duve and Baudhuin, 1966; Masters and Holmes, 1977; Lord and Roberts, 1980). However, recent investigations showed that the classical model of de Duve and Baudhuin (1966) which proposed that peroxisomal proteins are synthesized on membrane-bound ribosomes (rough endoplasmic reticulum), subsequently translocated in the lumen of the endoplasmic reticulum and transported into the developingperoxisome, is not generally valid. It has been shown that at least some of the enzymes present in the peroxisomalmatrix, for example catalase and uricase (Goldman and Blobel, 1978; Robbi and Lazarow, 1978), are not synthesized on the endoplasmic reticulum but on free cytoplasmic polysomes. Also certain matrix proteins of glyoxysomes which develop in germinating plant seeds, i.e. catalase and isocitrate lyase, are synthesized on free polysomes (Lord and Roberts, 1980) and translocated across the glyoxysomal membrane after their synthesis has been completed. As summarized (see Section IV.C), peroxisomes that develop in methylotrophic yeasts originate from pre-existing organelles. Recently, Lazarow (1980a) suggested that rat-liver peroxisomes may develop in a similar fashion. However, so far, experimental evidence is lacking regarding the synthesis of the peroxisomal membrane or the synthesis, subsequent transport and activation of the main peroxisomal matrix proteins in methanol-grown cells, catalase and alcohol oxidase. It is generally considered that in methylotrophic

56

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

yeasts, as in germinating plant seeds (Lord and Roberts, 1980), the peroxisomal membrane is most probably derived from the endoplasmic reticulum. However, Lazarow et al. (1979) recently showed that the protein composition of the membranes of peroxisomes in rat liver cells differed greatly from that of the endoplasmic reticulum. Such a difference might also be expected in methanol-grown yeast cells as judged by the great difference in membrane structure of peroxisome membranes and endoplasmic reticulum as has been observed after freeze-etch procedures (Veenhuis et al., 1976). With respect to the classical model of peroxisomal biogenesis, a major problem is encountered in the methylotrophic yeast Hansenula polymorpha, namely the apparent absence of membrane-bound ribosomes (rough endoplasmic reticulum; Fig. 27). The absence of this rough endoplasmic reticulum during growth of this yeast on methanol eliminates the possibility of translocation of peroxisomal matrix proteins by a cotranslational ribosomemediated process. However, the close associations of strands of endoplasmic

FIG.27. Detail of a sphaeroplast prepared from a methanol-grown cell of Humenulu polymorphu. Ribosomes are not observed in the peroxisomal matrix. Ribosomes are also not attached to the peroxisomal membrane or at the endoplasmic reticulum, visible in this section (arrow).

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

57

reticulum with developing peroxisomes invariably observed in methylotrophic yeasts, suggests some functional relationship. Since ribosomes are absent from the peroxisomal matrix and do not occur attached to the peroxisomal membrane (Fig. 27), apparently two possibilities remain for synthesis and transport of the main peroxisomal matrix proteins. The first includes synthesis of these enzymes on cytoplasmic polysomes, which are then discharged into the lumen of the endoplasmic reticulum and subsequently transported via reticular channels to their target, the developing peroxisome(s), in a manner similar, for instance, to the transport of hydrolases and secretory glycoproteins mediated by the endoplasmic reticulum in yeasts (van Rijn et al., 1975; Cortat el al., 1973). Although direct membrane continuity or open connections between endoplasmic reticulum and peroxisomes have never been encountered in methanol-grown cells of this organism, it cannot be ruled out that the translocated proteins are released into the peroxisome by temporary connnections of the endoplasmic reticulum and the organelles. Peroxisomes may mature in a relatively short time and may then lose their capacity for import of matrix proteins. It is not unlikely that this process of maturation is associated with the loosening of their association with the endoplasmic reticulum. In this respect it is of interest to note that such associationshave not been observed with the cuboid mature peroxisomescontained in old cells of H. polymorpha grown under methanol-limitation in the chemostat (see Section II.C, p. 17). The second possibility involves a post-translational membrane translocation of the matrix proteins after their synthesis on free polysomes. Such a vectorial translocation of proteins into peroxisomes may require “signal” or “leader” sequences in the polypeptide chains newly synthesized by cytoplasmic polysomes. In addition, specific receptor proteins may be present in the peroxisomal membrane, which guide the appropriate proteins to the correct membranes as in mitochondria and chloroplasts (Chua and Schmidt, 1979; Sehatz, 1979; Neupert and Schatz, 1981) and perhaps also an adenosine triphosphatase which may supply the energy required for the translocation of these proteins across the membrane. This may of course also occur after transport of proteins via reticular channels. It must be stressed that activities of the major peroxisomal enzymes outside these organelles have so far not been detected by cytochemical methods. If one of the above models is correct, this failure to detect peroxisomal enzyme activities in the cytosol of yeastsmay be explained by the fact that the transport system of these proteins is so effective that the concentration of enzymes in the cytosol is kept very low (in fact below the limit of detection) or, alternatively, that these enzymes are activated inside the peroxisomes. The latter mechanism has been described in the case of catalase in peroxisomes of rat liver cells (Lazarow and de Duve, 1973; Lazarow, 1978,1980b). These authors suggested that activation of this

58

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

enzyme, i.e. the addition of the haem-group to the catalase apo-monomer and the subsequent formation of the tetramer, occurs inside the peroxisomes.

V. Inactivation of Peroxisomal Enzymes and Degradation of Peroxisomes In methanol-grown cells of H. polymorpha the activity of alcohol oxidase that can be detected by cytochemical methods is strictly confined to peroxisomes (Veenhuis et al., 1978b, 1979a). However, biochemical and cytochemical data have been obtained which indicate that this enzyme is not invariably present in an active form in these organelles(Veenhuis et al., 1980d). In methanol-grown yeasts different modes of regulation of alcohol oxidase activity must exist. For instance, during bud formation a temporary inactivation of enzyme activity has been observed (Veenhuis et al., 1978b) while after transfer of cells into glucose-containingmedia in which the activity of alcohol oxidase is no longer required for growth, the subsequent inactivation of alcohol oxidase is irreversible (Bormann and Sahm, 1978; Veenhuis et af.,1978a). These aspects will be considered in detail below after a general discussion of inactivation as a mechanism for the regulation of enzyme activity.

A . REGULATION OF ENZYME ACTIVITY BY INACTIVATION

In recent years it has become appreciated that, apart from control of enzyme levels through regulation of the rate of enzyme synthesis and control of enzyme activity via non-covalent binding of ligands, organisms may also regulate enzyme activities via selective inactivation. In his review on this topic, Switzer (1977) recognized two types of inactivation, namely modiJication inactivation, in which the enzyme protein remains intact but loses activity due to a change in its physical state or to the attachment of a covalent modifying group, and degradatiue inactivation, in which at least one of the peptide bonds is cleaved as part of the inactivation process or subsequently to it. Protein turnover (defined as a total hydrolysis of the enzyme to the individual amino acids) may or may not follow this type of inactivation. Inactivation of enzymes as a means of regulation of their activity is a widespread phenomenon (Switzer, 1977;Holzer, 1978; Wolf, 1980).Generally the mechanism of inactivation is poorly understood and in most cases it is not known whether turnover of enzyme protein is involved. Inactivation of enzymesmay be a continuous process. There is a wealth of data, mainly based on isotope studies, indicating the general occurrence of protein turnover in animals (Schimke, 1973; Segal, 1976; Goldberg and Dice, 1974; Goldberg and

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

59

St. John, 1976) a process also occurring continuously in growing bacteria (Holzer et ul., 1975). In micro-organisms, inactivation of enzymes may also occur rapidly when they are exposed to a changed environment that no longer requires the activity of these proteins for growth. Holzer (1976) has termed this phenomenon “catabolite inactivation” which includes both reversible and irreversible processes. For example, in yeasts, cytoplasmic malate dehydrogenase, fructose 1,6-bisphosphataseYphosphoenolpyruvate carboxykinase and a-isopropylmalate synthase are rapidly inactivated following transfer of cells from acetate to glucose (Holzer, 1976). Very recently, Miiller and Holzer (1981) showed that the initial reversible inactivation of fructose 1,6-bisphosphatase was due to phosphorylation of serine residues in the enzyme protein. This phosphorylated enzyme was subsequently attacked by proteolytic enzymes. Selective protein inactivation has also been observed after transition of yeast cells from vegetative growth to spore formation (Betz and Weiser, 1976a). In most cases it is not known whether these phenomena, which also occur in bacteria, include protein turnover. Proteolysis may be involved in spore formation in yeasts since the activities of proteinase A and B drastically increased after transfer of cells into the sporulation medium (Betz and Weiser, 1976b). In some cases, evidence has been obtained that reappearance of activity of the inactivated enzyme is dependent on de novo synthesis of proteins (Gorts, 1969; Duntze et ul., 1968; Gancedo, 1971; Gancedo and Schwermann, 1976). More substantial evidence that protein degradation can underly a sudden inactivation has been obtained recently by immunochemical methods. Disappearance of fructose 1,6-bisphosphataseand malate dehydrogenase activities of acetate-grown Succhuromyces cerevisiue on transfer to glucose and of NADP-dependent glutamate dehydrogenase activity in Cundidu utilis after exhaustion of the carbon source is paralleled by the irreversible loss of enzyme antigen able to cross-react with specific antisera raised against these enzymes (Neeff et ul., 1978; Hemmings, 1978; Funayama et al., 1980). The mechanism of this process in uiuo, which apparently involves proteolysis, is not yet known. In yeasts, the vacuoles are the principal cellular compartments of proteolytic activity (Holzer, 1978; Wiemken et al., 1979) and most probably proteins, to be degraded in the process of protein turnover, have first to be transported into the vacuole. Six intracellular proteases have so far been detected in the yeast Sacchuromyces cerevisiue. Of these enzymes proteinase A, proteinase B and carboxypeptidase Y are located in the vacuole; the localization of the other enzymes, namely carboxypeptidase S and aminopeptidases I and 11, is not yet fully elucidated (Holzer, 1978; Wiemken et ul., 1979). Specific endogenous inhibitors of proteinase A and B and carboxypeptidase Y have been found in the cytosol (Wolf and Holzer, 1978). They show a high

60

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

specificity towards proteinases and most probably are present to prevent unwanted proteolysis, for instance due to leaky vacuoles (Holzer, 1978). As already mentioned, the different examples of inactivation of alcohol oxidase and catalase activities in methanol-grown yeasts known at present may be the result of quite different processes which will be discussed in detail below. They include various examples of modification inactivation, for instance during bud formation, during vegetative growth and after exposure of methanolgrown cells to excess methanol (Veenhuis et al., 1978b). A unique and intriguing example of catabolite inactivation resulting in the turnover of peroxisomes is seen after transfer of methanol-grown cells into glucose- or ethanol-containing media. Such a transfer produces drastic changes in the ultrastructure of the cell (Bormann and Sahm, 1978; Veenhuis et al., 1978a). Moreover, it is also unique in quantitative terms since alcohol oxidase and catalase may together form up to 30% of the protein content of the cell-free extract.

B. MODIFICATION INACTIVATION OF PEROXISOMAL ENZYMES

1. Inactivation of Alcohol Oxidase During Bud Formation

During vegetative growth of H. polymorpha on methanol the number of peroxisomes per cell generally increases by the separation of small organelles from mature ones (see Section IV.B, p. 48). This process is generally not associated with inactivation of alcohol oxidase in these newly formed organelles. However, in one particular case, namely when small peroxisomes are separated and migrate into developing buds (Veenhuis et al., 1978b) inactivation of alcohol oxidase, but not of catalase, is observed in these organelles (Fig. 28a,b). This inactivation of enzyme activity was not associated with alterations in the substructure of the organelles; they remained crystalline throughout this process indicating that alcohol oxidase was still present (see Section II.C, p. 16). Reactivation of alcohol oxidase in such organelles depended on the prevailing physiological conditions. In fast growing cells the peroxisomes in the developing buds again showed alcohol oxidase activity when the synthesis of crosswall between mother cell and bud was initiated. At low growth rates, however, alcohol oxidase activity was only regained after the crosswall between mother cell and bud had closed. 2. Inactivation of Alcohol Oxidase at Low Growth Rates The presence in H. polymorpha of peroxisomes harbouring inactive alcohol

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

61

FIG. 28. Micrographs of methanol-grown cells of Hansenula poiymorpha showing that alcohol oxidase activity (b) but not catalase activity (a) is inactivated during and after the process of division of peroxisomes in the neck between mother cell and bud. Note the small unstained peroxisomes in the developing bud shown in (b). Cells in (a) were stained with diaminobenzidine and hydrogen peroxide (from Veenhuis et ai., 1978b) and in (b) they were stained with diaminobenzidine and methanol (from Veenhuis et al., 1976).

oxidase is not necessarily a temporary process associated with bud formation. Cytochemical experiments indicated that, during cultivation in a methanollimited chemostat, cells may contain peroxisomes in which alcohol oxidase is present in an inactive form. Such peroxisomes are particularly found in old cells which are characterized by a large number of bud scars and which occur abundantly in chemostat cultures grown at low dilution rates. In cells produced in batch cultures, peroxisomes exhibiting this property have so far not been encountered. The number of peroxisomes carrying inactive alcohol oxidase, as well as the extent of inactivation of the enzyme in individual organelles, depend on cultivation conditions and increase with decreasing growth rate (Figs 5c, 29a,b). Therefore, in cultures grown under these conditions, the alcohol oxidase activity estimated in cell-free extracts (van Dijken, 1976) may not be a reliable measure of the amount of alcohol oxidase protein in such cells. Incubation of these cells with 3,3-diaminobenzidine and hydrogen peroxide showed that catalase activity was still present throughout these organelles (Veenhuis er al., 1976). However, the possibility that those peroxisomesin which part of the alcohol oxidase is present in an inactive form also contain lowered levels of catalase activity, has not yet been investigated. It is not known whether alcohol oxidasewas first active in these organellesand became inactivated during ageing of the cells, or was at no time active. A possible mechanism for the inactivation of alcohol oxidase during migration of peroxisomes into developing buds (see above) and in cells from methanol-

62

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

FIG.29. Micrographs of cells of Hunsenulupolymorphu, grown in a methanol-limited chemostat, demonstrating the overall staining pattern and the difference in staining intensity of individual peroxisomes, present in cells grown at D = 0.10 h-' (a) and at D = 0.03 h-' (b). Alcohol oxidase activity was demonstrated with diaminobenzidine and methanol. limited chemostat cultures could be the uncoupling of the prosthetic group from the enzyme protein.

3. Inactivation of Alcohol Oxidase and Catalase after Exposure of Cells to Excess Methanol When cells of H. polymorpha which contain large amounts of alcohol oxidase are transferred into fresh media containing excess methanol, a rapid decrease in alcohol oxidase and catalase activity is observed (Fig. 30). This is associated with the excretion of formaldehyde and formate into the culture fluid (Schwartz, 1978; Veenhuis et al., 1978b). The degree of enzyme inactivation and the proportion of cells which remain viable after exposure to excess methanol are probably related to the amount of alcohol oxidase initially present in these cells. Generally, transfer of chemostat-grown cells into media containing excess methanol did not result in growth. Instead the culture died within a few hours (M. Veenhuis and J. P. van Dijken, unpublished observations; Schwartz, 1978).However, cells obtained from batch cultures in the stationary phase of growth, which contained lower levels of alcohol oxidase activity, survived the transfer and growth generally started after a lag of two to three hours (Fig. 30). Electron microscopic observations showed that the decrease of enzyme activity during the lag was not associated with changes in the number and substructure of the peroxisomes in the cells (Fig. 30). Cytochemical staining experiments, however, revealed a decreased

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

63

f

e

L9

Time ( h )

FIG. 30. Growth, enzyme profiles and volume fraction of peroxisomes in Hansenula polymorpha, grown in batch cultures on 0.4% methanol. The cultures were inoculated

with methanol-grown cells in the stationary phase of growth..-., Growth; 0-0, alcohol oxidase;b w , catalase; A-A, volume fraction of peroxisomes. intensity of staining of the organelles when these were assayed for alcohol oxidase activity. If staining had occurred, the reaction products were located in the central part of the peroxisomal matrix (Fig. 3 1) probably indicating that

FIG. 3 1. Sections of methanol-grown cells of Hansenula polymorpha, incubated with excess methanol. After one hour of incubation alcohol oxidase, as demonstratedafter incubation with diaminobenzidineand methanol, is only detected in the central part of the peroxisomes (a) or is completely absent (b).

64

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

inactivation commences at the periphery of the organelles. Recent evidence has been obtained which indicates that the inactivation of alcohol oxidase and catalase in these experiments is most probably caused by an overproduction of formaldehyde and/or hydrogen peroxide (A. Douma, unpublished findings).

C. DEGRADATIVE INACTIVATION OF PEROXISOMAL ENZYMES

Transfer of methanol-grown cells of Candida boidinii and Hansenulapolymorpha into ethanol- or glucose-containing media showed that synthesis of both catalase and alcohol oxidase were completely inhibited following transfer. In addition, a rapid inactivation of pre-existing enzymes was also observed (Meisel et al., 1978; Bormann and Sahm, 1978; Veenhuis et al., 1978a). For instance, addition of 0.4% glucose to a batch culture of H . polymorpha growing exponentially on methanol, caused a loss of approximately 70% of the activities of both alcohol oxidase and catalase within three hours (Fig. 32).

- 140 - 120

-

-

._ W

c

- 100

2

a

-

-I

C ._

-80

R

-2 T 4

-60

.->

.-c 0

-40

a

0 c

- 20

0

I I

I 2

I

1

I

3

4

5

0"

JO

Time ( h )

FIG. 32. Effect of the addition of 0.4%glucose to an exponentially growing batch culture of Hansenula polymorpha on 0.5% methanol. 0-0, Growth; 0-0, alcohol oxidase activity; A-A, catalase activity. From Veenhuis et al. (1978a).

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

65

In addition, other peroxisomal enzymes like L-a-hydroxy acid oxidase and D-amino acid oxidase exhibited a pattern of inactivation similar to that displayed by alcohol oxidase (M. Veenhuis, unpublished observations). Similar observations have been made with methanol-grown C. boidinii, transferred from the late-exponential growth phase into ethanol-containing media; both catalase and alcohol oxidase activities decreased dramatically and six hours after the transfer the activity of alcohol oxidase was no longer detectable (Bormann and Sahm, 1978). Bormann (1980) recently showed that in the case of alcohol oxidase the decrease in specific activity, as estimated by assays in uitro, was due to a decrease in alcohol oxidase protein (Fig. 33). Different observations have been described by Yasuhara et al. (1976). These authors showed that addition of 5% glucose to a methanol-grown culture of Kloeckera sp. 2201 inhibited the synthesis of catalase, but the kinetics of the decrease in specific activity of catalase in the culture indicated a dilution of enzyme over newly formed cells, rather than a decrease in catalase protein. Experiments with cells of H. polymorpha have shown that the rate of inactivation of alcohol oxidase and catalase depends on environmental conditions extant during growth of the cells prior to the transfer. For instance, cells harvested from stationary phase batch cultures showed a lower rate of

FIG. 33. Polyacrylamide-gel electrophoresis (7.5% polyacrylamide) of cell-free extracts of methanol-grown Cundida boidinii (gels 1,4) and of cells, two (gels 2, S), four (gels 3, 6) and six (gels 4,8) hours after transfer into ethanol-containing media. From Bormann (1980).

66

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

loss of specific enzyme activity when transferred to glucose media than did cells taken from cultures growing exponentially on methanol (Veenhuis et al., 1978b). Also in chemostat-grown cells the rate of inactivation of alcohol oxidase and catalase increased with increasing growth rate (Table 7). Transfer of cells grown on methanol plus methylamine into glucose plus ammonium sulphate-containing media also leads to a rapid inactivation of alcohol oxidase, amine oxidase and catalase (Veenhuis et. al., 1981a). However, when such cells were transferred to glucose plus methylamine, the initial decrease in amine oxidase activity was followed by a subsequent sharp increase in activity of this enzyme, which started two hours after the transfer (Fig. 34). Peroxisomes which contain alcohol oxidase are also known to develop in cells of H . polyrnorpha under conditions which do not require the activity of this enzyme for growth (see Section IV.A, p. 42). Again addition of excess glucose leads to a rapid inactivation of both alcohol oxidase and catalase activities in these cells (Egli et al., 1980). With regard to the definition of Holzer (1976) this observed loss of alcohol oxidase and catalase activities must be considered a special case of catabolite inactivation, since inactivation of these enzymes occurred when they performed no apparent physiological function. Changes in the environment resulting in conditions wherein the activity of a particular peroxisomal enzyme was no longer required for growth, did not necessarily lead to its inactivation. For example, following transfer of methanol plus methylamine-grown cells into methanol plus ammonium sulphate-containing media, no inactivation of amine oxidase was detected. The observed decrease in specific activity could be explained by dilution of pre-existing enzyme activity throughout newly formed cells (Veenhuis et al., 198la). The findings of Bormann and Sahm (1980), made after transfer of methanol-grown cells of C. boidinii into ethanol-containing media, indicated that inactivation of alcohol oxidase and catalase was an energy-dependent process. Furthermore, they also showed (Bormann and Sahm, 1978) that inactivation of these enzymes following transfer of cells to ethanol media is irreversible and not dependent on protein synthesis. In this respect it must be mentioned that ethanol may not be a suitable substrate for use in studies of the inactivation of alcohol oxidase and catalase in methanol-grown yeasts. As has already been stated, addition of excess methanol to a methanol-grown culture of H . polyrnorpha leads to a rapid modification inactivation of alcohol oxidase, due to an overproduction of formaldehyde. Addition of excess ethanol to methanol-grown yeast cultures similarly will lead to an overproduction of acetaldehyde, which reacts in the same way as formaldehyde; this would, at least partially, account for the decrease of enzyme activities detected

TABLE 7. Decrease in relative specific activities of alcohol oxidase and catalase associated with the decrease in number and volume fraction of peroxisomes after transfer of cells of Hansenula polymorpha grown on methanol in batch and chemostat culture, into glucose-containing media Relative specific activities Chemostat Batch culture D=0.13 h-'

D z 0 . 0 2 h-'

&63=

1.8

Morphometrical analysis Chemostat Batch culture D=0.13 h-'

Dz0.02 hk'

&,3=1.8

100 50 20 19

100 33 18 17

100 90 81 40

100 81 70 42

100 30 20 10

100 30 25 18

3.3

36.9

3.4

53.8

1.0

19.6

1.4 0.5

19.3 8.1

2.1 1.6

34.1 15.2

0.2 0.2

1.8 1.6

-

-

-

-

-

3 rn

z

1

iz 0

r cn

3

alcohol alcohol alcohol volume volume volume Time oxidase catalase oxidase catalase oxidase catalase number fraction number fraction number fraction

0 2 4 6

-

cn

0

-

: 0

z

0

D m

a 0

z

0

0

The number of peroxisomes is expressed as average number per section, the volume fraction is percentage of cytoplasmic volume.

3 T

0

68

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

c

8 - 7c

-

-- c

-

/"

70

-60

-7 -

E

+ Q

-

I

I

-30

6

g ?

+ 0 0

c 0

0

2

4

6

8

1

0

1

2

Tlme ( h )

FIG. 34. Growth and enzyme profiles after transfer of cells of Hunsenulu polymorphu growing in chemostat culture on methanol and methylamine into batch cultures catalase supplemented with glucose and methylamine; 0- - - - -0, Growth; A-A, amine oxidase activity x lo-*. activity; b m , alcohol oxidase activity; 0-0, From Veenhuis et ul. (1981a).

in these cultures (J. P. van Dijken and M. Veenhuis, unpublished work; Bormann, 1980). Bonnann and Sahm (1980) and Bormann (1980) have studied different protease activities present in methanol-grown cells of C. boidinii. The activity of an acid protease (PH 3.0) was demonstrated together with that of a neutral protease (PH 7.0) and a carboxypeptidase.Experiments in vitro with proteases purified from Sacch. cerevisiae indicated that alcohol oxidase was rapidly degraded by the neutral proteinase 'B' and to a significantly lesser extent by the acid proteinase 'A'. The carboxypeptidase of Sacch. cerevisiae had no influence on the activity of alcohol oxidase. The extent and rate of degradation of peroxisomal enzymes in methanolgrown cells of H. polymorpha is dependent upon the nature of the carbon

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

69

4 W n

v?

Y

f

B

f (3

'i

0

a

0

Y

5

Y

0

1

2

3

4

5

6

Time ( h )

FIG. 35. Effect of addition of 0.5% glycerol to an exponentially growing batch culture of Hansenula polymorpha on 0.5% methanol. W-W, Growth; 0-0, alcohol oxidase activity; 0-0,

catalase activity.

source present in the media in which the methanol-grown cells are subsequently resuspended. The addition of 0.5% glycerol, a substrate which only partly represses alcohol oxidase synthesis, to a batch culture in the midexponential phase of growth on methanol, resulted in a small increase in growth rate associated with a decrease of the specific alcohol oxidase activity of approximately 40% in the first 90 minutes of incubation (Fig. 35; M. Veenhuis, unpublished observations). During subsequent growth the specific activity of this enzyme remained approximately constant. Similar observations were made after the addition to the culture of 0.5% dihydroxyacetone. Addition of 0.5% deoxyglucose, a substrate that cannot be metabolized, to a culture of H. polymorpha growing exponentially on methanol, resulted in an immediate cessation of growth. However, the presence of this compound had no influence on the levels of alcohol oxidase and catalase in the cells.

D . SUBCELLULAR EVENTS I N PEROXISOMAL DEGRADATION

Electron microscopic observations revealed that the inactivation of alcohol oxidase and catalase in methanol-grown yeast cells, occurring after the transfer of such cells into glucose- or ethanol-containing media, is paralleled by a decrease in number and volume fraction of the peroxisomes originally

70

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

present in these cells (Table 7; Meisel et al., 1977, 1978; Bormann and Sahm, 1978; Veenhuis et al., 1978a). The mechanism of degradation of the peroxisomes in methanol-grown cells has only partially been elucidated. Meisel et al. (1978) suggested that the peroxisomes are displaced into the central vacuole, whereas Bormann and Sahm (1978) described a decrease in size of the organelles. A more detailed ultrastructural study of the mechanisms of peroxisomal breakdown was performed with the yeast Hansenula polymorpha, grown on methanol under different conditions and subsequently transferred into glucose-containing media (Veenhuis et al., 1978a 1981a,c). These studies revealed that for individual peroxisomes the process generally can be divided in two consecutive events: (a) inactivation of alcohol oxidase and catalase (and probably also other peroxisomal enzymes) associated with a simultaneous disappearance of the crystalloidspresent in the peroxisomalmatrix; (b) “visible” degradation of the peroxisomal proteins as a result of the action of proteolytic enzymes originating from the vacuole. The involvement of the vacuole in the degradation process is not surprising since the initiation of the process is suggested to be independent of protein synthesis (Bormann and Sahm, 1978) and vacuoles are known to be the main compartments in which hydrolytic enzymes are located in yeast cells (Holzer, 1978; Wiemken et al., 1979). Electron microscopical observations on potassium permanganate-fixedmethanol-grown cells of H . polymorpha exposed to excess glucose showed that the first visible event was the formation of dark membranous layers around those peroxisomes destined to be degraded (Fig. 36a,b). Similar observations have been made after addition of excess glucose either to cells of H . poiymorpha growing on glycerol or to cells in the stationary phase of batch culture growth on glucose. Generally, in cells of exponentially growing methanol cultures initially only the large peroxisomes were surrounded by membranes, whereas in cells grown in a chemostat several individual peroxisomes became surrounded at the same time (Fig. 36a). Along with the synthesis of these membranes which are most probably derived from the endoplasmic reticulum (Fig. 36c), especially in older cells from chemostat cultures grown at low dilution rates which contained many peroxisomes, we frequently observed fragmentation of the vacuole into a number of smaller vesicles. By a mechanism as yet unclear, one or several of these vacuolar vesicles became incorporated within the dark membranes and thus were sequestered from the cytosol together with the peroxisome (Fig. 37a). Following sequestration the membranes of the vacuolar vesicles probably disrupt, thereby exposing the contents of the sequestered cell components to attack by the vacuolar hydrolases. At this stage of the degradation process the peroxisomal crystalloid has disappeared and the organelle, which may now be considered as an autophagic vacuole, has become spherical in shape. Cytochemical

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

71

FIG. 36. Cells of Hansenulu polymorpha, grown in methanol-limited chemostat culture, 20 minutes after the transfer of cells into glucose-containing media. (a) Shows the presence of dark membranes, partly or completely surrounding individual peroxisomes (P) present in cells of Hansenulupolymorpha. (b) Shows these membranes in more detail in a magnification of one cell from (a); the inset (c) shows the association of numerous layers of these membranes with a vacuolar vesicle (V). (d) Shows (after freeze-etching) the continuity between the endoplasmic reticulum and the typical, almost smooth membrane covering a peroxisome.

72

M. VEENHUIS, J.

P. VAN DIJKEN AND W. HARDER

staining experiments indicated that at least part of the vacuolar fluid really was present in these organelles since activities of vacuole-specific hydrolases, e.g. acid phosphatase and glucose 6-phosphatase, which are absent from intact peroxisomes, were now present in the disintegrating peroxisomes (Fig. 38a). In contrast, alcohol oxidase and catalase activities could no longer be demonstrated in these autophagic organelles (Veenhuis et al., 1978a). As degradation progressed, small areas of disintegration became visible which increased in volume and number. Instead of the electron-dense membranes originally present, the organelles were now surrounded by a single unit membrane of about 9 nm (90 A) in width (Fig. 37c), which probably originated from the outer layer of these dense membranes. Remnants of these membranes and of the incorporated vacuolar vesicles were frequently observed in the autophagic vacuoles (Fig. 37b). Subsequently the matrix of the degrading peroxisome became finely granular (Fig. 37b) and ultimately it turned into an organelle, which was considered to be a vacuole. A special case of peroxisomal breakdown was observed during studies on the generative reproduction of H. polymorpha (Veenhuis et al., 1980b). Cultivation of H. polymorpha in a sporulation medium containing methanol as the carbon source and urate as the nitrogen source, resulted in a decrease of alcohol oxidase and catalase activities after growth of the culture had ceased. Electron microscopic observations of such cells showed that the decrease of enzyme activity was associated with the fragmentation of the large peroxisomes, originally present in the cells, into several organelles of smaller size, followed by degradation of a number of the latter organelles. Enzyme inactivation, observed after the addition of excess glycerol or dihydroxyacetone to a culture growing on methanol was also associated with the degradation of peroxisomes, effected by a similar mechanism to that described above. However, as was expected from the kinetics of the inactivation of peroxisomal enzymes, the extent of the degradation was much less than that following addition of excess glucose. Addition of excess deoxyglucose also led to the formation of dark membranes around individual peroxisomes, as was observed after transfer to glucose. These membranes remained intact during the following hours of incubation. However, degradation of peroxisomes was not observed. Invariably, degradation of peroxisomes was initiated by their sequestration from the cytosol. Displacement of organelles into the vacuole by a process resembling pinocytosis was never observed. Also, peroxisomes to be degraded were sequestered individually and we have observed no instances in which more than one organelle was present in the compartments formed by the electron-dense membranes. As already indicated, in cells present in batch cultures growing exponentially on methanol, initially only the large peroxisomes were generally

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS

73

FIG. 37. Different stages of peroxisomal breakdown after the addition of glucose to batch cultures of Hansenula polymorpha, grown on methanol. (a) Detail of a cell 20 minutes after addition of glucose, showing the continuity of the dark membranes, surrounding a peroxisome, and a vacuolar vesicle. (b) Shows the extent of peroxisomal degradation after 90 minutes’ incubation in the presence of glucose. (c) The initial stage of degradation is visualized as it is observed after 30 minutes’ incubation in the presence of glucose. Note the increased width of the membrane of the degrading organelle compared to the membrane of the intact peroxisome (arrows); (b) and (c) are from Veenhuis et al. (1978a).

74

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

FIG. 38. Details of chemostat-growncells of Hunsenulu polymorpha 60 minutes after transfer into glucose-containingmedia. (a) Demonstration of glucose 6-phosphatase activity in the matrix of the degrading peroxisome (revealed by incubation with cerium chloride and glucose &phosphate). (b) Shows fusion of two degrading peroxisomes.

degraded, leaving the smaller ones unaffected (see Table 7). This observation may explain the results of Bormann and Sahm (1978) who described in C. boidinii a decrease in size of the organelles after transfer of cells from methanol to ethanol. In chemostat-grown cells of H . polymorpha, in which several peroxisomes may be subject to degradation at the same time, fusions of the different autophagic vacuoles have frequently been observed (Fig. 38b). The mechanism underlying the process of degradation resembles that of autophagic processes known to occur in plant cells (Matile, 1975). A major differencefrom the latter processes is, however, that in methanol-grown yeasts the vacuole is involved in degradation because this organelle supplies the hydrolytic enzymes required for degradation of redundant peroxisomes. The observations of Bormann and Sahm (1978) on cells of C. boidinii transferred from glucose into ethanol media showed that (a) inactivation of alcohol oxidase and catalase is an irreversible process and (b) de nouo protein synthesis is necessary for the recovery of the activity of these enzymes. These aspects were further investigated in cells of H. polymorpha placed under conditions in which a high rate of alcohol oxidase synthesis occurred. In experiments in which methanol-grown cells of this yeast after transfer to

PEROXISOME METABOLISM OF ONE-CARBON COMPOUNDS ~

75

IL"

- 100 al ul

D

s

-80 U 0

.I-

>.

.-c

-60

2

U

.-U ._

.I-

U

0

-40 > .e

-al 0

- 20 I

1

1

I

I

I

I

1

0

1

2

3

4

5

6

7

K

10

Time ( h )

FIG. 39. Growth and relative specific activity of alcohol oxidase in batch cultures of Hansenula polyrnorpha on 0.5% methanol, supplemented in the mid-exponential growth phase with 0.5% glucose and after 2 hours' incubation transferred back into fresh medium containing 0.5% methanol. 0-0, Growth; 0-0, relative specific activity of alcohol oxidase.

glucose for two hours were transferred into fresh methanol medium, enzyme studies indicated that in the first hours after transfer the rate of alcohol oxidase synthesis was approximately three times higher than during normal growth on methanol (Fig. 39). Electron microscopical observations demonstrated that along with the recovery of alcohol oxidase activity a number of small peroxisomes developed in the cells, and thereafter quickly increased in size during further cultivation. Together with the development of these organelles we observed in the same cell a continued degradation of organelles, which had already been subject to some degradation before the cells had been transferred into the final methanol medium. As has been described for the synthesis of newly formed peroxisomes after transfer of cells from methanol plus methylamine in to glucose plus methylamine media (see Section IV.B, p. 52), the enhanced rate of peroximal synthesis under these conditions suggests

76

M. VEENHUIS, J. P. VAN DIJKEN AND W. HARDER

that some at least of the amino acids, resulting from proteolysis, are incorporated into the newly synthesized peroxisomal enzymes.

V. Concluding Remarks The information summarized above on various aspects of the role of peroxisomes in the metabolism of one-carbon compounds in yeasts clearly shows that these unicellular organisms offer an almost ideal model system for the study of function, morphogenesis and turnover of these intriguing organelles. It is now beyond doubt that in H. polymorpha, and most probably in other yeasts also, peroxisomes originate from pre-existing organelles and develop by means of growth and division. Their ultimate shape, number, substructure and physiological function depend entirely on the environmental conditions to which the yeast cell is exposed. It follows that by manipulating these environmental conditions it is possible to prescribe the metabolic role of these organelles. Studies on the biogenesisand turnover of peroxisomes in yeasts have not yet progressed beyond the descriptive stage. This is mainly due to the fact that only comparatively recently have a number of physiological functions in which peroxisomes play a key role been discovered in these organisms. It is to be expected, however, that our knowledge of the molecular biology of the synthesis and turnover of yeast peroxisomes will rapidly expand in the years to come. Acknowledgement The authors are indebted to Mr. J. Zagers for his skilled technical assistance and to Marry Pras for her help in the preparation of the manuscript. REFERENCES

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Polyphosphate Metabolism in Micro-Organisms IGOR S. KULAEV and VLADIMIR M. VAGABOV Institute of Biochemistry and Physiology of Micro-organisms, Academy of Sciences of the U.S.S.R., Pushchino, Moscow Region, U.S.S.R. I Introduction

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B. Distribution in micro-organisms . . C. Methods of detection, identification and fractionation of inorganicpolyphosphates . . . . . . . . . . . . 11. High molecular-weightpolyphosphates . . . . . . A. Intracellular localization . . . . . . . . . B. Enzymes involved in biosynthesis and degradation of polyphosphates . C. Metabolism of polyphosphates in eukaryotes . . . . . . D. New data on polyphosphate metabolism in prokaryotes . . . . E. Concluding remarks on the physiological role of high molecular-weight . . . . polyphosphates in microbial metabolism 111. Inorganic pyrophosphate: new aspects of metabolism and physiological role . A. Utilization of pyrophosphate in phosphorylation reactions in bacteria . B. Energy-dependent synthesis of pyrophosphate during photosynthetic and oxidative phosphorylation . . . . . . . . . C. Relationship between pyrophosphate and polyphosphate metabolism in micro-organisms . . . . . . . . . . IV. Modem concepts about the role of high molecular-weightpolyphosphates and pyrophosphate in evolution of phosphorous metabolism . . . . V. General conclusions . . . . . . . . . . VI. Acknowledgements . . . . . . . . . . . References . . . . . . . . . . . .

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phates, among which the major role is obviously played by nucleoside di- and triphosphates, and in the first place by the ATP-ADP system. However, numerous findings reported in the last three decades show that cell energetics are not inconsiderably contributed to by compounds other than nucleoside polyphosphates. These include so-called condensed inorganic phosphates which are found primarily in micro-organisms and appear to be primitive energy accumulators in living organisms (Belozersky, 1959). They were first reported at the end of the last century (Liebermann, 1888); however, a thorough study of their structure and metabolism began only in the middle of this century, when the investigations carried out mainly by Jeener and Brachet (1 944), Wiame (1949), Ebel (1951), Belozersky ( 1958), Kulaev and Belozersky (1962), Lohmann and Langen (1956), HoffmannOstenhof and Weigert (1952), Hoffmann-Ostenhof (1962), Kornberg et al. (1956), Harold (1966) and some others (see Dawes and Senior, 1973; Kulaev, 1973a,b, 1979) laid the foundations of the biochemistry of inorganic polyphosphates. Ever-increasingattention has in recent years been paid to new aspects of the biochemistry of these extremely interesting phosphorus-containing compounds. Despite the availability of a number of reviews pertaining to this field of knowledge (Kuhl, 1960, 1974; Harold, 1966; Dawes and Senior, 1973; Kulaev, 1975) and the publication of a monograph on the biochemistry of high molecular-weight polyphosphates (Kulaev, 1979), it seems necessary today to revise certain concepts of the basic features of the metabolism and physiological role of these high-energy phosphorus compounds. Many problems of their biochemistry, extensively reviewed in the abovementioned publications, will not be discussed here. In this review, accent will be laid on the problems the study of which has recently yielded principally new data, and on the aspects which have not been discussed earlier in sufficient detail.

A . INORGANIC POLYPHOSPHATES

Inorganic polyphosphates are linear polymers in which orthophosphate residues are linked by energy-rich phospho-anhydride bonds (Yoshida, 1955; Flodgaard and Fleron, 1974; Fig. 1). The number of phosphate residues in these compounds, as identified in living organisms, may vary noticeably: from two in the simplest compound of this type, pyrophosphate, to several hundreds and thousands in high molecular-weight polyphosphates (Kulaev, 1979). The structure and properties of polyphosphates are described in a number of reviews and monographs (Van Wazer, 1958; Boulle, 1965; Ohashi, 1975; Kulaev, 1979).

POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS

r

85

1( n + 2 ) -

FIG. 1. Molecular structure of a linear polyphosphate. Me’ is a monovalent metal. From Thilo (1959).

B. DISTRIBUTION IN MICRO-ORGANISMS

Inorganic polyphosphates have been found in almost all tested representatives of living cells (Kulaev, 1979). They have been detected in eubacteria, fungi, algae, mosses, protozoa, insects, and in various tissues of higher plants and animals. Unfortunately, up to date no attempts have been made to detect these compounds in representatives of a new realm of living beings, namely the archaebacteria (woese and Fox, 1977; Steckenbrandt and Woese, 1979). The quantities of high molecular-weight polyphosphates detected hitherto in cells of higher plants and animals are small. According to published data, their phosphorus contents amount to tens, at most hundreds, of micrograms per gram wet tissue of these organisms. As to cells of micro-organisms, the situation is just the opposite. Yeast, for example, when grown in a medium with phosphate and glucose and certain cations (K+, Mg*+), after phosphorus starvation, may accumulate polyphosphates in amounts of up to 20% of the cell dry weight. Liss and Langen (1960) called this phenomenon “Polyphosphat ~berkompensation”which has been translated into English as “polyphosphate overplus” (Harold, 1964). Such a “polyphosphate overplus” occurs in cells in the absence of growth, i.e. when most of the energy has been released during glucose oxidation and the bulk of phosphate absorbed accumulated in polyphosphates. However, some intensively growing bacteria, e.g. Acinetobacter, during cultivation on butyrate, are capable of accumulating, besides a substantial amount of lipids, inorganic polyphosphates in quantities of 10-20% of the dry weight (Deinema et af.,1980). It is noteworthy that, in this case, uptake of a large amount of exogenous phosphate and its accumulation inside the cells in the form of polyphosphate granules are characteristic of normal metabolism of these bacteria. In contrast to the “polyphosphate overplus”, this phenomenon has been termed “luxury uptake” (Levin and Shapiro, 1965).

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Metabolism of polyphosphates in microbial cells has been studied in most detail since, in micro-organisms, these compounds accumulate in significant quantities. Nevertheless, inorganic polyphosphates are not vitally important for living organisms, and do not appear to be obligatory cell components. This has been demonstrated by Harold, who obtained mutants of Aerobacter aerogenes which were unable to synthesize and accumulate high molecularweight polyphosphates (Harold and Harold, 1963). Mutants of the cyanobacterium Anacystis nidulans, deficient in polyphosphates, were recently obtained by Vaillancourt et af. (1978). These mutants could grow, though very poorly, under certain cultivation conditions.

C . METHODS OF DETECTION, IDENTIFICATION A N D FRACTIONATION OF INORGANIC POLYPHOSPHATES

The oldest and most extensively used methods for determining condensed phosphates in biological materials, although of course the least accurate, are based on staining of cells and tissues by certain basic dyes, such as toluidine blue, neutral red and methylene blue. The presence of condensed phosphates in the organisms is judged from the appearance in cells of metachromatically stained granules, or, as they are also known, volutin granules (Kuhl, 1974; Kulaev, 1979). It must be said, however, that, despite the fact that in most cases the cytochemicaldetection of polyphosphate granules is associated with the actual presence of condensed phosphates in the organism, the use of such methods must nevertheless be attended with great caution (Martinez, 1963). This is primarily due to the fact that the basic dyes used to identify polyphosphate granules are also capable of metachromatically staining other polymeric compounds which are encountered in biological material. However, in recent years, primarily owing to the works of Jensen and his coworkers, cytological methods of detecting polyphosphate granules in situ have been significantly improved (Jensen, 1968,1969;Jensen and Sicko, 1974; Sicko-Goad et al., 1975, 1978; Lawry and Jensen, 1979; Baxter and Jensen, 1980a,b). In these works, very interesting results on the structure and formation of polyphosphate granules in cyanobacteria were obtained by electron microscopic and cytochemical methods. Particularly impressive data were furnished by electron microscopy combined with X-ray dispersion analysis (Coleman et al., 1972).This method gives the opportunity of reliably detecting phosphate in the electron-dense inclusions detected by electron microscopy, and also makes it possible to establish the nature of cations present in polyphosphate granules and to establish whether they contain organic components. Besides the above works of Jensen and his coworkers, in

POLYPHOSPHATE METABOLISM I N MICRO-ORGANISMS

a7

a number of studies this method was successful for detection and chemical analysis of polyphosphate granules in various organisms (Jones and Chambers, 1975;Wool and Held, 1976; Kessel, 1977; Hutchinson et al., 1977; Peverly et al., 1978; Adamec et al., 1979; Barlow et al., 1979; Tillberg et al., 1979; Doonan et al., 1979). All these works revealed that the composition of polyphosphate granules changed markedly depending on the chemical and, in the first place, ionic composition of the cultivation medium. However, strictly speaking, this method of polyphosphate detection is not universally appropriate. Firstly, in granules it identifies the mere presence of phosphate but not phosphoryl groups linked by anhydride bonds. Secondly, it does not detect polyphosphates in cells if their concentrations in the subcellular structures is not high enough. A more sensitive and convenient method of polyphosphate detection in situ is by fluorescencemicroscopy using fluorochrornesof the type 4‘,6‘-diamidin0-2-phenylindole-2HCl(DAPI; Allan and Miller, 1980). To date, high-resolution 31P nuclear magnetic resonance has proved to be efficient in detecting intracellular polyphosphates containing phosphate residues linked by anhydride bonds, or so-called “middle” phosphate groups (Glonek et al., 1971; Salhany et al., 1975; Burt et al., 1977; Navon et al., 1977a,b; Ugurbil et al., 1978; Ferguson et al., 1979; Sibeldina et al., 1980; Ostrovsky et al., 1980). Thus, at present, various physical methods are efficiently used for detection of polyphosphates in situ. Moreover, as stated above (Kulaev, 1979), a number of chemical methods are currently employed to identify exactly polyphosphates in extracts from biological material. Of these, the most widely used are chromatographic methods, particularly thin-layer chromatography (Kulaev and Rozhanets, 1973;Kulaev et al., 1974a;Ludwig et al., 1977;Lusby and McLaughlin, 1980; Guerrini et at., 1980; Solimene et af., 1980). However, these methods are applicable only for identification and analytical separation of low molecular-weight polyphosphates between two and seven residues, whereas higher molecular-weight polyphosphates are practically unresolvable. Our preliminary data (I. S. Kulaev, K. G. Skryabin, P. M. Rubtsov and V. D. Butukhanov, unpublished results) suggest that the chromatographic techniques widely used at present for fractionation of oligonucleotides (Maxam and Gilbert, 1977) may prove expedient for separating highly polymerized polyphosphates. As this method combines chromatography and radioautography of 32P-labelled products, it may be considered as a rather promising and sensitive method not only for detecting polyphosphates in biological material, but also for determinating their chain length. The most accurate, though a rather painstaking, method of identification of condensed polyphosphates by the specific product of their partial hydrolysis, cyclic trimetaphosphate, still remains important (Thilo and Wieker, 1957;

88

IGOR S. KULAEV AND VLADlMlR M . VAGABOV

Kulaev, 1979). It is regretable that only a few researchers resort to this method for strict identification of condensed polyphosphates in extracts from biological material. Enzymic methods for rigorous identification of inorganic polyphosphates in biological objects would surely prove to be less time-consuming and more reliable. However, up to the present, researchers do not yet have at their hands reliable preparative techniques for obtaining pure and stable enzymes of polyphosphate metabolism which could be used as specific reactants for the detection of inorganic polyphosphates. Special mention should be made of several specific methods of analytical determination of inorganic pyrophosphate reported in recent years (see e.g. Putnins and Yamada, 1975). As will be indicated later, use of these methods disclosed certain specific metabolic features of this simplest representative of inorganic polyphosphates in micro-organisms(Mansurova et af., 1975a, 1976; Shakhov et af., 1978; Ermakova et al., 1981). As to the methods of extraction of inorganic polyphosphates from biological material and their fractionation, no new techniques have been reported lately. Of all the available methods of polyphosphate fractionation (Kulaev, 1979) the most informative is that of Langen and Liss (1958), which proved to be expedient owing to the fact that it produced fractions characterized by different intracellular localization and physiological activity (Alking et af., 1977).This method consists in successive extraction in the cold with 5% trichloroacetic acid or occasionally with 0.5 M perchloric acid (acid-soluble fraction Polyp,), saturated sodium perchlorite solution (saltsoluble fraction PolyP2), dilute sodium hydroxide solution (pH 10; alkalisoluble fraction PolyP3) and a more concentrated solution of alkali (0.05 M sodium hydroxide; alkali-soluble fraction PolyP~).However, for a number of organisms, these sequential treatments do not ensure complete extraction of polyphosphates from the cells. Kulaev et al. (1966) suggested extracting the remaining polyphosphates with hot perchloric acid, thus hydrolysing them to orthophosphate. The use of less convenient schemes of fractionation for extraction of polyphosphates from biological materials has meagre prospects, since this is quite often connected with difficulties in interpreting data on polyphosphate metabolism. As indicated below, various fractions of polyphosphates have different pathways of synthesis and degradation. Before concluding the introductory part of this review, it should be pointed out that recent work (see e.g. Baltscheffsky and Stedingk, 1966; Mansurova et al., 1973a,b; Reeves, 1976; Wood, 1977) has unambiguoudy pointed to an essential difference in metabolic pathways and physiological activity between condensed polyphosphates and inorganic pyrophosphate. In this connection, it seems appropriate to discuss some peculiarities of their metabolism.

POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS

89

11. High Molecular-Weight Polyphosphates A. INTRACELLULAR LOCALIZATION

Not long ago, it was postulated that high molecular-weight polyphosphates are localized in the microbial cell within the so-called metachromatic granules or volutin granules (Wiame, 1949; Ebel, 1951; Kuhl, 1960; Kulaev and Belozersky, 1962). This concept applied to both prokaryotes and eukaryotes. A wealth of data reported in recent years shed new light on the intracellular localization of these compounds. The most profound studies in this field have been conducted in eukaryotes. Therefore, we shall begin the discussion with these organisms. 1. Eukaryotes Since the intracellular localization of polyphosphates in eukaryotes is best studied for yeast and fungi, let us in the first place consider data pertaining to these organisms. First indications that, at least in yeast, not all polyphosphates are present inside cells in volutin-like granules were obtained by Weimberg and Orton (1965), Weimberg (1970) and Souzu (1967a,b). Their data suggested that a portion of high molecular-weight polyphosphates were localized on the cell surface, in the region of cytoplasmic membrane. Further progress in this field was related to research conducted in a number of laboratories on polyphosphate metabolism in the fungi Neurospora crassa (Kulaev et al., 1966, 1970a,b; Krasheninnikov et al., 1967, 1968;Trilisenko el al., 1980) and Endomyces magnusii (Kulaev et al., 1967a,b; Afanas’eva et al., 1968; Skryabin et al., 1973; Ostrovsky et al., 1980), as well as in yeast (Indge, 1968; Vagabov et al., 1973; Urech et al., 1978; Diirr et al., 1979; Wiemken et al., 1979; Martinoia et al., 1979; Cramer et al., 1980; Tijssen et al., 1980; Lichko et al., 1982). Following Weimberg and Orton (1965), we studied localization of various polyphosphate fractions by a modified method of Langen and Lis (1958). For this purpose, protoplasts (sphaeroplasts) were isolated from mycelia of N. crassa and cells of E. magnusii, and pure and fairly intact nuclei and mitochondria were obtained from these organisms (Kulaev et al., 1970c,d;Skryabin et al., 1973). Table 1 summarizesthe data obtained by analysing the localization of various fractions of polyphosphates in cells of E. magnusii normally grown for 12 hours and in the same kind of cells cultivated for four hours under conditions of “polyphosphate overplus” after six hours of phosphorus starvation. It can be seen that, after removal of the cell wall, the amount of polyphosphates decreased by 2530% in both types of cells. The

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90

most condensed polyphosphates extractable by hot perchloric acid (PolyPs) disappeared in both cases. Concentrations of the alkali-soluble (Polyp3+ PolyP4) and salt-soluble (PolyPz) fractions were found to decrease significantly, whereas concentration of acid-soluble polyphosphates (PolyP~) slightly increased. The data obtained showed that an important portion of polyphosphates is localized on the surface of the cell either outside the plasma membrane or closely adjacent to it. Polyphosphates lost in the course of cell-wall lysis are hydrolysed therewith to orthophosphate. This suggests that they are apparently in close contact with polyphosphatases which hydrolyse them during protoplast preparation. The results in Table 1 clearly demonstrate that various fractions of polyphosphates differentially extracted from E. magnusii are localized in different cell compartments. Thus, these results show that different extractability of specific polyphosphate fractions is conditioned by different topographic and chemical compartmentation of polyphosphates in the cell. It follows from the data in Table 1 that high molecular-weight polyphosphates are absent from mitochondria of E. magnusii, and the only fraction detected in nuclei is the salt-soluble one. Similar data were obtained in our laboratory using the same methodological approach for mycelia of N. crassa (Kulaev et af., 1966, 1970d; Krasheninnikov et af., 1967, 1968). The presence of 30-35% polyphosphates in the peripheral parts of yeast cells was repeatedly shown in our laboratory for Saccharomyces carfsbergensis. As seen from Table 2, the most highly polymerized alkali-soluble TABLE 1. Contents of various polyphosphate fractions in Endomyces magnusii (whole cells, protoplasts, mitochondria, and nuclei; mg phosphorus (g dry cells-')). A are data for cells grown in phosphate-sufficient medium, B for cells grown in phosphate-rich medium. After Kulaev et al. (1967a), Afanas'eva et a f . (1968) and Skryabin er al. (1973) Conditions of culture growth A B Phosphorus-containing Whole MitoWhole compounds cells Protoplasts chondria Nuclei cells Protoplasts Acid-soluble PolyPl Salt-soluble PolyPl Alkali-soluble Polyp3 Polyp4 Hot perchloric acid extract PolyPs High molecular-weight polyphosphates (total)

+

0.2 0.7 0.9

0.7 0.4 0.4

0.0 0.0 0.0

0.4 0.0

11.2 3.4 3.3

12.1 1.4 0.8

0.2

0.0

0.0

0.0

2.5

0.0

2.0

1.5

0.0

0.4

20.6

14.3

-

PO LY PHOSPHATE M ETABOLlSM IN MICRO-0 RGANI SMS

91

TABLE 2. Contents of polyphosphate fractions in whole cells and protoplasts of Saccharomyces carlsbergensis. After Vagabov et al. (1973) Content (pg phosphorus (g wet cells)-’) Phosphorus-containing compounds Acid-soluble (PolyP1) Salt-soluble (PolyP1) Alkali-soluble (pH 8-10; Polyp,) Alkali-soluble (pH 12; Polyp4) High molecular-weight polyphosphates (total)

Whole cells

Protoplasts

706 516 208 356

728 299 23 0

1786

1050

polyphosphate fractions of this yeast are also removed on lysis of cell walls by the “snail” enzyme and are absent from resulting protoplasts (Vagabov et al., 1973). The fact that a substantial portion of highly polymerized polyphosphates is localized in yeast and fungi on the cell surface was directly or indirectly shown in works of other researchers. Van Steveninck and his associates (Jaspers and van Steveninck, 1975; Tijssen et al., 1980) provided cytochemical and biochemical evidence for the presence of highly polymerized polyphosphates on the surface of the yeast cell, outside the plasma membrane. It was also found that, in the logarithmic growth stage, this highly polymerized surface fraction of polyphosphates accounted for up to 40% of the total amount of these compounds in yeast, whereas in the stationary phase, it accounted for only 9%. The rates of turnover of two pools of inorganic polyphosphates detected by these authors (the intracellular pool and the one residing outside the cytoplasmic membrane) differed dramatically. The difference in the chain length and turnover rates of the yeast polyphosphates in these two fractions also supports, albeit indirectly, the concept of their dissimilar compartmentation. Recent data of Trilisenko et af. (1980) speak in favour of a common localization of high molecular-weight polyphosphates and polyphosphatase on the surface of cells of N. crassa. In this work, investigation of polyphosphate turnover in a “leaky” mutant of N. crussa showed that a drastic decrease in polyphosphatase activity in the mutant leads to a substantial accumulation of highly polymerized polyphosphates. This can be clearly seen from the data in Table 3. The results of studies on phosphorus metabolism in a slime mutant of N. crussa (Trilisenko et al., 1982), as well as in the plasmodia of Physurzun polycephalum (Sokolovsky and Kritsky, 1980), suggest the localization of the most polymerized fractions of polyphosphates outside the fungal plasma membrane. In both cases the most polymerized fractions (PolyP3, Polyp4and

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IGOR S. KULAEV AND VLADlMlR M . VAGABOV

TABLE 3. Contents of polyphosphatefractions in Neurospora crassa during maximum polyphosphatase activity. Data are shown for strain ad-6 and its leaky mutant 30, 19-3. After Trilisenko et al. (1980) Contents (jig (g dry wt-I)) Phosphorus-containing compounds

A ad-6

B 30, 19-3

B/A (%)

Acid-soluble (PolyP~) Salt-soluble (PolyPz) Alkali-soluble (Polyp3 +PolyP4) Hot perchloric acid extract (PolyPs) Orthophosphate Total (Polyp3 PolyP4+ Polyps) High molecular-weight polyphosphates (total)

2,370 1,920 1,700 200 700 1,900

1,550 2,080 3,000 1,200 720 4,200

65 108 176 600 102 220

6,190

7,830

126

+

Polyps) were absent from fungal protoplasts devoid of cell walls. These data prove the presence of these polyphosphate fractions in the periphery of the cell walls of fungi and yeast. As far as intracellular polyphosphates are concerned, they appear to occupy several cell compartments in these organisms. As already mentioned (see Table l), a portion of the salt-soluble polyphosphates PolyPz was detected in nuclear preparations of E. magnusii (Skryabin et al., 1973) and N . crassa (Kulaev et al., 1970a). The presence of specific fractions of high molecularweight polyphosphates in nuclei of different origin has been demonstrated by many researchers (Penniall and Griffin, 1964; Goodman et al., 1968, 1969; Sauer et al., 1969; Bashirelashi and Dallam, 1970; Mansurova et al., 1975b; Hildebrandt and Sauer, 1977; Sokolovsky and Kritsky, 1980; Offenbacher and Kline, 1980). Sokolovsky and Kritsky (1980) reported interesting findings confirming the occurrence in nuclei of Physarum polycephalum of salt-soluble polyphosphates and also of a certain portion of acid-solublepolyphosphates. Attempts were made to detect the localization of polyphosphates inside nuclei. As found in our laboratory (Mansurova et al., 1975b) in rat liver nuclei, the high molecular-weight polyphosphates exhibiting positive metachromatic reaction occur in fractions of nuclear globulin, histones and acid proteins (Zbarsky, 1970). On the other hand, Offenbacher and Kline (1980) showed that in the same organelle polyphosphates were linked to non-histone proteins. Hildebrandt and Sauer (1977) pointed to the occurrence of high molecular-weight polyphosphates in nucleoli of Physarum polycephalum, i.e. in the site of synthesis of RNA and ribosomes. As already indicated, thoroughly purified and fairly intact mitochondria from E. magnusii (Afana-

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s'eva et al., 1968; Kulaev et al., 1970a,c)and N.crassa (Krasheninnikov et al., 1968; Kulaev et al., 1967b, 1970a,c),unlike nuclei, lack high molecular-weight polyphosphates. Italian researchers (Solimene et al., 1980) recently reported that, in yeast possessing respiring mitochondria at the stage of exponential growth, a distinct peak was detected indicative of the accumulation of low molecularweight polyphosphates with a length of three to eight residues. No such peak could be revealed at this stage of growth in non-respiring yeast. However, these results may be interpreted to indicate that such polyphosphates form and accumulate in respiring yeast not in the mitochondria proper, but rather outside the organelle, from ATP and pyrophosphate (Mansurova et al., 1975a).It is noteworthy that high molecular-weight polyphosphate are absent not only from mitochondria but also from other structures related to energy generation in eukaryotic chloroplasts. This was shown for chloroplasts of Acetabularia mediterranea (Rubtsov et al., 1977) and higher plants (Valikhanov and Sagdullaev, 1979).On purification of A. mediferraneachloroplasts in a sucrose density gradient, the peak of metachromatically stained labile phosphorus compounds was quite distant from the chloroplast fraction (Fig. 2). Electron microscopy revealed that chloroplasts are well preserved under 1000

i3

< -

5.

u

c

.-c 500 'z u 0 .-0

0 0

0

0 Number of fractions

FIG. 2. Distribution of chlorophyll ( 0 pg fraction-'), radioactivity (A pulse min-' fraction-'), and compounds which give metachromatic staining (0)during centrifugation of Acetabularia mediterranea chloroplasts in a gradient of sucrose concentration (0.5-1.5 M, 50,00Og,60 minutes). After phosphorus starvation cells were transferred to a medium containing radioactive phosphate before chloroplasts were isolated. From Rubtsov et al. (1977).

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IGOR S. KULAEV AND VLADIMIR M. VAGABOV

these conditions. These results testify that metabolism and physiological role of high molecular-weight polyphosphates are not directly connected with the respiratory and photosynthetic phosphorylation which are known to operate in mitochondria and chloroplasts. In previous reviews (Kulaev, 1975, 1979) the problem of the occurrence of specific polyphosphate fractions in vacuoles and vesicles of the endoplasmic reticulum was not discussed in detail. However, this question is essential and is at present widely debated in the literature. Its importance stems from the fact that vacuoles and vesicles of the endoplasmic reticulum are specific compartments of eukaryotic cells. Therefore, it is of special interest to provide evidence for the occurrence of high molecular-weight polyphosphates in these subcellular structures of eukaryotes. Indge (1968) was the first to indicate the presence of high molecular-weight polyphosphates in yeast vacuoles. The next step in the investigation of polyphosphate metabolism in yeast vacuoles was initiated by the work of Matile and his associates (Matile, 1978; Urech et al., 1978; Durr et af., 1979; Wiemken el al., 1979; Martinoia et al., 1979; Huber-Walchli and Wiemken, 1979). Employment of methods developed by Wiemken and his colleagues for obtaining purified preparations of intact vacuoles and differential extraction of the cytoplasmic and vacuolar pools of ions and compounds from yeast protoplasts has confirmed the occurrence of inorganic polyphosphates in these cellular structures (Urech er al., 1978; Durr et af., 1979). Of great importance is the conclusion drawn by these authors that, in the course of fractionation, nearly all polyphosphates contained in yeast protoplasts are found in the “gross particulate fraction” which includes mainly vacuoles, nuclei and mitochondria. Though protoplasts contained only about 80% of cellular polyphosphates, the authors inferred that all or nearly all of these compounds were contained in the vacuoles of yeast cells. This conclusion seems to be somewhat erroneous, since polyphosphates disappearing from yeast cells during preparation of protoplasts could be readily degraded to orthophosphate by the closely localized polyphosphatase (see Section II.B, p. 110). Besides, it should be taken into account that nuclei which could also contain some polyphosphates were present in the “gross particulate fraction” and probably to some extent in the purified vacuolar fraction. However, with these reservations in mind, it should be accepted that in the yeast studied by Wiemken et al. (1979) most polyphosphates were present in vacuoles (and possibly in vesicles of the endoplasmic reticulum). Important data were obtained by Diirr et al. (1979) on the chain length of polyphosphates present in yeast vacuoles. As can be seen from Fig. 3, the polyphosphates contained in the vacuoles of yeast cells fall into two fractions on the basis of their chain lengths. The first fraction comprises polyphosphates with a chain length (if) of five units, and the second one had ii values of

PO LY PH0sPHATE M ETA6 0 LI S M IN MIC R 0- 0 R GA NI S MS

-

2.0

95

I-

In

c c W

0 5 .-

a U m W

c

c 0 n c 0

n

v/v,

FIG. 3. Sephadex G-75 filtration of vacuolar polyphosphate. The insert shows a calibration of the column with synthetic polyphosphate of known chain length. From Durr et al. (1979).

15 to 25. This fact, together with the well-known data of Langen and Liss (1958), suggest that in the work of Wiemken e f al. (1979) one part of the vacuolar polyphosphates of yeast may belong to the acid-soluble polyphosphate (Z=5) class and the second to the salt-soluble class (Z= 15-25). These data are in a good agreement with the results of Solimene et al. (1980) who detected substantial amounts of low molecular-weight polyphosphates (Z= 3-8) in stationary respiring and non-respiring yeast, tripolyphosphates being the most abundant. Lusby and McLaughlin (1980) recently detected large quantities of tripolyphosphate (1.8 pmol (lo4cells)-') in Saccharomyces cerevisiae. The concentration of polyphosphates decreased with increasing chain length. However, such a situation is far from being universal in yeasts. Quite recently, French researchers (Beckerich et al., 198I ) failed to detect appreci-

96

IGOR S. KULAEV AND VLADlMlR

M. VAGABOV

able quantities of low molecular-weight polyphosphates in Saccharomycopsis lipolytica. About 40% of the total polyphosphates accounted for had chain lengths of three or five units, and the remainder was salt-soluble. It is not excluded that {hese salt-soluble polyphosphates of S. IipoIytica were also localized preponderantly in vacuoles. The presence of polyphosphates in yeast vacuoles was also estabished with the help of a specific fluorochrome (Allan and Miller, 1980). We also attempted to detect polyphosphates in the vacuole pool of Saccharomyces carlsbergensis using Wiemken's method of differential extraction (Lichko et al., 1982). Our data, summarized in Table 4, suggest that the major portion of the polyphosphates is found in the vacuolar pool. It is noteworthy that polyphosphates are most abundant in vacuoles under conditions of polyphosphate overplus, preceded by a period of phosphorus starvation. Nonetheless, in all cases, some polyphosphates were also detected in other cellular compartments. However, in all studies carried out by the differential extraction method, it remained obscure which cellular compartments contributed to the so-called vacuolar pool. In other words, does the latter term imply only the vacuoles as such, meaning by this the lytic compartment of yeast or other eukaryotic cells as postulated by Matile (1975, 1978), or does it also include vesicles of the endoplasmic reticulum which perform a quite different function in cells of

TABLE 4. Influence of the composition of cultivation medium on the content of phosphorus compounds in Saccharomyces carlsbergensis. After Lichko et al. (1 982) Content (pmol (g wet cells-')) Yeast transferred from complete medium to fresh complete medium (5 hours' growth) Cytoplasm orthophosphate polyphosphate Vacuoles orthophosphate polyphosphate Total cell phosphate

Yeast transferred from phosphate-deficient to fresh complete medium (5 hours' growth)

Yeast transferred from phosphatedeficient medium to fresh phosphate deficient medium (5 hours' growth)

0.65 1.94

0.97 2.90

0.01 1.29

13.71 23.55 191.29

16.29 88.87 321.61

17.10 17.42 141.94

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yeast and other organisms? These vesicles were shown not only to contain polyphosphates (Shabalin et al., 1978,1979), but also to have a specificsystem for their biosynthesis related to formation of glycoproteins (see p. 123). Besides yeast, a substantial portion of polyphosphates was recently shown to occur in the form of intravacuolar polyphosphate granules in other eukaryotes. Cramer et al. (1980) found that, in the N. crassa strain studied, a prominent portion of polyphosphates (at least 50%) was contained in vesicles and vacuoles. The presence of polyphosphate granules was established by electron microscopic examination (see Fig. 4) in cells of the slime mould Dictyostelium discoideum (Gezelius, 1974) and in zoospores of a parasitic aquatic fungus Rosella allomycis during cyst formation (Wool and Held, 1976). Soon after

FIG. 4. Spore of Dictyostelium discoideum with numerous polyphosphate deposits, mainly in small vacuoles. Granules are also seen in elongated vacuoles (upper left) and in two crenate mitochondria (arrows). The section was held in the electron beam to evaporate some of the polyphosphate in the granules. The section was treated with glutaraldehyde, osmium tetroxide and uranyl acetate. Magnification x 66,000.From Gezelius (1974).

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this, the presence of polyphosphate granules in vacuoles of acquatic fungi was confirmed in Blastacladiella emersonii by Hutchinson et al. (1977). In this work, use was made of the X-ray dispersion micro-analytical detection of phosphorus in combination with electron microscopy. This method was first used by Coleman et al. (1972) for detecting polyphosphates in granules in Tetrahymenapyriformis. Using the same technique, polyphosphate granules were detected in vacuoles of Chlorellasp. (Atkinson et al., 1974;Peverly et al., 1978; Adamec et al., 1979)and in Scenedesmus sp. (Tillberg et al., 1979,1980). In the latter organism intravacuolar polyphosphate granules had been detected earlier by cytochemical methods (Sundberg and NilshammarHolmvall, 1975). It can, therefore, be stated that, in eukaryotic cells, a substantial portion of cellular polyphosphates is deposited in the form of polyphosphate granules. However, it remains obscure whether the cell sap of these organisms contains soluble polyphosphates which are isolated from the environment only by the plasma membrane. It appears probable that a portion of the least polymerized polyphosphates happens to be in a free state in the yeast protoplasm. However, such a suggestion should be given experimental support. Results of Ostrovsky et al. (1980), obtained by a 31Pnuclear magnetic resonance 145.78 MHz method of high resolution, point to the possible existence of such a mobile free pool of polyphosphates in Endomyces magnusii (see Fig. 5 ) . It should be noted that the 31Pnuclear magnetic resonance method of high resolution, which is widely employed nowadays for detecting various

I

I

-30

-20

I

-10

I

I

I

I

0

10

20

30

Chemical shift (p.p.m. relative to trimethylphenyl phosphoiadyl)

FIG. 5. A 145.87 MH~-~*P-nuclear magnetic resonance spectrum of cells of Endomyces mangusii (A) with integral intensity (B). Assignment of signals: 1, standard; 2, sugar phosphates; 3,4, intra- and extracellular orthophosphate of hydrocarbons; 5, 6, y-phosphate of nucleoside tri- and diphosphates; 7, a-phosphates of di- and triphosphates; 8, dinucleotides NAD+ and other compounds; 9, derivatives of nucleoside diphosphates; 10, #I-phosphate of nucleoside triphosphate; 11, pofyphosphate. From Ostrovsky et al. (1980).

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phosphorus-containing compounds in cells (Solhany et al., 1975; Burt et al., 1977; Navon et al., 1977a,b), has some limitations. It allows various phosphorus-containing compounds, such as ATP, sugar phosphates and polyphosphates, to be detected only when they occur in cells in a free state. If they are linked, as for example in polyphosphates, to other cellular components, such as proteins or nucleic acids, then they may not be detected by this method. Therefore, conventional chemical analysis should be conducted in parallel with investigations aimed at the quantitative assessment of intracellular polyphosphates by the 31Pnuclear magnetic resonance method of high resolution. Unfortunately no such results are available in the literature to date. 2. Prokaryotes Prokaryotic cells have much simpler structures compared with the simplest eukaryotes, such as yeast, fungi or algae. First of'all they have no nucleus enveloped by a membrane. Instead they have a nucleotid containing DNA strands and nucleoplasm. Moreover, prokaryotes, in particular eubacteria and cyanobacteria, do not contain vacuoles. As for autotrophic bacteria and cyanobacteria, they possess thylacoids (specialized protrusions of the plasma membrane, in which the photosynthetic apparatus of these organisms is localized) and the so-called polyhedral bodies, or carboxysomes, which harbour, according to Stewart and Codd (1975), the key enzyme of photosynthesis ribulose 1,5-diphosphate carboxylase. When discussing localization of high molecular-weight polyphosphates in prokaryotic cells, it should be noted that the latter do not possess the two compartments that eukaryotes have for storing the bulk of intracellular polyphosphates, namely the nucleus proper, limited by the nuclear membrane, and vacuoles. Then where are polyphosphates accumulated in prokaryotic cells? Polyphosphate-containinggranules, or volutin granules, have been unequivocally demonstrated in bacteria, in particular in Spirillum uolutans (Ebel et al., 1958; Drews, 1962; Hughes and Muhammed, 1962; Kulaev and Belozersky, 1962). Various cytochemical methods were elaborated for detecting volutin-like granules in different micro-organisms(Keck and Stich, 1957;Ebel et al., 1958; Talpasayi, 1963). Cytological methods for detecting polyphosphate granules were boosted by the use of the electron microscope (Niklowitz and Drews, 1957; Ebel et al., 1958; Ris and Singh, 1961; Drews, 1962; Jost, 1965; Voelz et al., 1966; Friedberg and Avigad, 1968;Jensen, 1968, 1969). For early references on detection of polyphosphate granules we recommend the reader to refer to Kuhl (1960, 1962, 1974), Drews (1962), Harold (1966), Shively (1974) and Kulaev (1979).

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In recent years, the most important and comprehensive data in this field were obtained by Jensen and his coworkers (Jensen, 1968, 1969; Jensen and Sicko, 1974; Sicko-Goad et al., 1975; Sicko-Goad and Jensen, 1976; Lawry and Jensen, 1979; Baxter and Jensen, 1980a,b). These works were carried out with cyanobacteria which at present are given much attention because of the necessity of solving a number of applied problems concerning water pollution. Special attention is paid to the possible use of blue-green algae as accumulators of substantial amounts of phosphates in the form of polyphosphates. This problem arose from severe pollution of inland water bodies, especially in industrialized countries, with various detergents of which sodium tripolyphosphate is the most abundant pollutant. Using electron microscopy with the cyanobacteria Nostoc prunifarme (Jensen, 1968), Plectonema boryanum (Jensen, 1969; Jensen and Sicko, 1974; Sicko-Goad and Jensen, 1976) and Anacystis nidulans (Lawry and Jensen, 1979), Jensen and his colleagues investigated the accumulation of polyphosphate granules under various cultivation conditions. In these experiments special emphasis was laid on the accumulation by cyanobacteria of polyphosphate granules under conditions approximating those of inland water bodies, i.e. conditions of phosphorus and sulphur starvation. Waters in such reservoirs are known to contain about 0.01 pg phosphorus I - ’ (Jensen and Sicko, 1974), and this creates conditions of phosphorus starvation for micro-organisms. When large amounts of industrial and domestic detergents, in particular tripolyphosphate, enter inland water bodies, an intensive “fluorescence” of cyanobacteria occurs leading to contamination of vast reservoirs of drinking water. This has become quite a serious problem, and many laboratories all over the world are endeavouring to solve the problem. From electron microscope studies on the localization of polyphosphate granules in Plectonema boryanum cultured in medium containing the normal content of phosphorus, as well as under conditions of phosphorus starvation followed by subsequent “phosphate overplus” in medium enriched with phosphorus, Jensen drew the following conclusions (Jensen and Sicko, 1974). In normal growth conditions, polyphosphate granules are found mainly on DNA fibrils and in a zone enriched with ribosomes. Under conditions of phosphorus starvation, in addition to these sites there was a zone of average electron density formed in the region of nucleoplasm, apparently as the result of degradation of a portion of nucleic acids. Under conditions of “phosphate overplus”, polyphosphates accumulated in the region of nucleoplasm, and polyphosphate granules appeared in the polyhedral bodies directly involved in the dark reactions of photosynthesis in cyanobacteria (Stewart and Codd, 1975). In certain cells, polyphosphate granules formed near thylakoids which in these organisms contain chlorophyll and perform phosphorylation reactions.

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Thus electron microscopy helped to establish that, in blue-green algae, polyphosphate granules are localized in most cases in the region of nucleoid (DNA fibrils and nucleoplasm) rich in ribosomes and near the subcellular structures participating in photosynthesis. Similar reports for cyanobacteria were made by other authors using the same approach (Vaillancourt et al., 1978; Barlow et al., 1979).The data obtained, at least those for localization of polyphosphate granules in the vicinity of the bacterial nucleoid, correlate well with previous findings using the same method on heterotrophic prokaryotes (Drews, 1962;Voelz et al., 1966; Friedberg and Avigad, 1968; Deinema et al., 1980). However, the data of Jensen and other cytologists have one shortcoming; there was no chemical determination of polyphosphates carried out in parallel with the electron microscope studies. In some studies, Jensen (Sicko-Goad and Jensen, 1976) attempted to compare the electron microscope picture with accumulation of total phosphorus in certain fractions. Still, this is evidently insufficient for allowing a rigorous conclusion to be made about the polyphosphate nature of granules detected in cells. As was referred to in the Introduction, Kessel (1977) and Jensen and his coworkers (Sicko-Goad et al., 1975; Baxter and Jensen, 1980a) combined electron microscopy with X-ray energyaispersion microanalysis(Colemanet al., 1972)to reveal the chemical nature of these granules. This method enables one to locate in situ phosphorus, sulphur, calcium, potassium and carbon dioxide in specific cellular compartments. However, this method does not provide information as to the forms of compounds of phosphorus, carbon and other elements occurring in the cellular inclusions. Nevertheless, the results obtained with this technique allow one to establish the phosphate nature of granules and to determine which cations may be present in these granules. In his recent investigations, Jensen (Baxter and Jensen, 1980a,b) showed that, under ordinary cultivation conditions, appreciable amounts of potassium and comparatively low quantities of calcium and magnesium are present, in addition to phosphorus, in polyphosphate granules of the cyanobacterium Plectonema boryanum. Under special conditions, when the medium contains an excess amount of a particular metal, such as Mg2+,Ba2+,Mn2+or Zn2+, they accumulate in large quantities in polyphosphate granules. Strontium is also known to be able to accumulate in considerable amounts in cells of these algae, not in polyphosphate granules, but in some inclusions containing, together with K + and Ca2+,sulphur instead of phosphorus. Useful information on localization of polyphosphates in bacterial cells may be provided by 31Pnuclear magnetic resonance of high resolution at 145.78 MHz (Ferguson et al., 1979; Ostrovsky et al., 1980). Ostrovsky et al. (1980), for example, believed that a marked increase in the intensity of a low-field 3'P nuclear magnetic resonance signal shift for polyphosphates, observed when cells of Mycobacterium smegmatis were treated with ethylene diamine

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tretra-acetic acid, points to localization of certain amount of mobile inorganic polyphosphates in the periplasmic region of Mycobacterium smegmatis, i.e. outside the cytoplasmicmembrane. These findings, though not yet confirmed by other investigators, appear to be extremely important, since in eukaryotes also a portion of polyphosphate is localized outside the cytoplasmic membrane, and only part of polyphosphates occur in the form of polyphosphate granules inside the cell. Some recent research on the chemical fractionation of polyphosphates in a number of bacteria also support, albeit indirectly, such a conclusion (Bobyk et al., 1980;Egorova et al., 1981; Nikitin et a)., 1979, 1983). Bobyk and his coworkers isolated and qualitatively assessed various fractions of polyphosphates from Bdellovibrio bacteriovorus, and showed that, in this bacterial parasite, most polyphosphates occur in the form of acid-insoluble highly polymerized fractions. A similar situation was revealed in prosthetic oligotrophic bacteria, namely Tuberoidobacter and Renobacter spp., studied by Nikitin et al. (1979). In contrast, in the thermophilic Thermusflavus growing at 6570°C most polyphosphates were detected in the form of low-polymeric fractions PolyP~and PolyP~.Comparison of these results with similar data obtained for eukaryotes suggests that Bdellovibrio sp. and prosthetic oligotrophic bacteria populating the atmosphere, under conditions of permanent starvation, contain predominantly outwardly localized highly polymerized polyphosphates, whereas in thermophiles, polyphosphates of relatively low molecular weight are mainly localized within the plasma membrane. Similar examples may be found in Kulaev’s (1979) monograph summarizing all previous publications in the field. This book provides data, though rather scanty to date, on isolation in a pure state of polyphosphate granules from cells of some micro-organisms. Recently, Jones and Chambers (1975) succeeded in isolating these granules from Desulfovibrio gigas (Fig. 6). The granules proved to be soluble only in 1 M HCI, and insoluble in water, 1 M NaOH, ethanol, ether and other organic solvents, and appeared to be tripolyphosphate of magnesium (Mg~(P301o)s). It is interesting that in this work, infrared spectroscopy (Corbridge and Lowe, 1955) was employed to establish the precise nature of these granules. Using this and other methods, it was rigorously proved that the granules are composed of magnesium tripolyphosphate. It should be noted that these granules formed only in sulphate-reducing bacteria Desulfovibrio spp. and after repeated inoculations of the culture. The possibility of accumulation of large amounts of magnesium tripolyphosphate in bacterial cells in the form of volutin granules is a novel and very interesting fact. These results correlate, to some extent, with the investigations of Rosenberg (1 966) and Simkiss (1981). In both of these studies (in the first one with the protist Tetrahymena puriformis, and in the second with the

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FIG. 6. Electron micrograph of isolated granules from Desulfovibrio gigus. From Jones and Chambers (1975).

hepatopancreas of the mollusc Helix aspersa) the granules isolated were composed almost entirely of Ca-Mg pyrophosphate.

B . ENZYMES INVOLVED IN BIOSYNTHESIS A N D DEGRADATION OF POLYPHOSPHATES

Investigations on the localization of high molecular-weight polyphosphates in cells of various organisms were carried out simultaneouslywith studies on the enzymes involved in their metabolism. 1. Polyphosphate: ADP Phosphotransferase

The enzyme polyphosphate: ADP phosphotransferase (EC 2.7.4. I) was the first to be identified. This enzyme catalyses the transfer of the high-energy

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IGOR S. KULAEV AND VLADlMlR M. VAGABOV

phosphate residue from ATP to polyphosphates and back from polyphosphates to ADP to form ATP. The history of the discovery of this enzyme and its occurrence in various organisms have been described in Kulaev’s monograph (1979). Polyphosphate: ADP phosphotransferase (polyphosphate kinase) was first isolated from Escherichia coli and purified as described by Kornberg et al. (1956). It catalyses the following reaction: polyphosphate

ATP+ (polyphosphate),,

ADP + (polyphosphate),+1

(1)

kinase

It was later isolated from other micro-organisms (Kulaev, 1979). After the discovery of the enzyme, some researchers (Hoffmann-Ostenhof, 1962; Y oshida, 1962) began to consider high molecular-weight polyphosphates as peculiar microbial phosphagens, i.e. as compounds that can be synthesized and utilized in micro-organisms only via the ADPctATP system, similarly to creatine and arginine phosphates in animal tissues. However, further studies of enzymic reactions involving high molecularweight polyphosphates have shown the limited nature of this hypothesis on their physiological role in micro-organisms. Moreover, our findings (Kulaev and Rozhanets, 1973; Kulaev et al., 1974a)indicate that both high molecularweight polyphosphates and polyphosphate kinase are present in animal tissues. They have been found in the rat brain, i.e. in the tissue for which the existence of the classic phosphagenic creatine phosphate-creatine kinase system has long been known. This fact, considered as such, suggests that the physiological role of high molecular-weight polyphosphates cannot be confined to the function of common “phosphagens”, particularly “microbial phosphagens”. This statement is corroborated by the detection of this enzyme in yeast vacuoles (Shabalin et al., 1977). Further, Schwencke (1978) reported the presence in these cellular structures of polyphosphate depolymerase (see below) which apparently also plays some role in metabolism of the vacuolar polyphosphate pool. However, it is not excluded that the function of this enzyme in vacuoles is basically connected with transport of polyphosphates through the tonoplast. In some prokaryotic organisms, polyphosphate kinase may possibly stand in the centre of the entire polyphosphate metabolism, and be the key metabolic enzyme. This idea is supported by the fact that, in mutants defective in polyphosphate kinase, such as Aerobacter aerogenes (Harold and Harold, 1963) and Anacystis nidulans (Vaillancourt et al., 1978), accumulation of high molecular-weight polyphosphates stopped. However, in fungi, for example Neurospora crassu, this enzyme probably does not occur at all (Kulaev et al., 1971). Nevertheless, this organism accumulates and metabolizes these compounds. Although polyphosphate: ADP phosphotransferase has been found in the

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slime mould Dictyostelium discoideum, its activity proved to be low at all stages of the differentiation of this organism (Gezelius, 1974). The enzyme did not materially contribute to the process of ATP formation which testifies convincingly against the phosphagenic function of polyphosphates detected in large amount in the fungus (Gezelius, 1974; Al-Rayess et al., 1979). On the other hand, it has been shown in our recent work (Butukhanov et al., 1979) that, in a Corynebacterium sp., in production of ATP from exogenous adenine, polyphosphate: ADP transferase contributes significantly to synthesis of ATP which is accumulated in large amounts (0.6-1 .O mg ml-I) in the culture medium. During the period of maximum synthesis of ATP from exogenous adenine in autolysed cells of the strain, the activity of polyphosphate: ADP phosphotransferase increased greatly. However, it does not follow from these data that the enzyme and its substrate, high molecularweight polyphosphates, are essential for ATP formation inside cells of a normally growing culture of Corynebacterium sp. 2. Polyphosphate: Adenosine Monophosphate Phosphotransferase Soon after detection of polyphosphate kinase in some micro-organisms, Winder and Denneny (1957) found another enzyme which suggested that metabolism of high molecular-weight polyphosphates in micro-organisms could, in many ways, be connected with that of adenine nucleotides. This enzyme, polyphosphate: AMP phosphotransferase, was isolated and partially purified from mycobacteria in Ebel’s laboratory (Dirheimer and Ebel, 1965). The enzyme is responsible for the reaction:

+

polyphosphate: A M P

+

AMP (polyphosphate), , A ADP (polyphosphate),phosphotransferase

1

(2)

Hitherto this enzyme had been found only in mycobacteria and corynebacteria (Kulaev, 1979). Recently, an unsuccessful attempt was made to detect polyphosphate: AMP phosphotransferase in the slime mould Dictystelium discoideum (Gezelius, 1974).On the other hand, the occurrence of this enzyme in corynebacteria and its involvement in ADP synthesis was shown indirectly by Butukhanov et al. (1979) in a Corynebacterium strain that produces ATP from exogenous adenine.

3. Polyphosphate (Metaphosphate)-Dependent NAD + Kinase Quite recently, another enzyme linking high molecular-weight polyphosphates (metaphosphates) with energy metabolism, i.e. polyphosphate (metaphosphate)-dependent NAD+-kinase was detected in some eubacteria by Murata et al. (1980). In the course of studies on the specificity of this enzyme

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for the phosphoryl-group donor, these authors found it to be specific for metaphosphate manufactured by Katayama (Japan). However, as pointed out by Kulaev (1979), chemists estabished that the so-called high molecularweight “metaphosphate” was no more than a linear high molecular-weight polyphosphate of the Graham salt type. Murata et al. (1980) did not investigate specifically “metaphosphate” from Katayama. In this connection, the Japanese researchers seem to have dealt with a preparation of linear polyphosphates. They found polyphosphate (metaphosphate)-dependent NAD+ kinase in species of Acetobacter, Achromobacter, Brevibacterium, Corynebacterium, and Micrococcus, but failed to detect it in species of Escherichia, Proteus and Aerobacter. This enzyme catalyses the following reaction: NAD

+

+polyphosphate (metaphosphate),eNADP + polyphosphate (metaphosphate),-

I

(3)

This enzyme differs from the ATP-dependent NAD+ kinase in pH optimum, thermostability and a number of other properties. 4. Polyphosphate: D-Glucose 6-Phosphate Phosphotransferase

The Polish scientist Szymona (Szymona, 1962;Szymona and Ostrowski, 1964) detected another enzyme of polyphosphate metabolism, polyphosphate: D-glucose 6-phosphotransferase (polyphosphate-glucokinase,EC 2.7.1.63) in mycobacteria, and showed that it catalysed the specific transfer of the phosphate group from high molecular-weight polyphosphates to glucose to form D-glucose 6-phosphate:

+

polyphosphate-

D-Glucose (polyphosphate), Y-_-D-glucose 6-phosphate + glucokinase

(polyphosphate),-

(4) 1

The discovery of this enzyme in this and related micro-organisms(Szymona et al., 1967; Uryson and Kulaev, 1968; Szymona et al., 1977; Szymona and Szymona, 1979; Eroshina et al., 1980; Ziizina et al., 1981) has shown that high-energy phosphate residues of polyphosphates can be utilized directly without the participation of the ADP-ATP system. This finding has also demonstrated that, in some cases, polyphosphates can perform the function that is normally carried out by ATP itself. The reaction described by Szymona and his colleagues is identical with the classical hexokinase reaction during which glucose undergoes phosphorylation caused by ATP. Szymona and his coworkers, in their recent work on purification and specific functions of polyphosphate glucokinase in Nocardia

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minima (Szymona and Szymona, 1979), Mycobacterium tuberculosis H37Ra (Szymona et al., 1977; Pastuszak and Szymona, 1980), and a number of other mycobacteria (Szymona and Szymona, 1978),showed that electrophoretically homogeneous preparations of polyphosphate glucokinase exhibited certain activity also with ATP as the phosphate donor. However, it is not yet clear whether they dealt in both cases with a mixture of two enzymes having polyphosphate hexokinase and ATP hexokinase activities, or a single protein possessing two different active centres one of which operates with ATP and the other with high molecular-weight polyphosphates. H. G. Wood (personal communication) reported his success in distinguishing between the two activities in the course of isolating polyphosphate hexokinase from Propionibacterium shermanii. However, his highly active preparation exhibited low ATP-hexokinaseactivity. It is noteworthy that, in the above works, Szymona and his colleagues have established the existence in various bacteria of isoforms of polyphosphate glucokinase having different molecular weights. Nocardia minima, for example, was found to have three isoenzymes with molecular weights of 59,000, 76,000 and 150,000, respectively (Szymona and Szymona, 1979).The prevailing fraction, which accounted for 80% of the total polyphosphate glucokinase activity, showed preference for polyphosphate but one of the minor fractions preferred ATP. Summing up, this very important enzyme of polyphosphate metabolism is now receiving the most detailed investigation and, before long, one should expect a breakthrough in understanding of the mechanism of its operation. It is of interest that, in recent years, researchers undertook a number of investigations aimed at detecting polyphosphate glucokinase activity in various organisms, of which special mention should be made of the progress in detecting this activity in a bacterial parasite Bdellovibrio bacteriovorus (Bobyk et al., 1980), as well as in the recently described oligotrophic bacteria Renobacter vacuolatum (Nikitin et al., 1983). Of importance also is the fact that, in Dictyostelium discoideum and other fungi (Kulaev, 1979),this enzyme was not detected (Gezelius, 1974). In conclusion, it should be recalled that Szymona and his coworkers also revealed the existence of a whole series of adaptive enzymes that are responsible for transfer of phosphate group from high molecular-weight polyphosphates to other sugars and their derivatives, namely fructose, mannose and glucuronic acid (Szymona and Szumilo, 1966; Szymona et al., 1969).

5 . I ,3-Diphosphoglycerate: Polyphosphate Phosphotransferase In addition to the enzymes already referred to, another enzyme catalysing synthesis of high molecular-weight polyphosphates was found (Kulaev et al.,

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1968; Kulaev and Bobyk, 1971; Kulaev, 1979). This enzyme, which was first detected in N. crassa and later in other micro-organisms including fungi and eubacteria (Kulaev et al., 1971), is involved in transfer of a high-energy phosphoryl group from 1,3-diphosphoglyceric acid to polyphosphate. Thus, the enzyme, 1,3-diphosphogIycerate:polyphosphate phosphotransferase (EC 2.7.4.17), participates in a reaction similar to the well known reaction of ATP formation during glycolytic phosphorylation.

+

+

1,3-Diphosphoglycerate (polyphosphate),~3-phosphoglycerate

(polyphosphate),+ I

(5)

In our work, it has been shown that this enzymic system is controlled by adenine nucleotides,principally ATP (Kulaev, 1979).This was first revealed in a mutant of N. crassa deficient in adenine synthesis (Kulaev et al., 1968; Kulaev and Bobyk, 1971). In all cases when this enzymic activity was found in micro-organisms, it proved to be fairly low and obviously did not provide for biosynthesis of all polyphosphate fractions (Kulaev e f al., 1971, 1973a). Apparently, in N. crassa, this enzyme participates primarily in metabolism of low molecularweight polyphosphates contained in polyphosphate granules and it was in these granules that it was detected by Kulaev and Konoshenko (1971a). In conclusion, it may be noted that, of all micro-organisms studied, highest activity of the enzyme has been detected in Bdellovibrio bacteriovorus (Bobyk et al., 1980).If one takes into account that polyphosphate glucokinasehas also been found in this organism, then an inference may be drawn about a close relation between polyphosphate metabolism and glycolysis. 6. Polyphosphate Polyphosphohydrolases In addition to the already mentioned phosphotransferases, two types of phosphohydrolases are involved in metabolism of high molecular-weight polyphosphates (Kulaev, 1979). Phosphohydrolases of one type split high molecular-weight polyphosphates within the chain into smaller fragments. These are the so-called polyphosphate polyphosphohydrolases (polyphosphate depolymerase, EC 3.6.1.10). They catalyse the reaction depicted below: (polyphosphate),+ water

polyphosphate

depolymerase

(polyphosphate),-,

+(polyphosphate),

In cells of eukaryotes, namely yeasts and fungi, several different polyphosphate depolymerases cleaving polymers of different lengths in the middle of the chain (Ingelman and Malmgren, 1947, 1948; Mattenheimer, 1956a,b,c;

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Kritsky et al., 1972) have been detected. Kritsky et al. (1972) showed that polyphosphate depolymerases which split specific fractions of polyphosphates exhibit their activities at various stages of growth of N. c r ~ ~ s a . Therefore, their action on specific fractions of polyphosphates is timed to a definite physiological state of this fungus. The problem of the intracellular localization of polyphosphate depolymerases appears to be of special interest. In our studies on localization of polyphosphate depolymerases which split high molecular-weight polyphosphates (n= 180and290)inN.crassa,itwasshownthat theiractivityismainlyexhibited at the cell periphery. In protoplasts, after removal of the wall, polyphosphate depolymerasesretain about 10-1 5% of their activity on the already mentioned substrates. As to specific cellular structures, activity was found in fractions of nuclei, as well as in vesicles of the endoplasmicreticulum and vacuoles (Kulaev et al., 1972a).These results were recently supplementedby those of Schwencke (1 978) who revealed the presence of polyphosphate depolymerase-splitting polyphosphates of lower molecular weight in yeast vacuoles. It may be suggested that polyphosphate depolymerases play an extremelyimportant role in polyphosphate metabolism linking the pools (fractions)of these compounds in cells, particularly in those of lower eukaryotes. According to Kritsky and Chernysheva ( I 973), polyphosphate depolymerases participate in translocation of specific polyphosphate fractions through cell membranes. The authors believe that the energy released during polyphosphate cleavage may be utilized for translocation of fragmented molecules through membranes. From this point of view, it is not surprising that polyphosphate depolymerase activities were revealed in regions of plasma membrane, nuclei and vacuoles, i.e. at the sites of localization of basic specific pools (fractions) of polyphosphates in cells of lower eukaryotes. 7. Polyphosphate-Phosphohydrolases Another group of polyphosphatases split one terminal phosphate residue from each polyphosphate molecule. Most investigatorsshare the opinion that these enzymes, called polyphosphate phosphohydrolases or simply polyphosphatases (EC 3.6.1.1 l), are responsible for the occurrence of the following reaction:

+

(polyphosphate), water-+(polyphosphate),- I +Pi

(7)

Thus, enzymic breakdown of polyphosphates occurs by processes similar to the enzymic degradation of other biopolymers, for example, proteins and polysaccharides. Polyphosphate depolymerase and polyphosphatase are, in other words, endo- and exopolyphosphate phosphohydrolases that catalyse

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cleavage of internal and terminal phosphoanhydride bonds. However, the results of numerous studies reported earlier (Kulaev, 1979), as well as quite recent experimental findings, suggest that both mechanism of action and physiological role of these enzymes may differ markedly. Polyphosphatases capable of hydrolysing high molecular-weight polyphosphates to orthophosphate were found in many organisms (Kulaev, 1979).The number of organisms in which these enzymes were found has increased markedly in recent time and includes, among others, Nocardia erythropolia and Brevibacterium sp. (Eroshina et al., 1980), Corynebacterium sp. (Butukhanov et al., 1979), Bdellovibrio bacteriovorus (Bobyk et al., 1980), Streptomyces levoris (Zuzina et al., 1981), Tuberoidobacter mutans and Renobacter vacuolatum (Nikitin et al., 1983), Dictyostelium discoideum (Gezelius, 1974) and Candida guillermondii (Kulaev et al., 1974b).It is noteworthy that a rather high polyphosphatase activity was revealed in many soil micro-organisms (Aseeva et al., 1981), for example, in bacteria of the genera Bacillus and Micrococcus, fungi of the genera Aspergillus and Penicillium and coryneforms of the genus Arthrobacter. The fungi Aspergillus wentii and Cladosporium herbarum released their polyphosphatase into the cultivation medium; in these organismsone would suggest that outside the cells of these fungi, the enzymes are capable of digesting polyphosphates available in soil into orthophosphate. Remarkably, Aseeva et al. (1981) revealed polyphosphatase activity in sterile soil. These authors showed also that many soil organisms could grow intensively on a medium with polyphosphates as the sole form of phosphate. These substrates are degraded therewith by polyphosphatase to orthophosphate which is assimilated. Other researchers succeeded in finding polyphosphatase activity (towards polyphosphate with n =40) in sterile roots of the cotton plant (Valikhanov and Sagdullaev, 1979; Igamnasarov and Valikhanov, 1980). Owing to this activity, cotton plants could assimilate the phosphorus of polyphosphates available in the isolation medium more efficiently than when the same amount of phosphate was available in the form of orthophosphate. These data point to an important part played by polyphosphates of micro-organisms and plants in the phosphorus cycle in the soil, as well as to promising prospects for the use of polyphosphates as phosphorus fertilizers. It may be of interest that we have not detected any polyphosphatase hydrolysingpolyphosphates of high molecular-weight in the phytopathogenic fungus Phytophthora infestans (Sysuev et al., 1978). This may be a specific feature of polyphosphate metabolism in microbe-parasite associations which proliferate in the cells and tissues of plants rather than the soil. The problem of intracellular localization of polyphosphatases appears to be very important. Previous reports (see Kulaev, 1979) as well as recent investigations (Nesmeyanova et al., 1975a, 1976; Severin et al., 1975;

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Trilisenko et al., 1980,1982)indicated that polyphosphatase hydrolysing high molecular-weight polyphosphates to orthophosphate was essentially localized on the outer side of the plasma membrane of eukaryotes and prokaryotes. It was shown that in both Neurospora crassa (Kulaev, 1979)and Escherichia coli (Nesmeyanova et al., 1975a, 1976; Severin et al., 1975) a significant portion of the polyphosphatase was membrane-bound and therefore could be detached only by treatment with detergents, like Triton X-100.However, the polyphosphatases of N. crassa and E. coli differ markedly. In bacteria, polyphosphatase, or rather its biosynthesis and secretion into the periplasm, is repressed by exogenous phosphate (Harold, 1966; Nesmeyanova et al., 1975a,b, 1976; Maraeva et al., 1979; Kulaev, 1979),whereas in fungi this does not occur (Umnov et al., 1974a; Kulaev, 1979; Trilisenko et al., 1981). Investigations of Nesmeyanova et al. (1976) and those of Maraeva et al. (1979) showed that, during derepression of polyphosphatase synthesis in E. coli under conditions of phosphorus starvation, a substantial portion of this enzyme was transferred from the membrane to the periplasm. It was also found that E. coli polyphosphatase had complex regulatory relations with other phosphohydrolases, and some membrane proteins, of the bacteria. This fact is discussed in greater detail on p. 136. When discussing polyphosphatases hydrolysing high molecular-weight polyphosphates to orthophosphate, mention should be made of the detection of multiple forms of these enzymes in eukaryotic micro-organisms. In particular, this was shown for Endomyces magnusii (Afanas’eva el al., 1976) and Neurospora crassa (Trilisenko et al., 1982). It should be noted too that, even in the “leaky” mutant of N. crassa marked by a five-fold decrease in polyphosphatase activity, the same two isozymes are preserved compared with the initial culture. All of the information presented in this section about polyphosphatases concerns only those that hydrolyse high molecular-weight polyphosphates. Our studies on the polyphosphatase from N. crassa showed that this enzyme hydrolysed, with fairly high and nearly the same rates, polyphosphates of different degrees of polymerization (Kulaev, 1979). However, N. crassa was found to possess a specific enzyme that hydrolysed tripolyphosphate to orthophosphate (Kulaev et al., 1972b; Umnov et al., 1974b; Egorov and Kulaev, 1976). It was called tripolyphosphate hydrolase (EC 3.6.1.25). Tripolyphosphatase activity was found in many organisms (Kulaev, 1979). It has also been detected in some bacteria: Nocardia erythropolus (Eroshina er al., 1980), Tuberoidobacter mutans (Nikitin et al., 1983), Renobacter vacuolatum (Nikitin et al., 1983), Escherichia coli (Nesmeyanova et al., 1975b), Streptomyces levoris (Ziizina et al., 198l), Bdellovibrio bacteriovorus (Bobyk et

112

IGOR S. KULAEV AND VLADlMlR M . VAGABOV

al., 1980),andinaparasiticfungus Phytophthora infestans(Sysuevetal., 1978). The intracellular localization of tripolyphosphatase was investigated in N. crassa (Kulaev et al., 1972b; Kulaev, 1979). This study provided convincing proof that the enzyme is mainly localized in the mitochondria of N . crassa. It should be underlined that, in this fungus, compartmentation of the enzyme under consideration is quite different from that of the polyphosphatase hydrolysing high molecular-weight polyphosphates. It remains obscure where and in which way tripolyphosphatase is localized in cells of prokaryotic organisms. Igamnasarov and Valikhanov (1980) reported a high extracellular tripolyphosphatase activity in sterile cotton seedlings in a medium deficient in phosphate. It is noteworthy that, in some instances, tripolyphosphatase may be absent from cells of micro-organisms. The data of Jones and Chambers (1975) point indirectly to this fact. Without special precautions and using simple techniques, they succeeded in isolating, from the bacterium DesuIfovibrio gigas, polyphosphate granules composed of pure magnesium tripolyphosphate. If tripolyphosphatase were contained in these bacteria, then under the conditions used for isolating the granules the tripolyphosphate would have undoubtedly been hydrolysed. 8. Variations in the Enzymes of Polyphosphate Metabolism in Micro-Organisms

Investigation of the metabolism of high molecular-weight polyphosphates in various organisms has revealed dramatic variations in the sets of enzymes involved (Kulaev, 1979). Polyphosphate hexokinase has so far been detected only in organisms that fall into the actinomycetes classified according to Krasil'nikov (1949; Szymona et al., 1967; Uryson and Kulaev, 1968, 1970; Kulaev et al., 1971, 1973a, 1976; Uryson et al., 1973, 1974; HoStalek et al., 1976; Eroshina et al., 1980; Murata et al., 1980; Ziizina el al., 1981). In addition to these organisms, polyphosphate hexokinase was recently detected in Bdellovibrio bacteriovorus (Bobyk et al., 1980) and an oligotrophic bacterium Renobacter vacuolatum (Nikitin et al., 1983). However, the systematic position of these exotic bacteria is not yet clear. The group of micro-organisms claimed by Krasil'nikov to be actinomycetes, in particular mycobacteria, corynebacteria and propionic bacteria, contain practically all known enzymes of polyphosphate metabolism (Kulaev, 1979). In contrast, Aerobacter aerogenes according to Harold (1966) and Murata et al. (1980), has only two enzymes of this set; these are polyphosphate kinase, involved in synthesis of polyphosphates, and a polyphosphatase, which hydrolyses them to orthophosphate. At the present time, not all

POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS

113

enzymes of polyphosphate metabolism have been detected in eukaryotes. As already indicated, polyphosphate kinase has not been found yet in N.crassa, and polyphosphatase has not been found in Phytophthora infestans. Nevertheless, in most other eukaryotic micro-organisms all enzymes are detectable. Moreover, owing to the more complex compartmentation of cells, and cellular metabolism as a whole, they apparently have more complex enzymic systems for both synthesis and utilization of polyphosphates. This will be discussed later (p. 114). So far, metabolism of high molecular-weightpolyphosphates, the enzymes involved, and their intracellular localization, have been best studied in heterotrophic eukaryotes. In this connection, Fig. 7 shows a schematic localization of different enzymes of polyphosphate metabolism in an abstract cell of a heterotrophic eukaryote. As for autotrophic eukaryotes, their metabolism, as well as enzymes involved in polyphosphate transformations, have not been sufficiently studied. Polyphosphate kinase, for example, has been found only in three representatives of autotrophic organisms; these are

1

4 5 9

2

3

6

10

7

8

FIG. 7. Localization of polyphosphates and enzymes ofpolyphosphate metabolism in a typical cell of heterotrophic eukaryotic micro-organisms. 1, indicates acid-soluble polyphosphates (Polyp& 2, salt-soluble polyphosphates (PolyP2); 3, acid-insoluble polyphosphates (PolyP3,d,s);4, polyphosphate: ATP phosphotransferase; 5, 1,3-phosphoglycerate: polyphosphate phosphotransferase; 6, biosynthesis of polyphosphate connected with mannan synthesis; 7, biosynthesis of polyphosphate connected with nucleic acid synthesis; 8, tripolyphosphatase; 9, polyphosphatase; and 10, polyphosphate depolymerase.

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IGOR S. KULAEV AND VLADlMlR M. VAGABOV

the green sulphur bacterium Chlorobium thiosulphatophilium (Hughes et al., 1963; Cole and Hughes, 1965), the green alga Chlorella sp. (Iwamura and Kuwashima, 1964) and Acetabularia mediterranea (Rubtsov and Kulaev, 1977). In the last organism, a polyphosphatase hydrolysing highly polymeric polyphosphates to orthophosphate was also detected in the cell-free extract. In A. mediterranea, activities of both of the enzymes detected, polyphosphatase and polyphosphate kinase, increased markedly during growth of this alga under conditions of phosphorus starvation.

C . METABOLISM OF POLYPHOSPHATES IN EUKARYOTES

As already mentioned, the metabolism and role of inorganic polyphosphates have been most completely studied in fungi and yeast. Therefore, the present section will be devoted to a detailed discussion of investigations conducted with these eukaryotic micro-organisms.

1. Yeasts and Fungi as Representatives of Heterotrophic Eukaryotic Micro-Organisms Yeasts are micro-organisms in which polyphosphates were not only first discovered (Liebennann, 1888) but in which their metabolism was also best studied (see reviews by Kulaev and Belozersky, 1962; Langen et al., 1962; Hoffmann-Ostenhof, 1962; Yoshida, 1962; Harold, 1966; Dawes and Senior, 1973; Matile, 1978; Kulaev, 1975, 1979; and articles by Weimberg, 1975; Ludwig et al., 1977; Diirr et al., 1979; Tijssen et al., 1980; Lusby and McLaughlin, 1980). Metabolism of polyphosphates has been fairly well studied in Neurospora crassa (Harold, 1966; Kulaev, 1979; Cramer et al., 1980; Trilisenko et al., 1980, 1982) and Aspergillus niger, Penicillium chrysogenum (Kulaev, 1979), Physarum polycephalum (Sauer et al., 1969; Goodman et al., 1969; Hildebrandt and Sauer, 1977; Sokolovsky and Kritsky, 1980), Dictyostelium discoideum (Gezelius, 1974; Al-Rayess et al., 1979) and in a number of parasitic fungi (Bennett and Scott, 1971; Wool and Held, 1976; Sysuev et a/., 1978; Umnov et al., 1981). Polyphosphate metabolism proved to be similar in all of the yeasts and fungi studied. Therefore, on the basis of all available data, it would be appropriate to attempt to draft a scheme of polyphosphate metabolism common to all these micro-organisms. Yet, early work on polyphosphate metabolism in fungi and yeasts (Kulaev and Belozersky, 1962; Langen et al., 1962; Harold, 1966; Kulaev, 1979) showed that a metabolic link existed

POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS

115

between different fractions of polyphosphates having, as we now know, different chain lengths and dissimilar intracellular localization. Experiments with radioactive phosphate indicated that the high molecular-weight polyphosphates closest to the surface may be degraded, at particular stages of growth of these micro-organisms, to less polymerized fractions localized inside the cells. Thus, the possibility of transformations: PolyP+PolyPd+ PolyPpPolyP2+PolyPI was demonstrated. However, many questions remained unclear. For example, how do the most polymerized fractions of polyphosphates produce further less polymerized fractions? Are there independent pathways for synthesis and utilization of each of these fractions? In recent years it has become clear that specific fractions of polyphosphates represent different pools of these compounds characterized by specific metabolic features and related by their functions and biogenesis to particular cellular compartments (Kulaev, 1973a,b; Kulaev and Konoshenko, 1971a,b; Kulaev et al., 1972a,b; Konoshenko et al., 1973). Only one enzyme of polyphosphate metabolism (1,3-diphosphoglycerate:polyphosphate phosphotransferase) was found in the cytoplasmic volutin-like inclusions of N. crassa (Kulaev and Konoshenko, 1971a).Therefore, it may be concluded that metabolism of the Polyp, fractions localized in the cellular inclusions of N. crassa is very closely related to glycolysis. Formation and utilization of polyphosphates in these volutin-like granules may well be one of the mechanisms for regulating glycolysis in this fungus. We showed that the activity of this enzyme increased drastically in N. crmsa when in these fungal cells ATP synthesiswas inhibited by 8-aza-adenine(Kulaev et al., 1968).Thus, it may be suggested that polyphosphates of the plasmic volutin-like granules are most actively involved in the functioning and regulation of glycolysis under conditions when, in cells of N. crassa and possibly in other microorganisms, owing to some factors, metabolism of adenine nucleotides is blocked. Besides volutin-like granules, high molecular-weight polyphosphates were found in the nuclei of N. crassa (Kulaev et al., 1970d). They were also detected in similar structures of other yeasts and fungi (Skryabin et ul., 1973; Hildebrandt and Sauer, 1977; Sokolovsky and Kritsky, 1980). It is not yet understood how biosynthesis and utilization of polyphosphates are carried out in these cellular structures. Still, there is an extremely important phenomenon revealed in N . c r a m and other fungi and yeast (Kritsky et al., 1968, 1970; Melgunov and Kulaev, 1971; Kulaev et al., 1970d, 1973b, 1977). Direct correlation was established between rates of accumulation of nucleic acids, namely RNA, and of salt-soluble polyphosphates (PolyPz; Fig. 8) localized, at least partially, in the cellular nuclei. On the basis of these data, we suggested that, in nuclei, a mechanism is operative for the synthesis of polyphosphates from pyrophosphate formed during RNA synthesis with the help of RNA-polymerase (Fig. 9). Such a mechanism of inorganic polyphos-

116

IGOR S. KULAEV AND VLADIMIR

0.3

M. VAGABOV

-

0.2 -

0.1

e e

01. 0

e e ee

I

I

0.f

0.2

Velocity of RNA synthesis

FIG. 8. Graph showing correlation between rates of formation of RNA and polyphosphate salt-soluble fraction PolyPz in Neurosporu crussa. From Kritsky et al. (1970).

phate (metaphosphate) formation was recently shown to operate in in uitro experiments on RNA biosynthesis with crude preparations of the DNAdependent RNA polymerase from E. coli (Volloch et al., 1979). Of evident interest are recent findings of Hildebrandt and Sauer (1977) who showed that, in nuclei of Physarum sp., polyphosphates are present in nucleoli, i.e. at the sites of rRNA synthesis and ribosome formation. They also revealed that in in vitro experiments this fraction is functioning as an inhibitor of RNA polymerase A which catalyses rRNA biosynthesis. Finally, in the process of differentiation of this fungus, the amount of this fraction of polyphosphates, referred to by the authors as “specific nucleolar initiation inhibitor”, varies strongly depending on the stage of differentiation. These results testify that polyphosphates may play an extremely important part in the life of organisms, regulating such an’important process as biosynthesis of nucleic acids. The paramount importance of this polyphosphate fraction for the function-

POLYPHOSPHATE METABOLISM I N MICRO-ORGANISMS

( n ) pyrophosphate

x

pyrophosphatase

(n

- I) pyrophosphate+orthophosphate

117

polyphosphate,

(7)

polyphosphate,

+I

FIG. 9. Proposed scheme for the interrelationship between biosynthesis of salt-soluble polyphosphatesand nucleic acids. From Kulaev er al. (1973b).

ing of any living cell is shown by the fact that they were found even in nuclei of higher animals (Penniall and Griffin, 1964; Bashirelashi and Dallam, 1970; Mansurova et af., 1975b; Offenbacher and Kline, 1980). In higher animals, high molecular-weight polyphosphates are found only in their nuclei. This indicates that polyphosphates are most important in these structures, being involved at all stages of development of living organisms. The third pool of polyphosphates occurring inside the plasma membrane is localized in vacuoles of fungal and yeast cells (Indge, 1968; Urech et af., 1978; Durr et af., 1979; Cramer et al., 1980; Okorokov et al., 1980; Lichko et al., 1982). Recently, numerous data have appeared pointing to the fact that polyphosphates having pronounced polyanionic properties participate in vacuoles primarily in the binding of considerable amounts of low molecular-weight compounds carrying a positive charge (Durr et al., 1979; Cramer et af., 1980; Allan and Miller, 1980; Okorokov et al., 1980; Lichko et al., 1982; Beckerich et af.,1981).According to the reports of Matile (1978), Durr et al. (1979), and Cramer et af. (1980), considerable amounts of arginine and lysine linked by ionic bonds to polyphosphates are accumulated in vacuoles of yeast and N. cassa. Generally, accumulation in vacuoles of these basic amino acids and polyphosphates in these organisms occurs simultaneously, though under certain extreme conditions they may be supplied to vacuoles separately. Another positively charged metabolite which accumulates in vacuoles and is bound in them to polyphosphates is S-adenosylmethionine (Allan and Miller, 1980). According to Okorokov et al. (1980) and Lichko et al. (1982), Mn2+and Mg2+may also accumulate in yeast vacuoles, being mostly bound

IGOR S. KULAEV AND VLADlMlR M. VAGABOV

118

to polyphosphates. By binding cations, polyphosphates participate in regulating the turnover of these most important cellular metabolites. Of the enzymes of polyphosphate metabolism, vacuoles were found to contain polyphosphate kinase (Shabalin et ul., 1977) and a polyphosphate depolymerase which hydrolyses high molecular-weight polyphosphates to low molecular-weight fragments, possibly to trimetaphosphate (Schwencke, 1978;Durr et al., 1979). Table 5 shows that in whole cells of Succh. carlsbergensis, as in vacuoles, “reverse” polyphosphate kinase activity prevails catalysing ATP synthesis by transfer of the terminal phosphate from polyphosphates to ADP. Thus, vacuolar polyphosphate kinase may be actively involved in maintenance of a certain ATP level and, through this, of the contents of other nucleoside phosphates in yeast cells. As a result, polyphosphate kinase may take part in the utilization of polyphosphates stored in vacuoles during nucleic acid synthesis. The consumption of polyphosphates that are soluble in acids for synthesisof nucleic acids in fungi and yeast was emphasized in several reports. From Table 5 it is clear that, though the role of polyphosphate kinase is not significant in synthesis of polyphosphates from ATP in whole cells and protoplasts of yeast, in vacuoles of these organisms the activity of ATP: polyphosphate phosphotransferase increases markedly to bring about synthesis of polyphosphates. If this enzyme is localized in the tonoplast of vacuoles (Y. A. Shabalin, personal communication), it is possible that it plays a key role in symport through the membrane of phosphate and positively charged ions (Matile, 1978). Polyphosphate depolymerase may also prove to be important in metabolism of vacuolar polyphosphates. If this enzyme is localized in the tonoplast, then it may be assumed that it is also involved in metabolism and transport of cations and positively charged compounds into the yeast vacuoles (Matile, 1978). Kritsky and Chernysheva (1 973) suggested that polyphosphate depolymerTABLE 5. Activity of polyphosphate kinases in some subcellular fractions of Saccharomyces carlsbergensis. After Vagabov and Shabalin (1 979) Activity (mE (mg protein)-’) ATP: polyphosphate phosphotransferase

Polyphosphate: ADP phosphotransferase

Vacuoles

1.24

4.10

Cell envelope

0.053 0.033

4.20 3.40

Fraction

Protoplast lysate

E indicates an International Unit of activity.

POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS

119

ase may participate not only in breakdown of highly polymerized polyphosphates to less polymerized ones, but also in translocation of the fragments formed through membranes at the expense of the energy released during cleavage of phosphoanhydride bonds. If this suggestion is correct then, during the translocation of short-chain polyphosphate, symport of positively charged ions and metabolites may occur. The interrelations between accumulation in yeast cells of short-chain polyphosphates and positively charged amino acids were investigated by Ludwig et al. (1977) and Lusby and McLaughlin (1980). These authors showed that addition of free L-amino acids, particularly arginine and lysine, to yeast culture media under conditions of nitrogen starvation resulted in an intensification of yeast growth and rapid intracellular accumulation first of tripolyphosphate and then of more polymerized chains, including tetrapolyphosphate and pentapolyphosphate. Experimentswith 32Pshowed that the tripolyphosphate formed originated not from orthophosphate supplied in the medium, but from a more polymerized polyphosphate fraction. Accumulation of tripolyphosphate and, to a lesser extent, of other smaller polyphosphates in cells of intensively growing yeast was also observed by Solimene et ul. (1980). These results are very interesting, although unfortunately they do not provide sufficient information about the localization of low molecular-weight polyphosphates in yeast cells during intensive growth. Here, one undoubtedly deals with the Polyp, fraction which accumulatesin yeast cells either in vacuoles or in the cytoplasm. Low molecular-weight polyphosphates accumulate in the cytoplasmic(vacuolar) granules of yeast (at least under conditions of an intensive L-amino acids uptake) as a complex with positively charged arginine and lysine residues. Such electroneutral complexes represent a pool of negatively charged phosphate and positively charged amino acids in a rather inert form convenient for the cell. The polyphosphate complex with arginine (or lysine) which accumulates in vacuoles and probably to some extent in the yeast cytoplasm, resembles cyanophycin, which is a copolymer of aspartic acid and arginine discovered by Simon (1971) in blue-green algae. This copolymer, in which the role of a negatively charged complex is performed by aspartic acid, accumulatesin cells of blue-green algae as granules and forms an intracellular reserve of the most important nitrogen-containing metabolites. Recalling the investigations of McLaughlin and his coworkers, one should take into account the formation of tripolyphosphate in yeast cells from high molecular-weight polyphosphate fractions in the presence of supplied L-amino acids. It is difficult to state which polyphosphate fraction is depolymerized to tripolyphosphate. It should be noted that polyphosphate depolymerase was found in cells of N. crussa (Kulaev et al., 1972a) both outside the cytoplasmic membrane

120

IGOR S. KULAEV AND VLADlMlR M . VAGABOV

TABLE 6. Intracellular localization of polyphosphate depolyrnerase activity in Neurosporu crussu. After Kulaev et ul. (1 972a) ~~

Activity (mE(mg protein-’)) Substrate

Intact cells

Protoplasts

Polyphosphate (ii=290) Polyphosphate (ii= 180)

8.3 9.4

0.6 1.3

Nuclei

Mitochondria

0.15 0.20

0.0 0.0

Microsomes Cytosol 2.2

-

0.0 0.0

E indicates an International Unit of activity and ri is chain length.

(major part) and in nuclei and the “microsomal” fraction into which vacuoles were undoubtedly fragmented during fractionation (Table 6). Schwencke (1 978) also detected polyphosphate depolymerase directly in yeast vacuoles. Its action on high molecular-weightpolyphosphates yielded tripolyphosphate as the final product. It is not precluded that, in the presence of L-amino acids, the depolymerase contained in nuclei hydrolyses high molecular-weight polyphosphate bound in nucleoli to RNA polymerase A thereby inhibiting this enzyme, thus contributing to RNA and protein synthesis, as well as culture growth. Tripolyphosphate formed during depolymerization may be a “primer” for synthesizing“de novo” high molecular-weight polyphosphates. It appears quite probable that tripolyphosphate may be the form in which polyphosphates could be translocated through the cellular membranes. This suggestion correlates well with the finding of Valikhanov and Sagdullaev (1979) indicating that tripolyphosphate uptake from the medium by cotton roots is more rapid compared with other phosphates, including orthophosphate. Thus, it appears probable at present that, in fungal cells and possibly in cells of other organisms, tripolyphosphate is, on the one hand, the form of polyphosphate appropriate for transport through membranes and, on the other hand, a “primer” of the biosynthesis “de novo” of high molecularweight polyphosphates. It appears probable that the repeatedly shown (Kulaev, 1979) reversible transformations: PolyPz$PolyP1 are connected with the existence in cells of fungi and yeast of the system:

-

High molecular-weight polyphosphates tripolyphosphate

polyphosphate synthases

plyphosphate

depo1ym erasc

high molecular-weight polyphosphates

Such transformations were clearly demonstrated recently during yeast dehydration followed by their reactivation (Kulaev et a/., 1977; Table 7).

121

POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS

Summarizing about the metabolism of polyphosphate fractions localized in cells of yeast and fungi inside the cytoplasmic membrane (cytosol, vacuoles, nuclei), it is appropriate to underline their multifunctional roles. They are not only reservoirs of phosphates and energy, but also most important regulators of cellular metabolism. They participate in the regulation of ATP, ADP, ortho- and pyrophosphate levels, in control of glycolysis and intracellular ionic fluxes and, finally, in regulation of nucleic metabolism and growth processes in general. TABLE 7. Contents (pg P (g dry wt cells)-') of orthophosphate and polyphosphate fractions of Succhuromyces cereuisiue-14 grown in a molasses medium in a fennentor then dehydrated or reactivated. After Kulaev et ul. (1977) Content (pg P (g dry wt cells)-') in Phosphorus compound of fraction

Initial cells

Cells dehydrated at 37°C for 24 hours

Cells reactivated at 37°C for 30 minutes

Orthophosphate Acid-soluble (PolyP,) Salt-soluble (PolyPz) Alkali-soluble (PolyPs) Hot perchlorate extract (PolyPs) High molecular-weight polyphosphates (total)

2150 350 4470 1340 1160

1830 3710 850 1570 210

2980 620 1910 1530 520

7230

6340

4580

Significant amounts of the most polymerized polyphosphates are localized in fungi at the cell periphery, close to the cytoplasmic membrane (Weimberg

and Orton, 1965; Kulaev et al., 1966, 1967a, 1970a,b; Souzu, 1967a,b; Weimberg, 1970; Vagabov et al., 1973). Studies on localization of polyphosphate metabolism enzymes in N.c r a m revealed that the cell periphery, in the proximity of the pool of polyphosphates, contains substantial amounts of polyphosphate depolymerase which hydrolyses polyphosphates in the middle of the chain (Kulaev et al., 1972a) and polyphosphatase splitting off terminal phosphate residues (Kulaev et al., 1972b;Trilisenko et al., 1980). Recently, Trilisenko et al. (1980,1982)isolated a mutant of N . crassa with a markedly low polyphosphatase activity. In this mutant, hydrolysis of high molecular-weight polyphosphates (i= 180) by polyphosphatase proceeded at a lower rate than with the same enzyme from the wild-type strain of N. crassa. The affinity of this polyphosphatase for high 180) proved to be two orders of molecular-weight polyphosphate (i= magnitude higher compared to low molecular-weight polyphosphate (i= 9).

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IGOR S. KULAEV AND VLADlMlR M. VAGABOV

This observation and accumulation of highly polymerized polyphosphates in the mutant with low polyphosphatase activity (Table 3) argue in favour of the participation of this enzyme in uivo in utilization of polyphosphates with a peripheral location in fungal cells. Data on the peripheral localization of highly polymerized polyphosphates were also obtained for Endomyces magnusii (Kulaev et al., 1967a;Kulaev, 1979), Sacch. carlsbergensis (Vagabov et al., 1973) and Sacch. cereuisiae (Tijssen et al., 1980). It should be stressed that, in N . crassa, polyphosphatases degrading polyphosphates at terminal phosphate residues are apparently firmly bound to the cytoplasmic membrane. This conclusion may be drawn from the fact this enzyme is removed from the protoplast surface only after treatment with the detergent Triton X-100(Kulaev ez al., 1972b;Kulaev, 1973b;Konoshenko et al., 1973; Krasheninnikov et al., 1973). Some indirect data are available (Kulaev, 1979) which indicate that high molecular-weight polyphosphates in the peripheral part of the cell are localized, in N . crassa and E. magnusii,close to the polyphosphatase hydrolysing them to orthophosphate, i.e. in the proximity of the cytoplasmic membrane. Under various conditions affecting the cytoplasmic membrane of these organisms, these are the fractions of highly polymerized polyphosphates that are subject to hydrolysisto form orthophosphate. Unsuccessful attempt of Wiemken and his co-workers (Diirr et al., 1979) and Davis and his coworkers (Cramer et al., 1980) to detect peripheral fractions of polyphosphates in yeast and N. crassa cells were, beyond any doubts, due to this cause. The methods they used for pretreatment and fractionation of cells caused a prompt and selective breakdown to orthophosphate of very highly polymerized fractions of polyphosphates localized in the cytoplasmic membrane near to polyphosphatase. The latter enzyme, and the above-mentioned polyphosphate depolymerase whose occurrence in yeast and fungi was reported in a number of studies (Malmgren, 1949, 1952; Mattenheimer, 1951; Kritsky et al., 1972), are responsible in fungi for utilization and degradation of the most polymerized polyphosphate fractions (PolyP3, Polyp4 and PolyPs). It seems that, in yeast and fungi, the depolymerase functions in transformation of highly polymerized polyphosphates localized outside the plasma membrane giving rise to less polymerized polyphosphates capable of being translocated through the ‘membrane (Belozersky and Kulaev, 1957; Kulaev et al., 1959; Langen et al., 1962; Kritsky and Chernysheva, 1973), while the polyphosphatase has quite a different function in metabolism of these compounds. Over the past 15 years, Van Steveninck, in collaboration with other Dutch researchers, has obtained convincing results pointing to the fact that highly polymerized polyphosphates, localized at the periphery of yeast cells, are involved as energy donors in the basic transport of sugars through the cytoplasmic membrane (Van Steveninck, 1963; Van Steveninck and Booij,

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123

1964; Dierkauf and Booij, 1968; Jaspers and Van Steveninck, 1975; Tijssen et al., 1980). Umnov et al. (1974a) and Trilisenko et al. (1980) showed that polyphosphatase plays an important role in utilization of these polyphosphate fractions for active transport of sugars through the cytoplasmic membrane in N. cra~saby hydrolysing them to orthophosphate. It is admitted that polyphosphatase may operate in vivo in yeast and fungi not only as a phosphohydrolase but also as a phosphotransferase. It may also transport activated phosphoryl derivatives of polyphosphates instead of water to some components of the system for active sugar transport, acting as energy donor for this process. The ability of some phosphohydrolases to carry out certain phosphotransferase reactions has been known for a long time (Nordlie and Arion, 1964; Stetten, 1964). Returning to investigations of localization of enzymes of polyphosphate metabolism in fungi, it should be noted that, in the region of localization of the most polymerized polyphosphates in the proximity of the cytoplasmic membrane, we detected only enzymes of degradation and utilization of polyphosphates. Until recently it was not known how polyphosphates of this fraction are synthesized. Certain progress in solving this problem has been achieved in our laboratory in the course of investigation of polyphosphate metabolism in yeast (Kulaev et al., 1972c,d; Vagabov et al., 1973; Tsiomenko et al., 1974a,b; Vagabov and Shabalin, 1979; Shabalin et al., 1978, 1979). These reports showed good correlation (Table 8) between rates of accumulation of polymerized polyphosphates localized in the cellular envelope and synthesis of polysaccharides of the cell wall (Kulaev et al., 1972c,d). The highest value for the correlation coefficient (0.8-0.9) was found for the Polyp4

TABLE 8. Correlation coefficients between the rates of formation of various polyphosphate fractions and polysaccharides in Saccharomyces carlsbergensk After Kulaev et al. (1972c,d) Polysaccharides

Polyphosphates

Correlation coefficients"

(Polysaccharides) Glycogen Glycogen Glycogen Glucan + mannan Glucan Glucan Mannan Mannan Mannan

(Polyphosphates) PolyP, PolyP2 (PolyP2, PolyP3, PolyP4, PolyPs) (PolyP2, PolyP3, PolyP4, PolyPs) PolyPz PolyP3 PolyP2 PolyP3 PolyP4

0.806f 0.068 0.077f 0.02 0.141 f 0.008 0.173f0.018 0.750f 0.087 0.291k0.180 0.615f0.122 0.136f 0.192 0.035f 0.196 0.813 0.098

The coefficients were calculated from the results of 36 determinations.

IGOR S. KULAEV AND VLADlMlR M. VAGABOV

124

fraction and mannan synthesis. The behaviour of these two fractions during normal growth, as well as during various impositions on culture growth, was a clear-cut parallelism. This can be seen from Fig. 10. These data suggested the existence of some specific interrelation between metabolism of these two compounds which, though quite different in their chemical nature, are nevertheless components of the same organelle, namely the cell envelope. Synthesis of mannoproteins from GDP-mannose is known to proceed not in the envelope itself but inside the cell, in the so-called microsomal fraction, and in vesicular membrane structures which bud off to the localization site in the cellular envelope(Matile, 1975; Lehle et al., 1977; FarkaS, 1979; Schekman et al., 1981). Experiments with [P-32P]GDP-['4C]mannose and the microsomal fraction of Sacch. carlsbergensis showed that GDP-mannose is not only the donqr of mannosyl groups during biosynthesisof mannoproteins (Brehrens and Cabib, 1968), but also acts as the source of phosphate in biosynthesis of polyphosphates (Shabalin et al., 1978; Vagabov and Shabalin, 1979; Kulaev et al., 1979). It was also established that syntheses of both mannan and polyphosphates require Mn2+, whereas Mg2+ inhibited both processes. Further investigations indicated that transport of phosphate groups from GDP-

I2O

c

loot

c

c

a3

c

c

0

u

50

60

70 00

Time (hours)

FIG. 10. Changes in the contents of mannan (0)and high molecular-weight polyphosphate fraction Polyp4 ( 0 ) under different growth conditions of Saccharomyces carlsbergensis in Rider medium in the presence (a) or absence (b) of nitrogen. From Kulaev et al. (1972c,d).

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mannose to polyphosphates involves a lipid intermediate identical in its characteristics to dolichol-pyrophosphate mannose (Vagabov and Shabalin, 1979; Shabalin et al., 1979). On the basis of the results obtained, the mechanism of biosynthesis of polyphosphates from GDP-mannose in yeast Sacch. carlsbergensis may be depicted as follows:

+

+

1. GDP-mannose dolichol phosphate4 GMP dolichol-pyrophosphatemannose 2. Dolichol-pyrophosphatemannose (mannan).+dolichol pyrophosphate (maman),,+I 3. Dolichol pyrophosphate +@olyphosphate)n-rdolicholphosphate+ @olyphosphate)n+I

+

+

Available data lead us to conclude that, in yeast, a new pathway for biosynthesis of high molecular-weight polyphosphates has been shown to be closely related to the synthesisof mannoproteins of the cell wall. Thus, it stems from the above data that both biosynthesis and utilization of the surfacelocated highly polymerized polyphosphates are closely connected with the biogenesis and functioning of a most important cellular compartment of yeast and fungi, namely their cellular envelope. It is noteworthy that not only biosynthesis but also degradation of polyphosphates and mannoproteins of yeast are closely interrelated. As shown in Fig. 11, practically synchronous changes in polyphosphatase and mannosidase activities occur during yeast cultivation (Tsiomenko et al., 1974a,b). Metabolism of these two biopolymers of the cellular envelopes seems to be closely co-ordinated. Data pointing to a relation between biogenesis and functioning of a polyphosphate fraction and that of the fungal cell wall were also obtained by Wool and Held (1976). Using ultracytochemical and X-ray dispersion analyses, these authors studied localization of polyphosphates in isolated zoospores of Rozella allomycis fungus parasitizing species of Allomyces. In this work, vesicles of the endoplasmic reticulum were shown to contain both polysaccharide precursors of the cyst cell wall and polyphosphates; in cysts, polyphosphates were found in the regions of the cytoplasmic membrane and cell wall. In this fungus, in addition to the above sites, a polyphosphate fraction was detected in vacuoles of cysts before germination. The authors believe that formation of these polyphosphates was connected with degradation of a nucleic acid fraction, whereas their utilization (hydrolysis to orthophosphate) occurred immediately before the start of cyst germination, thus creating the required osmotic pressure for the ‘‘explosion’’ of cysts and penetration of germ cells into the tissue of the host fungus. It should be recalled that in earlier literature on polyphosphates, similar mechanisms for production of specific polyphosphate fractions (during degradation of nucleic acids) as well as those of their utilization (for creating excessive osmotic pressure) were frequently demonstrated and discussed (Harold, 1966; Kulaev, 1979). In particular, Kritsky and his colleagues showed that the osmotic pressure developed during hydrolysis of

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-8

0.6

-6

0 0.04

0*40>L9 0.2

0

-

0

0

1

0

I

I

I

3

0.02

I

0

5

Time (hours)

FIG. 11. Variation in the activity of enzymes hydrolysing polyphosphates (a) and mannan (b) in Succharomyces carlsbergensis after their transfer into fresh medium. 0, Activities of polyphosphatase and a-mannosidase in (a) and (b) respectively; 0 , content of polyphosphate and mannan, in (a) and (b) respectively. E indicates one International Unit of Activity. From Tsiomenko et ul. (1974a,b).

polyphosphates in the lamellae of fruiting bodies of Agaricus bisporus is involved in dissemination of spores (Kulaev et al., 1960; Kritsky et al., 1965a,b). Data from Gezelius (1974) also argue in favour of the existence of quite different pathways for polyphosphate formation in various fungi. During investigation of polyphosphate metabolism in Dictyostelium discoideum, Gezelius showed that large amounts of polyphosphates were synthesized during the transition of D . discoideum from the amoeboid to the aggregated stage. Dictyostelium discoideum is known to produce cyclic AMP intensively

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from ATP during this period (Pastan et al., 1975). It may be that this fungus possesses an enzyme system that catalyses synthesis of polyphosphates from pyrophosphate formed during biosynthesis of cyclic AMP from ATP. Gezelius (1974) also pointed to the existence in this fungus of a mechanism for polyphosphate formation different from its synthesis with the participation of polyphosphate kinase. The occurrence of several mechanisms of polyphosphate synthesis was also confirmed genetically. Beckerich et al. (1981) isolated a number of mutants of Saccharomycopsis lipolytica (Table 9) which lacked certain fractions of

TABLE 9. Distribution of polyphosphate fractions in mutants of Saccharomycopsis lipolytica. After Beckerich et al. ( 1981)

Polyphosphate fraction (nmol K2HP04 equiv. (mg dry wt)-l) Strain

FI

F2

15901.7

0

2 8 0 10 25

PlY 1 PlY 2 PlY 3 ply 4 ply 5

0

PlY 7 PlY 9

0

F3

F4

Fs

33

7

5 0 5

0

0 0 4 0 0 0 0 0 0

14 0 0 0 0

1 0

5

0

3 5 0 4

0 0

7 6

0 0 0 1

FI indicates an acid-solublepolyphosphate(ri= I-5),F2 a perchlorate-soluble polyphosphate (R up to 20), F3 a polyphosphate with ii=20-50, F5 a polyphosphate with A= 50-250 and F4 nucleic acids.

acid-insoluble polyphosphates, while the PolyPl/PolyP2ratio differed greatly from that in the parental strain. It was suggested that, in the mutants, the pathway for polyphosphate biosynthesis related to formation of the fungal cell wall was impaired. That the acid-insoluble polyphosphate fractions are involved in formation of the cell wall is supported not only by these results (Vagabov and Shabalin, 1979; Shabalin et al., 1979) but also by the data of Sokolovsky and Kritsky (1980) and those of Trilisenko et al. (1982) on the absence of these very fractions from Physarum polycephalum and a slime mutant of N. crassa (Fig. 12).

IGOR S. KULAEV AND VLADlMlR M. VAGABOV

128

Strain ad-6

f -

4-:"1.5

o o r L.b:C 0 10 20 30

Slime mutant

30,lS-3

L/ o:;v 1: -.A=&

0

15

30

50

0

0 10 20 30

Time (hours)

FIG. 12. Time-course of changes in the contents of various polyphosphate fractions during growth of Neurospora crassa: strain ad-6; a leaky mutant in polyphosphatase (30,19-3) and a slime mutant devoid of the cell envelope. 0 Indicates the growth phase of N . crassa and 0 indicates phosphate content in the polyphosphate fraction.

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2. Algae as Representatives of Phototrophic Eukaryotes The autotrophic nature of algae affects their polyphosphate metabolism (Miyachi et al., 1964; Kulaev and Vagabov, 1967;Ullrich and Simonis, 1969; Kuhl, 1974). In numerous reports it was shown that formation of polyphosphates and polyphosphate granules in algae occurs with a markedly higher intensity in the light than in the dark (see Kulaev, 1979). The most fundamental and detailed investigation of polyphosphate metabolism in algae was conducted by Miyachi et al. (1964) with Chlorella ellipsoidea. They showed that only one of four polyphosphate fractions detected in C. ellipsoidea was formed in the light. Both biosynthesis and utilization of polyphosphates were shown to occur in the light. It was also established that, in C. ellipsoidea as in heterotrophs, the fraction of polyphosphates extracted with cold acid was a component of volutin. According to Atkinson et al. (1974), in C . ellipsoidea these granules may be localized, at least partially, in vacuoles. Accumulation of polyphosphates contained in these granules also depends, according to Miyachi et al. (1964), on photosynthesis, since they are formed from the fraction synthesized in the light. The biosynthesis and utilization of two other fractions detected by Miyachi et al. (1964) in C. ellipsoidea were totally independent of photosynthesis. Their metabolism depended on the presence of phosphate in the incubation medium. Similar results were obtained later by Kanai and Simonis (1968) for Ankistrodesmus braunii. It is interesting that, in these and other early studies of polyphosphate metabolism in algae, a close and fairly complex relationship between certain polyphosphate fractions and nucleic and metabolism was established and found to be similar to that observed in heterotrophic organisms (Kulaev, 1979). Bearing in mind the data of Richter (1966), who demonstrated polyphosphate synthesisin the nucleus-freecell halves of Acetabularia sp., one may draw an indirect conclusion that the presence of the cell nucleus is not obligatory for the replenishment of at least some polyphosphate fractions in algae. In general, studies on polyphosphate metabolism in this gigantic unicellular alga appear to be very promising. In our laboratory, for example, Rubtsov and his coworkers (Kulaev et al., 1975)found that in the early stages of growth of Acetabularia mediterranea and Acetabularia crenulata, in contrast to heterotrophic organisms, only acid-soluble (Polyp,) and salt-soluble(PolyPz) fractions-were present (Table 10). At the stage af cyst formation (stage 4), characterized by intensive synthesis of their cell wall components, the distribution of these compounds in fractions in Acetabularia cells does not differ qualitatively from that in heterotrophs. At this stage of growth, in

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IGOR S. KULAEV AND VLADlMlR M . VAGABOV

TABLE 10. Contents of inorganic polyphosphates and orthophosphate in Acetabularia crenulata at different stages of development. After Kulaev et al. (1975) Phosphate content &g phosphorus cell-I) Fraction Orthophosphate Acid-soluble (PolyP~) Salt-soluble (PolyP2) Alkali-soluble (PolyPs) Hot perchloric acid extract (PolyP~) Total polyphosphates

Stages of culture growth: 1 2 3 4 0.52 0.67 0.12 0.0

1.40 3.30 0.46 0.0

0.76 1.88 10.10 2.41 0.44 1.27 0.25 0.54

0.79

0.0 3.76

0.0 10.79

0.81 5.03

Stages of growth were as follows: 1, young cells 1.5-2 cm long; 2, cells 2.5-3.0 cm long, up to 2 mm in diameter; 3, cells with umbellulles filled with secondary nuclei; 4, cells with mature umbellulles filled with cysts.

addition to acid- and salt-soluble polyphosphates (as in active young photosynthetic cells), alkali-soluble (Polyp3 and PolyP4) polyphosphates and those extractable by hot perchloric acid appear. In studies with A. mediterranea, it was first shown that high molecular-weight polyphosphates do not occur in chloroplasts (Rubtsov et al., 1977). Structures connected with photophosphorylation could not be shown to be capable of polyphosphate biosynthesis. The absence of high molecular-weight polyphosphates from chloroplasts was also confirmed for higher plants such as cotton (Valikhanov and Sagdullaev, 1979). However, as shown on p. 149, light-dependent synthesis of inorganic pyrophosphate attended by electron transfer along the electron-transport chain was revealed in chloroplasts of A. mediterranea and pea (Rubtsov et al., 1976). Investigations of A. mediterrunea provided the answer to the question of whether light-dependent accumulation of polyphosphates in algae was directly connected with photosynthesis itself or whether their formation in the light was simply stimulated by increased synthesis of ATP or pyrophosphate at the expense of photosynthetic phosphorylation. Detection of ATP: polyphosphate phosphotransferase in A. mediterranea (Rubtsov and Kulaev, 1977), as well as the absence of high molecular-weight polyphosphates and their biosynthesis in chloroplasts of this alga, together with corresponding inhibitor analysis, point convincingly to the fact that high

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molecular-weight polyphosphates are not directly, but rather indirectly, connected with photosynthesis through formation of ATP (but not of pyrophosphate) in photosynthetic phosphorylation. Taking into account that, in algae, polyphosphates accumulate in the light essentially in vacuoles (Atkinson et af.,1974; Sundberg and Nilshammar-Holmvall, 1979, together with detection of polyphosphate hydrolase activity in A. mediterranea (Rubtsov and Kulaev, 1977), one can depict the basic metabolic pathways of the light-dependent synthesis and utilization of polyphosphates in algae as in Fig. 13. According to this scheme, by analogy with yeast (Shabalin et al., 1977), polyphosphate kinase is localized in vacuoles. However, this assumption still requires experimental support. When reviewing studies on polyphosphate metabolism in algae reported

light

FIG. 13. Major metabolic pathways for light-dependent synthesis and utilization of polyphosphates (polyp) in algae.

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IGOR S. KULAEV AND VLADlMlR M . VAGABOV

during the past years, reference should be made to the work of Peverly et al. (1978) and Adamec et al. (1979). These authors studied the influence of potassium ions on accumulation of polyphosphate granules in Chlorella pyrenoidosa. Peverly et al. (1978) found that K + ions stimulate polyphosphate granule formation in this alga. These authors also showed a correlation between accumulation of phosphate and potassium in cells which, after phosphorus starvation, were transferred to a medium containing adequate amounts of both components. Microscopic examination revealed intensive accumulation of polyphosphate granules in cells. Further, using X-ray dispersion analysis in combination with electron microscopy, Adamec et al. (1979) detected potassium in addition to phosphorus in polyphosphate granules. On the basis of these results, these authors believe that K + is the major cation of polyphosphate granules in Chlorellapyrenoidosa growing in a medium with a sufficient amount of potassium.

D. NEW DATA O N POLYPHOSPHATE METABOLISM IN PROKARYOTES

At the present time, polyphosphate metabolism in prokaryotic microorganisms has been most extensively studied in mycobacteria (Winder and Denneny, 1957; Mudd et al., 1958; Drews, 1962; Dirheimer, 1964; Szymona and Ostrowski, 1964), corynebacteria (Sall et al., 1958; Hughes and Muhammed, 1962),propionic-acid bacteria (Kulaev et al., 1973a),streptomycetes (Kulaev et al., 1976), Aerobacter aerogenes (Harold, 1966) and Escherichia coli (Nesmeyanova et al., 1973a, 1974a). Detailed information about the characteristic features of the metabolism of these and a number of other eubacteria can be found in Kulaev’s (1979) monograph. The most important and experimentally supported inference from this work was the close relationship between polyphosphate and nucleic acid metabolism. We have postulated one possible mechanism for the interrelation between these two pathways (Kulaev et al., 1973b; Kulaev, 1975). The suggestion, based on our own and literature data, was that conjugation of these two metabolic pathways may occur at the level of synthesis of specific polyphosphate fractions from pyrophosphate formed during biosynthesis of nucleic acids. Recently Volloch et al. (1 979) in Tummerman’s laboratory confirmed this experimentally using a crude preparation of DNA-dependent RNA polymerase and phage SV-40 as a DNA template in in vitro experiments. They found that pyrophosphate (PP) formed during RNA biosynthesis by the preparation was not accumulated as such but condensed to form some polymeric phosphorus compound. For some unclear reasons, this compound was termed “trimetaphosphate”, though this work provided no valid support for such a term. It is interesting that use of purified preparations of

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DNA-dependent RNA polymerase and highly purified preparations of DNA polymerase I enabled the authors to observe formation of only pyrophosphate. These data prompt one to think that, in such a system, the synthesis of polyphosphate from pyrophosphate is not carried out by RNA and DNA polymerases but by a pyrophosphate-polyphosphate phosphotransferase tightly coupled to them. In recent years, a series of cytological data obtained with cyanobacteria and using electron microscopy have appeared. For these prokaryotes, the above-mentioned results demonstrated a very close topological relation between polyphosphates and nucleic acids. The most detailed information in this respect was provided by research from Jensen’s laboratory (Jensen, 1968, 1969; Jensen and Sicko, 1974; Sicko-Goad and Jensen, 1976; Sicko-Goad et al., 1975,1978;Lawry and Jensen, 1979;Baxter and Jensen, 1980a,b)as well as from a number of other laboratories (Kessel, 1977; Vaillancourt et al., 1978; Barlow et al., 1979; Ferguson et al., 1979) engaged on studies of phosphate metabolism in cyanobacteria. It should be noted that, at present, investigationof phosphorus metabolism in cyanobacteria is a very urgent problem. This is due to the “fluorescence” of cyanobacteria in inland water bodies which receive large amounts of various detergents and other phosphate-containing effluents of industrial production. In detergents, the most frequently used compound is sodium tripolyphosphate. Inland water bodies, particularly those located in industrialized countries, normally contain low concentrations of phosphorus (about 10 pg 1-’ or less) and suffer a massive “fluorescence” of cyanobacteria when substantial quantities of tripolyphosphate are “dumped” into them in waste waters. Conditions are created known as “phosphate overplus”. After a long phosphorus starvation, microbes finding themselves in a phosphorus-rich medium start to grow and reproduce very intensively. Under such conditions, cyanobacteria and other micro-organisms (Drews, 1962; Harold, 1966; Kulaev, 1979) accumulate large amounts of polyphosphates essentially localized in polyphosphate granules. The above authors studied in detail intracellular localization and chemical composition of polyphosphate granules under conditions of normal growth as well as under a lack or excess of certain nutrients in the medium. Investigating localization of polyphosphates in Plectonema boryanum, Jensen (Jensen, 1969; Jensen and Sicko, 1974) found that they were localized in this blue-green alga in five cellular sites. These were sites of ribosome formation, DNA fibrils, near thylakoids, polyhedral bodies which harbour key enzyme of photosynthesis, including ribulose 1$diphosphate carboxylase, and the nucleoplasmic zone. This implies that accumulation of polyphosphates in P. boryanum is basically conditioned by photosynthesisas well as biosynthesis and degradation of nucleic acids. It is interesting that in

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IGOR S. KULAEV A N D VLADlMlR M. VAGABOV

the first study devoted to polyphosphate formation in P . boryanum, Jensen (1969) reported that these compounds are detected cytochemically inside cell walls. Generally, in cyanobacteria, intracellular localization of polyphosphates does not differ from that in eukaryotic micro-organisms, particularly in phototrophs. Therefore, it can be concluded that their metabolism in cyanobacteria is primarily connected with nucleic acids and cell wall components and, to some extent, with functioning of the photosynthetic apparatus. It is not clear so far whether, in cyanobacteria, polyphosphates are produced directly by photophosphorylation or are secondary products from ATP. Shady et al. (1976) found that, in the phototrophic bacterium Rhodospirillum rubrum, polyphosphates are formed in chromatophores in the light indirectly, via ATP with participation of polyphosphate kinase. The availability of polyphosphate kinase in cyanobacteria, in particular in Anacystis nidulans, was demonstrated by Vaillancourt et al. (1978) who isolated “leaky” mutants in this enzyme. Of interest also is the fact that these mutants did not have polyphosphate granules observable by electron microscopy. It may be inferred that, in blue-green algae, polyphosphate kinase plays a very important part in phosphate metabolism. In this respect, A. nidulans is apparently similar to Aerobacter aerogenes mutants which are deficient in polyphosphate kinase and are also devoid of the ability to accumulate polyphosphates. It is noteworthy that, in A. nidulans, judging from the data of Vaillancourt et al. (1978), polyphosphatase has not been detected. Failure to detect this enzyme is very rare in micro-organisms. Ferguson et al. (1979) carried out an interesting study of mechanisms of polyphosphate synthesis in Paracoccus denitrijicans using 31Pnuclear magnetic resonance. Using inhibitors of oxidative phosphorylation and different conditions of energy metabolism, the authors showed the extreme importance of polyphosphate metabolism in the energy metabolism of this bacterium. In contrast to Harold (1966), the authors drew a conclusion shared by most researchers (see Kulaev, 1979), namely that, in bacteria, polyphosphates are reserves not only of phosphates but of energy also. In this work, it was observed that polyphosphates Polyp,, though synthesized in P . denitrijicans at the expense of the energy of succinate oxidation (possibly via intermediate ATP formation), do not utilize orthophosphate as phosphate source but use some intracellular phosphorus-containing compounds (possibly nucleic acids or products of their degradation such as nucleoside monophosphates). In recent years, some results were reported on mechanisms of polyphosphate utilization in bacteria. Butukhanov et al. (1979) reported intensive ATP synthesis (0.6-1.0 mg (ml medium)-’) from exogenous adenine in autolysing cultures of Corynebacterium sp. VSTII-301.They also reported data indicating that high molecular-weight polyphosphates and inorganic pyrophosphate

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were phosphate donors for ATP synthesis. It was calculated that, during the 24 hour growth of the autolysing cells of this culture, 58.3 pmol of acid-labile phosphate of ATP and ADP were synthesized and, at the same time, 33.5 pmol of acid-labile phosphate of intracellular polyphosphates and pyrophosphate were utilized. The author succeded also in isolating and purifying from culture not only polyphosphate: ADP phosphotransferase but also a new enzyme, pyrophosphate: ADP phosphotransferase. Moreover, Butukhanov et al. (1979) revealed that, after addition of exogenous adenine to a culture of Corynebacterium sp. autolysed for 72 hours, the activity of polyphosphatase activity decreased and that of polyphosphate: ADP phosphotransferase increased. After 84 hours when most of the adenine added was utilized for synthesis of ATP, polyphosphatase activity increased again in cells. This work demonstrated a competitive relationship between two enzymes, namely polyphosphate kinase and polyphosphatase, involved in utilization of polyphosphates. Similar competitive relationships between two different polyphosphate-utilizing enzymes were reported by Ziizina et al. (1981). In this work, results were obtained supporting previous observations (Kulaev et al., 1976; HoStalek et al., 1976; Kulaev, 1979) that, during antibiotic production in prokaryotes, inorganic polyphosphates, but not ATP, are used as an energy source. Polyphosphate utilization during synthesis of the antibiotic levorin was brought to light by Ziizina et al. (1981). They showed that polyphosphate utilization proceeds under conditions of phosphorus starvation with the help of polyphosphate glucokinase. As seen from Fig. 14, enzyme activity is dramatically enhanced in a culture of Streptomyces levoris producing this antibiotic under conditions of phosphorus starvation, and its variation clearly correlates with levorin accumulation. At the end of the stationary phase of growth of Strep. levoris, polyphosphatase activity increased, whereas polyphosphate glucokinase activity decreased. Substitution of the enzymes of polyphosphate metabolism may possibly be due to a deceleration of antibiotic formation during this period. In connection with this work, which demonstrated an important physiological role for glucose phosphorylation at the expense of high molecular-weight polyphosphates, recent research carried out in Szymona’s laboratory should be cited (Szymona et al., 1977; Szymona and Szymona, 1978,1979; Pastuszak and Szymona, 1980) dealing with structure and function of an enzyme catalysing this process in Nocardia sp. and mycobacteria, micro-organisms closely related to streptomycetes. At present, the specificity and individual features of this enzyme remain unclear. Returning to the problem of the possible functions of high molecularweight polyphosphates in prokaryotes, it should be noted that the most important role for these compounds is the regulation of orthophosphate level

136

IGOR S. KULAEV AND VLADlMlR

M. VAGABOV

r

-

0 D

v)

c .-

c

5

m

-P 0 1

3

6

9

Time (days)

FIG. 14. Correlation between the activity of polyphosphate glucokinase in Streptomyces leuoris (a) and formation of levorin in culture medium (b). 0 Indicates use of a medium with 0.4 mM KH2P04;0 indicates a medium with 4.0 mM KH2P04.

in the cells. For bacteria, this was first demonstrated by Harold (1966), then in our laboratory (Nesmeyanova et al., 1973a,b; 1974a,b; 1975b;Maraeva et af., 1979) as well as by other researchers (Yagil, 1975; Zuckier et al., 1980; Tommassen and Lugtenberg, 1980; Argast and BOOS,1980). Investigations conducted with E. cofi showed that, when these bacteria are placed in a fresh medium without orthophosphate, the level of polyphosphates in cells drops drastically (Fig. 15), and the subsequent addition of orthophosphate to the culture starved of phosphorus restores the initial polyphosphate level. The involvement of polyphosphates in regulation of the intracellular orthophosphate concentration in E. cofi is also supported by the fact that synthesis of polyphosphatase participating in polyphosphate hydrolysis is induced during phosphorus starvation simultaneouslywith other phosphohydrolases including tripolyphosphatase and alkaline phosphatase (Nesmeyanova et al., 1974a) as well as acid phosphatase (Maraeva et al., 1978).

POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS

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P v

u0

In

L

C 0

E

.m

c

C 0 W c 0

c 111 n c

a 60

I80 Time (min)

300 Time (min)

FIG. 15. Effect of exogenous orthophosphate on the concentration of intracellular orthophosphate and polyphosphate in Escherichiu colt (a) describes behaviour in a medium with excess of orthophosphate;and (b) with a deficiency of orthophosphate. 1 describes culture growth, 2 polyphosphate concentration (pg (mg dry weight)-’), 3 intracellular orthophosphate concentration (pg (mg dry weight)-’) and 4 the concentration of orthophosphate in the medium.

An identical response of different phosphohydrolases to concentrations of exogenous orthophosphate points to a common character in their functions connected, apparently, with regulation of orthophosphate level in the cells of this bacterium. The unity of function of different phosphohydrolases manifested in the orthophosphate requirements of E. coli was also confirmed by genetic studies. Using E. coli mutants for regulatory genes for alkaline phosphatase, Nesmeyanova et ul. (1975b, 1978) and Maraeva et ul. (1978) showed that polyphosphate phosphohydrolases are controlled by the same regulatory genes as alkaline phosphatase, thus forming a common phosphate regulon together with a number of proteins involved in phosphate metabolism. Such proteins include a phosphate-binding protein (Willsky and Malamy, 1976), one binding glycerophosphate (Argast and Boos, 1980), and one of the proteins of the outer membrane of E. coli, namely protein “e” which is supposedly involved in phosphorylation of pores specific for orthophosphate and its polymers (Argast and Boos, 1980; Tommassen and Lugtenberg, 1980). Comparison of these results points convincingly to the fact that in

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IGOR S. KULAEV AND VLADlMlR M . VAGABOV

living organisms, even such primitive ones as bacteria, regulation of intracellular orthophosphate concentration is subtle even at the genetic level. Control of regulation of orthophosphate concentrations is also explained by the fact that the same cells can contain several “metabolic traps” transforming excess intracellular orthophosphate into a polymerized form. To cite one example, in the fungus P . chrysogenum (Okorokov and Kulaev, 1968) and in other organisms (Okorokov et al., 1970, 1973a,b), polymeric complex compounds of phosphorus with various divalent metal cations (Fez+,Mg2+,Caz+,Co2+and others) were detected together with condensed polyphosphates in which, as we have already seen, orthophosphate residues are linked by the energy-rich phosphoanydride bonds. Investigations of the properties of the complexes isolated suggested that their phosphate residues are linked not by covalent but by co-ordination bonds via metal ions. Many organisms proved to have notable amounts of such polymeric metal phosphates. They occur in cells frequently together with high molecular-weight polyphosphates (Okorokov et ul., 1973b). Hence, orthophosphate released by various biochemical reactions may be bound either through formation of polymeric metal phosphate complexes or with the help of reaciions leading to condensed phosphates (polyphosphates). However, it is important to note that the two pathways of orthophosphate polymerization in the cell differ markedly in their energy requirements, i.e. in contrast to formation of polymeric metal phosphate complexes, polyphosphate biosynthesis requires an additional energy supply to form the macroergic phosphoanhydride bonds. Therefore, under some conditions, regulation of free phosphate concentrations may proceed primarily at the expense of polyphosphate formation in cells. In other cells, polymeric metal phosphates may accumulate, especially those that are, at that time, in excess in the cells and their environment. All of the above considerations refer in the first place to eukaryotes and above all to fungi in which, in addition to polyphosphatases, metal phosphate complexes have been detected. In prokaryotes, in particular in blue-green algae, it is high molecular-weight polyphosphates that are mainly involved in regulating intracellular concentrations of both phosphate anions and many cell-absorbed cations. This conclusion is supported by the investigations already referred to and conducted in Jensen’s laboratory (Sicko-Goad et al., 1975, 1978; Baxter and Jensen, 1980a,b). Improved techniques of X-ray dispersion analysis combined with electron microscopy provided the most detailed information on this problem (Baxter and Jensen, 1980b). This work revealed the ability of the cyanobacteria studied to take up and concentrate in polyphosphate granules divalent metals, such as magnesium, barium and manganese. It is interesting that strontium is accumulated in cells of P . boryanurn not in polyphosphate bodies but in some other electron-dense

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granules containing, instead of phosphorus, sulphur as well as potassium and some calcium. In other experiments (Sicko-Goad et al., 1975, 1978), it was found that calcium ions are also often present in polyphosphate granules. The cation composition of polyphosphate granules in cyanobacteria was found to vary markedly depending on the content of specific cations in the environment. These data suggest that, in prokaryotes, polyphosphates play a very important part in regulating the concentration in cytosol not only of phosphate but also of various metals. It is interesting that in some bacteria, such as Desulfooibrio gigas, low molecular-weight polyphosphates appear to be important in binding excess cations. In particular, Jones and Chambers (1975) isolated from D. gigas granules contaning pure magnesium tripolyphosphate. The physiological significance of the accumulation in prokaryotic cells (as well as in those of eukaryotes) of a large amount of phosphate and, respectively, of specific cations in the form of granules secluded from cytosol, consists in the maintenance of stable, usually rather low, intracellular concentrations of monomeric phosphate and free cations. The question arises concerning the purpose of such a maintenance. In fact, excess accumulation in cells of low molecular-weight compounds, e.g. glucose, amino acids, phosphate, or some cations, may drastically affect the intracellular osmotic pressure and pH value. At the same time, such compounds as AMP, ADP, ATP, acetyl-CoA, Mg2+, NADP, phosphate and glucose are, in certain concentrations, potent effectors and regulators of the functioning of important enzymic systems in the cell. In this connection, in the course of evolution organisms have developed systems of neutralizing excess amounts of physiologically active monomers, i.e. specific “metabolic traps”. In our opinion, such traps function by the processes involved in polymerization of corresponding monomers to glycogen, polyphosphates, poly-8-oxybutyric acid, cyanophycin or polymeric metal phosphates. It seems probable that similar processes involved in detoxication of osmotically active compounds (acetyl-CoA, organic and amino acids) are pathways leading to secondary metabolites including polyphenols, isoprenoids, antibiotics and alkaloids. In concluding the discussion of polyphosphate metabolism in prokaryotes, we will dwell on two other aspects. Recently, data were published on bacteria populating specific ecosystems. Firstly, Bobyk et al. (1980) investigated some particular features of polyphosphate metabolism in Bdellouibrio bacteriouorus, a parasite living on E. coli and some other bacteria. It was found that, in these parastic bacteria, the amounts of polyphosphates are several times higher than that in the host cells, and B. bacteriouorus contained predominantly the acid-insoluble, i.e. surface-localized, fraction of polyphosphates. As already mentioned, certain enzymes of polyphosphate metabolism were detected in these parasites. The activity of 1,Iphosphoglycerate: polyphos-

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M. VAGABOV

phate phosphotransferase and polyphosphatase was much higher than in cells of E. coli, in the periplasm of which they generally live. The fact that the amount of polyphosphates and the intensity of their metabolism in the parasite appears to be higher than in host cells with the B. bacferiovorus-E. coli system brings to mind a similar situation observed by Bennett and Scott (1971) during studies of the wheat stem rust fungus infecting wheat leaves. Secondly, Egorova et al. (198 1) demonstrated substantial amounts of polyphosphates and ATP in the extremely thermophilic bacterium Thermus Jlauus 71, normally growing at 65-70°C. Their concentrations in Thermus flaws 71 exceeded several times those in E. colicells. It is interesting that, in T. Jlavus, polyphosphates are represented mainly by low molecular-weight fractions (PolyP~and PolyPz), i.e. fractions usually localized inside the cytoplasmic membrane. Thirdly, Nikitin et al. (1979, 1983) reported interesting results on analyses of polyphosphate and ATP contents in the oligotrophic bacteria Renobacter vacuolatum and Tuberoidobacter mutans which populate the atmosphere and exhibit very slow growth with the scanty nutrients available in the air. Little ATP and tremendous amounts of polyphosphates, mainly acid-insoluble, were detected in both cases. Also, in R . vacuolatum, 0.54.7 pmol of ATP (g dry wt)-’ and 220-280 pmol of polyphosphates were detected, i.e. the ATP concentration was one order of magnitude lower compared with common eubacteria (e.g. E. coli), whereas that of polyphosphates was one order of magnitude higher compared with concentrations usually detected in bacteria. These findings, as well as other information available in the literature (see, e.g., Sudyina et al., 1978; Kulaev, 1979),enable one to infer that the amounts of polyphosphate fractions and their importance in metabolism vary in bacteria and other organisms and depend greatly on ecological factors. Another aspect of polyphosphate metabolism, which should be given at least brief consideration, is the intensive accumulation of high molecularweight polyphosphates in Acinetobacter sp. isolated from sedimentation tanks containing waste waters of certain industries (Fuhs and Chen, 1975). These bacteria are able to take up from the medium tremendous amounts of phosphate without prior starvation of phosphorus. They can absorb phosphate from sewage waters containing large concentrations of phosphate and accumulate it in the form of polyphosphates. This phenomenon was called “luxury uptake” (Levin and Shapiro, 1965). Deinema et al. (1980) found that the bacteria which absorb phosphate from phosphate-rich media containing butyrate or acetate as a carbon source accumulate large amounts of highly polymerized polyphosphates and lipids. After 40 hours’ growth, the phosphate content was 10-20% and the lipid content was up to 25% of dry weight, and bacterial cells were literally stuffed with polyphosphate and lipid granules. Investigation of these bacteria is of exceptional interest in view of their

POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS

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possible use in extraction of phosphate from waste waters discharged into inland water bodies in industrialized countries.

E. CONCLUDING REMARKS O N THE PHYSIOLOGICAL ROLE OF HIGH MOLECULAR-WEIGHT POLYPHOSPHATES IN MICROBIAL METABOLISM

Summing up, in spite of all that has been stated about the possible physiological role of high molecular-weight polyphosphates in the activities of organisms, it should be underlined that they are regulators of the intracellular concentration of important metabolites including ATP, ADP, other nucleoside polyphosphates, and finally pyro- and particularly orthophosphate. Moreover, they represent a valuable pool of activated phosphate, which can be utilized in various metabolic processes, primarily in those connected with different stages of carbohydrate and nucleic acid metabolism; transport and oxidation of carbohydrates, biosynthesis of cell-wall polysaccharides, and biosynthesis, degradation and functioning of nucleic acids. It is in micro-organisms that high molecular-weight polyphosphatesplay an exceptional role. This is basically explained by two circumstances. First, unlike higher organisms, they do not have a well-developed system of hormonal and nervous regulation; second, micro-organisms depend very much on environmental conditions, resulting from direct contact of cells with the surrounding medium. An impoverished set of regulatory mechanisms in micro-organisms must obviously lead, under certain conditions, to insufficiently finely balanced biochemical reactions. Therefore, micro-organisms should have “metabolic traps” such as high molecular-weightpolyphosphates capable of maintaining their intracellular homoeostasis. The need for “metabolic traps” is also due to a very strong dependence of micro-organisms on environmental conditions. When growth and development of microorganisms depend directly on the environment, it appears very important for the organism to be able to enhance its vital activities immediately on creation of favourable conditions. The availability of sufficient amounts of such valuable endogenous pools as high molecular-weight polyphosphates makes micro-organisms,on the one hand, less dependent on external conditions and, on the other hand, capable, at any suitable moment, of initiating growth and reproduction without any considerable lag-period. In higher organisms, the role of such phosphorus compounds in metabolism is apparently less essential. This inference may be supported by the poor accumulation of polyphosphates in tissues of higher plants and animals and the availability of only a limited number of enzymes for polyphosphate metabolism. It may be assumed that, in highly organized organisms, polyphosphates perform some quite specialized functions, being donors of

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IGOR S. KULAEV AND VLADlMlR M. VAGABOV

activated phosphate only for quite specific biochemical or physiological processes. Such is, in general words, today's position regarding the physiological role of high molecular-weight polyphosphates in the vital activities of contemporary organisms.

111. Inorganic Pyrophosphate: New Aspects of its Metabolic and

Physiological Role Inorganic pyrophosphate is orthophosphate anhydride, which can be considered the least polymeric polyphosphate with an ii value of 2. The free energy of pyrophosphate hydrolysis is close to that released as a result of splitting terminal phosphate groups from ATP and ADP. However, in the presence of bivalent cations, the value of the free energy of pyrophosphate hydrolysis (AGO') is somewhat lower than that for ATP and ADP. According to Lawson et al. (1976), in the presence of 1 mM Mg2+,pH 7.0 and 38"C,the AGO' value of ATP hydrolysis to ADP is 31.8 kJ mol-I (7.6 kcal mol-I) and that of pyrophosphate hydrolysis is 22.1 kJ mol-I (5.27 kcal mol-I). Taking into account the cation concentration in Entamoeba histolytica, Reeves et al. (1974) obtained a AGO' value of 25.1 kJ mol-I (6.0 kcal mol-') for pyrophosphate hydrolysis. Under similar conditions, Flodgaard and Fleron (1 974), in experiments on liver cells, obtained lower values for pyrophosphate, i.e. about 16.7 kJ mol-I (4 kcal mol-I). Thus, from the thermodynamic point of view, pyrophosphate can be a potential source of energy in phosphorylation reactions. It was, however, believed for a long time that pyrophosphate is only a byproduct of numerous reactions of pyrophosphorolysis involved in biosynthesis of proteins, nucleic acids, lipids, polysaccharides and nuceloside coenzymes. It was thought that, due to the activity of cellular pyrophosphatases, pyrophosphate could not be accumulated in the cell and was hydrolysed to orthophosphate, thus ensuring that the above reactions were irreversible (Kornberg, 1957, 1959; Hoffmann-Ostenhof and Slechta, 1957).

A. UTILIZATION OF PYROPHOSPHATE IN PHOSPHORYLATION REACTIONS I N BACTERIA

The first reaction in micro-organisms in which pyrophosphate was shown to be utilized as a source of phosphorylation, thus replacing ATP, was that revealed by Siu and Wood (1962) in Propionibacterium shermanii. This

PO LY PH0 s PHATE METAB0LI S M I N M I C RO - 0 R G AN IS MS

143

reaction is catalysed by the pyrophosphate (PPi)-dependent phosphoenolpyruvate (PEP) carboxykinase (EC 4.1.1.38):

+

PPi oxaloacetate+PEP +Pi +COz

(1)

Pyrophosphate-dependent phosphoenolpyruvate carboxykinase reaction (1) is similar to the following reaction: ATP + oxa1oacetateePEP + ADP + CO2

(2)

which is catalysed by phosphoenolpyruvate carboxykinase (GTP) (EC 4.1.1.32) earlier discovered by Utter and his coworkers (Utter and Kurahashi, 1954; Utter et al., 1954) and widely found in nature (Scrutton and Young, 1972). As shown by Wood et al. (1966), pyrophosphate-dependent carboxykinase (EC 4.1.1.38) operates in P . shermanii as phosphoenolpyruvate carboxykinase (EC 4.1.1.32), especially in bacteria grown on lactate. In the protozoan E. histolytica, the polyphosphate-dependent enzyme could also substitute for the lack of phosphoenolpyruvate carboxykinase (GTP) (Reeves, 1970, 1976). Later, this enzyme was found in Rhodopseudomonas palustris (Chernyadyev et al., 1972) and Brevibacterium ammoniagenes (Baryshnikova and Loginova, 1979). In Rh. rubrum, pyrophosphate-dependent carboxykinases are active only when bacteria are grown in the light in the presence of malate, i.e. under conditions of active pyrophosphate synthesis by Rh. rubrum cells (Shady et al., 1975). Another reaction occurs in micro-organisms in which pyrophosphate is a phosphorylating agent. This reaction is catalysed by pyruvate, phosphate dikinase (EC 2.7.9.1) and proceeds as follows:

+

+

PPi AMP PEP+pyruvate+ ATP +Pi

(3) This enzymic pathway of pyrophosphate utilization has been found in propionic bacteria (Evans and Wood, 1968), E. histolytica (Reeves, 1968)and Bacteroides symbiosus (Reeves et al., 1968; Reeves, 1971). Later, this enzyme was found in Acetobacter suboxydans cultured on substrates of the tricarboxylic acid (TCA) cycle (Benziman and Palji, 1970; Benziman and Eisen, 1971) and in some photosynthetic bacteria (Buchanan, 1974). The pyruvate, phosphate dikinase reaction (3) is similar to the reaction catalysed by pyruvate kinase (EC 2.7.1.40): ADP +PEPepyruvate + ATP

(4)

In B. symbiosus and E. histolytica, pyruvate phosphate dikinase is involved in glycolysis (Reeves, 1968, 1976; Reeves et al., 1968; Wood, 1977; Wood et al., 1977) substituting for pyruvate kinase which these organisms lack. In this situation, pyrophosphate acts as a direct source of high-energy phosphate required for ATP biosynthesis. The functioning of these two enzymes depends

144

IGOR S. KULAEV AND VLADlMlR

M. VAGABOV

directly on the conditions under which micro-organisms are grown. In A. suboxydans, pyruvate phosphate dikinase can be found only when organisms are cultured in a medium containing pyruvate or TCA cycle substrates. A study of the regulation of pyruvate kinase and pyruvate phosphate dikinase in A. suboxydans showed that activities of the two enzymes are regulated in an opposite manner, depending on the energy state of the cell, particularly on the ratio of concentrations of adenine nucleotides, i.e. AMP, ADP and ATP. Pyrophosphate-dependent phosphofructokinase (EC 2.7.1.9.0), which is responsible for the reversible reaction:

+

Fructose 6-phosphate + PPiefructose 1,6-diphosphate Pi

(5)

was detected and studied in E. histolytica (Reeveset al., 1976)and P . shermanii (O’Brien et al., 1975). Recently this enzyme was found in marine organisms Alcaligenes sp. and Pseudomonas marina (Sawyer et al., 1977)and Bacteroides fragilis (Macy et al., 1978). The above reaction ( 5 ) is similar to that catalysed by ATP-dependent phosphofructokinase:

+

ATP + fructose 6-phosphateefructose 1,6-diphosphate ADP

(6)

It is interesting to note that activities of pyrophosphate-dependent phosphofructokinase in P . shermanii and E. histolytica are one order of magnitude higher than those of the ATP-dependent enzyme. Apparently, in propionic bacteria grown on glucose, pyrophosphate essentially replaces ATP in synthesis of fructose 1,ddiphosphate (O’Brien et al., 1975). The transformation of fructose 1,Qdiphosphate takes place mostly due to activity of phosphofructokinase to form pyrophosphate (by the reverse of reaction 5) compared to hydrolysis by fructose diphosphatase, as the activity of the latter is 1 5 2 0 times lower than that of pyrophosphate-dependent phosphofructokinase. Attempts to detect pyrophosphate-dependent phosphofructokinase in yeast failed. Konoshenko et al. (1979) demonstrated that pyrophosphate is a competitive inhibitor of ATP-dependent phosphofructokinase. In addition to the above enzymes, in P . shermanii, P . technicum and P. freudenreichii, an enzyme responsible for direct phosphorylation of serine to 0-phospho-L-serine (EC 2.7.1.80) (Cagen and Friedmann, 1968, 1972) has been detected:

+

PPi serine*phosphoserine

+Pi

(7)

A pyrophosphate-dependent acetyl kinase was found in E. histolytica to catalyse a reaction similar to the ATP-dependent acetyl kinase reaction (Reeves and Guthrie, 1975):

PPi + acetate$acetyl phosphate + Pi

+

ATP acetategacetyl phosphate +Pi

(8) (9)

PO LY PH0sPHAT E METAB0LISM I N MIC R 0- 0R GA NI S MS

145

It was then reported that two microsomal polypeptides can be phosphorylated at the expense of pyrophosphate (Lam and Kasper, 1980a,b). All of these data point to an important role being played by pyrophosphate as a compound which, in certain cases, can successfully replace ATP in phosphotransferase processes in micro-organisms. At the same time, a pyrophosphate-dependent glucokinase, which catalyses phosphorylation of glucose to glucose 6-phosphate with pyrophosphate, has not hitherto been detected in micro-organisms (Nordlie and Arion, 1964; Stetten, 1964; Stetten and Tafft, 1964; Nordlie, 1976). More detailed information on the enzyme involved in phosphorylation reactions can be found in the reviews by Reeves (I 976), Wood (1977), Wood et al. (1977) and Mansurova (1982), as well as in other recent publications (Milner et al., 1978; Moscovitz and Wood, 1978; Yoshida and Wood, 1978).

B . ENERGY-DEPENDENT SYNTHESIS OF PYROPHOSPHATE DURING PHOTOSYNTHETIC A N D OXIDATIVE PHOSPHORYLATION

The first evidence for an energy-dependentbiosynthesis of pyrophosphate was obtained with animal tissue homogenates (Cori, 1942; Cross et al., 1949). At the end of the 1950s, Klungsoyr and his colleagues (Klungsoyr et al., 1957; Klungsoyr, 1959) demonstrated intensive incorporation of [32P]orthophosphate into pyrophosphate by aerated cells of A. suboxydans, E. coli and Merrulins lacrimans. At the same time, Shaposhnikov and Fyodorov (1960), working on the green sulphur bacterium Chlorobium thiosulphatophilum, showed that under illumination in the absence of carbon dioxide [32P]orthophosphate was incorporated into polyphosphates of the acid-soluble fraction at a high rate. However, the authors did not identify the particular compound. In 1966, Baltscheffsky and his colleagues (Horio et al., 1966; H. Baltscheffsky et al., 1966; M. Baltscheffsky et al., 1966; Baltscheffsky and Stedingk, 1966) reported that chromatophores of the non-sulphur purple bacterium Rh. rubrum could synthesize pyrophosphate as an alternative to ATP and utilize it in reactions occurring at the level of the photosynthetic electron-transport chain. Almost simultaneously, light-dependent synthesis of pyrophosphate was found in cells of Chlorella sp. and spinach chloroplasts (Pedersen et al., 1966a,b). It should be noted that Rh. rubrum cells, grown in the light, contained pyrophosphate either in the same or far greater amounts than ATP. Pyrophosphate was not detected in an acid extract from the same cells grown in the dark (Shady et al., 1975). The experimental data available suggest that pyrophosphate-dependent energy metabolism is inherent not only in Rh. rubrum but also in Rh. palustris and Rh. viridis (Chernyadyev et al., 1972; Knobloch, 1975; Jones and

146

IGOR S. KULAEV AND VLADlMlR M. VAGABOV

Sanders, 1972) and probably in other photosynthetic micro-organisms (Shaposhnikov and Fyodorov, 1960). An outstanding contribution to a proper understanding of the important role that pyrophosphate may play in the bioenergetic processes of the cell was made by H. Baltscheffsky and his coworkers. They demonstrated that, in the absence of ADP, chromatophores of Rh. rubrum carry out light-dependent synthesisof pyrophosphate (Horio et al., 1966; H. Baltscheffsky et al., 1966; M. Baltscheffsky et al., 1966). It was shown that syntheses of both pyrophosphate and ATP depend on photosynthetic electron transport and are inhibited by antimycin and uncouplers. However, there is an important difference between synthesis of pyrophosphate and of ATP. Oligomycin does not inhibit synthesis of pyrophosphate and sometimes stimulates it slightly; moreover dicyclohexylcarbodiimide had no effect on synthesis of pyrophosphate, whereas Di 0-9 inhibits that of both compounds (Guillory and Fisher, 1972). The Baltscheffskys (H. Baltscheffsky et al., 1966, 1969, 1971; M. Baltscheffsky, 1969a,b; H. Baltscheffsky and M. Baltscheffsky, 1972;Baltscheffsky, 1977), Keister and his colleagues (Keister and Yike, 1967a,b; Keister and Minton, 1971a,b; Rao and Keister, 1978) and Skulachev and his colleagues (Isaevet al.?1970,1976;Kondrashinet al., 1980;Skulachev l971,1972a, 1975; Ostroumov et al., 1973) have described in detail the mechanism of pyrophosphate biosynthesis and utilization at the level of the electron-transport chain. The reaction sequence is presented in Fig. 16. Cyclic transport of electrons results in formation of a non-phosphorylated high-energy intermediate or a certain energized state of the membrane whose energy can be used to maintain ion transport, reverse electron transfer and transhydrogenase reaction of pyrophosphate-dependent NAD+ reduction, to modify the conformation of carotenoid molecules leading to changes in their absorption spectrum, and, finally, to synthesize ATP and pyrophosphate (H. Baltscheffsky et al., 1966; M. Baltscheffsky et al., 1966; M. Baltscheffsky, 1969a,b, 1971, 1974, 1977; Keister and Yike, 1967a,b; Keister and Minton, 1971a,b;Azzi et al., 1971; Fischer and Guillory, 1969a,b). A detailed survey of the problem can be found in a recent publication by Baltscheffsky (1978). Baltscheffsky and Stedingk (1966) hypothesized that the resultant pyrophosphate can be further used in biosynthesis of inorganic polyphosphates. The most important inference from this work is that pyrophosphate is a product of photophosphorylation in chromatophores as an alternative to ATP, and that membrane pyrophosphatase is a factor coupling electron transport and pyrophosphate synthesis. These conclusions are supported by studies in Saccharomyces cerevisiae and a yeast-like fungus Endomyces magnusii (Mansurova et al., 1975b, 1977a, 1978).It appears that mitochondria of these organisms synthesize, during oxidative phosphorylation, not only ATP but pyrophosphate as well. Inhibitors of the respiratory chain, such as antimycin

POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS

147

Light

Bacteriochlorophy ll

Cyclic electron- transport chain

-X ,

quinone

cytochrome b

Change in the spectrum of carotenoids

transhydrogenase reaction

PH

PPI

ATP

FIG. 16. Mechanism of light-dependent biosynthesis of polyphosphate in chromatophores of Rhodospirillum rubrum and its utilization in dark reactions. From Mansurova (1982).

and cyanide, and the uncoupler 2,4-dinitrophenol inhibit both synthesis of ATP and pyrophosphate. Oligomycin inhibits only ATP synthesis, whereas sodium fluoride inhibits only pyrophosphate synthesis. It can be concluded that, in mitochondria, pyrophosphate synthesis depends on the respiratory chain and that ATP does not act as pyrophosphate precursor. It appears that maximal synthesis of ATP and pyrophosphate in mitochondria and chromatophores requires different conditions. For instance, this operates with respect to the rate of electron flow along the photosynthetic and respiratory electron-transport chains, and to the redox potential of electron carriers (Horio et al., 1965; Horiuti et al., 1968; Nishikawa et al., 1973; Pullaiach et al., 1980). It can be postulated (Pullaiach et al., 1980) that phosphorylation yielding pyrophosphate takes place when ATP synthesis becomes restricted or impossible. As already mentioned, membrane-bound pyrophosphatase participates in energy-dependent synthesis of pyrophosphate. Rhodospirillum rubrum contains at least two pyrophosphatases; one cytoplasmic (Klemme and Gest, 1971a,b) and the other membrane-bound (M. Baltscheffsky et al., 1966;

148

IGOR S. KULAEV A N D VLADlMlR

M . VAGABOV

Fischer and Guillory, 1969a; Isaev et al., 1970; Keister and Minton, 1971a,b; Guillory and Fischer, 1972; Dutton and Baltscheffsky, 1972). The soluble enzyme has entirely different properties from the membrane one. The two pyrophosphatases were found in mitochondria of all organisms studied; including yeast, fungi and animal tissues (Irie et al., 1970; Kulaev et al., 1973c; Umnov et al., 1974b). It should be noted that the chromatophore pyrophosphatase activity can be manifested only in a lipid environment. Detergent extraction of the enzyme from membranes followed by removal of phospholipid results in complete inactivation. However, the enzyme can be reactivated in the presence of phospholipids (Isaev et al., 1970; Kondrashin et al., 1980). Kondrashin et al. (1980) showed that membrane pyrophosphatase isolated from chromatophores can be incorporated into liposomes and generate a membrane electrochemical potential on addition of pyrophosphate. It is so far unclear what is the nature of the common intermediate for ATP and pyrophosphate synthesis. Recent data suggest that the common intermediate in the synthesis of ATP and pyrophosphate is, in agreement with Mitchell’s views (Mitchell, 1961, 1966, 1968), an energized state of the membrane with an electrochemical potential across it. Inorganic pyrophosphate is one of the components of the common energy pool of the cell, and the energy of the phosphoanhydride bond can be transferred from ATP to polyphosphate and back through intermediate formation of the electrochemical potential. The validity of reaction sequences leading to ATP and pyrophosphate (Fig. 17) has been convincingly proved by the studies of pyrophosphate-dependent

Adenosine triphosphatase

AT P

A~u.H+

Pyrophosphatase

PP

FIG. 17. Mechanism for the energy-dependent synthesis of ATP and pyrophosphate (PP) in chromatophores, chloroplasts and mitochondria. A and B are components of the electron transport chain.

POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS

149

synthesis of ATP and ATP-dependent synthesis of pyrophosphate carried out by Keister and his colleagues (Keister and Minton, 1971a,b; Keister and Raveed, 1974;Baltscheffsky and Baltscheffsky 1972; Mansurova et al., 1973a, 1975a,c). Of great importance is the fact that these reactions took place with simultaneous involvement of ATPase and pyrophosphatase and when the respiratory chain was switched off. In this situation, only the energy of the phosphoanydride bond of ATP or pyrophosphate is used, and the orthophosphate residue is not transferred. For instance: 1. PPi

pyrophosphau

2. ADP +"Pi

+

,2pi + "

N

"

ATPase

"N "

-ATP3'

The data obtained indicate that, in chromatophores and mitochondria, the content of ATP and pyrophosphate are in equilibrium, established with participation of coupling ATPase and pyrophosphatase. In chromatophores, synthesis of one molecule of ATP is accompanied by hydrolysis of 10 molecules of pyrophosphate (Keister and Minton, 1971a,b). It should be mentioned, however, that, in addition to the coupling membrane pyrophosphatase, chromatophores contain an active cytoplasmic membrane which also contributes to pyrophosphate hydrolysis. The intensity of ATP and pyrophosphate production is, to a great extent, affected by the physicochemical state of the membrane. It was shown that the rate of ATP and pyrophosphate synthesis depends directly on the viscosity of the phospholipid component of the mitochondria1inner membrane (Kulaev et al., 1980; Mansurova ef al., 1982). Synthesis of ATP increases with decreasing membrane fluidity, and inversely, increasing membrane fluidity favours pyrophosphate synthesis. The literature indicates that light-dependent synthesis of pyrophosphate can be performed not only by photosynthetic bacteria but also by chloroplasts of algae and higher plants (Rubtsov et al., 1976). Both in chromatophores (Guillory and Fischer, 1972) and in chloroplasts (S.E. Mansurova, personal communication), maximal synthesis of pyrophosphate occurs at a lower illumination than that required for ATP. Thus, data obtained in recent years on pyrophosphate metabolism in micro-organisms and the occurrence of energy-dependent synthesis of pyrophosphate in both mitochondria of lower organisms (yeasts) and higher eukaryotes (mammals) (Mansurova et al., 1973a,b, 1975a,b, 1976, 1977a,b), as well as in chloroplasts of algae and higher plants (Rubtsov et al., 1976) enable one to regard pyrophosphate not only as a byproduct of pyrophosphorolysis reactions used in the bioenergetics of the most ancient forms of life but also as a high-energy compound similar to ATP, involved in storage and

150

IGOR S. KULAEV AND VLADlMlR M . VAGABOV

utilization of energy in contemporary highly organized micro-organisms. Investigating the role of pyrophosphate in metabolism has turned from a matter of elucidating the peculiarities of the energetics of the most ancient life forms and revealing the manifestations of “predeluvian metabolism” (Wood, 1977; Wood et al., 1977; H. Baltscheffsky, 1971) into a problem of general biological interest.

C . RELATIONSHIP BETWEEN PYROPHOSPHATE A N D POLYPHOSPHATE

METABOLISM IN MICRO-ORGANISMS

The Italian authors Ipata and Felicioli (1963) were the first to raise this question. They reported enzymic phosphorolysis of high molecular-weight polyphosphates to pyrophosphate in yeast. We attempted to reproduce these experiments and failed to demonstrate an enzymic character of the reaction (Mansurova et al., 1973; Kulaev and Skryabin, 1974):

+

(Polyp),+ 32Pi+32PPi (Polyp),-

1

We showed that, in the presence of divalent cations, significant quantities of radioactive pyrophosphate were formed nonenzymicallythrough phosphorylation of [32P]orthophosphateby high molecular-weight polyphosphate. Pyrophosphate can be synthesized enzymically from tripolyphosphate by means of a specific enzyme, tripolyphosphatase. This enzyme has been detected in many organisms, including Aspergillus oryzae (Neuberg er ul., 1950), Neurospora crassa (Kulaev and Konoshenko, 1971b), Phyrophrhoru infestans (Sysuev et al., 1978), yeast (Mattenheimer, 1956a,b,c; Felter and Stahl, 1970), Aerobacter aerogenes (Dawes and Senior, 1973), Bacillus sp. (Szymona and Zajac, 1969) and E. coli (Nesmeyanova et al., 1973a).A study of the intracellular localization of the enzyme in yeast and N . crussu demonstrated its presence in vacuoles (Schwencke, 1978), periplasmic space (Kulaev et al., 1972b; Konoshenko et al., 1973) and, to a large extent, in mitochondria (Kulaev et al., 1972b; Konoshenko et al., 1973; Umnov ef ul., 1974b). The enzyme from N . crussa was purified to homogeneity. Egorov and Kulaev (1976) convincingly demonstrated that tripolyphosphate hydrolysis by tripolyphosphatase takes place according to the equation: PPPi +PPi

+Pi

In recent years, relationships between the metabolism of pyrophosphate and that of acid-soluble polyphosphates were given special study (Mansurova, 1979; Ermakova et al., 1981). It was found that, during yeast growth, the pyrophosphate content varies drastically forming two maxima of accumulation at the beginning and at the end of the exponential phase of growth

POLYPHOSPHATE METABOLISM I N MICRO-ORGANISMS

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(Ermakova er al., 1981; Shakhov et al., 1978). This pattern was also seen during growth of the yeast hybrid strain Sacch. cerevisiae N.C.Y.C. 644 SU3 with and without aeration in media containing various concentrations of glucose, and during aerobic cultivation of Candida guilliermondii on glucose or petroleum hydrocarbon-containing media (Shakhov et al., 1978). Peak pyrophosphate accumulation was not related to changes in the rates of respiration or fermentation. The pyrophosphate content of the yeast Sacch. cerevisiue exceeded the ATP content at different stages by a factor of 10-1000, reaching 2-17 mg (g dry wt)-'. It can be seen that accumulation of such large amounts of pyrophosphate is not associated with bioenergetic processes, and it can be assumed that its major function, as with certain fractions of highly polymeric polyphosphates, is as a form of energy and phosphorus reserve in the cell. When the content of pyrophosphate reached a maximum, that of acid-soluble polyphosphates with higher molecular weights dropped to a minimum (Fig. 18). Microscopic examination of the yeast showed that marked variations in the contents of pyrophosphate and other acid-soluble polyphosphates during growth are associated with significant synchronization of cell budding (Ermakova et al., 1981). Pyrophosphate accumulation is particularly active in cells having a large number of small intensively growing buds. These data are in good agreement with the findings of Nurse and Wiemken (1974) who observed accumulation of low molecular-weight

2oFt tI t

--1 25

Growth (hours)

FIG. 18. Changes in the content of pyrophosphate ( 0 )and acid-soluble polyphosphates without pyrophosphate (0) during growth in aerated (a) and non-aerated (b) cultures of Succh. cerevisiue N.C.Y.C. 644 SU3. From Ennakova et al. (1981).

152

IGOR

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AND VLADlMlR M. VAGABOV

substances at the beginning of bud formation in the yeast. As the buds approached the size of the mother cell, the content of pyrophosphate decreased dramatically and that of acid-soluble polyphosphates increased. These polyphosphates are located within the cell, primarily in vacuoles (Indge, 1968; Urech et af., 1978; Schwencke, 1978; Cramer et al., 1980) and, in contrast to other polyphosphate fractions, their behaviour is opposite to that of pyrophosphate throughout the entire period of cultivation. It is unclear so far how acid-soluble polyphosphates are utilized. It is probable that their high-energy phosphate groups can be used in synthesis of ATP and other nucleoside triphosphates by means of polyphosphate: ADP phosphotransferase (EC 2.7.4.1) or be transformed into inorganic pyrophosphate through direct degradation by polyphosphate depolymerase (EC 3.6.1.10) and tripolyphosphatase. It should be noted that three enzyme activities are present in yeast vacuoles (Shabalin et af.,1977; Schwencke, 1978). It is likely that the resultant pyrophosphate is used as an energy source during cation transport through the tonoplast (Okorokov et af., 1980). In addition, degradation products of vacuolar polyphosphates, in particular tripolyphosphate, pyrophosphate and orthophosphate, may participate in transport of arginine and other positively charged molecules across the membrane, as conjectured for yeast (Diirr et af., 1979; Matile, 1978; Okorokov et af., 1980). Having discovered light-dependent synthesis of pyrophosphate in Rh. rubrum chromatophores, Baltscheffsky and Stedingk (1966) assumed that it may take part in the biosynthesis of high molecular-weight polyphosphates. However, our study of polyphosphate and pyrophosphate metabolism in this organism (Kulaev et al., 1974c; Shady et af., 1976) failed to support this concept. In fact, culturing Rh. rubrum in light led to accumulation of significant quantities of pyrophosphate and salt-soluble polyphosphates. Synthesisof high molecular-weight polyphosphates depended on the electrontransport chain and was inhibited by antimycin (Table 11). TABLE 11. Synthesis of ATP, pyrophosphate and salt-soluble polyphosphates by chromatophores of Rhodospirihm rubrum in the light Rate of synthesis (32Pcounts min-l) With ADP

Without ADP

Without

With

With

Compound

inhibitors

antimycin

oligomycin

Without inhibitors

antimycin

ATP Pyrophosphate Polyphosphate

99,490 570 19,110

11,570 160 1,370

27,460 570 3,690

200 2700 330

200 1360 000

With

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Oligomycin inhibition of ATP synthesis led to a parallel decline in accumulation of this fraction of polyphosphates (Shady et al., 1976). It can be assumed that synthesis of salt-soluble polyphosphates in these bacteria is realized at the expense of ATP by action of ATP: polyphosphate phosphotransferase (EC 2.7.4. l), an enzyme widely distributed among microorganisms, including photosynthetic bacteria (Kulaev, 1979). In this case, the pyrophosphate-polyphosphate relation can be mediated by the adenine nucleotide system. It is appropriate to mention here the work of Butukhanov et al. (1979) in which direct phosphorylation of AMP and ADP at the expense of pyrophosphate was reported: AMP + PPi +ADP

+Pi ADP + P P +ATP ~ +pi

The enzymes responsible for the two reactions differed from the corresponding polyphosphate-dependent enzymes. It may be postulated therefore that, in certain micro-organisms, the energy of the phosphoanhydride bond and sometimes the phosphoric acid residue can be readily transferred from pyrophosphate to polyphosphate and back via adenine nucleotides. However, as already shown, for example, for synthesis of nucleic acids and for some other biosynthetic processes, a direct transfer of orthophosphate residues from pyrophosphate to polyphosphates is not excluded. This process seems to be closely connected topologically with the functioning of those biosynthetic systems in which pyrophosphate is one of the end products. It may be thought that such conjugated systems are analogous to the recently detected multi-enzyme complex of adenylate translocase and creatine phosphokinase in animal mitochondria (Saks et al., 1977, 1980).

IV. Modern Concepts about the Role of High Molecular-Weight Polyphosphates and Pyrophosphate in Evolution of Phosphorus Metabolism At the present time, most geochemists and biologists hold that the earliest living beings on Earth were anaerobic micro-organisms which obtained energy from hexose fermentation to lactate and ethanol (Oparin, 1957). The further course of evolution is debatable. Some researchers believe the fermenting anaerobes were followed by “anaerobically breathing” organisms that had an incipient membranous electron-transport chain (Sagan, 1967; Margulis, 1970; Hall, 1971). In their opinion, mutations of the gene coding for the cytochrome prosthetic group led to emergence of chloro-

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phyll and anaerobic photosynthetic organisms. In contrast, most authors (Schlegel, 1972; Skulachev, 1972b, 1974; Broda, 1971, 1975; Gusev and Gokhlerner, 1980) share the opinion that the first electron-transport chains and the coupling mechanisms of electron transport and phosphorylation developed after anaerobic micro-organisms acquired photosynthetic abilities. Oxygen accumulation in the Earth’s atmosphere could be accounted for by the appearance of photosynthetic cyanobacteria capable of water photolysis (Oparin, 1957; Broda, 1971; Gusev and Gokhlerner, 1980; Wilson and Lin, 1980). The presence in the atmosphere of sufficient quantities of oxygen was responsible for emergence of aerobic organisms that utilized it as the terminal electron acceptor. It is hard to say whether anaerobic respiration was primary or secondary to photosynthesis. It is essential that, at a certain evolutionary stage, glycolytic phosphorylation occurring in solution was paralleled by membrane bioenergetics. It is possible that, on the early Earth, initial high-energy phosphates were represented by high molecular-weight polyphosphates synthesized at high temperatures during volcanic and other processes (Belozersky, 1959; Kulaev, 1971, 1979) and by inorganic pyrophosphate produced either from orthophosphates (Calvin, 1963, 1971; Miller and Parris, 1964; Beck and Orgel, 1965; Lipmann, 1965)or non-biologicallyfrom polyphosphates in an aqueous medium (Mansurova et al., 1973c; Kulaev and Skryabin, 1974). It can be assumed that, when the Earth was surrounded by a reducing atmosphere with a low concentration of oxygen, both high molecular-weight polyphosphates and pyrophosphate were major components of the energy system in primordial organisms. Calvin (1963) and Lipmann (1965) were the first to suggest the participation of pyrophosphate in accumulation and transfer of energy-rich bonds on the primeval Earth. These authors put forward the idea that the reactions typical of primitive forms of life evolved from prebiological systems, and that living organisms of today still have the ability to utilize pyrophosphate as a high-energy compound. This concept found support in our investigations (Mansurova et al., 1975a,c, 1976, 1977a,b; Rubtsov et al., 1976; Mansurova and Ibragimov, 1979) and in experiments carried out by Wood and his colleagues (Wood, 1977; Wood et al., 1977)and other workers (Baltscheffsky et al., 1966; Batscheffsky and Stedingk, 1966; Nordlie and Arion, 1964; Nordlie, 1976; Reeves, 1976). We have demonstrated that contemporary primitive organisms, including bacteria, actinomycetes and fungi, contain an enzyme catalysing transfer of activated phosphate from 1,3-diphosphoglyceratenot to ADP to form ATP but directly to high molecular-weight polyphosphates (Kulaev et al., 1968; Kulaev and Bobyk, 1971). Active synthesis of pyrophosphate at the expense

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of glycolytic phosphorylation has also been observed in yeast (Mansurova et al., 1976). Extensive studies on the distribution of polyphosphate hexokinase among micro-organisms performed in our laboratory have shown that this enzyme occurs only in phylogenetically very ancient and closely related micro-organisms forming a distinct class of actinomycetes according to Krasilnikov’s (1949) evolutionary systematics. It should be emphasized that, in the oldest representatives of these micro-organisms, e.g. micrococci, tetracocci and propionic-acid bacteria (Kulaev, 1979), activity of polyphosphate hexokinase is several times higher than that of ATP hexokinase, whereas in the newest representatives of this class, i.e. true actinomycetes,the activity of ATP hexokinase significantly exceeds that of polyphosphate hexokinase. These data, as well as the findings of Wood et al. (1977), give evidence that, in the best studied propionic-acid bacteria, glycolytic degradation of glucose takes place with participation of polyphosphates and pyrophosphate rather than the ADP-ATP system. It can be expected that, with increasing importance of membrane-bound energy processes in primitive orgaisms, the bioenergetic role of ATP and pyrophosphate will become more significant while that of polyphosphates will correspondingly decrease. It is far from incidental that high molecular-weightpolyphosphates, and enzymes of their metabolism, are absent from chloroplasts of algae and higher plants (Rubtsov and Kulaev, 1977; Rubtsov et al., 1977) and mitochondria (Kulaev et al., 1967b) which, according to the theory of symbiogenesis of eukaryotic cells (Sagan, 1967; Margulis, 1970), are of microbial origin. It should be recalled, however, that tripolyphosphate and tripolyphosphatase are present in Rh. rubrum and mitochondria (Kulaev et al., 1972b; Konoshenko et al., 1973; Umnov et al., 1974b). It is logical to raise the question whether this enzyme, like adenosine triphosphatase and tripolyphosphatase, can or could participate in biosynthesis of tripolyphosphate coupled with electron transport. Having essentially lost the role of primary energy acceptors and donors in the course of evolution, high molecular-weight polyphosphates began to perform new functions. They play a particularly important part in the life of contemporary micro-organisms serving as pools of activated phosphate groups, high molecular-weight ion exchangers, and regulators of many metabolic processes. However, even in today’s micro-organisms, these compounds can be synthesized during glycolyticphosphorylation and utilized together with pyrophosphate in substrate phosphorylation (Kulaev, 1979). No matter how great was the role of polyphosphates and pyrophosphates in primitive micro-organisms, it appears that they were able to synthesize ATP prior to the development of photosynthesis or anaerobic respiration. If this were not so, modern fermenting bacteria could hardly have the capacity to synthesize ATP, as Broda (1971, 1975) indicates.

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Estimating the role of membranes in bioenergetic processes, Mitchell (1970) wrote: “It is an evolutionary attractive proposition that the proton-translocating oxidoreduction loop system and the reversible proton-translocating ATPase may have arisen separately as alternatives for generating the pH difference and membrane potential required for nutrient uptake and tonic regulation via porter systems in primitive prokaryotic cells and may then have provided the means of storing free energy of oxidoreduction in ATP synthesized by the reversal of the ATPase, or in some other anhydride, such as pyrophosphate, produced by a similar mechanism”. A very similar composition and identical structure of coupling membranes, high similarity of coupling mechanisms in membranes of photosynthetic and aerobic bacteria, as well as in mitochondria and chloroplasts, allowed Skulachev (1972b) to suggest that the system by which electron transport and phosphorylation are coupled, once created, was used in every organism that has survived until today without any fundamental change. This seems to hold true for energy-dependent synthesis of pyrophosphate, which occurs not only in chromatophores of very old micro-organisms from the evolutionary point of view, i.e. Rh. rubrum (H. Baltscheffsky et af., 1966; M. Baltscheffsky et af., 1966; Baltscheffsky and Stedingk, 1966) but also in chloroplasts of algae and higher plants (Rubtsov et af.,1976) and in mitochondria of lower and higher eukaryotic organisms (Mansurova et af., 1973a, 1975a,b, 1977a). It is very likely that, after the appearance of pyrophosphate synthesis coupled with electron transport, pyrophosphate synthesized in one way or another could, performing other functions as well, participate in maintenance of the electrochemical potential across the membrane (Skulachev, 1978). Evolution of bioenergetic processes also involved evolution and sophistication of regulatory systems. This could have led to supersession of high molecular-weight polyphosphates and pyrophosphate as monotonically built compounds by a more complicated multifunctional and readily recognizable structure, i.e. ATP. Nevertheless, even in mammalian cells, sufficiently high amounts of pyrophosphate (Guynn et al., 1974; Mansurova et al., 1977a; Veech et af., 1980) and certain quantities of polyphosphates (Kulaev and Rozhanets, 1973; Mansurova et af., 1975b) can be detected. Moreover, specific enzymes utilizing pyrophosphate in phosphorylation reactions have been identified (Nordlie and Arion, 1964; Stetten, 1964; Stetten and Tafft, 1964; Nordlie, 1976; Mansurova and Ibragimov, 1979). It is obvious that, in higher plants and animals, the importance of high molecular-weight polyphosphates and pyrophosphate in bioenergetic processes decreased. However, it is still unclear what specific functions these compounds have retained in the course of evolution from primitive forms of life to the highiy organized living beings of today.

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V. General Conclusions The present review devoted to the physiological role of inorganic polyphosphates was conceived with the purpose of depicting the precise state of investigations of this yet vague problem. A wealth of new data have appeared since publication of a detailed monograph (Kulaev, 1979) devoted to various aspects of the biochemistry of polyphosphates. Some findings provide certain support or, on the contrary, refute earlier assumptions and postulates, whereas others open new aspects in studies of inorganic polyphosphate metabolism. The general conclusion drawn from these data is that inorganic polyphosphates can play an essential part in metabolism of organisms containing these compounds, in the first place as high-energy phosphorus compounds functionally alternative to ATP. Moreover, the ever-increasing amount of data point to an important role of polyphosphates in regulation of numerous biochemical processes. A major achievement of recent investigations consisted in the formation of the concept about the heterogeneity of different polyphosphate fractions not only as regards chain length but also in both intracellular localization of the pathways of their biosynthesis and utilization and their functional role in the vital activities of the cell. Finally, it has become evident that studies on the biochemistry of inorganic polyphosphates contribute not only to the progress of fundamental science but also have practical significance for the most burning problem of the present time, namely preservation of the environment. Many problems of the biochemistry of polyphosphates have been raised in this review and necessitate further investigations. They are: 1. What are the precise mechanisms of the relationship between polyphosphate and nucleic acid metabolism? 2. Are there any specific mechanisms of polyphosphate transport from one membrane structure to others? 3. What are the relationships between the metabolism of high molecularweight polyphosphates and pyrophosphate? 4. What is the practical significance of polyphosphates as biological ion-exchangers?

We hope that these and many other prcblems of the biochemistry of polyphosphates will be successfully solved in the near future.

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V. Acknowledgements The authors are very grateful to their colleagues from the Laboratory of Regulation of Biochemical Processes of the Institute of Biochemistry and Physiology of Micro-organisms of the USSR Academy of Sciences and from the Department of Molecular Biology of the Lomonosov State University, Moscow, whose work on the biochemistry of polyphosphates in microorganisms stimulated writing of this review. Special thanks go to Drs S. E. Mansurova, M. A. Nesmeyanova, M. A. Bobyk, M. S. Kritsky, D. I. Nikitin, L. A. Okorokov and D. N. Ostrovsky for the kind submission of new data and for fruitful discussion. The authors’ thanks are also due to A. V. Mudrik and L. G. Sergeyeva for the assistance in the preparation of the manuscript and to V. D. Gorokhov for its translation into English. REFERENCES

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Physiology of Acidophilic and Alkalophilic Bacteria TERRY A. KRULWICH AND ARTHUR A. GUFFANTI Mount Sinai School of Medicine of the City University of New York. New York, N.Y. 70029, U.S.A. 1. Introduction . . 11. Acidophilic bacteria

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I. Introduction

Five years ago, when Langworthy (1978) reviewed the physiology of acidophilic and alkalophilic micro-organisms, one of his major stated intentions was to remind the reader of “what little is known about these organisms and their means of survival”. The past half decade has been much more active than almost all the time before with respect to interest and activity directed towards the isolation, characterization, and, particularly, the comprehension of organisms that grow at extremes of pH value. To some extent, the burst of interest has been part of a generally heightened awareness ADVANCES IN MICROBIAL PHYSIOLOGY, VOL. 24 ISBN 0-12027724-7

Copyrlght 0 1983 Academic Press London All rights of reproduction in any form re6em-d.

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of microbial ecology and of biological activity in unusual environments. In connection with this awareness, there exists the possibility that a much fuller understanding of acidophily and alkalophily would facilitate an exploitation of the relevant organisms for use in specific processes and fermentation settings. Or, it might become possible to decrease the activity of interfering acidophiles in particular situations. Apart from considerations of potential applications of knowledge gained, increased interest has been focused on bacteria that grow at extremes of pH value because of the bioenergetic issues that they raise or exaggerate. Concomitant with the growing acceptance of Mitchell’s chemiosmotic hypothesis (Mitchell, 1961, 1966, 1968), it became clear that both acidophiles and alkalophiles were intrinsically puzzling. According to Mitchell’s formulation, oxidations uiu the respiratory chain (or primary photosynthetic events) lead to an extrusion of protons out across the bacterial cell or chromatophore membrane (or mitochondrial/chloroplast membrane, in eukaryotes). The gradient of protons thus established comprises a transmembrane gradient of protons, ApH, acid outside and a transmembrane electrical charge gradient, A$, positive outside. The sum of these two gradients is an electrochemical gradient of protons, variously called the proton-motive force or A&+. The core of the chemiosmotic hypothesis is that the A,&+ is the energy form that is generated during respiration or photosynthesis and that, in turn, energizes a variety of membrane-associated processes. Among the proposed AiiH+-dependent processes are: ATP synthesis via proton-translocating (FIFo) adenosine triphosphatase (ATPase), many solute-transport systems, bacterial motility, transhydrogenase activity and reverse electron transport (e.g. see reviews by Greville, 1969; Harold, 1977; Hamilton, 1977). For each such process there is a proposed or presumed mechanism whereby energization can occur via utilization of the A / i H + . For example, inward translocation of protons occurs in conjunction with oxidative phosphorylation as well as transport of certain solutes. In consideration of chemiosmotic principles, specific problems were clear with respect to either acidophilic or alkalophilic organisms. As outlined by Garland (1977), acidophiles growing at external pH values of 2-3 would probably have to maintain extraordinarily large pH gradients in order to maintain reasonable cytoplasmic pH values. This would in turn raise questions with respect to orientation of the A$ (poised in the opposite direction, perhaps?), proton pumping and/or barrier functions, and the number of protons that would be inwardly translocated during bioenergetic work. By contrast, alkalophiles growing at external pH values of 10 to 11, would be expected to require a relatively more acidic cytoplasm. If primary proton pumping were in the usual direction, outward, how could a net concentrative uptake of protons be achieved? Having generated such a

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“reversed” chemical gradient of protons, would an enormous A$ or some other set of mechanisms offset this ApH for the purposes of bioenergetic work? The interests of the authors are largely concerned with these bioenergetic questions. This emphasis will be reflected in the review. We will also, however, endeavour to survey the organisms that have been characterized, and the range of unusual physiological features that have been found. Finally, we will echo Langworthy (1978) in noting how little is known about the precise survival mechanisms of both acidophiles and alkalophiIes, even five years later.

11. Acidophilic Bacteria A . SPECIAL PROBLEMS OF LIFE A T LOW

PH

VALUES

As already noted, the greatest problem with respect to life at low pH values is the maintenance of a cytoplasmicenvironment far less acidic than the external milieu. Acidophiles could accomplish this in two ways, firstly, by pumping protons outward particularly effectively and/or secondly, by possession of a cell-surface barrier extremely impermeable to protons. At a typical external pH value of 3.0, where acidophiles thrive, a ApH as high as 4.0 units might exist. As discussed on p. 178, the actual basis for maintenance of extremely large ApH values is quite controversial. Garland (1977) perceptively predicted that a counter-vailing A$ (positive inside) would be generated due to the relatively low capacitance of the cell membrane. His prediction has been verified in all the instances in which the A,&+ in acidophiles has been studied. The mechanism whereby a A$, inside positive, is generated has not yet been elucidated. It is not clear whether the charge gradient is purely a Donnan potential, established by impermeable charged macromolecules, or is actively maintained. A positive intracellular charge presents a problem for cation accumulation. In neutralophilic bacteria, for example, potassium is accumulated to concentrations as high as 500 mM in response to the A$, interior negative. Epstein’s group (Rhoads and Epstein, 1977,1978)has characterized several potassiumtransport systems in Escherichiu coli. Four separate systems have been found, some of which depend on the A$, while at least one system is driven directly by ATP (Epstein et al., 1978; Laimins et al., 1978). Although cation-uptake systems have not yet been explored in acidophiles, it would be more likely that ATP would be the driving force than the “reversed” A$, which would be counter-productively poised. Although other bioenergetic processes such as H+/solute cotransport and ATP synthesis would presumably take advantage

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of the large driving force available as the ApH, it would be important that the H +-conductors involved be sufficiently “tight” to preclude wholesale uptake of protons into the cytoplasm. Or, the membrane would have to be unusually impermeable to counter ions, thus preventing H + uptake. If the cytoplasmic pH value of acidophiles is maintained near neutrality, then no special problems would be expected in connection with cytoplasmic enzymes or synthetic processes. However, extracellular enzymes, flagella, and all processes associated with the external membrane surface would have to function at extremely acid pH values. Unusual properties of the cell wall and membrane layer would be anticipated.

B. ORGANISMS DESCRIBED

Many bacteria are able to grow at pH values as low as 4.0, but the vast majority of such organisms can also grow at neutral pH values. More unusual are those bacteria that thrive at pH values below 3.0 and cannot grow at neutral pH. We will concentrate on these obligately acidophilic species, which were reviewed by Langworthy (1978) several years ago. They fall into four distinct genera of prokaryotes, namely Thiobacillus, Bacillus, Sulfolobus and Thermoplasma. We briefly describe each in turn. Both Thiobacillus thiooxidans, isolated many years ago by Waksman and Jaffe (l922), and Thiobacillusferrooxidans, isolated by Temple and Colmer (1951), oxidize elemental sulphur with the production of sulphuric acid. The latter organism has also been shown to oxidize ferrous ion to ferric ion, accompanied by acid production (Dugan and Lundgren, 1965). Both thiobacilli grow optimally at pH values near 2.0. Coal-mine refuse piles, mine effluents and solfataras (acid soils in which sulphur has precipitated out) are the natural habitats of T.ferrooxidans and T. thiooxidans. Unlike other genera of acidophiles described, all of which are thermophiles, Thiobncillus spp. do not grow at high temperatures (above 55’C). Bacillus acidocaldarius, an aerobic spore-forming rod, was first characterized by Darland and Brock (1971). This bacterium grows between 45°C and 70°C (optimum 60°C) and pH values from 2.0 to 6.0 (optimum pH 3.0). The natural habitat is acidic hot springs. Both Belly and Brock (1974) and Uchino and Doi (1967) isolated acidophilic Bacillus coagulans species from hot springs, but Darland and Brock (1971) have grouped the most acidophilic bacilli under B. acidocaldarius. The two remaining primary genera of thermoacidophiles have features that set them apart from species of Thiobacillus and Bacillus, both of which are clearly eubacteria. Such features as their unique membrane lipids, the sequence of their 16s ribosomal RNA and their unusual or totally absent

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cell-wall structure have prompted Woese et al. (1978) to classify Sulfolobus acidocaldarius and Thermoplasma acidophilum as archaebacteria. Brock (1978) has very extensively described these two organisms in his highly readable book about thermophiles. We shall, therefore, only attempt to summarize the salient characteristics of each organism. Sulfolobus acidocaldarius was first isolated from hot sulphur acid springs (Brock et al., 1972). Brock points out that he has found S. acidocaldarius in such diverse areas as Yellowstone Park, Iceland and Italy. Others have isolated it from New Zealand (Bohlool, 1975)and Japan (Furuya et al., 1977). The name Sulfolobus is derived from the organism’s ability to oxidize elemental sulphur to sulphuric acid and its unusual lobed shape. Growth can be autotrophic (whereby carbon dioxide is fixed into organic compounds) or heterotrophic (whereby organic compounds are supplied in the medium). Like T.ferroxidans, S. acidocaldarius can oxidize ferrous ion (Brierly and Brierly, 1973). Growth is optimal at about 70°C and between pH 2.0 and 3.0. The cell wall does not contain the peptidoglycan characteristic of eubacteria, but appears to be a protein-lipid complex (Weiss, 1974). Pili-like structures appear to attach Suffolobus to crystals of sulphur (Weiss, 1973). Thermoplasma acidophilum was isolated from a coal-refuse pile by Darland et al. (1970). Growth was optimum between pH 1.0 and 2.0, and at a temperature of 59°C. Belly et al. (1973) have found Thermoplasma sp. only in self-heated coal refuse piles, and were not successful in isolating it from thermal springs (Brock, 1978). Primarily due to its lack of any cell wall, T. acidophilum has been classified as a Mycoplasma sp. Vancomycin, an inhibitor of peptidoglycan synthesis, had no effect on T. acidopilum, but novobiocin, an inhibitor ofmycoplasmas, killed the organism. The work of Christiansen et al. (1975) and Searcy and Doyle (1975) established the genome size of Thermoplasma sp. to be less than lo9daltons, the smallest DNA content ever reported for a non-parasite. Although lacking a cell wall, T. acidophilum is not particularly osmotically fragile. Unlike other mycoplasmas, it is stable in triple-distilled water (Belly and Brock, 1972). Such osmotic stability may be due to the peculiar membrane of this organism (see p. 184). When grown in culture, Thermoplasma sp. requires yeast extract (Smith et al., 1975). The active component appears to be a polypeptide of eight to ten amino-acid residues which, it is speculated, may help protect the organism from the high ambient concentration of protons. Other possible roles for the polypeptide may be to supply essential amino acids or to sequester ions to facilitate transport. Brock has speculated that coal-refuse piles probably contain low molecular-weight organic material derived from pyrolytic reactions on coal and complex organic molecules. A factor similar to that in yeast extract may thus exist in the natural habitat. In fact, Bohlool and Brock (1974) have shown that an extract of coal refuse could support growth of Thermoplasma

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acidophilum. Brock (1978) has pointed out that, although coal-refuse piles are man-made, similar natural habitats for T . acidophilum might have arisen by exposure of coal seams to the atmosphere through erosion or earthquakes.

C.

PHYSIOLOGICAL ADAPTATIONS THAT MEET THE PROBLEMS

1. The Transmembrane p H Gradient and its Usefor Bioenergetic Work All acidophilesmust maintain an intracellular environment far less acidic than the exterior. There is a good deal of controversy in the literature with respect to whether the transmembrane pH gradient, ApH, is actively or passively maintained in acidophiles. A survey of the results so far obtained, and the methods used to determine them, may help clarify the possible sources of discrepancies. Hsung and Haug (1975) reported an internal pH value of 6.4 to 6.9 for Thermoplasma acidophilum. Their conclusion was based on the following evidence: (I), cells of T. acidophilum, washed with distilled water and then sonicated, produced suspensions with pH values of 6.3 to 6.8; (2) a titration curve of cells with sodium hydroxide showed an inflection point between 6.5 and 6.9; (3) cytoplasmically derived malate dehydrogenase had a pH profile for activity with an optimum between pH 8.5 to 10.0; (4) the distribution of the weak acid, [14C]dimethyloxazolidine-2,4-dione (DMO), measured using a centrifugation assay, indicated an intracellular pH value near neutral. The same internal pH value was obtained in cells suspended at 56°C or 24°C at extracellular pH values of 2.0,4.0and 6.0. Hsung and Haug (1975) concluded that active metabolism is not necessary to maintain a ApH of as large as 4.5 units; boiling cells or treating with 100 p~ 2,4-dinitrophenol, 10 mM sodium azide or 10 mM iodoacetate had no effect on the internal pH value measured by DMO distribution. It was postulated that the pH gradient was primarily due to a Donnan potential across the membrane. Searcy (1976), on the other hand, either by titrating cells with sodium hydroxide or by rupture with a French pressure cell, concluded that the cellular pH value of 5.5 was not maintained after boiling the cells. What might explain such a discrepancy? Searcy (1976) speculated that DMO might be non-specificallyabsorbed by cells or somehow dissolved in the membrane lipids, thus leading to an overestimation of the internal pH value. Another explanation seems more plausible. As Cox et al. (1979) have pointed out, at an external pH value of 2.0, the DMO would be nearly 100% protonated because it has a pK value of 6.5. Therefore, the differencebetween a ApH of 4.5 units and no pH gradient is only a two-fold change in accumulation, values probably too similar to distinguish by the

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method employed. Until other acid probes of lower pK values are used in T. acidophilum, the question of whether the ApH is actively or passively maintained in this organism remains open. Further findings that T. acidophilum, unique among mycoplasmas, possesses menaquinones (Langworthy et al.,1972; Hollander et al., 1977), c-type cytochromes (Belly et al., 1973) and b-type cytochromes plus cytochrome d (az)oxidase (Hollander, 1978)indicate an array of respiratory components in this acidophile. Such a respiratory mechanism could play an important role in establishing the transmembrane pH gradient. Detailed studies of the role of respiration in the bioenergetic profile of this and other acidophiles have yet to be conducted. Thomas et al. (1976), using DMO and a fluorescent dye (fluorescein diacetate), found an internal pH value for Bacillus acidocaldarius of 5.6 to 5.8. In an organism similar to B. acidocaldarius,Yamazaki et al. (1973) also found an internal pH value of about 5.8. Oshima et al. (1977) showed that DNA extracted from B. acidocaldarius was rapidly decomposed by hot acid, indicating that the internal pH value is probably neutral. Moreover, a cytoplasmic enzyme, glyceraldehyde 3-phosphate dehydrogenase, exhibited optimal activity and stability near neutral pH values. The intracellular pH value, measured by DMO distribution, was approximately 6.0. However, accumulation of DMO was unchanged by the addition of uncouplers (such as carbonylcyanide p-trifluoromethoxylphenylhydrazone or carbonylcyanide m-chlorophenylhydrazone), gramicidin A, a i d e or cyanide, whereas carbonylcyanide p-trifluoromethoxylphenylhydrazone or gramicidin A did inhibit proline transport in B. acidocaldarius at acid external pH values. Again, DMO may have been an inappropriate probe. Oxygen uptake by cells of B.acidocaldarius at pH 3.4 and 50°C was inhibited by 10 mM azide. Krulwich et al. (1978) reported that the internal pH value of B. acidocaldarius at 50°C was 5.85 to 6.13 over an external pH range of 2.0 to 4.5. The intracellular pH value was measured in a flow dialysis assay by distribution of ['4C]acetylsalicylicacid. The pH optimum of B-galactosidase, an intracellular enzyme, was between 6.0 and 6.5, correlating well with the internal pH value determined by acetylsalicylic acid distribution. Moreover, 2,4-dinitrophenol and nigericin abolished the pH gradient. Transport of thiomethylgalactoside, coupled to H + uptake, was inhibited when the ApH was collapsed. Once again, there appears to be a discrepancy in the literature as to whether the ApH in an acidophile, B.acidocaldarius in this case, is actively or passively established. Perhaps the differences between Oshima et al. (1977) and Krulwich et al. (1978) are again due to the use of a weak acid with a pK near neutrality (the pK value of DMO is 6.5) by the former, whereas the latter group employed acetylsalicyclic acid (pK 3.5), a more sensitive indicator of ApH values in the acidic range in which B. acidocaldarius thrives. By lysing cells with sodium dodecyl sulphate, DeRosa et al. (1975)

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determined an internal pH value of 6.3 in Sulfolobus sp. growing at pH 3.0 outside at 70°C. Cooling the cell suspension at pH 3.0 to ambient temperature .or starvation for nutrients led to proton leakage in the Sulfolobus sp., resulting in death of the organism and acid-coagulation of the cytoplasm (J. D. Bu’Lock, personal communication). The same phenomena were observed when B. acidocaldarius was treated at pH 3.5 with 0.2 PM nigericin (Guffanti et al., 1979b). The effect of nigericin diminished with increasing external pH value. Acid-congealing of the cytoplasm in Sulfolobus acidocaldarius and Bacillus acidocaldarius on dissipation of the ApH, by cessation of metabolism in the former or exchange of H + for K + in the latter organism, strongly indicates that the pH gradient in these organisms is actively maintained and is not predominately a Donnan effect. In the fourth genus of obligate acidophiles, Beck’s (1960) demonstration that Thiobaciffusferroxidans can tolerate long periods at acid pH values under non-respiring conditions has led to the conclusion that the barrier to H is of a passive nature. Ingledew et a f .(1977) proposed that, at an external pH value of 2.0, T.ferrooxidans maintains an intracellular pH value near neutral by the internal removal of H + uia reduction of oxygen to water. Cox et al. (1979) reported a constant internal pH value of 6.5 in T. ferrooxidans cells over external pH values ranging from 1.0 to 8.0. Distribution of either [I4C]acetate or [3H]methylamine was followed using a filtration assay. Boiling the cells completely collapsed the ApH and A$ (see p. 183), but 2,4-dinitrophenol and 1 mM azide, known to inhibit respiration at pH 2.0 (Ingledew et al., 1977), did not significantly affect ApH, although the A&+ was nearly abolished because of a dramatic increase in the transmembrane electrical charge gradient (inside positive). Such results seem contradictory for, if 2,4-dinitrophenol is raising the internal positive charge by conducting protons inward, why did Cox et al. (1979) not see an effect on the ApH? One possible explanation might be a mechanism (antiporter?) whereby protons brought in by 2,4-dinitrophenol are rapidly exchanged for another cation. Such secondary ion movements in acidophiles remain to be elucidated. Matin’s group, working with Thiobacillus acidophilus a close relative of T. ferrooxidans (Ma0 et al., 1980; Martin et al., 1981), obtained similar results. Their ApH measurements were made with acetylsalicylic acid. Wilson and Matin (1982) showed that H+/2-deoxy-~-glucose symport (cotransport) in T. acidophilus was inhibited by protonophores and respiratory-chain inhibitors which did not, however, abolish the A,&+. Interestingly, the H+/solute ratio was consistent with a higher APH+ than was found. Moreover, when T. acidophilus was depleted of ATP (Zychlinsky and Matin, 1982) and treated with respiratory inhibitors (Matin et al., 1982), a substantial ApH was still measured at an external pH value of 3.0. It will be of interest to follow further studies of this puzzling system. Or particular importance will be studies of ion +

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fluxes, respiration and membrane permeability. The possibility that probes become trapped in the membrane or in some localized gradient (unstirred layer?) near the inner or outer membrane surface will have to be considered. As noted in several instances above, the ApH energizes several documented transport systems in obligate acidophiles via a H +/solute symport mechanism. Not surprisingly, ATP synthesis also utilizes the large ApH as a driving force. Apel et ul. (1980) have demonstrated ATP synthesis driven by an artificially imposed pH gradient (acid outside) in membrane vesicles of T. ferrooxiduns loaded with ADP and phosphate. The optimal internal pH value was 7.8 and the optimal external pH 2.8. Uncouplers (2,4-dinitrophenol and pentachlorophenol) completely prevented ATP synthesis. The ATPase was presumed to be similar to other bacterial FIFOenzymes in that ATP synthesis was Mg2+-dependent and strongly inhibited by N,N’-dicyclohexylcarbodiimide. Addition of valinomycin enhanced ATP synthesis, presumably by allowing K + efflux to balance the charge movement on proton influx through the ATPase. There is some evidence that, in environmental situations, T. ferrooxiduns utilizes a low external pH value that is established by another organism. Walsh and Mitchell (1972) showed that an acid-tolerant organism, Merullogonium sp., may lower the pH value in coal refuse so that T. ferroxiduns can grow. At pH 4.5 or above, abiotic iron oxidation proceeds, while at pH 3.5 or lower T. ferrooxiduns catalyses appreciable iron oxidation. Dugan and Apel (1978) postulate that ferrous sulphate is oxidized by T. ferrooxiduns as follows: Fe2++Fe3++eS-S2-

+ 302 +2H20+2(SO~~-)+16e- +4H+

(1)

(2)

Summing: FeS2+302+2H20+2HzS04+Fe3+

The Fe3+ can abiotically react to form more acid: Fe3++3H20+Fe(OH)3+3Hi Apel and Dugan (1978) have demonstrated that the external pH value of T. ferrooxiduns, suspended at pH 2.4, rose when ferrous sulphate was added. This is presumably due to an initial influx of protons. After an additional 20 minutes’ incubation, the pH value began to drop. The magnitude of H + uptake correlated directly with the ferrous ion concentration. Apel and Dugan (1978) have postulated that T. ferrooxiduns is an obligate acidophile because oxidation of Fe2+ to Fe3+ “results in the removal by the cell of one electron, and in order to maintain balance of electrical charge, a proton obtained from the cell’s environment may ultimately be utilized for the

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reduction of C02 and 0 2 ” . It was concluded that H + is a essential nutrient for T.ferrooxidans. Notwithstanding several very puzzling results in acidophiles that indicate a ‘‘passively’’-derived and/or maintained ApH, we are inclined to expect that substantial conformation to chemiosmotic principles will be found on more detailed investigation with, perhaps, new probes and methods for ApH measurements. Experiments to measure proton fluxes during, or secondary to, respiration by these bacteria are critical. Some proton-pumping mechanism should exist if bioenergetic work involves proton uptake. Also important with respect to the ApH are determinations of the passive proton (and other cation) permeability of acidophile membranes. 2. Transmembrane Electrical Charge Gradient, A$ Information about the transmembrane electrical charge gradient in acidophiles is just starting to accrue. As Garland (1977) predicted, in the face of a ApH across the membrane of the order of four pH units or more, the electrical charge gradient appears to be counterpoised. In those acidophileswhere it has been measured, the A$ value is positive inside relative to the external milieu. The mechanism for generation of such a gradient remains unknown. There is speculation that, in some instances, the internal positive charge may be due to charged macromolecules to which the membrane is impermeable. Such macromolecules have not yet been identified, although the histone-like protein associated with the DNA of Thermoplasma acidophilum (Searcy, 1975) is a candidate. Another possibility is that some unusual pattern of ion fluxes is responsiblefor the “reversed” A$. No relevant information on ion fluxes is yet available for acidophiles. It is known, however, that valinomycin-mediated K + efflux from B. acidocaldarius leads to a rapid secondary collapse of the ApH (Krulwich et al., 1978). In accord with their results for ApH measurements in Thermoplasma acidophilum, Hsung and Haug (1977a) have shown, by measuring accumulation of [I4C]thiocyanateions in a centrifugation assay, that the A$ value was approximately 120 mV, positive inside, at pH 2.0 and 56°C. As the external pH value was raised the A$ value decreased to only 10 mV at pH 6.0. Neither 10 mM sodium wide nor 1 mM 2,4-dinitrophenol had an effect on the transmembrane potential. Thus, the authors concluded that the A$ value is a passively derived Donnan potential of unknown origin. Hsung and Haug (1977b) also measured the 5 potential or the cell-surface potential of T. acidophilum, by microscopic electrophoresis. The potential was 8 mV, negative relative to the bulk medium. A negative surface charge was deduced from several lines of evidence: (1) cells migrated from the negative toward the

<

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positive electrode; (2) if the external pH value was raised above 6.0, the mobility increased, presumably by exposing more negative changes; (3) calcium ions (at concentrations of 1 mM and higher) lowered the electrophoretic mobility at pH 6.0, because these ions probably bind the negative surface charges; (4) the surface charge is probably due to the phosphate head groups of phospholipids; ( 5 ) at pH 2.0, where most binding sites are protonated, calcium ions had little effect on cell mobility. Hsung and Haug (1977b) present a model in which the total charge of the cell is negative, with the bulk gradient (A$) across the cytoplasmic membrane being positive inside and the surface highly negatively charged. Some indirect evidence is presented to support a Donnan potential as the source of the bulk charge gradient. Between external pH values of 7.0 and 10.5, the cells increase dramatically in mobility, possibly due to leakage of positively charged proteins (Ruwart and Haug, 1975). In Bacillus acidocaldarius,at an external pH value of 3.5,the A$ was at least 35 mV, positive inside, as determined by the uptake of [14C]thiocyanateions. Either 5 mM sodium thiocyanate or 2 p~ valinomycin (through K+ efflux) dissipated the A$ value and abolished the ApH value (Krulwich et al., 1978). Hence, the relatively positive interior appears to be necessary for maintenance of the large pH gradient. Furthermore, 20 mM tetraphenylboron (a lipophilic anion) dissipated the A$ while 10 PM carbonylcyanide m-chlorophenylhydrazone increased the A$, presumably by mediating proton influx. Cox el al. (1979) measured the A$ value of Thiobacillusferrooxidans cells, with either [14C]thiocyanateor [3H]dibenzyldimethylammonium(plus tetraphenylboron) in a filtration assay, over a range of pH values from 1.O to 8.0. At the lowest pH value, the A$ value was 70 mV (interior positive). The A$ decreased to 10 mV (interior positive) at pH 2.0 and rose to 50 mV (interior negative) at pH 3.0. The value increased to 170 mV (interior negative) at an external pH value of 8.0. Boiled cells did not exhibit any A$ value. As already noted, both 2,4-dinitrophenol and sodium azide had little effect on the ApH value (at pH 2.0 outside), but both inhibitors caused a large change in A$, increasing it from only 10 mV (interior positive) to values as high as 190 mV (interior positive). Thus, the A j H + value was lowered by either inhibitor from 256 mV to only approximately 55 mV. The authors concluded that the A$ value, but not the ApH value, in T. ferrooxidans was energy-dependent. In summary, the orientation of the A$ value in obligate acidophiles has been universally found to be positive inside. Moreover, the general range in the magnitude of the A+ value and its pH dependence do not generate appreciable controversy. However, the nature of the A$ value is not at all clear, be it macromolecular/Donnan or the result of active ion fluxes. At least in some species, ion movements can increase or decrease the observed A$ value and cause apparently secondary effects on the ApH value. Further studies of the A$ value in acidophiles will be of considerable interest.

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3. Recent Findings on Surface Properties of Acidophilic Bacteria

Langworthy (1977a,b, 1978) reviewed the large body of work on the cell-surface structures of acidophilic bacteria. We will, therefore, discuss only those features that have been elucidated since Langworthy’s survey. As will be evident, almost all the work has been focused on membrane lipids. Moreover, properties that might specifically relate to acidophily have not been clearly delineated. This is a particularly cogent problem since many of the species studied are thermophiles as well as acidophiles. The ability of concanavalin A to bind a-D-mannopyranosyl residues was employed by Mayberry-Carson et al. (1974, 1978) to detect cytochemically a unique membrane-associated lipopolysaccharide in Thermoplasma acidophilum. Binding of concanavalin A was inhibited by a-methyl-D-glucopyranoside. It was speculated that the lipopolysaccharide, like that of other mycoplasmas, may impart antigenic specificity (Sugiyama et al., 1974). Yang and Haug (1979a) demonstrated, by gel electrophoresis, 21 to 22 protein bands from the plasma membrane of Thermoplasma acidophilium. A major band that stained for carbohydrate with the periodic acid-SchiftTreagent had a molecular weight of 152,000. This glycoprotein accounted for 32% (w/w) of the total membrane proteins, and contained less than 10% (w/w) carbohydrate. The carbohydrate portion contained mostly a 1-2-linked mannose residues. Yang and Haug (1979a) speculated that the glycoprotein, in this wall-less organism, may provide a protective coat for the plasma membrane. They predicted that the branched pattern of the non-reducing ends of the glycoprotein may envelop the membrane. A correlation between the hydrophobic nature of the glycoprotein (62 mol % hydrophobic residues) and the survival of T. acidopilum at high temperatures and low pH values is plausible. The authors propose a model whereby the mesh-like carbohydrate coat may immobilize water molecules. Interactions between carbohydrates and water may further account for the well documented rigidity of the T. acidophilum membrane (Smith et al., 1974), a rigidity that may be an adaptation to both high temperature and low pH values. Several recent studies have employed fatty-acid spin probes and electron paramagnetic resonance (e.p.r.) to measure the membrane fluidity of T. acidophilum. Weller and Haug (1 977), using 2(3-carboxydecy1)4,4-dimethyl-2tridecyl-3-oxazolidinyloxylas the spin label, demonstrated that the temperature at which a low-temperature phase transition occurred in T. acidophilum membrane lipids was raised with increasingcalcium ion concentration, i.e. the lipid membrane fluiditydecreased. Vierstra and Haug (1978) also showed that A13+ions substantially increased the low-temperature lipid phase transition at pH values above 2.0; at or below pH 2.0, these ions had no effect presumably

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because the membrane phospholipids were not appreciably ionized at this pH value. Even in the presence of calcium chloride, at pH 4.0, A13+ ions still increased membrane rigidity. When T. acidophilum was grown at 37"C, rather than its optimum temperature of 56"C, it exhibited membrane lipids with 42% more pentane cyclization (Yang and Haug, 1979b). Electron paramagnetic resonance showed increased membrane lipid fluidity in 37°C-grown versus 56°C-grown cells, with the upper transition temperature downshifted 10°C to 35°C. The membrane-bound ATPase of T. acidophilum, a Mg2+-dependent enzyme, showed similar differences between cells grown at 37°C and 56°C; thus: (I) the optimum pH value for ATPase activity in 56'C-grown cells was 6.8 as opposed to 9.0 for 37°C-grown cells; (2) the ATPase activity in 5 6 ° C - g r ~ ~cells n showed &scontinui
15°C and 42.5"C, whereas the

activity in 37°C-grown cells showed discontinuities at 14°C and 35°C. It was concluded that the ATPase functions properly as long as the membrane is in a fluid state. Further e.p.r. studies by Strong and Haug (1980) demonstrated discontinuities at 22°C and 56°C in experiments with whole cells, membrane vesicles or dispersed lipids of T. acidophilum. Discontinuities at 22°C and 56°C presumably reflect the onset and end of the lipid-phase separation between gel and liquid. Between these two temperatures, the authors assumed that lipids exist in a mixed state. The large temperature range, 22°C to 56"C, may be due to the methyl-branched character of T. acidophilum membrane lipids, a characteristic that allows less co-operativity between lipids (Yang and Haug, 1979b). Additionally, a fluid discontinuity at 40°C was found only in whole cells, probably due to lipid-protein interactions. This corresponds well to the lower temperature limit for growth of 38°C. At the time of Langworthy's (1978) review, he stated that the precise structure of the glycerol diethers in Sulfolobus acidocaldarius was not firmly established. Recent results from Bu'Lock and DeRosa's groups have greatly expanded our knowledge of lipid structure in the Caldariella sp. group (a genus convergent with Sulfolobus (DeRosa et al., 1975; Millonig et al., 1975)). In addition to the diglycerol tetra-ethers, DeRosa et al. (1980a)have described a macrocyclic tetra-ether in which one of the hydrophobic units is a glycerol residue and the second is a calditol residue, a branched-chain nonitol unique to Caldariella sp. Moreover, the two 16,16'-biphytanyl chains bridging the glycerol and calditol residues can each contain up to four cyclopentane rings (DeRosa et al., 1980b). The degree of cyclization increases with growth temperature (DeRosa et al., 1980~). Cyclization raises the transition temperature (melting point), reflecting increased membrane rigidity. The authors compared membrane lipids of Caldariella acidophila with those of other archaebacteria. Species of Halobacterium, Methanosarcina and Methanococcus have diphytanyl glycerol diether, not biphytanyl lipids. Methanobacterium and Methanospirillurn species have both diphytanyl diethers and

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biphytanyl tetra-ethers; however, the tetra-ethers are only acyclic. Among the theromoacidophilic archaebacteria, namely species of Caldariella, Thermoplasma and Sulfolobus, only biphytanyl tetraethers are found. The groups of Bu’Lock and DeRosa (DeRosa et al., 1980d) have postulated that the tetra-ethers of Caldariella sp. are synthesized from di-0-(geranylgerany1)glycerol diether. Direct reduction would result in diphytanylglycerol diether, whereas competition between complete reduction and 16,16-coupling would result in a mixture of di- and tetra-ether lipids typical of some methanogens. Coupling without reduction would, along with cyclization, result in the tetra-ethers typical of Caldariella sp. DeRosa et al. (1977) presented evidence that C. acidophila has neither menaquinones nor ubiquinones, but possesses a unique benzo(b)thiophen-4,7-quinone, which they named “caldariellaquinone”. This terpenoid quinone is structurally analogous to the menaquinones and is believed to play a respiratory function in this acidophile. Langworthy (1978) pointed out that the long branched hydrocarbon chains of lipids in species of Thermoplasma and Sulfolobus may be an adaptation to high temperature, allowing proper membrane fluidity. The unique ether linkages may confer some stability to acid hydrolysis, although this needs to be explored further. Unfortunately, no new evidence has been elucidated since Langworthy (1 978) concluded that “it appears difficult to relate any features of cell wall structure or membrane composition to acid stability in the thiobacilli”. There has been some conjecture that the o-cyclohexyl and branched-chain fatty acids of B. acidoculdurius help maintain membrane fluidity at high temperatures (Langworthy, 1978). It has been proposed (Langworthy et al., 1976) that the acid-resistant C-S bond of the sulphonolipid found in B. acidocaldarius may play some role in proton exclusion. Little has been done since Langworthy’s (1978) review to characterize the cell wall of B. acidocaldarius, a structure that Langworthy predicted may have unusual acid-resistant features. The flagella of acidophiles remain to be studied. It is known that the unusually shaped pili of Sulfolobus sp. are very acid- and heat-stable (Weiss, 1973). Another, as yet unexplored, but potentially, fruitful topic is the production, stability and activity of extracellular enzymes produced by acidophiles.

D . W H Y CAN’T OBLIGATE ACIDOPHILES GROW AT NEUTRAL p~ VALUES?

One of the fundamental problems of the physiology of acidophiles is their inability to grow at neutral pH values. We wondered whether this might be due to lack of proton/cation antiporters which, by their action in neutralo-

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philes (Beck and Rosen, 1979; Brey et al., 1978) and alkalophiles (Mandel et al., 1980), can convert a ApH into a A$? We have attempted to grow Bacillus acidocaldarius at neutral pH values in the presence of low concentrations of monensin, an ionophore that will exchange protons for sodium (Haney and Hoehn, 1967), or nigericin, a K + / H +exchanger (Harold and Papineau, 1972). No growth was detected, indicating either that monensin and nigericin did not function properly as antiporters or, more likely, that the inability of B. acidocaldarius to grow at pH 7.0 has some other or additional basis. Smith et al. (1973) speculated that the protonic environment of Thermoplasma acidophilum results in membrane carboxyl groups being uncharged. Raising the pH value to neutral may cause ionization of carboxyl groups and repulsion of negative charges, resulting in membrane disruption and cell lysis. Treatment of membranes with glycine methylester stabilized them at pH 7.0, presumably by removing negative charges from free carboxyl groups. An analogy has been drawn to Halobacterium sp. in which N a + , rather than H+ , neutralizes negative membrane charges (Brown, 1964; Kushner, 1968). In contrast to the results of Smith et al. (1973) with Thermoplasma acidopilum, Guffanti et al. (1979b) failed to demonstrate loss of viability when Bacillus acidocaldarius was suspended at pH 6.0 for 10 minutes at 50°C. Thus, inability of acidophiles to grow at neutral pH values may depend on other unusual characteristics of these organisms. Some possibilities include: (1) their unique membrane lipids; (2) the pH-dependent characteristics of membrane-bound porters, enzymes or respiratory chain components; (3) peculiar properties of the membrane-bound proton-translocating ATPase; (4) proton pumping and other ion-flux mechanisms which become counter-productive at neutral pH values. Exploration of these and other structural/functional properties of acidophiles has just begun.

111. Alkalophilic Bacteria A . S P E C I A L P R O B L E M S OF LIFE A T HIGH pH V A L U E S

For extreme, obligately alkalophilic bacteria, the optimal external pH value for growth on most substrates is between pH 10.0 and 11.0. Growth at even higher pH values is also observed. The most obvious and pressing physiological problem is regulation of cytoplasmic pH value. It is clear, a priori, that optimal external pH values would not be permissible cytoplasmic pH values since RNA would be unstable and it is unlikely that protein synthesis could occur. Thus, there has to be an effective organelle or other protective mechanism for alkali-labile cellular processes, or the cytoplasmic pH value

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must be acidic relative to the external milieu. If, as is most likely, the cytoplasm is more acidic than the outside medium, then a mechanism for the net inward translocation of protons must exist. As will be described on p. 192, at least some alkalophileshave been shown to extrude protons into the medium during respiration, i.e. they generate a conventional ApH, acid out, as a primary event. However, there exist porters whose activity, secondary to primary proton pumping, results in net proton accumulation in the cytoplasm. The physiological necessity of a “reversed” ApH-a chemical gradient of protons, acid in-raises an additional problem of great importance. That is, do the organisms generate enormously high transmembrane electrical potentials, A$ values, so that the total A&+ value reaches normal levels? Or, do the organisms have special mechanisms for carrying out bioenergeticwork such as ATP synthesis, motility and solute transport under conditions in which the total A P H + is rather low? In the former case, the mechanism for A$ generation would be extremely interesting; in particular, the structural/functional properties of the respiratory chain might be unusual. In the latter instance, the mechanisms for bioenergetic work at low ApH+ values would involve intriguing adaptations. Of course, it is quite possible that alkalophiles possess somewhat high A$ values, and some special capacity to generate them, as well as adaptations of bioenergetic processes to allow them to function at relatively kow A&+ values. In connection with ATP synthesis, the possible problem of a low driving force is further exacerbated by the requirement for somewhat higher energy at alkaline pH values simply because of the high pH-related increase in the energy of hydrolysis of the terminal phosphate bond of ATP (Rosing and Slater, 1972). Maintenance of a relatively acidic cytoplasm, i.e. nearer to neutral pH value than the highly alkaline pH value of the medium, might bypass any need for unusual pH optima or stabilities for cytoplasmic enzymes or macromolecules. However, extracellular enzymes, as well as all activities and structures that face the medium would have to be stable and functional at pH 10 to 11. It is possible, therefore, that the properties (lipid composition?) of the membrane might be unusual, and that the organisms might have special cell-wall layers or macromolecules. Moreover, the flagellar proteins and extracellular enzymes would be expected to differ from those found in neutralophiles with respect to structural properties related to pH optimum and stability.

B . ORGANISMS DESCRIBED

1. DeJinitions

For the purposes of our own investigative work, as well as this review, we

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distinguish between alkaline-tolerant bacteria and bacteria that are obligate alkalophiles. Alkaline-tolerant organisms grow well at pH 8.5 to 9.0, generally do not grow above pH 9.5, but grow at neutral pH values. By contrast, obligate alkalophiles generally cannot grow below pH 8.5, do not grow at neutral pH values but often exhibit optimal growth at pH 10.0 or above. Clearly the qualitative and quantitative problems facing these two groups are reasonably distinct. Nonetheless, great care is not always taken in characterizing new alkalophilic strains. The most important, and most often neglected problem, is whether the pH value at which growth commences corresponds to the initial pH value of the medium. Very few characterizations of alkalophiles have been conducted using chemostat conditions in which a constant pH value is maintained. In the batch-culture conditions usually employed, it is essential that the pH value be monitored during growth so that the actual pH during the initial phase of growth is known; this is not always done. A second concern is that alkalophiles, like any other bacteria, can exhibit somewhat different pH profiles on different growth substrates. For example, Bacillus alcalophilus, which was described originally by Vedder (1934), has a more alkaline pH optimum as well as a higher minimum pH values for growth on L-malate than for growth on lactose (Guffanti et af., 1978, 1979a). 2. Bacillus species Alkaline-tolerant species of Bacillus are probably quite common. In 1959, Kushner and Lisson (1959) described a strain of Bacillus cereus which had “developed” the capacity to grow at high pH values after repeated transfer at gradually increasing pH values. In the absence of specific information about the pH value at which growth commenced, the organism appears to be alkaline-tolerant rather than obligately alkalophilic. Similar alkaline tolerance is evident in several strains of Bacillus circulans (Chislett and Kushner, 1961; Guffanti et al., 1979c) and Bacillusfirmus (Guffanti et al., 1980). The ureaclastic species Bacillus pasteurii is an alkalophilic organism which may lie somewhere between the alkaline-tolerant species and the more extreme obligate alkalophiles. Bacilluspasteurii requires ammonium ions and alkaline pH values (8.0 or higher) for growth (Gibson, 1934). Wiley and Stokes (1963) suggested that the alkaline pH values allowed conversion of ammonium ions to free ammonia which, in turn, played a role in substrate uptake. Although the original description indicated that B. pasteurii grows well up to pH 11.0, the strains obtained from the American Type Culture Collection showed in our hands only modest growth above pH 9.5 (A. A. Guffanti and T. A. Krulwich, unpublished observations). Among the clearly obligately alkalophilic bacteria, Bacillus species are the

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most common. In addition to Vedder’s (1934) B. alcalophilus, several obligately alkalophilic Bacillus species have been described by Horikoshi (1971a,b, 1972) and Horikoshi and Atsukawa (1973a,b) as well as other workers (e.g. Boyer and Ingle, 1972; Ando et al., 1981a) whose major initial interest was in alkaline-stable extracellular enzymes. Horikoshi and Akiba (1982) have recently reviewed their work and that of many other groups. Although some alkalophilic bacteria have been isolated from specialized environments such as indigo dye balls or alkaline springs, many obligate alkalophiles are paradoxically found in conventional soil, e.g. Bacillus A-007 (Ando et al., 1981b) and Bacillusjirmus RAB (Guffanti et al., 1980). These latter species grow poorly below pH 9.0, and grow well up to pH 11. The presence of obligate alkalophiles in a variety of soil environments might relate to the alkaline clay particle content of such soils. 3. Other Non-Photosynthetic Bacteria In addition to Bacillus species, several Gram-positive alkalophiles in other genera have been described. Akiba and Horikoshi (1976) reported finding an alkalophilic strain of Micrococcus, and Souza et al. (1974) isolated an alkalophilic Clostridium sp. from an alkaline spring. Souza and Deal (1977) described an aerobic, rod-shaped, motile, non-sporulating alkalophile that was isolated from a high-pH value spring; they suggested that the strain, designated A-1, was a coryneform bacterium. Subsequently, Gee et al. (1980) isolated five alkalophilic strains from potato-processing effluent. These strains possessed an orange cellular pigment and resembled, in many respects, the A- 1 alkalophile. The few reports of Gram-negative non-photosynthetic alkalophiles include a Pseudomonas species (Hale, 1977), a Flavobacterium sp. (Souza et al., 1974), and a red-pigmented species of Halobacterium (Tindall et al., 1980). The alkalophilic Halobacterium sp. was isolated from the alkaline-soda lakes and ponds at Lake Magadi in Kenya; the organism required alkaline pH values, high osmolarity, as well as particularly high concentration of magnesium ions. 4. Photosynthetic Organisms

Alkaline tolerance appears to be common among the blue-green algae, the cyanobacteria. Indeed, most cyanobacteria have pH optima for growth that are in the alkaline range (Fogg, 1956). This preference has been the basis for certain physiological studies of Microcystis aeruginosa (McLachlan and Gorham, 1962) and Synechococcus sp. (Kallas and Castenholz, 1982a,b).

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These organisms grow in the neutral range of pH values and are not clearly found to grow at extremely alkaline pH values. They are therefore alkalinetolerant rather than obligately alkalophilic. Grant et al. (1979) isolated a species of Ectothiorhodospira from an alkaline mud sample from Lake Hannington in Kenya. This photosynthetic bacterium grew in a pH range from 7.5 to 11.0, with optimal growth at pH values from 9.0 to 9.5. The internal membrane and many physiological characteristics, except for the pH profile, were typical of other members of the genus.

C . PHYSIOLOGICAL ADAPTATIONS TO MEET THE PROBLEMS

1. Regulation of Cytoplasmic pH Value Two alkaline-tolerant strains, one of Bacillus circulans and one of B.Jirmus, have been characterized with respect to their electrochemical proton gradient, A ~ H +as, a function of the external pH value. Both strains exhibited a conventional ApH, acid out, at an external pH value of about 6.5 (Guffanti et al., 1979c, 1980). As the external pH value was increased, the magnitude of the ApH value decreased until, at pH 8.0 to 9.0, the value was zero. Neither alkaline-tolerant strain exhibited an ability to acidify the cytoplasm relative to the external medium. In these experiments, the ApH was assayed by following accumulation of the weak acid DMO, or the weak base methylamine, using the flow dialysis assay of Ramos et al. (1976). Using different techniques,cells of Escherichia coli have been found to maintain a cytoplasmic pH value that is slightly more acidic than the external milieu at media pH values from 8.0 to 9.0 (Padan et al., 1976; Slonczewski et al., 1981). Obligately alkalophilic bacilli, by contrast, maintain very large gradients of pH value in the reverse of the usual direction, i.e. acid in, over a broad range of alkaline pH values. Studies of the alkaline-tolerant and obligately alkalophilic bacilli suggest that the highest cytoplasmic pH value that is compatible with viability is pH 9.0 to 9.5 (Krulwich et al., 1981). For alkaline-tolerant species, this corresponds to the maximum pH value for growth, at which the cytoplasmic pH value is equal to the external value, ApH = O (Guffanti et al., 1979b, 1980). Obligate alkalophiles continue to maintain their cytoplasmic pH value at pH 9.5 or below when the external value is raised as high as pH 1 1 to 11.5. Bacillus alcalophilus grown on L-malate thus exhibits ApH values of almost 2.5 units (equivalent to + 151 mV) at pH 11.5; at its optimum pH value for growth (PH 10.5) the cytoplasmic pH value was about 9.5, representing +60 mV (Guffanti et al., 1978). Lactose-grown B. alcalophilus also generated ApH values, acid in, although the cytoplasmic and optimal growth pH values were lower than observed for growth on L-malate (Guffanti et al., 1979a).

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Lactose-grown cells maintained a cytoplasmic pH value within the range of 7.3 to 9.2 over a range of external pH value from 7.0 to 11.0. Another obligately alkalophilic species, B. firmus RAB, showed a ApH pattern quite similar to that found in B. alcalophilus grown on L-malate (Guffanti et al., 1980). Again, the method used in all the above studies was assay of methylamine accumulation. Using a very different method, namely inhibition of oxygen uptake by azide as a function of pH value, Hoddinott et al. (1978) found that B. pasteurii maintained its cytoplasmicpH value at 9.5 or below up to an external pH value of 10.0; whether their method is of comparable reliability with other methods is unclear. However, all of the obligate or near obligate alkalophiles examined can maintain a relatively acidified cytoplasm. These findings are consistent with the observation that pH optima for cytoplasmic enzymes generally fall in a conventional, broad, neutral-to-low alkaline range (Ohta et al., 1975; Ando et al., 1981a). Comparative studies of obligate alkalophiles and their non-alkalophilic mutant derivatives, using both whole cells and isolated membrane vesicles, have strongly implicated an electrogenic Na+/H+ antiporter in the vital physiological role of pH homoeostasis. It appears that subsequent to primary pumping of protons out of the cells during respiration, net proton accumulation is achieved by exchange of protons for Na+. Non-alkalophilic mutants were readily isolated from both B. alcalophilus (Krulwich er al., 1979) and B. firmus RAB (Guffanti et al., 1980)by plating suspensions of mutagenized cells on L-malate-containing media at pH 7.0. The mutants thus obtained had acquired the ability to grow from pH 5.5 to 8.5, but had lost the ability to grow above pH 9.0. Although, as described on p. 198, non-alkalophilic strains exhibit certain pleiotropic changes, the mutation and reversion patterns and frequencies are most consistent with a single mutation (Lewis et al., 1982). Starved 22Na+-loadedcells of the two wild-type strains, B. alcalophilus and B. firmus RAB, exhibit energy-dependent valinomycin-sensitive 22Na+efflux; their non-alkalophilic mutant derivatives, strains KM23 and RABN, do not (Krulwich et al., 1979; Guffanti er al., 1980).Cells of the two non-alkalophilic strains also fail to exhibit methylamine accumulation (acidification of the cytoplasm relative to the medium) at any pH value in a range from 5.5 to 10.5. Right side-out membrane vesicles have been prepared from B. alcalophilus and B.firmus RAB by a slight modification of the lysozyme method of Kaback (1971). At pH 9.0, vesicles from B. alcalophilus generate a conventional ApH value, acid out, on energization, if both sodium and potassium ions are omitted from the buffer (Mandel et al., 1980). Vesicles from B. firmus RAB generate a ApH value, acid out, as long as Na+ is omitted from the buffer (Krulwich et al., 1982). Thus, in the absence of Na+ (and K + for B. alcalophilus), vesicles from alkalophiles pump protons in the usual outward direction. For both species, addition of Na+ to the vesicle suspension resulted

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in establishment of a “reversed” ApH value, acid in. Only Na+ allowed generation of a reversed ApH value; K + and other cations did not (Mandel et al., 1980; Krulwich et al., 1982). The concentration of Na+ required to produce net accumulation of protons on energization was much higher for B. firmus RAB than for B. alcalophilus. The apparently lower affinity of the porter for Na+ correlated with a strong dependence of growth of B. jirmus RAB, but not B. alcalophilus, on added Na+ (Krulwich et al., 1982).This is an important point, since the absence of a growth requirement for added Na+ can accordingly reflect operation of high-affinity Na+-coupled porters rather than the irrelevance of the porters to the physiology of the organism. Importantly, right side-out vesicles from non-alkalophilic strains failed to accumulate protons on the addition of Na+ (Mandel el al., 1980; Krulwich et al., 1982). Sodium and hydrogen ion movements have been further studied using everted membrane vesicles from B. alcalophilus and its non-alkalophilic mutant derivative. Energy-dependent valinomycin-sensitive accumulation of 22Na+,and Na+-dependent extrusion of protons, are demonstrable in everted vesicles from the wild type but not the non-alkalophilic mutant. Taken together, studies of Na+ and H + movements in cells and vesicles indicate that obligately alkalophilic bacilli pump protons out of the cell during respiration as do other bacterial species. A secondary Na+/H+antiporter then catalyses inward movement of protons in exchange for Na+ such that the following gradients are established in cells or right side-out vesicles: Na+out > Na+in and H+in> H+out. Non-alkalophilic mutants lack the antiporter and cannot maintain a relatively acidified cytoplasm and, hence, have lost the ability to grow above pH 9.0. Antiporters specific for Na+/H+ have been described in numerous tissues and organisms. In E. coli, this antiporter has been proposed to have a role in pH value homoeostasis that is qualitatively, if not quantitatively, similar to that found in the alkalophiles (Schuldiner and Fishkes, 1978; Zilberstein et al., 1980). Considerable evidence indicates that the Na+/H+ antiporter of B. alcalophilus is indeed a secondary antiporter, and is not directly energized by ATP. Harold and his associates (Heefner and Harold, 1980; Heefner et al., 1980) have implicated ATP in direct energization of Na+ fluxes in a Streptococcus faecalis. Everted vesicles from B. alcalophilus accumulate 22Na+ when energized by ATP, but only when the ATP is hydrolysed and can generate a AjiH+ (Guffanti, 1982). There is also evidence that B. alcalophilus, but not B. jirmus RAB, possesses an electroneutral K+/H+ antiporter (Mandel ef al., 1980).No role for this activity has been demonstrated in this species. In E. coli, a K+/H+ antiporter has been suggested to play a role in pH homoeostasis (Brey et al., 1980; Plack and Rosen, 1980), albeit not a dominant role (Slonczewski et al., 1981).

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2. The Magnitude of the A$ and Total A&+ Values

An assessment of the magnitude of the transmembrane electrical potential, and the total electrochemical proton gradient APH+, is quite problematic. These measurements are extremely important. If, in the presence of a “reversed” ApH value, the A&+ value is still quite high (e.g. greater than - 150 mV), then there would be few special problems with respect to bioenergetic work but very fascinating questions about generation of such high A$ values. Alternatively, the A$ values may be in a usual or only somewhat high range, opening issues both with respect to generation of A$ values and the performance of bioenergetic work at low A&+ values. Unfortunately, the methods employed are, necessarily, indirect. Although some attempt has been made to make direct electrophysiological measurements on giant bacterial cells (Felle et al., 1980),such methods are not likely to be generally applicable. The most commonly employed indirect methods include use of potential sensitive fluorescent probes, distribution of K + or s6Rb+ions in the presence of valinomycin, and distribution of radiolabelled lipophilic cations such as triphenylmethylphosphonium (TPMP+), tetraphenylphosphonium (TTP+) and dibenzyldimethylammonium (DDA+)ions (Maloney et al., 1975). Use of lipophilic cations has developed from the pioneering efforts of Skulachev (1971) and the application of probes to bacterial-cell and vesicle systems by many investigators. In general, the A$ values measured by TPMP+ or TPP+ distributions are in close agreement with theoretical values for valinomycin-induceddiffusion potentials or values obtained by other methods up to A$ values of - 125 to - 150 mV; at higher A$ values, the lipophilic cations do not seem to measure the full A$ value (Schuldiner and Kaback, 1975; Kashket et al., 1980; Shioi et al., 1980; Guffanti et al., 1981a). Thus, if alkalophile A$ values are very high, they may be underestimated by these probes. Moreover, some whole cell measurements with Bacillus species may be subject to A$-unrelated effects with use of lipophilic cations (Zaritsky et al., 1981). The A$ values of alkaline-tolerant Bacillus species have been examined in whole cells using the distribution of TPMP+ or TPP+. In both B. circulans (Guffanti et ai., 1979c) and B. jirmus (Guffanti et al., 1980), the A$ was approximately -66 to - 77 mV at pH 6.5 and increased to - 1 15to - 137mV at pH 9.0. The total A P H + value was relatively stable over the pH range (because of the decreasing ApH, acid out), at - 1 15 mV for B. circulans and between - 145to - 170 mV in B.firmus. These are not dramatically low A&+ values. Similarly, using DDA+ in the presence of tetraphenylboron, Hoddinott et al. (1978) measured sufficiently high A$ values in cells of B. pasteurii to maintain A,&+ values close to - 180 mV over a range of pH up to 10.0.

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Thus, in those cells, a very high A+ value may largely compensate for the reversed ApH value up to pH 10.0. It should be noted, however, that the B. pasteurii system is not as highly alkalophilic as some species, is complicated by the presence of ammonium ions under growth conditions, has only been assessed with respect to the A+ values using methods that require the problematic presence of tetraphenylboron, and relies on an unusual method for ApH value determinations. In cells of the obligately alkalophilic B. alculophilus grown on L-malate, the A$ values, as measured from distribution of TPMP+, increased from -84 mV at pH 9.0 to - 152mV at pH 11.5(Guffanti et ul., 1978).These substantial increases did not, however, offset the progressively reversed ApH value. Therefore, the total AjiH+ value at pH 10.5and 11, where growth is abundant, was determined to be only - 80 mV and - 15 mV, respectively. Similarly, the A$ value measured using TPP+ in ceIIs of B.Jirmus RAB grown on L-malate increased from - 89 mV at pH 8.0 to - 145 mV at pH 11.0 (Guffanti et al., 1980).Nevertheless, the A j i H + value declined to about - 50 mV in the optimal highly alkaline range of pH values. Unless the A$ values are grossly underestimated, the low AjiH + values pose a considerable bioenegetic problem. Moreover, consideration must be given to how the A$ value, positive out, is generated at all, under circumstances of net proton accumulation. As shown in Fig. 1, with entirely hypothetical stoicheiometry, the operation of outward proton pumping via respiration followed by an electrogenic Na+/H+antiport could produce both a ApH value, acid in, and a A$ value, positive out. In Fig. 1, the circulation of Na+ is shown to be completed by inward movement of Na+ in symport (co-transport) with solutes.

Respiration

-

Antiporter

Not /Solute Syrnporter

1

\

FIG. 1. A model for hydrogen and sodium ion fluxes across alkalophile membranes showing hypothetical stoicheiometry.

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TERRY A. KRULWICH AND ARTHUR A. GUFFANTI

Studies on the generation of A$ and A,&+ values in obligate alkalophiles + although the possibility of certainly suggest that there is a low A ~ H problem, an under-estimation of the A$ value exists. Assays of A$ and A j H + values have also been conducted on energized right side-out membrane vesicles from B. alcalophilus. On energization with ascorbate/NNN’N’-tetramethyl-pphenylenediamine, Na+-loaded vesicles prepared from cells grown on L-malate generated a A$ value that increased from - 125 mV to - 135 mV over a range of external pH values from 8.0 to 11.5 (Mandel et al., 1980). It is interesting that the A$ values in vesicles exhibit a less dramatic increase with increasing pH value than observed in cells. Importantly, however, the range in which these A$ values fall are within the area of greatest confidence with respect to the lipophilic cationic probes. Thus, when bioenergetic studies are conducted using such preparations, and A$ values by several methods are in agreement, it is likely that the A$ values are at least a reasonable approximation of the full A$ value (Guffanti et al., 1981b). 3. Solute Transport and Motility Solutes are actively transported into bacterial cells by three main mechanisms, namely (1) phosphoenolpyruvate: hexose phosphotransferase systems, which are group-translocation processes that translocate certain hexoses, primarily in facultative or anaerobic species, (2) transport systems that directly utilize ATP or some other derivative thereof, and (3) A$- or &,+-dependent transport systems in which neutral or anionic solutes are co-transported with a cation. The last type of transport mechanism is probably the most commonly employed, and the vast majority of such “symport” systems in bacteria employ protons as the coupling ion (West and Mitchell, 1972; Harold, 1977; Kaback et al., 1977). Thus a problem is raised in alkalophilic bacteria, with generally low A&+ values and ApH value equal to zero, or even reversed ten-fold or more with respect to H+in/H+out.A H+/solute symport would apriori seem an unlikely transport mechanism to function efficiently in such organisms. Indeed, studies to date indicate two general ways in which alkalophiles bypass the A ~ H problem + for purposes of solute transport. First, certain carbohydrates are transported by some mechanism that employs ATP (or a metabolite thereof) directly, without the A$ or A j H + as intermediary. In lactose-grown cells of both an alkaline-tolerant strain of B. circulans (Guffanti et al., 1979c) and B. alcalophilus (Guffanti et al., 1979a), transport of 8-galactosidessuch as the non-metabolizable analogue thiomethyl 8-D-galactopyranoside correlated with cellular ATP levels and did not correlate with the A$ values. Similar observations have been made with respect to 2-deoxyglu-

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cose transport by glucose-grown cells of B.jirmus RAB (A. A. Guffanti and T. A. Krulwich, unpublished observations). The second, and more important, way in which the low A,&+ value problem is bypassed for purposes of solute transport by alkalophilic bacteria, is the use of Na+/solute symport rather than H+/solute symport. Use of sodium as coupling ion takes advantage of the A$ values, which tend to be high, and makes the reversed chemical gradient of protons irrelevant. Whereas Na+/solute symporters are commonplace in eukaryotic tissues that possess Na+ K+-activated ATPase activity (Crane, 1977), they are employed sparingly by conventional bacteria (see e.g. Stock and Roseman, 1971; Tokuda and Kaback, 1977; Lopilato et al., 1978). Only bacteria that require or thrive in environments containing high concentrations of sodium ions have previously been found to use these ions as the major or sole coupling ion for solute symport (Lanyi et al., 1978; Lanyi, 1979; Thompson and MacLeod, 1973). Several different investigators have now reported, however, the prevelance of Na+/solute symporters in alkalophiles. There has also been a report of a Na+/Ca2+antiporter in Bacillus A-007, whose role is presumably in calcium ion efflux (Ando et al., 1981b). Sodium ion-dependent transport of the amino acid analogue a-aminoisobutyric acid has been demonstrated in whole cells of four different alkalophilic bacilli (Koyama et al., 1976; Kitada and Horikoshi, 1977; Guffanti et al., 1978, 1980); in most of these systems, other cations, including lithium, could not substitute for sodium. Transport was optimal at highly alkaline pH values. Kitada and Horikoshi (1977) showed that increasing concentrations of sodium ions, within an appropriate range, caused a decrease in the K , value for transport of a-aminoisobutyrate without an effect on the V,,, value. Guffanti et al. (1978) showed that transport of a-aminoisobutyrate was an electrogenic process, i.e. that the solutes transported bear a net positive charge. Accordingly, transport was inhibited by compounds that dissipated the A$ value. Kitada and Horikoshi (1979) demonstrated Na+-dependent uptake of glutamate by whole cells of the alkalophile Bacillus No. 8-1, and Na+-dependent transport of 13 amino acids (including a-aminoisobutyrate and glutamate) in membrane vesicles from that species (Kitada and Horikoshi, 1980). Transport of a-aminoisobutyrate, L-malate and three different amino acids was also shown to be Na+-dependent in membrane vesicles of B. alcalophilus (Guffanti et al., 1981b,c) and additional amino acids, whose transport was not demonstrable in vesicles, were found to be transported in a Na+-dependent manner in whole cells (A. A. Guffanti and T. A. Krulwich, unpublished observations). Studies of uptake of a-aminoisobutyrate by energized membrane vesicles from both B. alcalophilus (Bonner et al., 1982)and B.firmus RAB (Krulwich et al., 1982) showed that the concentration of sodium ions affected the K,,,value for transport of a-aminoisobutyrate rather than the V,,, value. Passive efflux of

+

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L-malate, L-aspartate and a-aminoisobutyrate from unenergized membrane vesicles from B. alcalophilus has also been shown to be Na+-dependent (Guffanti et al., 1981~).Methods for studying solute efflux and exchange, developed by Kaback and his colleagues (Kaczorowski and Kaback, 1979; Kaczorowski et al., 1979; Cohn and Kaback, 1980), were applied to B. alcalophilus vesicles to elucidate further mechanistic aspects of solute translocation. The rate of efflux of a-aminoisobutyrate from unenergized vesicles is equal to, or slightly greater than, the rate of exchange; therefore, “return of the unloaded carrier’ or its real mechanistic equivalent is not the rate-limiting step. This was also the result with Na+/melibiose symport, but not H+/lactose symport, in E. coli (Kaczorowski and Kaback, 1979; Cohn and Kaback, 1980). Both efflux and exchange of a-aminoisobutyrate from vesicles of B. alcalophilus are markedly inhibited by generation of a A,&+ value across the membrane (Bonner et al., 1982).This suggests that the ternary complex between Na+, a-aminoisobutyrate and porter bears a positive charge. As expected for the alkalophile, the rates of efflux of a-aminoisobutyrate and exchange at pH 9.0 are greater than or equal to those at pH 7.0; very low rates are observed at pH 5.5. Most interesting has been the findings with respect to solute transport by non-alkalophilic mutant derivatives of B. alcalophilus and B.firmus RAB. On isolation of strains KM23 and RABN, the respective non-alkalophilic mutants, we found the expected absence of Na+/H+ antiport activity in both whole cells and vesicles (Krulwich et al., 1979;Mandel el al., 1980;Guffanti et al., 1980; Krulwich et al., 1982). Less expected was the absence of Na+-coupled transport of a-aminoisobutyrate which was also found in both cells and vesicles (Krulwich et al., 1979, 1982; Guffanti et al., 1980; Bonner et al., 1982).An inability to mediate Na+-coupled uptake of a-aminoisobutyrate could, of course, be secondary to loss of the antiporter, since that loss would prevent normal efflux of sodium ions. However: (a) in both cells and vesicles, the very initial rates of a-aminoisobutyrate uptake were affected and Na+-coupling for other solutes was pleiotropically lost (Krulwich et al., 1979; Guffanti et al., 1981~);(b) the non-alkalophilic mutant of B. alcalophilus, strain KM23, exhibited uptake and efflux of a-aminoisobutyrate at pH 7.0, but now in apparent symport with protons (Krulwich et al., 1979; Bonner et al., 1982); (c) whereas passive efflux of a-aminoisobutyrate, L-malate and L-aspartate were all markedly Na+-dependent in wild-type B. alcalophilus, they were Na+-independent in strain KM23 (Guffanti et al., 1981~);(d) various concentrations of monensin (whose ability to catalyse Na+/H+ exchange under the given experimental conditions was confirmed) failed to restore Na+/solute symport or Na+-dependent efflux. Many independently isolated non-alkalophilic strains all exhibit the same properties, and the wild-type phenotype is completely restored on reversion to alkalophily (Lewis

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et al., 1982). Moreover, several lines of evidence indicate that the Na+-coupled porters undergo a change in coupling ion to protons on mutational loss of the Na+/H+ antiporter and loss of the alkalophilic phenotype, but that the H+-coupled porters that now function are not a set of distinct porters that are simply repressed in the wild type (Bonner et al., 1982). For example, mutations of the wild type in a specific porter are still expressed on further mutation of the porter-minus strain to non-alkalophily. We have interpreted all of these observations to indicate a direct relationship between the Na+/H+ antiporter and Na+/solute symporters in certain obligately alkalophilic bacteria. A working hypothesis, and only one possibility, is that these Na+-translocatingporters share a common sequence or subunit (Krulwich et al., 1979; Guffanti et al., 1981b). A model depicting this hypothesis is shown in Fig. 2. We speculate that the porters possess the latent capacity for coupling solute transport to protons, a capacity that is suppressed in the wild type Na+-coupled state. On mutational loss of the Na+-translocating element, the Na+/H+ antiport activity is lost, and the porters are now H+-coupled, functioning at neutral pH values at which + feasible. This model is consistent with the data energization by a A ~ H is already summarized, with additional data on solute efflux and exchange in non-alkalophilic mutants (Bonner et al., 1982), and with the observation that a greater Na+ affinity for Na+/solute systems in B. alcalophilus versus B. firmus RAB correlates with the greater Na+ affinity of their respective

NO+

minoisobutyrate

Alkolophill

Non-olkolophllr

FIG. 2. A model for ion-translocatingactivitiesin the membranes of alkalophilesand their non-alkalophilic mutant derivatives.The clover-leaf shaped sodium ion-translocating subunit of Na+/H+antiporter and Na+/solutesymporters are shown to be lost on mutation to non-alkalophily. The non-alkalophile accordingly retains the porters which are now coupled to hydrogen-ion translocation. The respiratory chain of the alkalophile is shown as a more effectiveproton pump than that of the non-alkalophilic mutant.

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Na+/H+ antiporters (Krulwich et al., 1982). A very similar model has been proposed for Na+-translocating porters in E. coli by Padan and Schuldiner and their associates (Zilberstein et al., 1980;Schuldiner and Padan, 1982).The model is still, however, only one possible explanation for the data obtained by both groups of investigators. For example, the mutation that results in pleiotropic loss of sodium ion-translocating functions could be in a regulatory protein or an enzyme involved in covalent modification of porters. Future structural and genetic studies may clarify the precise nature of the mutation. An interesting, but only possibly relevant, observation with respect to alkalophiles has been the finding of a membrane-associated chromophore (absorbance peak at 526 nm in absolute oxidized-versus-water absorbance spectra; Lewis et al., 1982; Krulwich et al., 1982). This chromophore is absent from non-alkalophilic mutants; these mutants, on the other hand, invariably produce a soluble brown pigment in the growth medium by the end of the growth cycle. Concentrations of the chromophore are unaffected by light or dark conditions. With respect to another bioenergetic process, namely motility, obligate alkalophilesappear to employ the same solution to the low APH+ value as used for most solute systems. Motility in E. coli, B. subtilis and other bacteria is coupled to the Aj&+ value (Larsen et al., 1974; Ordal and Goldman, 1975). For B. subtilis, there is a threshold for the AiiH+ value of at least - 30 mV for motility (Khan and MacNab, 1980).There is now a growing body of evidence that a “sodium-motiveforce” (A&+) could be the driving force for motility in obligately alkalophilic bacteria. At least some obligate alkalophiles are highly motile at very alkaline pH values (e.g. Ohta et al., 1975), at which the A j i ~ +is quite low. Hirota et al. (1981) found that Na+ must be present in the medium in order to allow motility of alkalophilic Bacillus sp. YN-1 and No. 8-1. Agents which abolished the A# value completely inhibited motility, even in the presence of Na+. These findings are quite exciting because the adaptation involved in utilizing Na+ rather than H for energization of the flagellar rotor should be of great interest, and influence on any general hypotheses with respect to the energization of motility. Hirota et al. (1981) did not, however, rule out the slight possibility that the Na+ was required to generate the A# value rather than for flagellar motion per se. In studies in our laboratory (Kitada et al., 1982),cells of B.Jirmus RAB have been rapidly washed with and suspended in potassium carbonate buffer, pH 9.0. Under these conditions, they retain a A# value of about - 140 mV for at least 10 minutes and this A# value is essentially unaffected by addition of 10 mM sodium chloride. Motility, however, is only observed on addition of Na+. Moreover, in preliminary experiments,imposition of a sudden gradient of Na+ (Na+,,* >Na+in)did not result in motility in the absence of a A# value (valinomycin-treated cells in potassium carbonate buffer). Thus, a sodium-motiveforce, consisting of Na+ +

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and a A$ value, may indeed be involved in energizing motility in obligately alkalophilic bacteria.

4. Synthesis of A TP via Oxidative Phosphorylation

If oxidative phosphorylation occurs via a strictly chemiosmoticmechanism in which two protons are translocated inwardly through the ATPase (BFIF~) for each molecule of ATP synthesized,then a A ~ H value + of approximately -200 mV would be required to achieve [ATP]/[ADP] ratios of unity when the concentration of inorganic phosphate is 10 mM (Greville, 1969). Unless the A&+ measurements made on alkalophilic cells and vesicles are very gross underestimations, the A ~ +Hvalues of obligate alkalophiles fall greatly below the required force for ATP synthesis. This apparent discrepancy between the AjH+ value observed, and the required driving force, would be irrelevant if the organisms possessed either special organelles in which ATP synthesis occurs or employed a synthase which did not depend on A j H + values. Neither of these two solutions appears to obtain. Fine structural studies do not indicate the presence of intracytoplasmic organelles that could be the true site of ATP synthesis (Krulwich, 1982). Nor was any Na+-dependence found for either ATP synthesis by vesicles loaded with ADP and inorganic phosphate from B. alcalophilus (Guffanti et al., 1981b) or for ATP hydrolysis by everted vesicles (A. A. Guffanti, unpublished observations). Thus it appeared that the problem of a low AjH+ value is not bypassed for purposes of ATP synthesis by utilization of a sodium-motive force as is the case for solute transport and motility. When it became clear, however, that Na+-dependent processes in B.firmus RAB generally required higher concentrations of sodium ions than corresponding processes in B. alcalophilus, it seemed worthwhile to re-examine the possibility of Na+-dependent ATP synthesis in the former species. Right side-out membrane vesicles from B. jirmus RAB grown on L-malate were prepared so that the intravesicular space was loaded with 5 mM ADP (potassium salt) and 10 mM inorganic phosphate. The vesicles were energized by addition of ascorbate/phenazine methosulphate in the presence or absence of 50 mM Na+ (as carbonate salt at pH 9.0 and phosphate salt at pH 7.0). Initial rates of ATP synthesis were independent of sodium ion concentration at both pH 7.0 and pH 9.0 (Fig. 3). Moreover, as had been found with a similar vesicle system from B. alcalophilus (Guffanti el al., 1981b), ATP synthesis was sensitive to A"'-dicyclohexylcarbodiimide and conditions (e.g. valinomycin, uncouplers at pH 7.0 and nigericin) that dissipated the A j H + value. Thus, a A,&+ value appears to be used for ATP synthesis by alkalophiles. This conclusion is supported by several pieces of evidence in addition to the

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TERRY A. KRULWICH AND ARTHUR A. GUFFANTI

6.0 c

a

% c

0

-

4.0

c

+

2 +

5 V

A

0 .-

2.0

c

s

A - 0 0

1

2

3

A

4 5 0 1 Time (minutes)

A

A

A-

2

3

4

5

FIG. 3. The effect of inhibitors and sodium ions on the synthesis of ATP by membrane vesicles of Bacillus firmus RAB. Right side-out membrane vesicles of Bacillus firmus RAB were prepared in either 100 mM potassium carbonate, 10 mM magnesium sulphate, and 10 mM potassium phosphate (pH 9.0), or 100 mM potassium phosphate and 10 mM magnesium sulphate (PH7.0). The vesicles were loaded with 5 mM ADP (potassium salt) as described previously (Guffanti el al., 1981b). For assays in the presence of sodium the basic buffer contained 50 mM potassium carbonate together with 50 mM sodium carbonate @H 9.0; b) or 50 mM potassium phosphate together with 50 mM sodium phosphate (pH 7.0; a). Vesicles were added to 1.0 mg protein ml-I and incubated at 30°C under a constant stream of water-saturated oxygen. When present, 10p~ valinomycin or 500 p~ NN’-dicyclohexylcarbodiimidewere added to vesicles ten minutes before initiation of the reaction. Oxidative phosphorylation was initiated by addition of 20 mM potassium ascorbate together with 0.1 mM phenazine methosulphate. Samples were removed to cold perchloric acid and assayed for ATP by the luciferin-luciferase method (Stanley and Williams, 1969). 0 indicates behaviour of suspensions supplemented with sodium ions, 0 not supplemented with sodium ions, A supplemented with NN’dicyclohexylcarbodiimide,and A supplemented with valinomycin; 0 indicates behaviour of suspensions in which energy was not generated.

absence of a Na+ effect and the inhibitory effects of NN’-dicyclohexylcarbodiimide and dissipators of the A j H + value. First, Koyama et al. (1980) purified the BFI polypeptide from an ATPase of an alkalophilic Bacillus sp. and found conventional structural/functional properties. Secondly, Guffanti et a f . (1 98 1 b) demonstrated inward, NN’-dicyclohexylcarbodiimide-inhibitable,

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203

proton movements concomitant with ATP synthesis by vesicles from E . alcalophilus loaded with ADP and inorganic phosphate. If, then, a proton-translocating BF, FOpolypeptide complex catalyses ATP synthesis in alkalophiles, what are the dimensions of the bioenergetic problem, and what are the possible solutions? For ATP synthesisby vesicles of B. alcalophilus loaded with ADP and inorganic phosphate, phosphorylation potentials (AGp=AGo+RT In [ATP]/[ADP] [Pill) were calculated to be 11 and 12 kcal mol-' (50.4 kJ mol-I) at pH 10.5 and 9.0, respectively, whereas the A j i ~ +values in vesicles at these two pH values were quite different (-40k20 mV at pH 10.5 and - 125+20 mV at pH 9.0; Guffanti et al., 1981b). The mV equivalents of the AGp values are - 23 1 mV at pH 10.5 and -252 mV at pH 9.0 assuming a H+/ATP ratio of two; the values may be doubled to assess mV equivalents without stoicheiometric assumptions. Clearly, the discrepancy between the ATP synthesized, as reflected by the AGp values, and the AjiH+ is very great, especially at pH 10.5, the optimal pH value for growth. Two possible solutions, not mutually exclusive, may be considered, in addition to the re-iterated caveat that gross misestimations of the A&+ values or the AGp values would account for the apparent problem. Instead of two or three protons being translocated for each molecule of ATP, alkalophiles might translocate more protons for each ATP, e.g. four at pH 9.0 and eight at pH 10.5, to account for the vesicle data. Alternatively, the bulk + might be less directly transmembrane gradients (measured as the A j i ~ value) coupled to ATP synthesis than some microscopic or more localized gradient which could be larger than the A&+ value (Williams, 1978; Gould and Cramer, 1977; Rottenberg, 1979; Kell, 1979). Importantly, the alkalophiles represent an exaggeration of a major current bioenergetic problem that goes way beyond this specialized group of organisms. Indeed, the exaggerated experimental system that they offer for study of this and other fundamental bioenergetic issues represents a valuable investigative opportunity.

5 . The Respiratory Chain Several aspects of alkalophile physiology that have been part of all the foregoing discussion lead to the expectation that the respiratory chain of these organisms might be of particular interest. First, the A@ values generated by obligate alkalophiles, even during operation of the energy-consuming Na+/H+ antiporter, are fairly high. Secondly, both the energy cost of the maintenance of a relatively acidified cytoplasm and the somewhat greater energy required to make ATP at high pH values (Rosing and Slater, 1972) represent unusual energy demands, and yet obligate alkalophiles grow very well. Stouthamer (1969) conducted major studies of bacterial growth as a

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function of energy generation for non-alkalophilic bacteria using molar growth yields as one of the parameters. The obligate alkalophiles, B. alcalophilus and B. jirmus RAB, exhibit molar growth yields on L-malate (Ymal)that are comparable with growth of bacteria under optimal conditions; the values are 42 and 39 g dry wt. (mol malate)-' for the two species, respectively, at pH 10.5(R. J. Lewis, unpublished observations). Notably, the Y,,, for the non-alkalophilic derivatives of the two alkalophiles, measured at the growth optimum pH value of 7.0, is no more than half that for the parental strains. This greatly decreased growth yield could reflect the compromised function of some physiological process that is adapted to perform and/or be expressed optimally at highly alkaline pH values. The respiratory chain and ATP synthase are both excellent candidates for such a process. It was therefore particularly interesting to note the intensely red colour of membranes isolated from the obligate alkalophiles. Moreover, the non-alkalophilic mutant derivatives of B. alcalophilus and B. firmus RAB exhibited a yellowish membrane colour by comparison. Indeed, membranes of the two alkalophilic species contain extraordinarily high contents of cytochrome haem, at least 5.5 nmol (mg membrane protein)-' (Lewis et al., 1980). Membranes from the non-alkalophilic strains KM23 and RABN contain much lower concentrations of haem, with especially low levels of b- and c-type cytochromes. Interestingly, the respiratory rates of whole cells of the two alkalophiles and their non-alkalophilic mutants are comparable, at their respective pH optima, with one another and with respiratory rates of other bacteria. A characterization of the respiratory chain components of B. alcalophilus, in comparison with derivative non-alkalophile strain KM23, has been conducted (Lewis et al., 1981). The alkalophile was found to have, both absolutely and relatively, a large number of distinguishable redox components. Cytochromes with the following mid-point potentials can be identified in membrane vesicles from B. alcalophilus when titrations were conducted at pH 9.0 (a typical cytoplasmic pH value of whole cells): cytochromes a and a3, + 240 mV; cytochrome b (or spectrally similar species), + 20 mV, - 120 mV, - 240 mV and - 320 mV; and cytochrome c, +70 mV. One of the a-type cytochromes and at least one of the b-type cytochromes exhibit a pH-dependent midpoint potential, which could be a property associated with proton pumping. Vesicles from B. alcalophilus also contain an unusually low potential Reiske iron-sulphur protein (e.p.r. signal at g 1.90) as well as numerous iron-sulphur clusters. Evidence from studies of mitochrondria (Trumpower and Edwards, 1979; Trumpower et al., 1980) and other bacteria (Bowyer et al., 1979,1980)supports a role for the Rieske protein as an immediate electron donor to cytochrome c. The isopotential midpoint potentials of the B. alcalophilus Rieske protein and cytochrome c are

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consistent with such a function and, importantly, that coupling would be lost, because of potential shifts, in alkalophile membranes at pH 7.0 (Lewis et al., 1981). In vesicles from non-alkalophilic strain KM23 at pH 7.0 (a typical cytoplasmic pH value for this strain), fewer redox species are found: a cytochrome a, +lo0 mV; one cytochrome b species, -120 mV; and a cytochrome c, + 140 mV. The Rieske protein was not clearly distinguished although the two strains contained comparable amounts of other iron-sulphur clusters as well as ubiquinone. The finding of ubiquinone in a Bacillus species is unusual; bacilli have been reported to have only menaquinone (Hess et al., 1979). However, conditions apparently used by Hess et al. (1979) to grow B. alcalophilus may not allow growth of this species. In addition to the quantitatively and qualitatively high complement of respiratory-chain components, alkalophile membranes (as well as cells) have respiratory rates that are similar to those of non-alkalophiles (Lewis et al., 1980), and alkalophilic organisms have normal growth yields on L-malate. Taken together, those observations and the high energy cost of life at high pH values, suggest that the respiratory chain of the alkalophiles may function particularly well with respect to energy conservation. In the model in Fig. 2 (p. 199), we indicated this speculation by showing the alkalophilic respiratory chain as a longer chain which might pump more protons per oxygen atom or malate consumed than that of the non-alkalophile. Jones et al. (1975, 1977) suggested that the efficiency of respiration-linked proton translocation (H +/0 ratios) may vary greatly from bacterium to bacterium depending on the specific respiratory-chain components found. Clearly, any chain that may exhibit particularly high H +/0ratios will be of special bioenergetic interest. Some preliminary data with respect to alkalophiles are encouraging (e.g. Krulwich et al., 1982)and further studies are under way. In the same context, the respiratory-chain complexes of the alkalophiles, especially complex I11 (the ‘‘k~”, complex) should be quite interesting. Finally, with respect to the respiratory chain, the quantitative and qualitative decrease in the number of redox species on mutation to non-alkalophily is dramatic. These decreases, and the likely concomitant loss of at least one energy-coupling site, could account for the poorer growth yield. But how does a single mutation, probably in the Na+/H+ antiporter, cause the observed effects on the respiratory chain? One of several possible hypotheses is that mutational loss of the antiporter alters the cytoplasmicpH profile in such a way as to affect expression of the genetic information (or membrane assembly) of the cytochromes. Changes in cytochrome levels on shifts in growth pH value in B. alcalophilus (lower cytochrome levels at lower pH values) and strain KM23 (elevated cytochrome levels at higher pH values) lend some support to this hypothesis (Lewis et al., 1982).

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6. Alkaline Stability of Surface Structures and Extracellular Enzymes By contrast with the extensive work conducted on membrane lipids of acidophilic bacteria, this area of important work is virtually void in connection with alkalophilic bacteria. Clearly, the lipid composition, cell-wall structure and flagellin structure will all be of interest, but have not yet been investigated. Some limited fine-structure studies of B. alcalophilus and B. firmus RAB suggest that these organisms may possess complex cell-wall layers (Krulwich, 1982), but nothing is known about the possible composition or role of such layers. On the other hand, production and properties of extracellular enzymes by alkalophilic bacteria have been extensively studied, especially by Horikoshi and his colleagues (Horikoshi, 1971a,b, 1972; Horikoshi and Atsukawa, 1973a,b; Kitada and Horikoshi, 1976a,b). In a large series of reports, these investigators demonstrated that obligately alkalophilic bacteria produce a spectrum of extracellular enzymes and products, including proteinases and pectinases (also found by Kelly and Fogarty, 1978). Production of extracellular enzymes,is markedly dependent on the growth pH value, with optimal enzyme production in the highly alkaline range. Similarly, both the stability and optimum for activity of the extracellular enzymes from alkalophiles are found to be highest at quite alkaline pH values, often as high as pH 10 or 11 (e.g. Kitada and Horikoshi, 1976a,b).

D. WHY CAN’T OBLIGATE ALKALOPHILES G R O W AT NEUTRAL pH

VALUES?

The characteristics of non-alkalophilic mutants of alkalophiles (Krulwich et al., 1979; Guffanti et al., 1980) and the apparent activity of the Na+/H+ antiporter in membrane vesicles at pH 8.0 (Mandel ef al., 1980) led to the proposal that obligately alkalophilic bacteria cannot grow at pH 7.0 because continued operation of the Na+/H+ antiporter results in lethal acidification of the cytoplasm (Krulwich et al., 1979). Recent evidence from two different alkalophiles suggests that this proposal is in error. First, Kallas and Castenholz (1982a,b) studied a variety of bioenergetic properties of the cyanobacterium Synechococcus sp. during growth in a range of pH values as well as on shift of actively growing cells from pH 8 or above to pH 5-6. Under both conditions, they found no impairment in cytoplasmic pH homoeostasis, i.e. there is no pronounced acidification of the internal pH value. The ATP/(ADP+ATP) ratio similarly did not seem to be a relevant factor in the poor growth at, or lack of sustained growth on transfer to, low pH values. The

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authors proposed that some membrane-associated function, perhaps solute transport, might be the neutral-labile process. Recent studies have been conducted on B. jirmus RAB, the obligate alkalophile with a requirement for at least 10 mM Na+ for growth, Na+/solute symport and operation of the Na+/H+ antiporter (Krulwich et al., 1982). Cells were grown at pH 10.5 and then transferred to non-nutrient or L-malate-containing buffers, at different pH values, containing either Na+ or K + . As expected, on transfer to buffer at pH 10.5 lacking Na+, cellsrapidly lost viability, exhibiting a rapid rise in cytoplasmic pH value. By contrast, after transfer to pH 7.0, the loss in viability was far less pronounced (although no growth occurred in nutrient media at pH 7.0). Strikingly, viability was somewhat better, even in non-nutrient media, in the presence of Na+. Acidification of the cytoplasm was not observed and, indeed, the data indicate that the Na+/H+ antiporter largely ceased to function (Kitada et al., 1982). The capacity of the cells to transport a-aminoisobutyrate is greatly diminished at pH 7.0, and this may well be secondary to a marked decrease in the All/ value. Quite possibly, the primary impediment to growth at neutral pH values may be a poor proton-pumping capacity (and hence low All/) of the alkalophile respiratory chain at this pH value. A sodium-coupled cell would be highly dependent on the All/ value even though a ApH value, acid out, is now produced. Non-alkalophilic mutants, now proton-coupled for transport, might just manage using the total ApH+ value. This very speculative scenario, however, is far from proved.

IV. Concluding Remarks Enormous progress has been made toward an understanding of the special physiological problems and adaptations of acidophilic and alkalophilic bacteria. Nonetheless, almost nothing at all is known about the intermediary metabolism and regulation of storage products of these organisms. Nor have developmental studies, e.g. of sporulation of the many acidophilic and alkalophilic bacilli, been reported. No genetics are yet available; indeed there have been no genetic applications of lysogenic phages, no mapping, and no cloning efforts. At neither pH extreme is it clear what precludes growth at neutral pH values, although this is a fundamental part of any general understanding of the problems. Membrane lipids of the acidophiles have been extensively examined, but other cell-surface layers are not well characterized. If motile species exist, their motility and flagellar properties have not been reported. Moreover, the perplexing problem of the ApH value, the nature of the A+ value, inside

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positive, the properties of the respiratory chain (proton pumping?) and the mechanisms of bioenergetic work by acidophilic bacteria are in the very initial stages of exploration. By contrast, nothing is known about either the membrane lipids or other surface layers of alkalophiles, while many more bioenergetic details have been elucidated. Even in the last area, fundamental questions with respect to structure-function of the respiratory chain, ATPase and Na+-coupled porters remain. It is particularly important to repeat, in closing, that in addition to their intrinsic ecological and industrial importance, bacteria that grow at extremes of pH value offer particularly exaggerated and stressed experimental systems in which to study bioenergetic problems of general importance.

V. Acknowledgements The unpublished studies described in this review were conducted with the support of research grants PCM78 10213 from the National Science Foundation and GM28454 from the National Institutes of Health, and of contract DE-AC02-81ER-10871 from the Department of Energy. We are grateful to our colleagues Richard Lewis and Makio Kitada for sharing their ideas and their preliminary results with us.

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Weiss, R. L. (1974). Journal of Bacteriology 118, 275. Weller, H. and Haug, A. (1977). Journal of General Microbiology 99,379. West, I. C. and Mitchell, P. (1972). Journal of Bioenergetics 3, 445. Wiley, W. R. and Stokes, J. L. (1963). Journal of Bacteriology 86, 1152. Williams, R . J. P. (1978). Biochimica et Biophysica Acta 505, 1. Wilson, B. and Matin, A. (1982). American Society for Microbiology 82nd Annual Meeting Abstracts, p. 139. Woese, C. R., Magrum, L. J. and Fox, G. E. (1978). Journal of Molecular Evolution 11, 245. Yamazaki, Y., Koyama, N. and Nosoh, Y. (1973). Biochimica et Biophysica Acta 314, 257. Yang, L. L. and Haug, A. (1979a). Biochimica et Biophysica Acta 556, 265. Yang, L. L. and Haug, A. (1979b). Biochimica et Biophysica Acta 573, 308. Zaritsky, A., Kihara, M. and MacNab, R. M. (1981). Journalof Membrane Biology63, 215. Zilberstein, D., Padan, E. and Schuldiner, S. (1980). Federation of European Biochemical Societies Letters 116, 177. Zychlinsky, E. and Matin, A. (1982). American Society for Microbiology 82nd Annual Meeting Abstracts, p. 139.

Note Added in Proof Matin et al. (1982) have proposed that, in Thiobacillus acidophilum, the failure of either uncouplers alone or bioenergetic work to collapse completely the ApH may result from a rapid buildup of the AY, inside positive, after just a small percentage of the protons move inward. The force thus established would impede further proton influx (thus retaining a APH) as long as the membrane did not allow efflux of some other, compensatory, cation. The recent discussion by Ingeldew (1982) similarly points towards this kind of explanation of the “uncoupier-insensitive” ApH of acidophilic thiobacilli. This possibility, however, leaves the nature of initial “reversed” BY still unknown. Moreover, a special membrane impermeability to cations may obtain in some acidophiles without being necessary for obligate acidophily. The ApH value of Bacillus acidocaldarius is completely abolished by uncouplers, and only a modest increase in the AY (inside positive) is concomitantly observed. Perhaps the membrane of that species allows efflux of some cation(s) during rapid proton influx. In our laboratory, we have recently shown that isolated membrane vesicles from B. acidocaldarius generate a conventionally oriented A,iiH+, with AY, outside positive, on addition of an electron donor at either pH 6.0 or 3.0. Importantly, the proton permeability, protonophore sensitivity and ionophore sensitivity of the membranes appear to be similar to those of non-acidophiles. Ingeldew, W. J. (1982). Biochimica et Biophysica Acta 683, 89.

Metabolism of One-Carbon Compounds Chemotrophic Anaerobes J. G . ZEIKUS Department of Bacteriology, University of Wisconsin, Madison, Wisconsin 53706, U.S.A. I. Introduction . . . . . . . . . . . A. Definitions . . . . . . . . . . . . . B. History and scope . . . . . . . . 11. Transformation of one-carbon metabolites by anaerobes . . . . A. Production of one-carbon compounds . . . B. Consumption of one-carbon compounds . . . . . 111. One-carbon transformations in methanogens . . . . . A. General physiology and species properties . . . . . . . B. Catabolism . . . . . . . . C. Cell carbon synthesis. . . . . . . . . D. Unification and regulation of metabolism . . . . . . . . . IV. One-carbon transformations in homo-acetogens . A. General physiology and species properties . . . . . . . B. One-carbonmetabolism . . . . . . V. General metabolic perspectives on unicarbonotrophy . . . . A. Relation of substrate-product thermodynamics to growth efficiency . B. Relation of chemotrophic anaerobes to phototrophs and aerobes . VI. Trends in the significance of one-carbon transformations . . . A. Environmental . . . . . . . . . . B. Evolutionary . . . . . . . . . . C. Biotechnological . . . . . . . . . VII. Acknowledgementsand dedication . . . . . . . References . . . . . . . . . . .

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I. Introduction Understanding the cellular properties and significance of micro-organisms that grow on one-carbon substrates as sole sources of carbon and energy has ADVANCES IN MICROBIAL PHYSIOLOGY, ISBN 0-12-027724-7 VOL. 24

Copyright 01983 Academic Press London All righrs of reproducrion in any form reserved.

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been a most interesting topic in metabolic biology during the past 15years. By and large, the general impression gathered from reading the past literature is that Cl-metabolizing species possess novel physiologies that are distinctive in many ways from the autotrophic and heterotrophic micro-organisms which seemingly are more widespread in the microbial world. Notwithstanding, both aerobic and anaerobic microbes can proliferate on the same one-carbon substrates; however, general metabolic concepts which unify the microbiology, physiology and biochemistry of CI metabolism as a whole have been limited to comparisons in aerobic microbes. The aim of the present review is to integrate what is known about the metabolism of Cl compounds by anaerobes into a common conceptual picture. Hence, the specific metabolism of one-carbon compounds by aerobic micro-organisms, which has been well reviewed elsewhere, will not be addressed here except to gain a general comparative metabolic perspective to that occurring in anaerobic bacteria.

A . DEFINITIONS

CI compounds refer to any oxidizable one-carbon substrate that contains carbon-bound electrons. A CI substrate differs noticeably from the C1 compound carbon dioxide because, in addition to being able to be reduced or assimilated, it can also be oxidized and, hence, can provide electrons for use in energy metabolism or cell synthesis. The term chemotrophic anaerobe refers to any obligately non-oxygen-catabolizingmicrobial species whose growth is solely dependent on the generation of metabolic energy from chemical substrates. The definition of anaerobiosis sensu strictu, that is, fermentation or life without air, is applied here. Thus, the term “anaerobic respiration” will not be used because respiration itself implies life with air (i.e. oxygen catabolizing). Different kinds of fermentations will be identified by the major types of reduced end-products formed via CI metabolism, and not on the basis of electron acceptors. Consequently, sulphidogenic fermentation is employed in lieu of “sulphate respiration”, and acetogenic or methanogenic fermentation instead of “carbon dioxide respiration”. A methanogen is a microorganism that produces methane as the major reduced end-product of cellular metabolism. This kind of designation is also useful because it eliminates confusion in microbiology. For example, the term “methane bacteria” does not allow a distinction to be drawn between CI-metabolizing species which consume or produce methane. Within a very diverse anaerobic group such as the acidogens, subdivisionssuch as the homo-acetogens are clearly recognized to be of prime importance. Homo-acetogens are unique because these species form acetyl-CoA as a consequence of CI metabolism, regardless of whether the substrate is glucose or methanol, and whether the product is acetic or

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butyric acid. However, homo-acetogens in analogy with homolactic acid bacteria can produce acetic acid as the only detectable end product when grown under the appropriate cultural conditions. For the sake of comparing nutritional distinctivenessamong CI-metabolizing chemotrophic anaerobes, the following terms are utilized: autotrophy, growth with carbon dioxide as the sole carbon source metabolized; heterotrophy, growth with substrates containing carbon-carbon bonds as carbon sources for growth; merhylotrophy, growth with R-CH3 as the sole carbon source where R is either not metabolized or is not linked to the methyl group by a carbon-carbon bond; unicarbonotrophy, growth with one-carbon compounds as the sole carbon and energy source. Unicarbonotrophy is specifically useful in eliminating confusion because this term can apply to species that grow on either hydrogen/carbon dioxide or methanol, irrespective of whether the former is capable of methylotrophy or the latter of autotrophy. Likewise, unicarbonotrophs can include species that do not grow autotrophically or methylotrophically, but are capable of proliferating with carbon monoxide or formate as the sole carbon and energy source. B. HISTORY AND SCOPE

The origins of CI metabolism in anaerobes began in the 1940s with the discovery of methanogenic fermentation of methanol, when Methanosarcina barkeri was isolated and described by Schnellen (1947). Notably, this species also formed methane from hydrogen/carbon dioxide mixtures, acetate or carbon monoxide (Kluyver and Schnellen, 1947). Prior to this, Weringa (1936) had discovered Clostridium aceticum, which utilized hydrogen as an electron donor for growth by reduction of carbon dioxide to acetic acid. Thus, the roots of understanding one-carbon metabolism in chemotrophic anaerobes belong to Dutch microbiologists. H. A. Barker (1956) was the first to provide a common metabolic concept that unified the carbon transformation reactions associated with methanogenesis, and his review on the biological formation of methane is of prime historical significance for the archives of microbial physiology. Since these early investigations, it is apparent to me that studies in two other laboratories clearly pioneered our present-day understanding of the microbiology and biochemistry of CImetabolism in chemotrophic anaerobes. Namely, R. S. Wolfe for his studies on the biological features of methanogens (see Balch er al., 1979), and H. G. Wood for his elucidation of the path of one-carbon metabolism in homo-acetogens (see Ljungdahl and Wood, 1982). At first glance, CI metabolism in anaerobes appears to be without any unifying features because it is widely displayed by sulphidogenic,acidogenic and methanogenic bacteria which themselves are extremely diverse and

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phylogenetically unrelated. However, a common metabolic theme has recently emerged from the blossoming, but still limited, research on this subject. That is, homo-acetogens produce acetyl-CoA/acetate from CI metabolism, and methanogens produce, via CI metabolism, either acetylCoA/acetate or carbon dioxide and methane, from one-carbon compounds, or methane and carbon dioxide from acetate. It is in the synthesis or degradation of acetate/acetyl-CoA via CI metabolism that the unique attributes of anaerobic species which grow on one-carbon compounds are united. The purpose of this review is to examine the microbial physiology that accounts for unicarbonotrophy in anaerobes. Emphasis is placed on what is known and how it was shown. This review is not intended to be encyclopedic, but rather to draw together the pertinent reference material which forms the foundation for this topical focus. Undoubtedly, some general references to more specific features of methanogens, homo-acetogens and sulphidogenswill not be incorporated here, but this information can be gathered from the more specific reviews cited with regard to each of these groups.

11. Transformation of One-Carbon Metabolites by Anaerobes A. PRODUCTION OF ONE-CARBON COMPOUNDS

The CI metabolites enter anaerobic ecosystems of the biosphere either as pollutants from aerobic environments, volcanic or deep subsurface emanations, or via chemical transformation reactions performed by anaerobic micro-organisms. The formation of CI metabolites often necessitates their removal in biological elemental cycles because they can accumulate and either alter normal carbon and electron flow within a cell, or become toxic and result in cell death. The biological formation of CI metabolites requires special biochemical transformation reactions (i.e. a specificmechanism).The rate and amount of CI metabolite formed generally depends on whether the onecarbon compound is produced as a result of either a catabolic, anabolic, exchange, or non-fortuitous reaction. Table 1 summarizes the biochemical transformations associated with generation of CImetabolites by anaerobes. Formate is produced by different species of anaerobic bacteria. Both homo-acetogenicand hetero-acidogenic Clostridium species generate formate (Thauer et al., 1976). Clostridium kluyeri, C. butyricum and C . butylicum generate formate via a CoA-dependent pyruvate formate lyase reaction (Thauer et al., 1972). In these species, formate serves as the CI unit required for anabolic reactions associated with amino acid and purine synthesis (Jungermann et al., 1968,1370).Streptococcus and Rhodospirillum also utilize

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TABLE 1. Formation of one-carbon compounds via chemotrophic metabolism of anaerobic bacteria C I compound Formate (HCOOH) Carbon monoxide KO)

Organism

Biochemical transformation

Rhodospirilum rubrum Streptococcus species Streptococcus mitis Butyribacterium methylotrophicum Clostridium species Acetobacterium woodii Escherichia coli Clostridium sporogenes Desulfovibrio species Methanosarcina barkeri

Carbon dioxide reductase, pyruvate formate lyase Pyruvate formate lyase Pyruvate formate lyase Haem cleavage (aerobic) Carbon dioxide/carbon monoxide exchange Pectin methylesterase Aromatic acid demethoxylation Protein methylesterase Methionine demethiolation Unknown Trimethylamine cleavage

Methanobacterium species Methanosarcina barkeri Clostridium pasteurianum Desulfovibrio species

Methyl-CoM reductase Methyl-CoM reductase Pyruvate cleavage Pyruvate cleavage

Clostridium species

pyruvate formate lyase to generate this CI compound (Lindmark et al., 1969; Jungermann and Schon, 1974). Clostridium thermoaceticum, C . pasteurianum, C . acidiurici and C .formicoaceticum produce formate by direct reduction of carbon monoxide via carbon dioxide reductase (Thauer, 1972, 1973; Thauer et al., 1970, 1976; Andreesen et al., 1973). The physiological functions proposed for carbon dioxide reductase (i.e. formate dehydrogenase, EC 1.2.1.2)includes formation of the methyl group of methionine in C. pasteurianum, and in catabolic formation of acetate by C . thermoaceticum and C . acidiurici. Formate is also a fermentation end-product of Ruminococcus albus (Miller, 1978). In short, the generation of this C1 compound by anaerobes can be viewed as a widespread property. Interest in microbial carbon monoxide metabolism has intensified recently (Uffen, 198 l), and especially in relation to aerobic hydrogen-oxidizing bacteria (Bowien and Schlegel, 1981). The latest review by Kim and Hegeman (1 982), which appears primary to understanding microbial carbon monoxide metabolism, indicates that carbon monoxide formation by anaerobes awaits documentation. It is worth noting that Streptococcus mitis, an acidogen,

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formed carbon monoxide via haem degradation, but only in the presence of air (Engel et al., 1972). Recently, Lynd et al. (1982) showed that Butyribacterium methylotrophicum actively formed [I4C]carbon monoxide from [14C]carbon dioxide via an isotopic exchange reaction during growth with carbon monoxide as sole energy source. The discovery of an active carbon dioxide/carbon monoxide exchange reaction, and the proposed functions for carbon monoxide dehydrogenase activity in homo-acetogens and methanogens (see Sections 111 and IV) suggest that carbon monoxide may be an intermediate, or formed as an atypical product of CI-metabolizinganaerobes. At present it is clear that consumption of carbon monoxide by anaerobes parallels that of hydrogen in anaerobic ecosystems where organic matter is vigorously decomposed since the partial pressures of both gases are maintained at extremely low values (Zeikus, 1977) and a wide variety of hydrogen and carbon monoxide consuming species exist (acidogens, methanogens and sulphidogens). Methanol is a major reduced fermentation end-product of pectin decomposition by both mesophilic and thermophilic Clostridium species (Sehink and Zeikus, 1980).Interestingly, all aerobic, facultative and anaerobic pectinolytic bacteria examined so far formed methanol via pectin methylesterase, but lacked the ability to consume methanol. Hence, pectin metabolism in part establishes the niche for methylotrophic anaerobes in nature. To date, no evidence exists to suggest that methanol is a direct end-product of lignin biodegradation (Zeikus, 1981). However, a variety of soluble methoxylated aromatic acids were metabolized by Acetobacterium woodii, and the methanol formed was concomitantly consumed, while the aromatic rings were left intact (Bache and Pfennig, 1981). Undoubtedly, more mechanisms of methanol generation from methoxylated substrates remain to be discovered in anaerobes.In this regard it should be noted that Escherichia coli forms methanol via a protein methylesterase activity (Toews and Adler, 1979).There is no reason to assume that this enzyme would not function under anoxic conditions but this point remains unproven. Methanthiol (methylmercaptan) is a significant CI compound formed via anaerobic digestion of proteins and glycoproteins. Facultative organisms (e.g. Proteus vulgaris) and obligate anaerobes (e.g. Clostridium sporogenes) form methylmercaptan via demethiolation of methionine (Segal and Starkey, 1972; Wisendanger and Nisman, 1953). Sulphate-reducing species also form traces of methylmercaptan during growth on lactate; however, the biochemical transformation reaction that accounts for this activity has not been reported (Hatchikian et al., 1976). Methylamine is formed as an intermediary metabolite during fermentation of trimethylamine by Methanosarcina barkeri (Hippe et al., 1979). In nature, trimethylamine is formed by the bacterial reduction of N-oxide present in

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marine fish (Kim and Chang, 1974) or by degradation of plant-derived components such as choline (Neil1 et al., 1978). Desulfouibrio species form trimethylamine from the metabolism of choline (Hayward and Stadtman, 1959; Walther et al., 1981a,b). Methane is the most abundant CIcompound generated in the biosphere. It represents the major reduced fermentation product of CI metabolism in methanogenic bacteria. Apparently, all methanogens form methane via the methyl-CoM reductase reaction (Wolfe and Higgins, 1979). Other anaerobic species also generate traces of methane. For example, Desulfooibrio species form methane ( < 0.1 % culture head space) as well as ethane and propane when grown on lactate (Hatchikian et al., 1976). Both Desulfooibrio sp. and C. pasteurianum produce methane from pyruvate in reaction mixtures that contained cell extract and vitamin B12 (Postgate, 1969). Notably, under these conditions, the methane was derived from the C-3 of pyruvate, and this suggests methyl reduction and not carboxyl reduction as the route for mini-methane production.

B. CONSUMPTION OF ONE-CARBON COMPOUNDS

A variety of physiologically diverse bacteria transform CI substrates during anaerobic growth (see Table 2). Both obligate anaerobes, including methanogenic, acidogenic and sulphidogenic species, and facultative photosynthogenic and dinitrogenogenic bacteria, can consume CI substrates. The anaerobic utilization of a C1 substrate by a given species also has a very high degree of correlation with its ability to metabolize hydrogen and carbon dioxide as a source of carbon and electrons. At present, the physiological functions known for CI consumption during anaerobic growth include use of the substrate as either a source of cellular carbon or electrons, or both. In addition, consumption of C1 compounds can also be non-utilitarian in anaerobes, and result from an exchange or non-fortuitous biochemical reaction. Thus, it is important to note that consumption of CI substrates by chemotrophic anaerobes is not always associated with its utilization as sole carbon and energy (i.e. electrons) source of growth. Two general trends concerning utilization of CI substrates by anaerobes are worth a brief mention. First, more species will be described that utilize C1 substrates, because this metabolic feature has been poorly examined in the microbial world. Second, facile utilization of certain substrates via CI metabolism, in obligate anaerobes, requires a prolonged cultural adaptation period. This latter feature is important in understanding the mechanism of methanogenesis from acetate by M. barkeri (Stadtman and Barker, 1951; Weimer and Zeikus, 1978b; Winter and Wolfe, 1979), and formation of

TABLE 2. Functional comparison of one-carbon compound transformation and hydrogen-dependent carbon dioxide utilization during anaerobic growth of bacteria

h)

!2

Physiological function Representative species Methanogen Methanosarcina barkeri Methanobacterium thermoautotrophicum Methanococcus vannielii Methanobacterium formicicum Acidogen Clostridiwn thermoautotrophicum Clostridium thermoaceticum Clostridium pasteurianum Acetobacterium woodii Butyribacterium methylotrophicwn

Hydrogen/ carbon dioxide Carbon &onoxide Formate Methanol Methylamine Methane 1

I

1 1 1

1

NR NR

NR

1

4

1

NR NR NR 1 3

1

1 NR 1 1

1 NR NR NR 1

NR NR 1 1

1

NR NR NR NR NR NR NR NR

4 4

NR 4

NR NR NR NR NR

L

n

Ex

C

cn Sulphidogen Desulfovibrio desulfuricans Desulfovibrio vulgaris Thermodesulfotobacteriumcommune Dinitrogenogen Paracoccus denitrificans Wotasynthogen Rhodopseudomonas acidophilia Rhodopseudomonas capsulatus

NR NR NR

4

NR

2 NR NR

NR

1

1

NR

NR

1

NR 1

1 1

1

1

NR NR

NR NR

4

1

NR NR

1

1

1

NR NR

Please note more species consume CI substrates than listed. Abbreviations: NR, not reported; 1, utilized as a carbon source and electron donor; 2, utilized as an electron donor; 3, utilized as a carbon source; 4, exchange or non-fortuitous reaction (i.e. anabolic or catabolic function not established).

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acetate from carbon monoxide by B. methylotrophicum (Lynd et al., 1982). By and large, general knowledge concerning consumption of C1 compounds by methanogens has been limited to utilization of these substrates as the sole carbon and energy sources for growth (Zeikus, 1977; Balch et al., 1979). Many methanogenic bacteria described to date can grow with hydrogen and carbon dioxide as the sole carbon and electron donor. Growth on formate is displayed by fewer species, like M . formicicum (Schauer and Ferry, 1980) and M . vanneili (Jones and Stadtman, 1976). Methanol and methylamine serve as the sole carbon and energy source for the growth of Methanosarcina species, including M . barkeri (Weimer and Zeikus, 1978a), M . vacuolata and M . mazei (C. A. Zavarzin, personal communication). The utilization of methylamine as a methanogenic precursor was first reported by Zhilina and Zavarzin (1973). In fact, this citation is a classic because it noted (C. A. Zavarzin, personal communication) the utilization of methylamine as an energy source for M . barkeri, and hydrogen/carbon dioxide for D. vulgaris. Unfortunately, these interpretations were lost in the English translation. Carbon monoxide serves as a carbon and electron donor for many methanogenic species (Kluyver and Schnellen, 1947; Daniels et al., 1977). However, the metabolism of carbon monoxide as sole carbon and electron donor for growth is very slow and requires a prolonged cultural adaptation period with respect to both M . thermoautotrophicum (Daniels et al., 1977)and M . barkeri (R. Kerby, unpublished findings). Growth of methanogens on formaldehyde, or methylmercaptan, has not been demonstrated. Notably, a variety of methane-producing bacteria were reported to oxidize methane to carbon dioxide during growth under high pressures of methane and with a utilizable energy source present (Zehnder and Brock, 1979b). Currently, this finding is not interpreted as a net consumption of methane because the data suggest that this transformation probably occurred via an active exchange reaction present in the CImetabolizing anaerobes (see Sections I11 and IV). This conclusion is also supported by their finding that [14C]methyl-CoMwas not labelled by [I4C]methane;however, an unidentified intermediary metabolite which chromatographically migrated near methylcobalamin was observed. The specific metabolic details of one-carbon transformations in methanogens is covered in Section 111. Recognition of the abilities of acidogenic anaerobes to grow on CI substrates as sole carbon and energy sources is quite recent. Utilization of hydrogen/carbon dioxide as carbon and electron donor for the growth of homo-acetogens is widespread, and occurs in physiologically diverse species including Acetobacterium woodii (Balch et al., 1977), Butyribacterium methylotrophicum (Zeikus et al., 1980b) and Clostridium thermoautotrophicum (Wiegel et al., 1981). This metabolic character in anaerobes is poorly studied

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as witnessed by the fact that C . thermoaceticum grows readily on hydrogen/ carbon dioxide or carbon monoxide as carbon and energy source (Kerby and Zeikus, 1983);however, this species was generally regarded as being incapable of growth on this substrate or on carbon monoxide (Thauer et al., 1977; and personal communication). Growth of C . thermoaceticum on carbon monoxide as sole carbon and energy source does require a prolonged period of adaptation. Utilization of carbon monoxide as the sole energy source for growth by homo-acetogens was first established by Lynd and Zeikus (1981). Growth of homo-acetogens on carbon monoxide may be as widespread as that on hydrogen/carbon dioxide. Genthner and Bryant (1982) established that Eubacterium limosum-B. rettgeri grows on carbon monoxide, and both our laboratories have demonstrated this feature for A . woodii (R. Kirby, unpublished findings). Clostridium thermoautotrophicum also grows on carbon monoxide alone (J. Wiegel, personal communication). The major discovery which led to these studies, however, was the demonstration that carbon monoxide was oxidized and served as an electron donor during the growth of C . pasteurianum and of various homo-acetogenic clostridia (Fuchs et al., 1974; Thauer et al., 1974; Diekert and Thauer, 1978). Less is known about formate utilization by homo-acetogens. Acetobacterium woodii (Balch et al., 1977)uses it as a sole carbon and energy source and B. methylotrophicum utilizes it as carbon source in place of carbon dioxide or acetate (R. Kerby, unpublished findings). Other hetero-acidogens, such as Vibrio succinogenes- Wolinella succinogenes, utilize formate as an electron donor for fumarate reduction, but C1 metabolism per se has not been examined (Kroger, 1976). Methanol is utilized as an electron donor and carbon source for the growth of B. methylotrophicum (Zeikus et al., 1980b),E. limosum-B. rettgeri (Genthner et al., 1981), A . woodii (Bache and Pfennig, 1981) and C . thermoautotrophicum (Wiegel et al., 1981). Utilization of other CIsubstrates such as methylamine, formaldehyde, methanthiol or methane, during the growth of acidogens, has not been demonstrated. The specific metabolic details of one-carbon transformations in homo-acetogens is covered in Section IV. At present, it is too early to generate specific physiological concepts concerning CI metabolism in sulphidogenic anaerobes. Indeed, the extreme metabolic diversity in catabolic substrate range of sulphate-thiosulphate-sulphur reducing species that display heterotrophic or autotrophic nutrition is just now being established (Pfennig and Widdel, 1981). Nonetheless, certain trends in the literature suggest that C1 metabolism also may occur in sulphidogenic anaerobes. The utilization of one-carbon substrates by sulphidogens was also reviewed by Barton (1981). Utilization of hydrogen/carbon dioxide as an electron donor and carbon source for sulphate-reducing bacteria was well established by the early 1970s

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(Sorokin, 1966; LeGall and Postgate, 1973). At present, chemolithotrophic growth on hydrogen and carbon dioxide plus acetate as sole electron and carbon sources has been shown to occur with Desulfovibrio vulgaris (Badziong et al., 1978), Thermodesulfotobacterium commune (Zeikus et al., 1982), D. desulfuricans and D . gigas (Brandis and Thauer, 1981). Notably, it was mentioned by the authors that growth on hydrogen by the latter three organisms required a prolonged period of cultural adaptation. Hydrogen metabolism in sulphidogens has become a controversial topic because alternative mechanisms have been proposed for hydrogenase function and localization (Hatchikian et al., 1978; Tsuji and Yagi, 1980; Glick et al., 1980; Badziong and Thauer, 1980; Odom and Peck, 1981), carbon dioxide assimilation (Alvarez and Barton, 1977; Badziong et al., 1979) and the mechanism of coupling sulphate reduction to ATP synthesis (Thauer and Badziong, 1981; Odom and Peck, 1982). Also two major discoveries concerning the mechanism of sulphate reduction are worth noting. First, sulphate reduction is coupled to electron transport-mediated phosphorylation in Desulfovibrio but not Desulfotomaculum species (Liu and Peck, 1981). In the latter genus, an additional substrate level phosphorylation is associated with pyrophosphatase, acetate phosphotransferase and acetate kinase activities. Second, Vainshtein et al. (1980) demonstrated thiosulphate as a free intermediate during sulphate reduction by Desulfovibrio sp. This finding is of significance in supporting free sulphur intermediates during sulphate reduction and may, in part, establish the niche for thiosulphate-reducing anaerobes in nature. The presence of a carbon monoxide oxidizing enzyme in chemotrophic anaerobes was first established in cell extracts of D. desulfuricans (Yagi, 1958, 1959).However, utilization of carbon monoxide as carbon and electron donor for the growth of Desulfovibrio sp. has not been demonstrated. Notably, both D. desulfuricans and D . vulgaris species utilize formate as an electron donor, and presumably as a carbon source for growth (Postgate, 1979). Methanol was reported as an electron donor for sulphate reduction in D. desulfuricans (Howard and Hungate, 1976).Methylotrophic sulphate reducers remain to be documented in pure culture, but methanol-consuming sulphate-reducingand acidogenic enrichment cultures can be obtained from aquatic sediments (T. Phelps, unpublished findings). Utilization of methylamine, methanthiol or formaldehyde during the growth of sulphidogenic bacteria has not been reported. Interestingly, Davis and Yarbrough (1966) showed that methane was oxidized by cultures of D. desulfiricans. A more recent report (Panganiban et al., 1979) suggests that a sulphidogenic bacterium can grow with methane as the electron and carbon donor. This result, however, remains controversial (see Zehnder and Brock, 1979b, 1980) and enigmatic. The anaerobic utilization of C,substrates by facultative species, which also

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can employ oxygen as electron acceptor for C1 oxidation, is of general interest; however, the physiological mechanisms of substrate metabolism are quite different from those found in obligately anaerobic, chemotrophic bacteria (see Section V, p. 276). Both dinitrogenogenic and photosynthogenic bacteria, which under anaerobic growth conditions form respectively nitrogen and photosynthate as the major reduced end-products, display C1 metabolism. Paracoccus denitrijicans can consume hydrogen/carbon dioxide, methanol or formate as carbon and electron donor during anaerobic growth, with nitrate as electron acceptor (Willison and John, 1979; Buchanan and Gibbons, 1975; Bamforth and Quayle, 1978). Photosynthetic bacteria, such as Rhodopseudomonas capsulata and R . acidophila, can utilize methanol, formate or hydrogen/carbon dioxide as carbon and electron donors in the light (Quayle and Pfennig, 1975; Siefert and Pfennig, 1979). Carbon monoxide serves as carbon and electron donor for the growth of R. capsulata in the dark (Uffen, 1976, 1981). Notably, methane has not been demonstrated to serve as a carbon and electron donor for the growth of facultative methylotrophs in the absence of oxygen.

111. One-Carbon Transformations in Methanogens It is obvious that methanogens are of considerable interest because almost as many reviews have been written since 1977 on aspects of methanogen physiology as E. coli has pili (Zeikus, 1977, 1980a,b;Thauer et al., 1977;Mah et al., 1977; Zehnder, 1978;Thauer and Fuchs, 1979; Balch et al., 1979;Wolfe and Higgins, 1979; Wolfe, 1979, 1980; Kandler, 1979;Smith et al., 1980; Mah and Smith, 1981; Prkvot, 1980a,b; Taylor, 1982; Zehnder et al., 1982). It is apparent from these reviews that the two words which most adequately describe the physiology of methanogens are unique and diverse. At the macromolecular levels, methanogen uniqueness shares the common attributes of the kingdom Archaebacteria, a taxonomic grouping of both aerobic and anaerobic procaryotes which itself is composed of metabolically unrelated halophilic, thermoacidophilic and methanogenic species (Woese, 1981). The shared macromolecular properties of this taxonomic group, that are distinct from the kingdom Eubacteria, include (i) a common 16s ribosomal RNA homology (Woese and Fox, 1977), (ii) cell walls devoid of muramic acid (Kandler, 1979), (iii) lipids comprising ether-linked polyisoprenoid chains (Tornabene et al., 1979), (iv) conserved genome organization (Sapienza and Doolittle, 1982; Mitchell et al., 1979; Searcy and Doyle, 1975; Moore and McCarthy, 1969) and (v) a novel protein synthesizing apparatus (Best, 1978; Douglas et al., 1980; Stetter et al., 1980). On the metabolic level,

METABOLISM

OF

ONE-CARBON COMPOUNDS

227

methanogen uniqueness also shares the common attributes of obligately anaerobic unicarbonotrophs, a metabolic grouping composed of taxonomically unrelated methanogenic, sulphidogenic and homo-acetogenic species. The discovery of any evolutionary relatedness within this metabolic group awaits comparative macromolecular analysis of common enzymes. The point here is to examine the physiological diversity that exists among methanogens whose metabolism is mechanistically unified by common CI transformation pathways. Methanogen diversity has been best demonstrated by comparison of the macromolecular relatedness of the 16s ribosomal RNA sequences (Fox ef af., 1980) and the cell wall compositions (Kandler, 1979). These studies indicate that 16s RNA sequence and wall composition varies tremendously among different methanogens; however, both macromolecular approaches support the subdivision of methanogens into three orders, four families and at least eight genera (Balch et af., 1979).

A. GENERAL PHYSIOLOGY A N D SPECIES PROPERTIES

Methanogens are anatomically diverse (Zeikus and Bowen, 1975). Major differences in morphology and wall ultrastructure corresponds to unique wall chemical compositions (Kandler and Hippe, 1977; Kandler and Konig, 1978). The following are presented as examples of this feature: Methanobacterium species, rods that contain a pseudomurein wall; Methanococcus species, regular coccus that have a protein subunit wall; Methanospirillwn species, regularly curved rods that have an inner wall composed of protein subunits and an external sheath; and Methanosarcina species, spherical packets that have a heteropolysaccharide wall. The neutral and polar lipid composition (Tornabene and Langworthy, 1978) and the DNA G + C content (Zeikus, 1977; Balch et af., 1979) also vary considerably in these genera. The specific wall chemical composition also varies among species of a given genus (Konig and Kandler, 1979). The unique cell wall and lipid compositions account in part for the resistance of methanogens to many antibiotics which commonly inhibit growth of eubacteria (Hilpert et al., 1981). The chemical composition of the cell wall of most methanogenic species also poses difficulties for investigators who need to prepare sphaeroplasts or protoplasts, because common enzymes that normally lyse cell walls, such as lysozyme, are not effective. However, techniques have been established for the generation of sphaeroplasts from M. hungatii (Sprott et af.,1979) and protoplasts from M. bryantii (Jarrell et af., 1982). Unicarbonotrophy unites the physiologicallydiverse methanogenicspecies. Several biochemical components, which appear widespread in methanogens,

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have been found to function in the expression of this unique metabolic potential. Corrinoids were first identified in M. barkeri and as a unique cobalamin form, factor 111, which contains 5-hydroxybenzimidazole as a-ligand (Stadtman and Blaylock, 1966a,b; Stadtman, 1967). Corrinoids abound in all methanogens so far examined (Krzycki and Zeikus, 1980), and M. barkeri contained the highest concentrations, but the amount varied with the carbon and electron source (Krzycki and Zeikus, 1980;Scherer and Sahm, 1981a,b). Cobalamins function in methyl transfer reactions in methanogens (Stadtman, 1967; Taylor and Wolfe, 1974b; Wood et al., 1982; Kenealy and Zeikus, 1981, 1982a). The presence of coenzyme M or 2-mercaptoethane sulphonic acid (Taylor and Wolfe, 1974a,b) appears to be unique to methanogens (Balch and Wolfe, 1979). Methyl-CoM is an intermediary metabolite of one-carbon metabolism in methanogens (McBride and Wolfe, 1971; Daniels and Zeikus, 1978; Shapiro and Wolfe, 1980) and functions in methyl transfer reactions (McBride and Wolfe, 1971; Taylor and Wolfe, 1974a,b;Gunsalus and Wolfe, 1980). Factor F430 (discovered by Gunsalus and Wolfe, 1978a) contains nickel (Whitman and Wolfe, 1980; Diekert et al., 1980a). This coenzyme is a tetrapyrrole (Diekert et al., 1980b) and is found in methanogens but not homo-acetogens (Diekert et al., 1981). The nickel tetrapyrrole is a component of methyl-CoM methyl reductase (Ellefson and Wolfe, 1981a; Vogels et al., 1982) and functions in methanogenesis (Ellefson and Wolfe, 1981b; Keltjens and Vogels, 1981; Thauer, 1982). A pteridine, coenzyme Y FC (yellow fluorescent compound), was discovered as an intermediary metabolite of one-carbon metabolism in diverse methanogens (Daniels and Zeikus, 1978). Recently, coenzyme-YFC was identified as a unique carboxylated pteridine, carboxydihydromethanopterin (Keltjens and Vogels, 1981). A novel deazoflavin, factor F.tzO,is abundant in methanogens (Eirich et al., 1978). Factor F420 is a low redox electron carrier that functions in both catabolic (Tzeng et al., 1975a,b; Daniels et al., 1977) and anabolic (Zeikus et al., 1977; Weimer and Zeikus, 1978a; Kenealy et al., 1982) oxidoreductase reactions. The presence of oxidized factor F420 in methanogens is of considerable practical significance in their tentative identification by fluorescence microscopy (Mink and Dugan, 1977; Doddema and Vogels, 1978). This procedure has been of value in my laboratory for the past seven years in noting methanogens in natural samples from digestors, sediments, wetwoods and thermal springs. In short, methanogens contain novel biochemical components associated with their unicarbonotrophy, and more await discovery. To establish some taxonomic unity in methanogen diversity, Balch et al. (1979) proposed a new taxonomic scheme based primarily on data obtained from analysis of 16s RNA oligonucleotide sequence homology. The new molecular taxonomic treatment included definition of the orders,

METABOLISM OF ONE-CARBON COMPOUNDS

229

Methanobacteriales, Methanococcales and Methanomicrobiales, the families, Methanobacteriaceae, Methanococcaceae, Methanomicrobiaceae and Methanosarcinaceae, and the genera, Methanobacterium, Methanobrevibacter, Methanococcus, Methanomicrobium, Methanogenium, Methanospirillum and Methanosarcina. Prbvot (l980a,b) also proposed a new classical scheme for taxonomic treatment of methanogens, based primarily on cell shape, Gram-staining, DNA G + C content and motility. The PrCvot scheme identified four families (Micrococcaceae, Ristellaceae, Bacteriaceae and Spirillaceae) and seven genera (Methanococcus, Methanosarcina, Methanogenium, Methanobacterium, Methanopoieticum, Zeikusella and Methanospirillum). It seems reasonable to evaluate these proposals and render an opinion which is common to the speculative nature of taxonomic assignments. My evaluation leads to the conclusion that the orders of families and genera proposed by Balch et al. (1 979) should be used. This conclusion is based on the understanding that the primary structure of 16s ribosomal RNA is highly conserved in species and adequately reflects the suprageneric relatedness of bacteria (Woese et al., 1979, whereas the more classical characterization of cell morphology, staining and DNA G C content means little by itself. The Balch scheme thus provides the groundwork on which to base taxonomic relatedness among methanogen diversity. However, it does not provide an adequate scientific basis for speciation or practical nomenclature. Utilization of 16s ribosomal RNA analysis in speciation is too difficult and requires a tremendous amount of expertise to identify oligonucleotides (C. R. Woese, personal communication). Most importantly, 16s RNA homology, like DNA G C content, cell shape or methanogenesis, independently or in combination with these characters, are not valid criteria for speciation. Strains having all the above features identical may be quite distinct and separate species when examined at the level of protein homology. The species and strain relatedness proposed by Balch et al. (1979) can be questioned because 16s RNA is coded by only a small part of total DNA. Future studies call for more specific macromolecular procedures including DNA-DNA/DNA-RNA hybridization and analysis of total and specific cell proteins. The inadequacy of methanogen speciation at present has been apparent in the recent literature. Perhaps the best example of this situation is presented by the taxonomic confusion associated with thermophilic methanogens. Methanobacterium thermoautotrophicwn was isolated from a heat exchanger (> SOOC) in a municipal sewage-treatment facility, and was the first extreme (i.e. grows optimally above 60°C) thermophilic methanogen described (Zeikus and Wolfe, 1972). At the time, T. D. Brock suggested the selection of a different genus name that would incorporate the thermal properties, but this advice was not followed. It is now clear from the literature that obligate thermophily requires multigenetic differences and that primary structural

+

+

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G. ZEIKUS

changes in proteins, including the presence of more hydrophobic amino acids, accounts in part for the thermal stability and activity of enzymes present in extreme thermophiles (Zeikus, 1979). The genus question now raised is whether M . thermoautotrophicum is more related to M . bryantii (Balch et al., 1979)than to the new thermophilic species Methanothermusferidus (Stetter et al., 1981)? All three species utilize hydrogen/carbon dioxide, have a pseudomurein wall, and the DNA G + C contents (in mole %) are 52,38 and 39 for the type strains of M . thermoautotrophicum (Zeikus and Wolfe, 1972),M . bryantii (Balch et al., 1979)and M .feridus (Stetter et al., 1981)respectively.The species question that is now raised about thermophilic methanogens was eloquently described by Brandis et al. (1981). These investigators compared the relatedness of Methanobacterium thermoautotrophicum type strains AH (Zeikus and Wolfe, 1972) with a new isolate, the Marburg strain, that was previously assumed to be the same species (Fuchs et al., 1978). These authors demonstrated the following in the two strains: (i) only 42% DNA homology by hybridization; (ii) different molecular weight DNA-dependent RNA polymerases; (iii) different amino and uronic acid sugar components in the pseudomurein wall; and (iv) membrane-bound adenosine triphosphatase (ATPase) was present in strain AH but absent from strain Marburg. My answer to this species question, based on the data, is that the two strains are not the same species. Actually, we never could achieve the amount of acetate incorporation found by Fuchs et al. (1978) with our M . thermoautotrophicum type strain AH, which assimilated acetate (Zeikus et al., 1975),but only 10%of the cell carbon came from acetate. Hopefully, this discovery of Brandis et al. (1981) will make colleagues who are sceptical of the results of others both more patient and more aware of the exact methanogenic strain used as an experimental model. Other thermophilic methanogenic strains have been isolated (Zinder and Mah, 1979; Zeikus et al., 1980a; Marty and Bianchi, 1981; Binder et al., 1981; Ronnow and Gunnarsson, 1981) and the species designation of all remain an open question. Speciation of Methanosarcina strains is another focal point of taxonomic confusion. Zhilina (1976) was the first to recognize different biotypes of Methanosarcina at a time when some colleagues questioned whether the gas vacuoles present in strains might suggest contamination by blue-green algae. Balch et al. (1979), considers Methanosarcina strains 227, MS and W as M . barkeri, although the DNA G + C content varies from 38.8 to 43.5 in these strains (Weimer and Zeikus, 1978a; Balch et al., 1979). Major differences in the regulation of C1 metabolism have been reported in these strains, and this will be described below (pp. 233 and 255). Both immunological (Macario et al., 1981)and very significantdifferencesin ribosomal protein pattern (Douglas et al., 1980)are displayed by Methanosarcina strains. Zhilina and Zavarzin (1979, and personal communication) recognize four species of Methanosarcina: M.

METAB0L IS M 0F 0NE- CAR B0N CO MPOU NDS

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barkeri, M . methanica, M . vacuolata and M . mazei. The last species name is contested by Mah (1980) who proposes Methanococcus as the genus name. The recognition of a new taxon, Methanobrevibacter arboriphilus (Balch et al., 1979) for Methanobacterium arbophilicum (Zeikus and Henning, 1975) was based on valid macromolecular differences observed between Methanobacterium and Methanobrevibacter. However, the consideration of Methanobacterium strain AZ (Zehnder and Wuhrmann, 1977) as a strain of M . arboriphilus (Balch et al., 1979) appears superficial. The basic problem with methanogen speciation is that their wide taxonomic divergence is associated with a very specialized metabolism (i.e. unicarbonotrophy). Two strategies can be used, either to recognize a limited number of species by classic tests (a la PrCvot) or to recognize a large number of species by expanding the macromolecular approach initiated by Balch et al. (1979), but with more precise analysis of nucleic acid and protein homology and immunochemical analysis. Two other new species described since 1979 include Methanothrix soehngenii (Zehnder et al., 1980) and Methanoplasma elizabethii (Rose and Pirt, 1981). The former is an important acetate-fermenting species that is architecturally distinct and appears widespread in anaerobic digestors (Zehnder et al., 1980), but it grows extremely slowly in pure culture. Notably, this species has not been found to grow on hydrogen/carbon dioxide or formate although it contains both hydrogenase and formate dehydrogenase activity (A. J. B. Zehnder, personal communication). Methanoplasma elizabethii is of interest as it was first noted as a glucose fermenting methanogen; but on cultural purification it was documented as the first methanogen devoid of a cell wall (Rose and Pirt, 1981). Growing a methanogen is, in itself, often an inherently difficult task, not because cultural techniques are lacking (see Balch et al., 1979)but because the organisms’ nutritional needs often are not understood or optimized. Feelings of amusement and hindsight have been expressed with, for example, an account of the discovery of a novel nickel tetrapyrrole coenzyme which was based on the growth-dependent increase associated with insertion of a stainless-steel gassing needle into an all-glass fermentor (Thauer, 1982). Also, the initial failure to isolate or grow M . vanneili on formate in places other than California, was traced to the absence of selenium in source waters and its requirement for formate dehydrogenase activity (Jones and Stadtman, 1977). And lastly, recognizing that the failure to grow M . barberi on acetate as a sole carbon and energy source was the result of a requirement for a prolonged cultural adaptation period. When the organism was routinely maintained on hydrogen/carbon dioxide in a basal medium for several years, or on complex medium with acetate, it took months of repeated culture transfer for them to grow readily on acetate alone (Zeikus et al., 1975; Weimer and Zeikus, 1978b; Krzycki et al., 1982). One prerequisite in studies of the growth physiology of

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methanogens is to understand their unique nutritional requirements. It is simply not known why some species grow slowly, and the growth of other species stops when electron, carbon or nitrogen are limiting nor why sulphur sources are not limiting in a pH-stat fermentation. Not all methanogens are capable of autotrophic growth. For example, M. uoltae requires isoleucine, leucine and acetate for growth on hydrogen/carbon dioxide, and these organic compounds are assimilated (more than 20% for each) into total cell carbon (Whitman et al., 1981). Detailed studies of methanogen growth physiology have been limited primarily to M . thermoautotrophicum, the most prolific species, and to M . barkeri, the most metabolicallyversatile species.The only substrates known to serve as sole carbon and electron donor for growth of M . thermoautotrophicum with phosphate, ammonium and sulphide as phosphate, nitrogen and sulphur sources, are hydrogen/carbon dioxide (Zeikus and Wolfe, 1972) and carbon monoxide (Daniels et al., 1977). The maximal cell growth rate, in pressure tubes, on carbon monoxide alone was only 1% of that on hydrogen/carbon dioxide and the strain used was not adapted to grow on carbon monoxide concentrations (in the head space) of greater than 30%. Taylor and Pirt (1977) reported that in the initial medium, iron and nitrogen sources used for growth of Methanobacterium thermoautotrophicum (type strain AH) on hydrogen/carbon dioxide were limiting, and reported a doubling time of 3 hours in batch culture. Schonheit et al. (1979,1980) showed that growth of strain Marburg, at 65"C, was dependent on nickel, cobalt and molybdenum, and achieved a doubling time of 1 to 1.8 hours and a cell density of 2 g cells (dry weight) litre-', in batch culture. In addition, growth of strain Marburg is dependent on sodium (Perski et al., 1981). Kenealy et al. (1982) demonstrated a doubling time of 2.6 hours for M . thermoautotrophicum (type strain AH) in an ammonium-limited chemostat and established that growth yield was directly dependent on the ammonium concentration of the medium. Brandis et al. (1981) reported a doubling time of 2.2 hours for this strain. We have obtained a doubling time of 60 minutes (flo%), near culture washout, during chemostat growth of M. thermoautotrophicum (T. Thompson, unpublished observations). Thus, thermophilic methanogens like M . thermoautotrophicum, are to date, and to my best knowledge, the most prolific methanogens, chemolithotrophs or autotrophs documented in the microbial world. In general, M . thermoautotrophicum type strain AH appears to be a chemolithotroph, but it is not an obligate autotroph. A significant portion of cell carbon can be derived from acetate (Zeikus et al., 1975). Cysteine can serve as a sulphur source (Kenealy et al., 1982) and glutamine as nitrogen source (Institut Pasteur). Sprott and Jarrell(l981) showed that the cell membrane is permeable to phenylalanine.

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The metabolic versatility of M . barkeri is based on its use of a variety of C I compounds, polymethylamines and acetate either as sole carbon and energy source during unitrophic growth or in combination with these substrates during mixotrophic growth. Weimer and Zeikus (1978a) demonstrated that M. barkeri (type strain MS) consumed either hydrogen/carbon dioxide, methanol or methylamine as sole carbon and electron donor for growth, and grew mixotrophically via simultaneous metabolism of hydrogen/carbon dioxide methanol or methanol/methylamine. The growth yield (mg cells (mmol methane formed)-') obtained at the end of batch cultivation was higher on methanol than on hydrogen/carbon dioxide. Hippe et al. (1979) established that M . barkeri grew on tri-, di- and monomethylamine as sole carbon and nitrogen source. Polymethylamines are an interesting substrate because metabolism of the methyl group mechanistically parallels utilization of the methyl group in acetate. In this regard it is interesting to note the recent report of Walther et al. (1981a,b) that demonstrated formation of 7943% CD3H and 1418% CD2H2 from highly deuterated methylamines. Studies of acetate fermentation by Methanosarcina strain 227 (Mah et al., 1978; Smith and Mah, 1978) and M . barkeri neotype strain MS (Weimer and Zeikus, 1978b; Winter and Wolfe, 1979) all agree that a cultural adaptation period (i.e. a lag period of more than 4 weeks) is required to establish active methane formation when stocks continuously maintained on either hydrogen/carbon dioxide or methanol are used as the inoculum. Continued transfer of stock cultures on acetate results in an acetate-adapted strain (i.e. one that grows much more readily). It is worth noting here that M . barkeri grows as clumps of cells and, while investigators claim cultural purity, the organism has not been verifiably cloned. Thus, the suggestion by Krzycki et al. (1982), that adapted strains are metabolic mutants, remains to be proven. Zhilina (1978) demonstrated that Methanosarcina biotype 2 grew on acetate as the sole carbon and energy source. This result was also shown for Methanosarcina strain 227 by Smith and Mah (1980) and for M. barkeri by Hutten et al. (1980), Scherer and Sahm (198 la) and Krzycki et al. (1982). In my laboratory, it took over a year to adapt M . barkeri cultures to grow with a doubling time of less than 40 hours with acetate as the sole energy source. Notably, Smith and Mah (1978) reported that either hydrogen/carbon dioxide or methanol inhibited acetate fermentation by a mechanism resembling catabolite repression in Methanosarcina strain 227. However, acetate and either hydrogen/carbon dioxide or methanol were simultaneously consumed during the growth of M. barkeri (Weimer and Zeikus, 1978b). N o evidence for catabolite repression of methanogenesis from acetate has been presented with respect to M . barkeri neotype strain MS (Hutten et al., 1980; Scherer and Sahm, 1981a,b;Krzycki et al., 1982). Figure 1 illustrates a typical batch growth curve of M . barkeri during

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' W

Time (hours)

FIG. 1. Comparison of (a) methanol (0)and (b) acetate (0)as energy sources for growth of Methunosurcinu burkeri (0) in batch culture.

fermentation of sodium acetate or methanol in 15-litre glass carboys without pH control. It is important to note that growth rate is correlated with the rate of substrate consumption. The majority of methane produced and acetate consumed by M. barkeri, under these conditions, occurred after growth had ceased. Comparison of cell yields of M. barkeri, during exponential growth on acetate or methanol, indicate similar metabolic efficiency (about 3.99 g cell (mol substrate)-') although methanol fermentation is thermodynamically favoured over acetate. Thus, our former conclusions, which indicated greater metabolic efficiency on methanol than on acetate, were erroneous because they were determined at the end of growth (Weimer and Zeikus, 1978b). It has become apparent from studies in progress that most growth parameters (i.e. ptmax, K,, YCH4,Ys) reported in the literature (Weimer and Zeikus, 1978a,b; Smith and Mah, 1978, 1980; Scherer and Sahm, 1981a,b), for growth of Methanosarcina strains on hydrogen/carbon dioxide, methanol or acetate,

METABOLISM OF ONE-CARBON COMPOUNDS

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may not be valid in continuous-flow culture. Detailed nutritional studies aimed at growth improvement of M.barkeri strains have been reported by Scherer and Sahm (198 la,b). Cobalt, molybdenum, selenium and nickel were required as trace elements for growth, certain strains required riboflavin, and methionine (but not cysteine) could replace sodium sulphide as sulphur source.

B. CATABOLISM

The overall mechanism for the generation of cellular energy via CI transformation reactions in methanogens is complex, and at present it is a very speculative science. Two major conceptual problems have been examined;the pathway of carbon and electrons from energy-substrate to methane and the less studied coupling of these reactions to ATP synthesis. Barker (1956) first proposed a carbon flow pathway that unified consumption of energy sources by methanogens. Today, Barker’s scheme (see Fig. 2) still serves as a useful model because the data at hand are in agreement. The thermodynamically most favoured reaction, which is common to methanogenesisfrom all energy substrates used for growth, is the reduction of a methyl-level intermediate to methane. By and large, establishing the exact biochemistry of this transformation has been the “holy grail of methanogenologists”. In the beginning, cobalamin biochemistry was considered essential for methyl reduction to

CO,

t

X H +XCOOH

l’.’.“.

XCHO CH,OH

CH,COOH

+

XH

\

+ XH

-2H -co,*

XCH,

1+2H +

XH

CH,

FIG. 2. Barker scheme for carbon and electron flow during methanogenesis. X represents unknown carbon carriers. From Barker (1956).

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methane (Stadtman, 1967).The next era focused on coenzyme M biochemistry as being primary to methanogenesis (Gunsalus et al., 1976). The latest quest concedes the importance of a nickel tetrapyrrole in the terminal methyl reductase reaction associated with methanogenesis (Vogels et al., 1982; Thauer, 1982). In general, two approaches have been utilized to understand the pathways of carbon and electrons in methanogenesis; namely, I4Clabelling studies in uiuo and enzymological characterization in uitro. The results of both approaches are incorporated below to indicate present knowledge on the biochemistry of methanogen catabolism. 1. Methanogenesis from HydrogenlCarbon Dioxide and Methanol

Detailed studies on the mechanism of methanogenesis from hydrogen/carbon dioxide primarily involve M . thermoautotrophicum as a model, whereas M . barkeri is used for studies on methanol. Daniels and Zeikus (1978) utilized short-term (below 60 seconds) labelling studies, with either [I4C]carbon dioxide or [I4C]methanol,to establish that one-carbon carriers and various intermediates of cell synthesis were products of one-carbon fixation;however, free C1 ihtermediates (methanol or formaldehyde) were not detected. Interestingly, [14C]formate was detected, but this was interpreted as a byproduct of the cell-extraction procedure because it, and other CI metabolites, are not detected in the culture broth of either species. Also, neither M. thermoautotrophicum nor M . barkeri grow on formate. Two coenzyme M derivatives (i.e. methyl-CoM and CIXT)were labelled by methanol or carbon dioxide, and this established a role in uiuo for methyl-CoM in CI metabolism. Notably, most of the label fixed from methanol, by M . barkeri, appeared in methyl-CoM; whereas [14C]carbondioxide often was incorporated at higher concentrations into a new intermediate YFC (yellow fluorescent compound) by either M. barkeri or M . thermoautotrophicum. Chemical and physical characterization of intermediate YFC indicated that it had the properties of a unique pteridine coenzyme. Recently, Keltjens and Vogels (1981) have identified coenzyme YFC as carboxy-7,8-dehydromethanopterin.Notably, [2-I4C]acetatewas not assimilated into detectable CI carriers by cell suspensions of M . thermoautotrophicum, which accords with the inability of this species to catabolize acetate. The short-term labelling approach failed to provide evidence for a role of methyl-B12in CI metabolism. This approach is limited to the detection of free intermediates and thus may not detect an enzyme-bound methyl-corrinoid. The discovery of coenzyme M (McBride and Wolfe, 1971; Taylor and Wolfe, 1974a,b)initiated a chemical-enzymic approach that focused on CoM derivatives and ignored the role of other CI carriers such as carboxy-7,8-

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dihydromethanopterin (Wolfe, 1979). Gunsalus et al. (1976) proposed a modification of the Barker scheme with coenzyme M itself replacing X (unknown carbon carrier) in all C1 transformations associated with carbon dioxide or methanol. This conclusion was based on initial studies involving general properties of the methyl reductase system in Methanobacterium and Methanosarcina. Gunsalus and Wolfe (1977, 1978a,b) discovered that addition of methyl-CoM stimulated methanogenesis from hydrogen/carbon dioxide some 30-fold in crude cell extracts of M . thermoautotrophicum, and that ATP was required for reduction of methyl-CoM. The methyl-CoM reductase of M. thermoautotrophicum was purified into three components, A-C (Gunsalus and Wolfe, 1980). Component A contained oxygen-sensitive hydrogenase activity, component B was an acidic cofactor, and component C another protein. The methyl reductase system displayed an absolute specificity for methyl-CoM, and methyl-Bl2 was not reduced to methane (Gunsalus et al., 1978; R. S . Wolfe, personal communication). Component C was later identified as the methyl reductase and component A was replaced by an NADPH
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that (i) it requires cobalamins as activators in methanogenesis from methylCoM, (ii) carbon dioxide reduction to methane is not stimulated by methyl-CoM and (iii) formaldehyde and HOHzC-CoM are not detectable intermediates. Wood et al. (1982) isolated a vitamin B12-containingprotein from M . barkeri which was methylated by [I4C]methyliodide and active in methanogenesis. Hence, they proposed a direct role for methyl-BI2 in methanogenesis via methylation of CoM by the methyl transferase described by Taylor and Wolfe (1974b). All these suggestions stated above require in viuo evidence and the use of metabolic mutants to determine the actual pathway of carbon flow in methanogenesis. The path of electron flow in methanogenesis, from hydrogen/carbon dioxide or methanol, is more poorly studied than that of carbon flow. The electron acceptor for hydrogenase, and its transfer to NADPH, was first documented in Methanobacterium extracts by Tzeng et al. (1975b). The hydrogenase in crude extracts of both M . thermoautotrophicum (Zeikus et al., 1977) and M . barkeri (Weimer and Zeikus, 1978a) couples to the reduction of the 5-deazaflavin F420 and not to ferredoxin or cytochromes. Reduced factor F420 is then linked to NADPH formation via F42rNADP oxidoreductase. Hence, both reduced F420 and NADPH serve as electron donors for reductive C1 transfonriations in methanogens. Other electron carriers remain to be identified, and hydrogenase itself remains to be purified. Methanol oxidation has not been characterized enzymically in M . barkeri but has been suggested to occur via C, transformation reaction in the reverse direction of CH3X generation from carbon dioxide/hydrogen (Zeikus, 1980a,b). Kenealy and Zeikus (1982b) demonstrated that carbon and electron flow are interconnected between methanol, methyl-CoM and carboxy-Y FC. A cytochrome was detected in Methanosarcina strain Fusaro, but its significance in methanogen metabolism is not known (Kuhn et al., 1979). In short, interpretations of the data reviewed here support the following path for carbon and electron flow, in methanogens, from methanol or hydrogen/carbon dioxide: CH30H or H2/CO2+CH3-S-CoM+CH4. But please do not interpret the single arrows as indicating that only one CI transformation reaction, or carrier, is employed. The role of CoM as methyl carrier is supported by the in viuo identification of this CI carrier (Daniels and Zeikus, 1978) and the specificity of the CoM methyl reductase (Gunsalus et al., 1978; Shapiro and Wolfe,. 1980; Ellefson and Wolfe, 1981a,b). 2. Methanogenesis from Other Substrates The intermediary pathways of carbon and electron flow during methano-

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genesis from carbon dioxide, formate, methylamine or acetate have not been documented. Carbon monoxide and formate are considered to be oxidized to carbon dioxide and reducing equivalents and the former then being reduced to methane. However, evidence has not been presented which rules out the following carbon flow path: CO or HCOOH-+OHCX+CHd. L+H+

co2 Carbon monoxide is oxidized during the growth of methanogenic bacteria on hydrogen/carbon dioxide/carbon monoxide gas mixtures (Daniels et al., 1977). The activity of carbon monoxide dehydrogenase in cell extracts of M . thermoautotrophicwn grown on hydrogen/carbon dioxide is 200-fold less active than hydrogenase, and the in vivo electron acceptor is the 5-deazoflavin, factor F420 (Daniels et al., 1977). Notably, carbon monoxide dehydrogenase activity increases five-fold and in M. barkeri is equivalent in activity to the hydrogenase of M. thermoautotrophicum when M . barkeri is grown on acetate (Krzycki et al., 1982). Likewise, synthesisof acetate from carbon monoxide by cell extracts of M . barkeri (Kenealy and Zeikus, 1982a) also suggests that the function in uivo of this CI-transforming enzyme is in the synthesis of a two-carbon metabolite. Nonetheless, carbon monoxide was converted into methane during the growth of a M . barkeri strain that was adapted to grow, albeit slowly, on 100% carbon monoxide in the gas phase as sole energy source (see Fig. 3). These results agree with conversion of carbon monoxide into methane, as demonstrated with cell suspensions of M. barkeri by Kluyver and Schnellen (1947), but differ from the results of Uffen (1976) who was not able to demonstrate carbon monoxide consumption. Formate dehydrogenase activity has been characterized in M . ruminantium, M. vunneili and M . formicicum. Tzeng et al. (1975a) indicated that formate oxidation was coupled to F420 reduction, and subsequent NADPH generation, in crude cell extracts of M. ruminantium. This route of formate metabolism was substantiated in M . vanneili by Jones and Stadtman (1980, 1981) who separated the system into a formate dehydrogenase, a 5-deazaflavin cofactor, and a 5-deazoflavin-dependentNADP+ reductase. Yamazaki and Tsai (1980) purified the 8-hydroxy-5-deazaflavin-dependentNADP reductase of M. vunneili, and showed that, at pH 7.0, NADPH generation was 24 times greater than its oxidation. Two separate formate dehydrogenaseswere documented in M. vanneili (Jones et al., 1979; Jones and Stadtman, 1981). One purified enzyme activity contained molybdenum, iron and acid-labile sulphide, whereas the other was a complex comprising a seleno-cysteine protein and molybdo-iron-sulphur protein subunits. Schauer and Ferry (1982) purified formate dehydrogenase 71-fold from M . formicicum and this 288,000

240

J. G. ZEIKUS 4400 4000

3600

3200

2800

2400

0

I00

200

300

400

Tlmc (hours 1

FIG. 3. Relationship between carbon monoxide (0)consumption and methane (0) production during growth of a Methanosarcina barkeri strain adapted .to grow on carbon monoxide (100% of the gas phase) as sole carbon and electron donor (R. Wolkin, unpublished observations).

molecular weight protein reduced factor F420. Notably, a fluorescent pteridine was associated with the purified protein. Figure 4 presents a carbon-flow model for conversion of acetate into methane by M . barkeri that extends Barker's scheme. This revised scheme is

FIG. 4. Scheme for carbon and electron flow proposed for methanogenesis during growth of a Methanosarcina barkeri strain adapted to grow on acetate as sole carbon and electron donor. Abbreviations: BIZ,a corrinoid; Y,unknown formyl carrier; CoA, coenzyme A; CoM, coenzyme M; YFC, methanopterin; numbers refer to moles of metabolite. This model suggests that carbon monoxide dehydrogenase is a key cleavage enzyme in oxidoreduction of acetate. Modified from Zeikus (1982).

METABOLISM OF ONE-CARBON COMPOUNDS

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based on multiple observations and it represents a hypothesis in need of further testing. Krzycki et al. (1982) compared the ratio of [14C]methaneand ['4C]carbondioxide formed during growth of an acetate-adapted strain of M. barkeri on differentially labelled acetate. Notably, 15% of the C-2 of acetate was oxidized to carbon dioxide and 15% of the C-1 of acetate was reduced to methane. Previously mixed-culture studies of Stadtman and Barker (195 l), results of Smith and Mah (1978) with Mefhanosarcina strain 227, and of Weimer and Zeikus (1978b), with a strain of M . barkeri that was not adapted to grow on acetate alone, all showed only a minor oxidation of C-2 of acetate to carbon dioxide. Notably, the only oxidoreductase activity shown to increase in the acetate-adapted strain of M. barkeri was carbon monoxide dehydrogenase (Krzycki et al., 1982). The role of acetate transformation of this enzyme in the proposed carbon flow path is also analogous to the function of carbon monoxide dehydrogenase in homo-acetogens (i.e. as a transcarboxylase activity which functions in the reverse direction). Kenealy and Zeikus (1982a) reported that cells grown on acetate had high concentrations of acetate kinase and phosphotransacetylase which could function in conversion of acetate into acetyl-CoA. In addition, methanogenesis and growth of M . barkeri on acetate were inhibited 50% by the presence of 4 PM iodopropane (Kenealy and Zeikus, 198I), a known antagonist of corrinoiddependent metabolism (Wood and Wolfe, 1966). Detailed studies of shortterm intermediary metabolites that were formed during acetate fermentation was not reported, but methyl-CoM was detected (J. Krzycki, unpublished observations). The importance of this scheme is to illustrate a redox model for methanogenesis from acetate which is based primarily on coupling formyl oxidation (i.e. YCHO) to methyl-CoM reduction. The generation of electrons (i.e. H) from oxidation of carrier-bound CI units can account for the effects of of methanol or hydrogen on influencing carbon and electron flow during methanogenesis from acetate. Thus, if methanol enters the pathway as methyl-CoM, then methane formed from acetate could still continue but with more carbon dioxide generated from [2-I4C]acetate,as suggested by Krzycki et al. (1982). This model is also consistent with the observation that methane formed from [ l-14C]acetate,in cell suspensions,is greatly diluted by unlabelled carbon dioxide, indicating an active exchange reaction (J. Krzycki, unpublished observations). This scheme does not deal with the mechanism of coupling of acetate transformation to ATP synthesis; however, it implies that a common exergonic redox reaction is coupled to methanogenesis (i.e. methyl-CoM reductase) from acetate. Also it assumes that any energy required for conversion of acetate into acetyl-CoA is maintained by the favourable net thermodynamic force of the overall redox process (i.e. CH&OOH+COz+CH4).

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3 . ATP Synthesis The mechanism of coupling methanogenesis to ATP synthesis remains at present more speculative than the pathways of carbon and electron flow in C I metabolism. Indeed these studies should be viewed in combination with the fact that methane production and growth can be readily uncoupled in cultures; hence so too, presumably, may methanogenesis and ATP synthesis in whole cells and membranes. Nonetheless, several interesting reports have established the background studies needed on this general topic, and this literature has been reviewed by Kell et a f . (1980, 1981). Thauer et a f . (1977) concluded that ATP synthesis occurred via electron transport phosphorylation in methanogens because ATP synthesis via substrate level phosphorylation was not demonstrated. On the other hand, Zehnder and Brock (1979a) hypothesized that energy production from acetate involved neither electron transport nor substrate level phosphorylation, and was not associated with a redox reaction. Wolfe and Higgins (1979) proposed that neither hydrogen nor acetate enters into the cell cytoplasm of a methanogen but a protonmotive force is generated (i.e. interior alkaline) via substrate oxidation. However, no experimental evidence has yet been presented to support the above speculations. The chemi-osmotic energy coupling hypothesis of Mitchell (1 974) suggests that the force driving ATP synthesis is an electrical gradient of protons across the cell membrane. Mountfort (1978) showed that shifting the external pH value from 8.2 to 2.5 in whole cells of M . barkeri resulted in ATP synthesis. This result implies an interior alkaline pH value for M . barkeri cells. By using whole cells, membrane-dialysis techniques and hydrogen peroxide to release incorporated ['4C]formate,a ApH of 2.2 units (interior alkaline) was estimated to be extant in methanol-grown M . barkeri (R. Wolkin, unpublished observations). On the other hand, Jarrell and Sprott (1981) estimated an internal pH value of about 6.7 in M . thermoautotrophicum, and suggested an outwardly directed proton pumping across the cytoplasmic membrane without the establishment of a ApH (interior alkaline). Kell et al. (1981) indicated that a pH gradient was not established with membrane vesicles of M . thermoautotrophicum, and the direction of transport-driven proton translocation was outward. Thus, the role of a membrane-bound adenine nucleotide translocase was considered essential in ATP synthesis (Kell et al., 1981). Sauer et al. (1980a) demonstrated that valinomycin inhibited synthesis of ATP and methane, but did not act as a membrane ionophore in M . thermoautotrophicum. Establishment of a proton gradient across the cell membrane of M . thermoautotrophicum is supported by the finding that the hydrogen present in methane comes from water, but this would imply an interior alkaline pH value (Daniels et af., 1980).

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Adenosine triphosphatase was detected in membranes of M . thermoautotrophicum type strain AH (Doddema et al., 1978, 1979), but not in strain Marburg (Brandis et al., 1981). Methanosarcina barkeri contains an active ATPase but its location was not determined (Kenealy and Zeikus, 1982a). Membrane fractions of M. thermoautotrophicum and M . ruminantium are active in methanogenesis without the addition of ATP (Sauer et al., 1979, 1980a; Doddema et al., 1979). The internal membranes of M . thermoautotrophicum were suggested as the site for methanogenesis (Zeikus, 1977) and this proposal is still supported by Spencer et al. (1980), Doddema et al. (1979) and Kell et al. (1981). In short, the mechanism of coupling ATP synthesis to methanogenesis is uncertain and controversial.

C . CELL CARBON SYNTHESIS

In parallel with the elucidation of the pathways of carbon and electron flow to methane, detailed studies on methanogen cell carbon synthesisare also limited to those that occur in M . thermoautotrophicum and M . barkeri. Fortunately, considerably more is known about the specific carbon transformations associated with anabolism than with catabolism, in methanogens. However, at the forefront of this research there still remains a clearer elucidation of the specific C1 transformation reactions involved in either autotrophic or meth ylotrophic growth. 1. Anabolism in Methanobacterium thermoautotrophicum Please note that no distinction is drawn in this section between M . thermoautotrophicum type strain AH (Zeikus and Wolfe, 1972) and strain Marburg (Fuchs et al., 1978). Taylor el al. (1976) published the first report aimed at understanding intermediary metabolism in M . thermoautotrophicum. These investigators compared the incorporation of 14C-labelledcarbon dioxide, CIsubstrates, carboxylic acids, amino acids and glucose into various cell components and into methane. Only carbon dioxide, acetate, formaldehyde and formate were significantlyincorporated (more than 2% of label) into methane and cell carbon. The labelling patterns observed suggested the absence of hydroxypyruvate reductase, hexulose phosphate synthetase and ribulose bisphosphate carboxylase. Acetate was implicated as a key intermediate because of its assimilation into amino acids. However, the data cast doubt on the homo-acetogenic path, present in clostridia, and favoured acetate synthesis via the reductive carboxylic acid cycle of phototrophs.

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Zeikus et ai. (1977) analysed various oxidoreductases associated with the tricarboxylic acid (TCA) cycle and other key anabolic enzymes of M. thermoautotrophicum.These studies indicated that a complete TCA cycle was not functional because several key oxidative enzymes were not detectable (e.g. isocitrate dehydrogenase and citrate lyase). Nonetheless, anabolic enzyme levels of two key reductive carboxylation reactions were present (that is, pyruvate synthase and 2-oxoglutarate synthase). The pyruvate and 2-oxoglutarate oxidoreductase activities were both coenzyme A acylating and required 5-deazaflavin as electron donor/acceptor. In addition, NAD+-linked malate dehydrogenase, NADP-linked glyceraldehyde 3-phosphate dehydrogenase, fumarate reductase, 5-deazaflavin-linked hydrogenase and Sdeazaflavinlinked NADP reductase activitieswere quantified and suggested to function in cell carbon synthesis. Daniels and Zeikus (1978) pulse-labelled cell suspensionsof M. thermoautotrophicum with [I4C]carbon dioxide, for short time periods, and determined where the label was incorporated. At 60 seconds the label was incorporated predominately into one-carbon carriers (i.e. CoM derivatives and YFC) and the amino acids alanine, aspartate and glutamate. Analysis of the percentage of total label distributed into different intermediary metabolites with time indicated that pyruvate (as alanine) was an early intermediate of net carbon dioxide fixation, and that the ribulose bisphosphate, serine or hexulose phosphosphate pathways of CI assimilation were not operative. Kenealy and Zeikus (1982b) reported on the kinetic and catalytic features of phosphoenolpyruvate carboxylase and showed that it was present in M. thermoautotrophicumat concentrations sufficient to account for the synthesis of oxaloacetate from carbon dioxide. The phosphoenolpyruvate carboxylase activity in partially purified cell extracts was insensitive to air and showed an absolute dependence on phosphoenolpyruvate and magnesium chloride. Thus, this enzyme functions in the net synthesis of a C4 unit which is required for cell synthesis when M. thermoautotrophicum is growing autotrophically. These studies also indicated that the enzyme which converts pyruvate into phosphoenolpyruvate requires characterization, but only low concentrations of pyruvate carboxylation were detected, and this activity was not inhibited by Avidin. Kenealy et al. (1982) provided evidence for the mechanism of alanine, aspartate and glutamate syntheses from 0x0 acids, and the mechanism of ammonia assimilation during growth on hydrogen/carbon dioxide. Cell extracts of M. thermoautotrophicum which aminated 0x0 acids contained glutamine synthetase, glutamate synthase, alanine dehydrogenase, glutamate oxaloacetate transaminase and glutamate-pyruvate transaminase, but glutamate dehydrogenase was not detected. The high K , value for NH4+ displayed by alanine dehydrogenase correlated with maximal cell enzyme activities

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when the organism was grown in continuous culture with excess ammonia. Likewise, the low K,,,value for NH4+ displayed by glutamine synthetase correlated with maximal cell enzyme activities in ammonia-limited chemostat cultures. Fuchs and his coworkers used 14C radiotracer analysis to provide more evidence for the function of pyruvate and 2-oxoglutarate synthase, and for the absence of a complete TCA cycle during growth of the Marburg strain on hydrogen/carbon dioxide. Analysis of the distribution of label in alanine, aspartate and glutamate, formed during growth of the Marburg strain on hydrogen/carbon dioxide and [U-I4C]acetate,supported the following activities: (i) conversion of acetate into acetyl-CoA; (ii) formation of pyruvate via pyruvate synthase; (iii) formation of oxaloacetate via phosphoenolpyruvate carboxylase; and (iv) formation of 2-oxoglutarate via 2-oxoglutarate synthase (Fuchs et al., 1978).Fuchs and Stupperich (1978) confirmed the absence of key oxidative enzymes of the TCA cycle by showing that ['4C]succinatewas only incorporated into glutamate. Notably, degradation of glutamate revealed that the C-1 of this amino acid was derived from carbon dioxide, and C-2 to C-5 from succinate, indicating that 2-oxoglutarate was formed by reductive carboxylation of succinyl-CoA by 2-oxoglutarate synthase. Stupperich and Fuchs (198 1) repeated short-term [I4C]carbondioxide labelling studies with the Marburg strain and confirmed the same patterns of incorporation into cell carbon intermediates previously demonstrated with M. thermoautotrophicum by Daniels and Zeikus (1978). Also, as expected, the position of the label in alanine, after a two second pulse, was 61% in C-l,23% in C-2 and 16%in C-3. Another point in the Stupperich and Fuchs (1981) paper was to show that the [I4C]acetateincorporation studies reported ealier (Daniels and Zeikus, 1978), which showed succinate and a carboxylic acid as assimilation products, were not valid because the commercial [I4C]acetate used was contaminated. We independently confirmed this finding, but unfortunately it was not discovered until 1979during the course of pulse-labelling cell suspensionsof M. barkeri to determine the pathway of acetate reduction to methane (J. Labos and J. Krzycki unpublished findings). The exact two-carbon intermediate of autotrophic carbon dioxide fixation (i.e. acetate or acetyl-CoA), in M. thermoautotrophicum, is a subject which needs further study. Oberlies et al. (1980) concluded that it was acetyl-CoA since acetate was not detected during the growth of strain Marburg, and cell extracts lacked acetate kinase and phosphotransacetylase. Kenealy and Zeikus (198 1) demonstrated that growth of diverse methanogens, including M. thermoautotrophicum and M . barkeri, on hydrogen/carbon dioxide, was inhibited by low concentrations of iodopropane, a corrinoid antagonist. Thus, 40 ,UM iodopropane inhibited growth of M. barkeri by 80% whereas methanogenesis was inhibited by only 30%. In the presence of acetate,

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iodopropane was no longer an inhibitor of methanogen growth on hydrogen/ carbon dioxide. This finding suggested that a cobalamin (i.e. vitamin B,z)-dependent step was involved in the autotrophic synthesis of acetyl-CoA or acetate in methanogens. R. K. Thauer (personal communication) has suggested that acetate can never be a two-carbon intermediate of autotrophic carbon dioxide fixation in C . thermoautotrophicurn. This may well be inherent if carbon monoxide dehydrogenase functions anabolically in methanogens in a way analogous to its proposed function in homo-acetogens (see Section IV). Other reports on cell-carbon synthesis in M . thermoautotrophicurn strain Marburg include the presence of L-alanine in its walls (Schoheit and Thauer, 1980) and the synthesis of factor F430 from the incorporation of 8-succinate (Diekert et al., 1980a,b).This last fact, and the inhibition of F430 synthesis by laevulinic acid (Jaenchen et al., 1981),an analogue of S-aminolaevulinicacid, was used as evidence that the nickel-containing F430 was a tetrapyrrole. Undoubtedly this will be proven in the near future with absolute chemicalphysical characterization data. 2 . Anabolism in Methanosarcina barkeri Daniels and Zeikus (1978) showed that [I4C]methanoland [14C]carbondioxide were fixed into similar short-term intermediary metabolites (i.e. CoM derivatives, YFC, alanine, aspartate and glutamate) by cells grown on either methanol or hydrogen/carbon dioxide as sole carbon and energy source. Notably, radioactivity in amino acids was only detected at longer labelling periods with [I4C]methanol,and label in glutamate from [14C]carbondioxide fixation was not detected immediately, as observed with M. thermoautotrophicum. Weimer and Zeikus (1978a) showed that during growth on hydrogen/ carbon dioxide, 48% of the cell carbon was derived from methanol indicating that equivalent amounts of cell carbon were derived from carbon dioxide and a more reduced organic intermediate. Cell extracts lacked dye-coupled methanol dehydrogenase activity, as well as activity for key cell-carbon synthesis enzymes of the Calvin cycle, serine pathway or hexulose pathway. The studies cited above indicated a unique mechanism of autotrophy and methylotrophy in methanogens. Also, Weimer and Zeikus (1978a) provided the first evidence to suggest that synthesis of a two-carbon intermediate, via unicarbonotrophy in methanogens, involved a C,-CI condensation reaction. Weimer and Zeikus (1978b) demonstrated that about half of total cell carbon was synthesized from [I4C]acetate,when cells were grown on either methanol or hydrogen/carbon dioxide. Kenealy and Zeikus (1981) showed that, during mixotrophic growth on hydrogenfcarbon dioxide/methanol/acetate, the percentage weight of cells derived from [I4C]carbon dioxide,

METABOLISM OF ONE-CARBON COMPOUNDS

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[I4C]methanol and [U-I4C]acetate, respectively, were 36.0, 13.5 and 50.8. Notably these studies clearly demonstrated that addition of acetate to cells growing on hydrogen/carbon dioxide/methanol greatly diminished the contribution of carbon dioxide and methanol to cell carbon, but addition of iodopropane did not significantly alter the distribution of incorporated label. Hence, acetate appeared as a direct precursor to cell carbon and its synthesis from one-carbon compounds is iodopropane sensitive. Kenealy et al. (1982) provided evidence that M. barkeri assimilated ammonia via glutamine:2-oxoglutarate aminotransferase, and documented the mechanism of 0x0-acid amination. Cell extracts contained glutamine synthetase, glutamate synthase, glutamate-oxaloacetate transaminase and glutamate pyruvate transaminase. Alanine dehydrogenase and glutamate dehydrogenase were not detected. Hydrogen-reduced 5-deazaflaviq or flavin mononucleotide, was used as the electron donor for glutamate synthase. Weimer and Zeikus (1979) established the major carbon-transformation reactions associated with the syntheses of C3, C4 and CSunits involved in the methylotrophic-autotrophiccarbon dioxide fixation pathway of M. barkeri. Cell extracts of M. barkeri contained anabolic levels of 5-deazaflavin-linked pyruvate synthase, citrate synthase, aconitase, NADP-linked isocitrate dehydrogenase and NAD +-linked malate dehydrogenase. The functioning of these enzymes in cell carbon synthesis was supported by analysis of the position of the label in alanine, aspartate and glutamate when cells were grown on hydrogen/carbon dioxide in the presence of [I4C]acetate. The specific radioactivity of glutamate from cells grown on [1-14C]-or [2-14C]acetate were approximately twice that of aspartate. The methyl and carboxyl carbons of acetate were incorporated into aspartate and glutamate to similar extents. Degradation studies revealed that acetate was not significantly incorporated into the C-1 of alanine, the C-1 or C-4 of aspartate, or the C-1 of glutamate. The C-5 of glutamate, however, was partially derived from the carboxyl carbon of acetate. In short, these studies indicate that the oxidative reaction of the TCA cycle were used, in M. barkeri, to synthesize a CSunit (i.e. glutamate), and this is quite different from M. thermoautotrophicum. Another major difference between these two methanogens is the absence from M. barkeri of phosphoenolpyruvate carboxylase. The enzymic activity that accounts for oxaloacetate synthesis from a C3 unit remains to be elucidated. Cell suspensions of unicarbonotrophically grown M. barkeri synthesized acetate and alanine from [14C]carbondioxide. The addition of iodopropane totally inhibited acetate synthesis but not incorporation of carbon dioxide into alanine, indicating inhibition of a corrinoid-dependent reaction but not the carbon dioxide exchange function of pyruvate synthase. Cell extracts of cells grown on hydrogen/carbon dioxide catalysed the synthesis of [I4C]acetate from [I4C]carbon monoxide (- 1 nmol min-I (mg protein)-') and an

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isotopic exchange between carbon dioxide or carbon monoxide and C-1 of pyruvate. Acetate synthesis from [14C]carbonmonoxide was stimulated by methyl-B12 but not methyl-tetrahydrofolate or methyl-CoM. Both methylCoM and CoM inhibited acetate synthesis. Cell extracts of M. barkeri, grown on hydrogen/carbon dioxide, methanol or acetate, contained high concentrations of phosphotransacetylase (> 6 pmol min-' (mg protein)-') and acetate kinase (>0.14 pmol min-l (mg protein)-'). Thus, it was not possible to distinguish between acetate or acetyl-CoA as the immediate product of two-carbon synthesis via autotrophic cell carbon fixation. In short, the evidence provided above indicates that (i) acetate/acetyl-CoA is synthesized via a CI-CI condensation reaction, and (ii) this condensation reaction is inhibited by iodopropane and associated with carbon monoxide dehydrogenase activity, and this reaction is not required when acetate is provided as the carbon source. 3. Autotrophic Pathways Figure 5 summarizes the findings reviewed above and presents a conceptual model that incorporates the data and illustrates the unique path of carbon during autotrophic growth of both M. thermoautotrophicum and M . barkeri. Please note that the conversion of hydrogen/carbon dioxide into acetyl-CoA, the direct precursor for pyruvate in both species, requires more biochemical characterization to determine the exact CI units employed, and to distinguish whether acetate or acetyl-CoA is the end-product of the net CI-CI condensation reaction. Also, note that synthesis of CSin M. barkeri (MB) occurs via the oxidative TCA cycle enzymes, whereas in M. thermoautotrophicum (MT) the reductive TCA cycle enzymes function in this role. Neither species has, or needs, a complete TCA cycle for anabolism or catabolism. Autotrophy in methanogenic unicarbonotrophs appears uniquely different from that described for aerobic species that employ the Calvin cycle because (i) acetyl-CoA is the direct precursor of cell carbon, not 3-phosphoglycericacid, (ii) the key enzyme is catalysed by a CI-C, condensation reaction and is suggested, but not proven, to have carbon monoxide dehydrogenase activity, and (iii) the key carboxylase is a highly reductive transformation catalysed by pyruvate synthase and not via an oxidative ribulose bisphosphate carboxylase activity. It is doubtful that non-methanogens will use this pathway because of the unique one-carbon and electron carriers employed in the initial CI transformation reactions. Analysis of stable carbon isotopic fractionations to decipher the autotrophic pathway in M. thermoautotrophicum strain Marburg (Fuchs et al., 1979) has been controversial because it leads to the general interpretation that

METABOLISM OF ONE-CARBON COMPOUNDS

249

Hydrogen/carbon dioxide

Methyl B,2

Acetyl-CoA "

Lipids

I

co," Pyruvate

Alonine sugars

Oxaloacetote

C i tra te

Aspartote

Molate

Fumarote Isocitrate

Oxoglu tarate

JI

Succinate

Glutomate

FIG. 5. Autotrophic carbon flow pathways proposed for cell carbon synthesis in Methanosarcina barkeri (MB) and Methanobacterium thermoautotrophicum (MT). This model indicates that initial reactions involve carbon dioxide reduction on CI carriers. The key biosynthetic reaction is shown as a methyl-corrinoid-dependentC1 condensation catalysed by carbon monoxide dehydrogenase. Please note that different TCA-cycle enzymes are used to synthesize glutamate in these methanogens. Modified from Zeikus (1980a).

J. G. ZEIKUS

250

pathways for synthesis of methane and cell carbon are not linked (Kell et al., 1981). R. K. Thauer (personal communication) suggests that these data (Fuchs et al., 1979) only indicate that most of the cell carbon formed by M. thermoautotrophicum is derived from reactions other than those involved in methanogenesis. This is to be expected if the data of Zeikus et al. (1977), Daniels and Zeikus (1978), Fuchs et al. (1978), Fuchs and Stupperich (1978), Daniels (1978), Stupperich and Fuchs (1981), Kenealy (198l), Kenealy et al. (1982) and Kenealy and Zeikus (1982a,b) are accepted as proof of the mechanism of cell carbon synthesis from carbon dioxide. Table 3 compares the stable carbon isotope fractionations of M. thermoautotrophicum and M . barkeri found during growth on hydrogen/carbon dioxide. A I2C/l3C fractionation analysis, during growth, is useful in indicating the general differences in the C, metabolism of the two species, but these data can clearly not be used to suggest that some CI transformation reactions are not common to methanogenesis and cell carbon synthesis. Notably, Kenealy (198 1) used these data to show clearly that acetate replaced carbon from hydrogen/carbon dioxide during growth of M. barkeri, and that carbon in both lipids and total cells showed similar fractionation profiles in the presence or absence of acetate. This latter finding suggests that lipids of TABLE 3. Stable carbon isotope fractionation of Methanobacterium thermoautotrophicum and Methanosarcina barkeri during growth on hydrogen/carbondioxide as energy sourceo

6I3C (ml-')b

Sample Carbon dioxide

Methanobacterium Methanosarcina barkeri strain MS thermoaut otrophicum strain AH Without acetate With acetate -9.7

- 9.9

- 5.0

Methane

- 32.0

- 56.2

- 24.0 - 46.0

Cells

-29.7 -31.9

-41.6 -46.3

-26.5 -29.5

Acetate

Lipids

a Both organisms were grown on phosphate-buffered basal medium at 65°C (strain AH) and 37°C (strain MS) in 12 1 fermentors. No difference was detected for carbon dioxide in the inlet and outlet gases, thus assuring the carbon dioxide pool was infinite. Data from Kenealy (1981) in collaboration with M. R. DeNiro at U.C.L.A. S ' ~ values C were calculated from:

613C (m1-I)

=

13c/12c sample 13C/12Cstandard

METABOLISM OF ONE-CARBON COMPOUNDS

251

methanogens are directly synthesized from a two-carbon intermediate (acetyl-CoA and not pyruvate).

D . UNIFICATION A N D REGULATION OF METABOLISM

Detailed studies on the regulation of metabolism, and the interrelatedness of C, transformations in catabolism and anabolism, have focused on M . barkeri as the model methanogen. Methanobacterium thermoautotrophicum is not a suitable organism for these research objectives because its facile growth is limited to only one carbon and energy source (hydrogen/carbon dioxide). The general impression gained from investigationson M . barkeri is that its extreme metabolic versatility is regulated by the nature of the specific electron donors or acceptors used in both catabolism and anabolism; and that catabolic, anabolic, methylotrophic and autotrophic metabolism share common CI transformation reactions. However, considerably more research is required to substantiate these general metabolic concepts.

1. Unification of Metabolism

Figure 6 represents a general carbon flow scheme for M . barkeri which illustrates CI transformation relationships between cell carbon synthesis, methanogenesis, autotrophy and methylotrophy. The metabolic interpretations presented are based on data already reviewed and on the experimental findings given below. Kenealy and Zeikus (1982a) examined the flow of carbon from [14C]methanoland [I4C]carbon dioxide into methane and cell components when M . barkeri was grown mixotrophically on hydrogen/carbon dioxide/methanol. Either ['4Clmethanol or [I4C]carbon dioxide was equivalently incorporated into the major cellular components (lipids, proteins and nucleic acids). The [I4C]methanol was selectively incorporated into the C-3 of alanine, with decreased amounts fixed in the C-1 and C-2 positions, whereas [14C]carbondioxide was selectively incorporated into the C- 1 moiety, with decreasing amounts assimilatedinto the C-2 and C-3 atoms. Notably, the specific activity of [I4C]methane and [3J4C]alanine, synthesized from [14C]methanolduring growth, shared a common specific activity distinct from that of carbon dioxide or methanol. These results indicated that common intermediates and carbon transformations linked both autotrophic (carbon dioxide) and methylotrophic (methanol) assimilation pathways of M . barkeri, and that the methyl (i.e. C-3) of alanine and the methyl of methane share a common precursor. The debate as to whether this common intermediate is methyl-CoM or methyl-B12will continue, although there is really no direct

J. G. ZEIKUS

252

Carbon dioxide

I

Carboxy- Y FC

Methanol-Methyl

J.

I

-X-Methyl-

Methyl-

CoM-

Methane

B,,

Methanol Carbon dioxide Carbon monoxide

Acetate

Carbon dioxide

+

5

Acetyl- COA

4 \1

Pyruvate

Amino acids

Alanine

Sugars

FIG. 6. Unification of carbon flow pathways for methane and cell synthesis in Methanosarcina barkeri. This model predicts shared CI carriers for initial carbon transformations leading to methane and cell synthesis. Autotrophy and methylotrophy is accounted for by use of common carbon carriers and a common CI-CI condensation reaction. Acetyl-CoA or acetate is the immediate biosynthesis product of unicarbonotrophic metabolism. From Kenealy and Zeikus (1982).

METABOLISM OF ONE-CAREON COMPOUNDS

253

evidence for methyl-vitamin BIZper se. Nonetheless, this represents the first evidence to suggest a biochemical unification between catabolic and anabolic CI transformations in methanogens. The biochemical mechanism that accounts for methylotrophy in M . barkeri appears quite distinct from that described in aerobic unicarbonotrophs which assimilate carbon either via carbon dioxide fixation, in the reductive pentose phosphate path, via formaldehyde fixation in the hexulose phosphate path, or through carbon dioxide and formaldehyde fixation in the serine path. Methanosarcina barkeri assimilates cell carbon at three different one-carbon oxidation states; that is (i) at the methyl level, (ii) a CI unit more oxidized than methyl, and (iii) as carbon dioxide. The key enzymic activity in C3 synthesis, during unicarbonotrophic carbon assimilation, thus appear to be methyltransferase, carbon monoxide dehydrogenase and pyruvate synthase. 2. Regulation of Metabolism Methanosarcina barkeri neotype strain MS readily catabolizes energy sources mixotrophically (Weimer and Zeikus, 1978a,b; Hutten et al., 1981; Scherer and Sahm, 1981a,b; Krzycki et al., 1982). However, the pathways of carbon and electron flow to methane and carbon dioxide were altered significantly during mixotrophic, compared with unitrophic, metabolism of a given substrate. Weimer and Zeikus (1978a) showed that when hydrogen/carbon dioxide/methanol was the energy source, considerably more methane was formed from methanol and less from carbon dioxide than was observed during unitrophic metabolism of these substrates. That these unicarbonotrophic transformations occurred simultaneously was confirmed by Kenealy and Zeikus (1981, 1982a). Notably, cells actively formed methane, but did not grow significantly on hydrogen/methanol alone (Weimer and Zeikus, 1978a). The biochemical basis for this phenomena was explained by Kenealy and Zeikus (1982a). Cell suspensions of M . barkeri that were grown on hydrogen/carbon dioxide/methanol readily incorporated [I4C]methanol or [14C]carbon dioxide into methyl-CoM and carboxy-YFC; however, label in the carboxy-YFC from methanol was only detected in the absence of hydrogen. This supports a role for carboxydihydromethanopterin in carbon dioxide and methanol metabolism, and suggests that carbon and electron flow between methyl-CoM and carboxy-YFC is reversible, and the direction of carbon flow is regulated in part by hydrogen. The same concentration of hydrogenase were present in cell extracts when M . barkeri was grown on methanol or on hydrogen/carbon dioxide alone (Weimer and Zeikus, 1978a). Weimer and Zeikus (I 978a,b) showed that methanol and hydrogenlcarbon dioxide greatly influenced catabolic carbon and electron flow during acetate

254

J. G . ZEIKUS

fermentation. Either hydrogen/carbon dioxide or methanol greatly decreased the contribution of [2-I4C]acetate converted into methane, and methanol greatly increased the amount of [14C]carbon dioxide produced from [2-I4C]acetate. These studies also suggested that M . barkeri could catabolize hydrogen/carbon dioxide, methanol and acetate mixotrophically. However, these investigations used a medium with yeast extract, and detailed kinetic studies showing relationships between substrate consumption and product formation were not performed. Krzycki et al. (1982) demonstrated simultaneous catabolism of methanol and acetate by a strain of M . barkeri adapted to grow on acetate alone. In the presence of 50 mM methanol and 50 mM acetate, 5% of the methane came from the C-1 of acetate, and 26% from the C-2, whereas 52% of the carbon dioxide came from C-1 of acetate and 18% from the C-2 position. During mixotrophic growth on equimolar concentrations of methanol and acetate, the rate constants for both [I4C]methaneand [14C]carbondioxide production from [2-14C]acetateincreased over the values observed during unitrophic metabolism of either 50 mM methanol or 50 mM acetate. Notably, the rate constant for [I4C]carbondioxide production from [2-I4C]acetate, during mixotrophic metabolism, increased 3-fold over the value observed during unitrophic acetate metabolism. The carbon flow scheme presented in Fig. 4 (p. 240) provides a biochemical hypothesis to explain this latter phenomena. Namely, the methyl group of acetate and methanol share a common intermediate, but enter the carbon flow path via different reactions. It is suggested here that carbon flow via the proposed acetate fermentation path also should be regulated in part by low redox reactions controlled by hydrogenase, such that hydrogen would reduce electron carriers also involved with acetate oxidation. The only documented oxidoreductase activity that increases in response to acetate catabolism of M . barkeri is carbon monoxide dehydrogenase (Krzycki et al., 1982). This activity is reversible in homo-acetogens and functions in transformylation (H. G. Wood, personal communication): Acetyl-CoA-CH3X

+[HCOOH]

The carbon monoxide dehydrogenase activity in homo-acetogens is also regulated in response to the carbon and electron donors used in catabolism (Lynd et al., 1982). Although reports exist (see Winter and Wolfe, 1979; McInnery and Bryant, 1981) that M . barkeri does not consume acetate, when hydrogen oxidation is coupled to methanogenesis from carbon dioxide, this appears as a regulatory phenomena dealing with intermediary concentrations of hydrogen/carbon dioxide. Schink and Zeikus (1982) clearly demonstrated that hydrogen, methanol and acetate were simultaneously consumed in C . butyricum-M. barkeri co-cultures that completely degraded pectin to methane

METABOLISM OF ONE-CARBON COMPOUNDS

255

and carbon dioxide. Interestingly, the intermediary levels of hydrogen, methanol and acetate produced were inversely related to the rate at which M. barkeri consumed these electron donors for methanogenesis (i.e. the rate of hydrogen consumption > methanol > acetate). The anabolic carbon and electron flow pattern in M. barkeri is also significantly controlled by the initial fermentation substrates. This organism can grow on hydrogen/carbon dioxide or methanol as the sole carbon and energy source, but both hydrogenlcarbon dioxide and methanol contribute to cell carbon during mixotrophic growth on these substrates (Weimer and Zeikus, 1978a; Kenealy and Zeikus, 1981, 1982a). Likewise, during mixotrophic substrate consumption, acetate replaces cell carbon synthesized from either hydrogenlcarbon dioxide or methanol, regardless of whether acetate is a significant methane precursor (Weimer and Zeikus, 1978b; Kenealy and Zeikus, 1981, 1982; Krzycki et al., 1982). Also, acetate is consumed and forms a significant portion of cell carbon during growth on hydrogen/carbon dioxide/methanol as energy source (Kenealy and Zeikus, 1981). In summary, it should be noted that regulation of CImetabolism as a whole in methanogens appears quite unique, and it is controlled in part by the specific amounts of carbon and electron donors present during growth and their specific rates of consumption. To date, no clear evidence for the induction or repression of a specific CI transformation reaction has been presented. It appears at this preliminary stage that the CI transforming enzymes in both catabolism and anabolism are expressed as a whole, but the activity levels of some methanogen enzymes vary with the given substrate conditions. For example, the levels of glutamine:2-oxoglutarateaminotransferase and alanine dehydrogenase were shown to be regulated by the concentration of NH4+ in cultures of M. thermoautotrophicum (Kenealy et al., 1982), and the levels of carbon monoxide dehydrogenase in M . barkeri increase when cells are grown on acetate as carbon and energy source (Krzycki et al., 1982). Also, acetate kinase activity decreases when cells are grown unior mixotrophically on methanol (Kenealy and Zeikus, 1982b). The mechanism for regulation of enzyme activity levels in methanogens is not known. Although reports exist which suggest that hydrogen or methanol exert catabolite repression over acetate metabolism in Methanosarcina strains, in a manner analogous to that of glucose over lactose utilization in E. coli (Smith and Mah, 1978), this conclusion can be debated on the basis of physiological mechanism. Diauxie in E. coli depends on the saccharide concentration, but glucose acts as a catabolite repressor by preventing expression of lactose uptake and activation enzymes. However, this is not the case with M. barkeri which always assimilates acetate, regardless of the energy source (hydrogen/

256

J. G. ZEIKUS

carbon dioxide, methanol or acetate) and the carbon monoxide dehydrogenase is constitutive. What changes in our acetate-adapted M. barkeri strains is the rate of acetate consumption which increases in parallel with the concentration of this dehydrogenase. Because C, assimilation enzymes appear expressed as a whole, the amount of methane formed, during mixotrophic growth on multiple substrates, will depend on intermediary metabolite levels of electron donors, and their rates of conversion. Thus, the observations (Winter and Wolfe, 1979; McInnery and Bryant, 1981) that acetate is utilized only after hydrogen is depleted is not interpreted here as catabolite repression. One could measure the percentage contribution of acetate to cells before and after hydrogen depletion to assess whether hydrogen does in fact inhibit acetate consumption. Mechanistically, the rate of methane formation depends first on the amount of methanogenic catalyst present, second on the concentrations of electron donor and acceptors present, third on the individual consumption rates of these substrates, and fourth, on the influence of these substrates on altering carbon and electron flow during methanogenesis from a given substrate. It can be assumed that hydrogen can act as a metabolic effector and can channel electrons to carriers involved with the oxidoreduction of acetate, and thus decrease the rate of acetate consumption, but prevention of enzyme synthesis at the molecular level as exists for catabolite repression in E. coli appears a remote possibility. The long incubation time required for cultural adaptation of M. barkeri to growth on acetate as the sole carbon and electron donor should point methanogenologists toward genetic awareness. At present, because of our inability to clone this species, this feature may be the result of there being two subpopulations of Methanosarcina sp. present in the cultures. However, the procedures used by various investigators to achieve good growth on acetate reflect standard industrial practices to obtain mutants of bacteria with altered metabolic features. Recently, the nature of spontaneous mutations for gaining substrate utilization was elegantly shown in E. coli with strains (i.e. lac-) that contained a large deletion in the lactose operon (Hall and Zuzel, 1980). Namely, a new gene function for lactose utilization evolved spontaneously, merely by selection on lactose nutrient broth medium plates. The mutation occurred in a different part of the E. coli gene map and coded for new lactose-metabolizing enzymes which enabled growth. Because of the common laboratory practice of continuous transfer and growth of methanogen stocks in liquid medium, other strain characters may change with time as well as those associated with acetate fermentation. Undoubtedly, the mysteries of methanogen intermediary metabolism will be clarified when a combined biochemical-genetic approach is developed.

METABOLISM OF ONE-CARBON COMPOUNDS

257

IV. One-Carbon Transformations in Homo-acetogens A . GENERAL PHYSIOLOGY A N D SPECIES PROPERTIES

In homo-acetogens, the general mechanism of CI transformation is characterized by the net synthesis of an acetyl-CoA from either heterotrophic or unicarbonotrophic modes of growth. Two major routes of one-carbon metabolism are displayed by chemotrophic anaerobes that make acetic acid as a principal end-product of one-carbon metabolism. The homo-acetogenic pathway involves a CI-C, condensation reaction for synthesis of a two-carbon unit (Schulman et al., 1972), whereas the glycine decarboxylase pathway for synthesis of a two-carbon unit occurs via the coupling of a CI-C~condensation reaction with a C-3 cleavage to account for net acetate synthesis (Weber and Wood, 1979). The total synthesis of acetate from carbon dioxide through heterotrophic anaerobic metabolism has been reviewed by Ljungdahl and Wood ( 1969, 1982). One-carbon transformations of glycine decarboxylasetype acetogens in species like Clostridium acidiurici, C . cylindrosporum and Peptococcus glycinophilus, will not be reviewed here because they to do not display the appropriate mechanism of CI transformation. As a group, homo-acetogens display considerable physiological diversity (Table 4). The DNA G + C content varies from 33 to 58 mol%. This variation is as broad a range as observed in methanogens. Their metabolic versatility is quite outstanding and, unlike the methanogens, homo-acetogens are not limited to one-carbon compounds and acetate as principal energy sources. Rather, homo-acetogens proliferate unicarbonotrophically on hydrogen/carbon dioxide, formate, methanol or carbon monoxide, and/or on hexose, lactate and a diversity of other substrates which appear highly species specific. All species form acetate as a fermentation product but formate, butyrate, propionate and hydrogen/carbon dioxide can be significant end-products in certain species. Acetobacterium woodii (Balch et al., 1977) ferments hydrogen/carbon dioxide, glucose, lactate and formate (after cultural adaptation). Methanol utilization, as an energy source, appears to be either strain specific or requires cultural adaptation. Bache and Pfennig (1981) reported that this species ferments methanol or cleaves and ferments the methoxyl moieties from a variety of aromatic acids. Balch et al. (1977) and Imhoff (1981) reported the absence of a methanol fermentation by A. woodii. Cultures can be adapted to grow on carbon monoxide as the energy source (B. Genther, personal communication; R. Kerby, unpublished observations). Acetate was the only recognized end-product of A. woodii fermentation (Balch et al., 1977), until

N

ul

OJ

TABLE 4. Comparison of DNA G + C contents, representative energy sources and fermentation end-products of described homo-acetogens” Energy sources DNA-G+C (molx)

Organism Acetobacterium woodii Butyribarterium merhylolrophicum Clostridium fhermoaceticum Eubacterium limosum (“Buryri6ocferiwn reftgeri”) Clostridium barkeri Clostridium formidoaceticum Clostridium thermoaulotrophicwn Closrridium oceficum Acetogeniwn kiwi

Hydrogen/ carbon Carbon Hexose dioxide Formate Methanol Lactate monoxide

++ +

39 49 45

+ + + + + +

46-48 45-58 34 53-55 33 38

Abbreviations:

Other vannillate glycerol

++

NR NR

NR

+

++

L

NR

pcntose

+

+

NR

NR

P

+

+

betaine

+

NR

NR

NR

x C

+ + + + +

NR

+

+

+-

++

++

+

NR

NR

+/-

+/-

+

NR NR

hypoxanthine glutamate

+

+

pentose

+ +

NR NR

ethanol pyruvate

+, used as energy source or major product;

Hydrogen/ carbon Acetic Formic Butyric dioxide

++ +

+

~~~~~~~~~~

a

End-products

v)

+

NR

NR NR

NR

NR

NR

NR

NR NR

NR

+

~

-, not used as energy source; +/ -, contrasting reports; NR, not reported.

E

+

METABOLISM OF ONE-CARBON COMPOUNDS

259

Braun and Gottshalk (1981) reported that hydrogen was evolved in amounts up to 0.1 mol (mol substrate oxidized)-’. Methylotrophy in homo-acetogens was first estabished in Butyribacterium methylotrophicum which readily ferments hexose, lactate, hydrogen/carbon dioxide or methanol/carbon dioxide/acetate (Zeikus et al., 1980b).The ability of homo-acetogens to grow on carbon monoxide as energy source was first reported by Lynd and Zeikus (198 1). Butyribacterium methylotrophicum can be adapted to grow rapidly (about 12 hours doubling time) on 100% carbon monoxide in the culture headspace (Lynd et al., 1982). Cultural adaptation required growth in methanol and carbon monoxide medium with the concentration of the latter being progessively increased on repeated transfer. When growth was proceeding on methanol plus 100%carbon monoxide, the methanol was removed and continued transfer on 100% carbon monoxide resulted in selection of a prolific carbon monoxide-utilizingstrain that differed from the wild-type strain Marburg which does not grow on 100% carbon monoxide as energy source. The principal end-products of B. methylotrophicum fermentation depend on the substrate and pH value of the medium. This species can produce either acetate or butyrate as sole end-product or can form mixtures of hydrogen/carbon dioxide, butyrate and acetate. Clostridium thermoaceticum was generally considered to be an obligate heterotroph (Fontaine et al., 1942; Ljungdahl and Wood, 1969; Thauer et al., 1977). However, its metabolic potential was not well examined. Andreesen and Ljungdahl (1973) reported formate as a fermentation end-product. Clostridium thermoaceticum type strain Fontaine can grow readily on hydrogen/carbon dioxide or it can be culturally adapted to grow on carbon monoxide as energy source (Kerby and Zeikus, 1983). At present, Eubacterium limosum is considered the type species of the non-spore-forming genus Eubacterium (Cato et al., 1981). This species name (Buchanan and Gibbons, 1975)replaced “Butyribacterium rettgeri” (Breed et al., 1957)which was metabolically recognized as a unique acetogen by Barker et al. (1945). Barker (1956)concluded that E. limosum was not equivalent to B. rettgeri, but should be considered as a Butyribacterium species (Breed et al., 1957).Eubacterium limosum has a fermentative metabolism that is very similar to B. methylotrophicum. Genthner et al. (1981) reported growth of E. limosum on hydrogen/carbon dioxide and methanol but not on formate as energy source. This organism also can metabolize carbon monoxide as an energy source under low (below 50% carbon monoxide) partial pressures (Genthner and Bryant, 1982). In addition to growth on hexose and lactate, E. limosum can ferment betaine (Miller et al., 1981). Some strains of E. limosum ferment formate but not hydrogen/carbon dioxide (Imhoff, 1981). Clostridium barkeri (Stadtman et al., 1972),like B. methylotrophicum, forms acetic and butyric acids. However, Imhoff (1981) was unable to demonstrate

260

J. G. ZEIKUS

spores, and considers C . barkeri to be taxonomically similar to E. limosum on the basis of DNA/DNA hybridization (102% homology observed). Nonetheless, Imhoff (198 1) reported that C . barkeri formed propionate as a significant end-product and it fermented nicotinic acid and hypoxanthine, but not formate, hydrogen/carbon dioxide or methanol. Clostridium formicoaceticum (Andreesen et al., 1970) and C . aceticum (Weringa, 1936; Braun et al., 1981) appear similar in DNA G C content but vary in substrate range and end-products formed. Clostridium formicoaceticum produces formate and ferments glutamate, and various uronic sugars, whereas C . aceticum forms only acetate and does not ferment methanol or lactate. Clostridium thermoautotrophicum (Wiegel et al., 1981) differs from these species in temperature range and is more similar to C. thermoaceticum in DNA G + C (Matteuzzi et al., 1978). Only C . thermoautotrophicum has been shown to grow readily on formate, hydrogen/carbon dioxide, methanol or carbon monoxide (J. Wiegel, personal communication). Acetogenium kiwi (Leigh et af., 1981) is a non-spore-forming thermophile with a fermentative metabolism and DNA G + C content more similar to C .formicoaceticum than to either C . thermoaceticum or C . thermoautotrophicum. Several other isolated homo-acetogens are not yet identified and some described species are probably homo-acetogens. Clostridium lactoacetophilum (Bhat and Barker, 1947) and C . sticklandii (Stadtman and White, 1954) are no longer in vogue but appear as likely candidates. Mesophilic clostridria, whose morphology differ from C . aceticum, but which ferment hydrogen/carbon dioxide, were isolated by Ohwaki and Hungate (1977) and by Zeikus (1980a). Samin et al. (1982) described a non-sporing, Gram-negative, mesophilic species that ferments hydrogen/carbon dioxide. Anaerobic spore-formers that ferment methanol to acetic acid, but which are quite distinct morphologically from B. methylotrophicum, have been isolated (Adams and Velzeboer, 1982; G. A. Zavarzin, personal communication). Notably, the spore morphological properties vary considerably among homo-acetogenic Clostridium species and B. methylotrophicum. Clostridium aceticum, C . thermoautotrophicum and C . formicoaceticum form terminal swollen phase white-refractile spores (Andreesen et al., 1970; Wiegel et al., 1981; Braun et al., 1981). Spore ultrastructure of these species is typical of other published Clostridium species and it includes a phase-bright cortex and multiple spore coat layers (Braun et al., 1981). However, we have not detected phase white-refractile spores in C . thermoaceticum type strain Fontaine (R. Kerby, unpublished observations). Rather, spores appear like swollen-clubs as in C . barkeri (Stadtman et al., 1972), and with limited phase-bright refractility, but with a definite dark external outline that delineates the spore shape. Spores of B. methylo trophicum are quite distinct morphologically from vegetative cells, they remain viable for a long time in spent culture fluid, and

+

METABOLISM OF ONE-CARBON COMPOUNDS

261

are resistant to heat treatment at 7580°C for 10 minutes (Zeikus et al., 1980b). The low heat resistance of B. methylotrophicum spores resembles that reported for Bacillus cereus, which does not even resist pasteurization (Han et al., 1976). Figure 7 illustrates the pleomorphic nature of B. methylotrophicum and shows the phase-contrast evidence for endospore formation in this species. The appearance of swollen cells and refractility alone are not sufficient to document spores. The appearance of cellular differentiation within a sporangium is also required because vegetative cells swell as a result of accumulating amylopectin as storage material (T. Thompson, unpublished observations). Notably, spores have an oval shape, are terminal and display phase blue-bright refractility. Figure 8 illustrates typical vegetative and sporulating cells. The endospores of B. methylotrophicum in thin sections (see Fig. 8) are not similar to those described in Clostridium or Bacillus species because they lack a spore cortex and spore coat layers. A considerable amount of taxonomic confusion resides in the homoacetogen group as a result of personal bias by investigators. However, this is common to the extremely speculative nature of this research discipline. Tanner et al. (1981) compared the 16s RNA oligonucleotide sequence homology of several homo-acetogens and non-homo-acetogens. They concluded that E. limosum-B. rettgeri, C . barkeri and A . woodii formed a group distinct from C. lituseburense, E. tenue and C . butyricum. This conclusion is quite in line with the value of using these results to forecast suprageneric relatedness among bacteria. However, they suggest that E. limosum and B. rettgeri are the same species, based on almost identical DNA G + C content, murein analysis and 16s RNA oligonucleotide sequence. This seems an erroneous conclusion because these procedures alone are not definitive in speciation, and the same approach when applied to other clearly different species (e.g. E. coli and Salmonella typhl) would also suggest a similar identity. It is clear that spore-forminganaerobes are quite distinct when the chemical composition of their wall is examined (Kandler and Schleifer, 1980; Weiss et al., 1981). These investigators indicate a separate phylogenesis for C. thermosaccharolyticum, C. barkeri, C. ramosum and C. pasteurianium. In the absence of available evidence, Genthner and Bryant (Seminar 1982 ASM meeting) concluded that E. limosum is identical to B. methylotrophicum, although they previously reported that E. limosum did not sporulate (Genthner et al., 1981). I disagree with their conclusion because E. limosum differs from B. methylotrophicum in that (i) B. methylotrophicum does not form detectable slime (indeed “limosum” means full of slime) and (ii) B. methylotrophicum has different vitamin requirements and nutritional versatility (T. Thompson and Tom Moench, unpublished observations). Clearly, a more detailed macromolecular analysis is necessary to distinguish properly the taxonomic differences, and similarities, of these two species. Nonetheless,

262

J. G. ZEIKUS

FIG. I. Phase-contrast photomicrographs of Butyribacterium methylotrophicum showing life cycle-dependent morphological differentiation. A, Logarithmic phase cultures showing branching (arrowed); B, mid-stationary phase cells illustrating swollen cells and refractility; C , late-stationary phase cells showing endospore formation; D, mature sporangium. From Zeikus et al. (1980). Bars denote 4 pm in A and B and 1.6 pm in C and D.

METABOLISM OF ONE-CAREON COMPOUNDS

263

FIG. 8. Electron photomicrographs of Butyribacterium~e~hylofrophicum illustrating vegetative and sporulating cell types. A, Vegetative cells grown on carbon monoxide as sole energy source; B, sporulating and non-sporulating cells. Abbreviations: RM, amylopectin reserve material in old vegetative cells; SI,initiation of spore formation showing unequal cell division; SM, spore maturation showing endospore development and mother-cell lysis. From unpublished observations of T. Thompson.

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the name suggested here for “E. limosum” is B. limosum. This follows from the results of Tanner et al. (1981) that “ E . limosum” is not clearly related to other Eubacterium species (the physiological diversity being cited above) and, most importantly, the finding that “E. limosum” has the same peptidoglycan cross-linkages present in B. methylotrophicum (0.Kandler, personal communication). It is worth stating again here that Barker noted the metabolic similarity between B. rettgeri and “E. limosum”, and suggested the inclusion of the latter species into the genus Butyribacterium (Breed et al., 1957). However, because of the discovery of unique spore structures in a new isolate, the Marburg strain (Zeikus et al., 1980b), the genus Butyribacterium was amended and a new species proposed, B. methylotrophicum. Whether these interpretations are accepted by the taxonomic community, however, is not clear because of the dissimilarity in logic employed in both the macromolecular and the classical approach to this scientific artform. The CI metabolism of homo-acetogens, like methanogens, is associated with high intracellular concentrations of corrinoids (Tanner et al., 1978; Lamm et al., 1980; Zeikus et al., 1980b). The only other carbon carriers implicated in homo-acetogen metabolism, at present, are pteridines which all derive from tetrahydrofolic acid as chromophore (Parker et ul., 1971; Tanner et al., 1978). By and large, the electron carriers involved in homo-acetogen metabolism have only been characterized in C. thermoaceticum and C . formicoaceticum (Ljungdahl et al., 1976) and include ferredoxin (Yang et al., 1977), rubredoxin (Yang et al., 1980), cytochrome b and menaquinone (Gottwald et al., 1975). Unlike methanogens, the carbon or electron carriers involved in homo-acetogen metabolism appear common to other similar substituents present in species from both the eubacterial and archaebacterial kingdoms.

B. ONE-CARBON METABOLISM

1. Clostridium thermoaceticum Metabolism Detailed knowledge on the biochemistry of CI transformation is limited to studies of C. thermoaceticum grown on hexose and not on one-carbon compounds. Wood (1952) proved that C. thermoaceticum accomplished a synthesis of acetate totally from carbon dioxide by showing that some of the acetate formed by glucose in the presence of [I3C]carbondioxide was two mass units heavier than unlabelled acetate. The first clues to the path of acetate formation from carbon dioxide came from the observation that [I4C]formate was selectively incorporated into the methyl group of acetate (Lentz and Wood, 1955). Tetrahydrofolate-bound intermediates were implicated by the

METABOLISM

OF ONE-CARBON COMPOUNDS

265

findings that methyl-tetrahydrofolate was converted into acetate by cell extracts (Ghambeer et al., 1971), and that methyl-tetrahydrofolate and 10-formyl-tetrahydrofolate became highly labelled after pulsing cells with ['4C]carbon dioxide (Parker et al., 1971). Formate was assumed to be an intermediate since formate dehydrogenase-carbon dioxide reductase activity was found in extracts which coupled NADPH oxidation to carbon dioxide reduction (Li el al., 1966; Thauer, 1972). The enzymes necessary to convert formate into methyl-tetrahydrofolate all required folate intermediates as carbon carriers (Andreesen et al., 1973). The most elusive part of the homo-acetogenic pathway is the mechanism of acetate synthesis from methyl-tetrahydrofolate; and this aspect is still not certain. Poston et al. (1964) showed that cell extracts converted [I4C]methyl-BI2into [2-I4C]acetatein the presence of pyruvate. Ljungdahl et al. (1965) demonstrated that ['4C]methyl-corrinoidsacquired a very high specific activity when cell extracts were exposed to [I4C]carbon dioxide and that [14C]acetatewas formed in the presence of pyruvate from either [14C]methylor [2-14C]acetyl-corrinoids. Corrinoids were further implicated in acetate synthesis by the finding that transmethylation via Co-methylcobalamin was inhibited by alkylhalides such as propyliodide (Ghambeer et al., 1971). Schulman et al. (1973) presented considerable evidence that carboxylation of methyl-corrinoids occurred via transcarboxylation of pyruvate, and not by direct fixation of carbon dioxide. This conclusion was based, in part, on observations that an a-0x0 acid was obligately required in this reaction and carboxylation was not dependent on the carbon dioxide concentration. Welty and Wood (1978) purified a corrinoid enzyme from C. thermoacetium, to 80% homogeneity, that catalysed the transmethylation of methyl-tetrahydrofolate to a methyl-enzyme complex and participated in acetate synthesis from methyl-tetrahydrofolate and pyruvate. Other homo-acetate pathway enzymes that have been purified from extracts include acetate kinase (Schaupp and Ljungdahl, 1974), and a formate dehydrogenase which contains selenium, tungsten and molybdenum (Ljungdahl, 1980). Figure 9 summarizes the present scheme prepared by Ljungdahl and Wood (1982) for glucose conversion into acetate. Although details of this scheme require further proof, it is clear that synthesis of acetate occurs via carrier-bound one-carbon reduction and transmethylation reactions. More emphasis should be placed on understanding CI metabolism at the level of oxidation of formate in C. thermoaceticum. For example, transcarboxylation may involve transcarbonylation, and the role of free formate needs better documentation. Thermodynamically, formate is not as favourable as a formyl intermediate, and it is known that a CI unit at the formyl oxidation state (i.e. carbon monoxide) can be directly assimilated into acetyl-CoA without oxidation to carbon dioxide. Recently, Drake et al. (1981) partially purified five components from crude

J. G. ZEIKUS

266

GLUCOSE

2H

2H

FIG. 9. Homo-acetogenic pathway proposed for Clostridium thermoaceticum grown on glucose. From Ljungdahl and Wood (1982). Abbreviations: THF, tetrahydrofolate; enz, enzyme; Co, coenzyme.

extracts of C. thermoaceticum that catalyse acetate synthesis from pyruvate. These components include (i) FI, a phosphotransacetylase, (ii) F2, a methyltransferase, (iii) F3, a carbon monoxide-dehydrogenase complex, (iv) F4, a pyruvate cleavage activity, and (v) ferredoxin. Of note, component F3 was purified 14-fold and shown to contain nickel (Drake et al., 1980). This finding supported the suggestion of Diekert and Thauer (1978, 1980) that carbon monoxide dehydrogenase activity required nickel. However, the partially purified carbon monoxide dehydrogenasecomplex was not inhibited by propyliodide, which is not in agreement with the results and conclusion of Diekert and Thauer (1978) that carbon monoxide dehydrogenase activity in clostridia is vitamin Bl2-dependent. Hu et af. (1982) demonstrated that acetyl-CoA was synthesized by component F3 and F2 according to the following reaction: [I4C]carbonmonoxide+methyl-tetrahydrofolate+ Coa ATP

-[

+

1-I4C]acetyl-CoA tetrahydrofolate

Thus, the direct role for methyltransferase (F2) and carbon monoxide dehydrogenase (F3) in acetate synthesis was established. The F3 enzyme complex requires further purification and elucidation; however, it appears to contain a corrinoid methyltransferase activity as indicated by the finding that propyliodide inhibited the exchange between carbon monoxide and [ 1-I4C]-

METABOLISM OF ONE-CARBON COMPOUNDS

267

acetyl-CoA. As a result of these findings, Hu et al. (1982) proposed a scheme for unicarbonotrophic growth of C. thermoaceticum on carbon monoxide. The carbon monoxide dehydrogenase activity functions in the generation of formate, carbon dioxide and reducing equivalents. A formyl intermediate is then reduced to methyl-tetrahydrofolate and transmethylated to methylcorrinoid by component F3. Acetyl-CoA results from the transcarboxylation of the methyl-cornnoid by factor F3. In short, acetyl-CoA, synthesized via carbon monoxide metabolism, becomes the precursor for further anabolic and catabolic reactions. However, their scheme fails to include a role for pyruvate which has not yet been eliminated as a key intermediary metabolite of homo-acetogens grown on one-carbon compounds. The following carbon flow scheme is an alternative proposal: 2CoA Methyl-B12c[HCOOH]tCO

CO+[HCOOH]

Lpyruva" 3c Acetyl-CoA

Acetyl-CoA

This speculative scheme is also novel in the context of energetic efficiency as it implies a formyl intermediate in lieu of free formate per se for methyl-tetrahydrofolate and/or pyruvate synthesis, and a substrate level phosphorylation site associated with net acetyl-CoA synthesis. 2. Acetobacterium woodii and Butyribacterium methylotrophicum Metabolism Very little is known about CI metabolism in homo-acetogens other than C. thermoaceticum. Schoberth (1977) showed that acetate was synthesized from hydrogen and bicarbonate in cell extracts of A. woodii, and reported that methylene blue was an artificial electron acceptor for hydrogenase. The C. thermoaceticum homo-acetate pathway was implicated in the synthesis of acetate by A. woodii because extracts contained high concentrations of corrinoids, formate dehydrogenase, formyl hydrofolate synthetase, methenyl hydrofolate cyclohydrolaseand methylene hydrofolate dehydrogenase (Tanner et al., 1978). Notably these studies also showed that methyl viologenlinked formate dehydrogenase activity decreased 20-fold when cells were grown on hydrogen/carbon dioxide in place of fructose. Finally, Braun and Gottschalk (198 1) showed that A. woodii was able to grow mixotrophicallyon hydrogen/carbon dioxide plus fructose. Butyribacterium methylotrophicum grows on hydrogenlcarbon dioxide, methanol/carbon dioxide/acetate, glucose or carbon monoxide as energy source (Zeikus et al., 1980b; Lynd, 1981; Lynd et al., 1982).The organism also grows mixotrophically on glucose or methanol in the presence of carbon

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monoxide, which replaces both carbon dioxide and acetate as the electron acceptor, and inhibits both hydrogen and butyrate production (Lynd et af., 1982). Table 5 illustrates fermentation balances and growth-yield values of B. methylotrophicum during unitrophic substrate metabolism. Notably, the formation of hydrogen from glucose and butyrate from hydrogen/carbon dioxide, occurs as late fermentation products but formate is not detectable. Yeast extract, present in the medium, was not a significant carbon or electron donor for the growth of B. methylotrophicum which can be grown on the above substrates in defined medium with a limited vitamin and trace metals mixture (T. Moench and R. Kerby, unpublished observations). It is important to note that the order of growth efficiency on C1 substrate was methanol> carbon monoxide > hydrogen/carbon dioxide. Carbon monoxide dehydrogenase, hydrogenase and formate dehydrogenase levels were compared in B. methyfotrophicum that had been grown on glucose, methanol or carbon monoxide as electron' donor. Notably, methyl viologen-linked carbon monoxide dehydrogenase activity was higher in cells grown on methanol than in cells grown on carbon monoxide, whereas formate dehydrogenase concentrations were higher in cells grown on carbon monoxide and lowet in cells grown on methanol. This finding, and the absence of free formate as a detectable intermediate, suggested the carbon and electron flow model indicated in Fig. 10. According to this speculative C1 transformation scheme for conversion of carbon monoxide into acetate, the principal role for TABLE 5. Comparison of net fermentation stoicheiometry and growth yields of Butyribacrerium methylotrophicum on different electron donors

Electron donor Glucose Methanol

Fermentation stoicheiometry (Pmol) 495 glucose-754 acetate+ 112 butyrate+ 117 Hz+136 COz+808 cell C 2742 methanol + 548 C02 + 197 acetate735 butyrate + 832 cell C 4431 CO-2403 C02+816 acetate+500 cell C

Carbon monoxide Hydrogen 3260 H2+ 1620 C02-+693 acetate+ 18 butyrate+ 174 cell C

Growth yield (g cell dry wt. (mole donor)-') 42.7 8.2 3.0

1.7

Experimental conditions: B. methylotrophicum was grown on basal salts medium that contained 0.05% yeast extract and either 100%carbon monoxide alone or a nitrogen/carbon dioxide gas atmosphere and growth-limiting amounts of either glucose, methanol/acetate/carbon dioxide or hydrogen. Data are from Lynd (1981). Growth yields were calculated during exponential growth.

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269

FIG. 10. Carbon and electron flow scheme proposed for conversion of carbon monoxide into acetic acid in homo-acetogens. The model predicts that carbon monoxide is converted into a formyl intermediate which is either oxidized to carbon dioxide via formate dehydrogenase activity, reduced to the methyl level or transmethylated via carbon monoxide dehydrogenase to yield acetate. Please note that any one arrow can represent more than one enzyme activity.

a formate dehydrogenase activity would be to couple formyl oxidation to methyl synthesis via a formyl reduction and carbon monoxide dehydrogenase would function primarily in transformylation of a methyl intermediate. Please note that this scheme does not eliminate pyruvate or acetyl-CoA as intermediary metabolites of acetate synthesis, as detailed above for C. thermoaceticum.The conversion of a formyl intermediate into carbon dioxide, methyl or acetate is viewed as a system of highly co-ordinated, multiple enzymes. Figure 11 is a model proposed here to illustrate the unification of one-carbon metabolism in B. methylotrophicum. By analogy with that which occurs in M. barkeri, both catabolism and anabolism in B. methylotrophicum would appear to share common CI transformations for the synthesis of acetyl-CoA (the direct precursor to acetate), butyrate and cell carbon. One-carbon substrates with different oxidation states presumably enter the path in different carriers, which is required to explain the growth efficiencies on these substrates. By analogy with that which occurs in C. thermoaceticum, synthesis of acetyl-CoA would appear to occur via a CI-CI condensation reaction by the reductive carbonylation of methyl-)<. The cells certainly contain high concentrations of corrinoids and carbon monoxide dehydrogenase, but detailed enzymic and I4C-isotope incorporation studies are required to establish that acetyl-CoA synthesis in B. methylotrophicum is parallel to that in C. thermoaceticum. It is worth noting that, unlike other methylotrophs that have been described (i.e. Methylococcus or Methanosarcina species), B. methylotrophicum requires an electron acceptor to grow on methanol because the fermentation products are all more oxidized than the electr6n donor (methanol). During mixotrophic

J. G. ZEIKUS

270

HOOC-X

f--,

COZ

OHC-X -CO

5 3

OH,C-X

H3C-XMCH30H

T CH,CO-S COA

BUTY RATE

ACETATE

CELL CARBON

FIG. 11. Unification of carbon flow pathways for catabolism and anabolism in Butyribacteriummethylotrophicum.The proposed model predicts C1 units are fixed on different carbon carries or enzyme-active sites (X). The key biosynthetic reaction involves acetyl-S-CoA synthesis via carbon monoxide dehydrogenase. Unicarbonotrophy is linked by a common carrier system for metabolism of different one-carbon substrates.

growth on methanol, carbon monoxide replaces the need for acetate or carbon dioxide as an electron acceptor (Lynd et al., 1982).The suggested biochemical basis for this phenomena is also indicated by analysis of the carbon flow scheme illustrated in Fig. 11. During growth on methanol/acetate/carbon dioxide, methanol must be oxidized to synthesize acetyl-CoA, to reduce acetate and carbon dioxide to butyrate, and to generate cell carbon. However, on methanol/carbon monoxide, both substrates could be assimilated directly to acetyl-CoA without the need for methanol oxidation. Thermodynamically, carbon monoxide oxidation is favoured over methanol oxidation for the generation of reducing equivalents (i.e. reduced electron carriers) at the redox state needed for both catabolic acetyl-CoA synthesis and anabolic pyruvate synthesis.

METABOLISM OF ONE-CAREON COMPOUNDS

271

3. Adenosine Triphosphatase Synthesis

Little is actually known, or even speculated, about the mechanisms of coupling ATP synthesis to one-carbon transformation in homo-acetogens. Nonetheless, it would appear that ATP is derived from both substrate level phosphorylation and electron transport-mediated phosphorylation, and perhaps by novel mechanisms. The high cell yield of C . thermoaceticum (40-50 g (mol glucose)- I ) shown by Andreseen et al. (1973) indicated a higher ATP gain (mol glucose)-' than via other anaerobic glycolytic pathways; and they postulated involvement of electron transport-mediated phosphorylation in addition to substrate level reactions. The growth yield of homo-acetogens on glucose is higher than for other glycolytic anaerobes (Stouthamer, 1979). The growth yield of B. methylotrophicum on glucose (i.e. 42.7 g (mol glucose)-') also supports the conclusions of Andreesen et al. (1973). However, the finding of Hu et al. (1982) that two mol of acetyl-CoA are derived from pyruvate and methyltetrahydrofolate invites the speculation already cited (p. 266) that the transcarboxylase reaction in homo-acetogens may be cyclic and may represent a new mechanism from substrate level phosphorylation (i.e. a transcarboxylase-dependent acetyl P synthesis). Thauer et al. (1977) indicated that the only non-nitrogenous, one-carbon transformation linked to substrate level reactions is the hydrolysis of formyl-tetrahydrofolate. Thus when homo-acetogens are grown on glucose, ATP synthesis via substrate phosphorylation may be associated with production of 2 mol ATP (mol glucose)- via glycolysis, 1 mol ATP (mol acetate)- I via transcarboylaseacetate kinase, 1 mol ATP (mol formate)- I via formyl-tetrahydrofolate synthetase, 1 mol ATP (mol acetate)-' via pyruvate dehydrogenase-acetate kinase, and 1 mol ATP (mol butyrate)-' via butyrate kinase. Thus, transcarboxylase can account for the high growth yield on glucose and electron transport-mediated phosphorylation may not necessarily be required. However, the ability of homo-acetogens to obtain ATP equivalents via a proton-motive force associated with product excretion, as shown in lactic-acid bacteria (Michels et al., 1979), also implies more than subs&ate level phosphorylation on glucose. The mechanisms of ATP synthesis, during growth of B. methylotrophicum on one-carbon substrates, is another matter, and it is clear that energy conservation must involve electron transport-mediated phosphorylation. Lynd et al. (1982) reported that growth yield on carbon monoxide (3.0 g mol-I) was consistent with the synthesis of 0.5 mol ATP (mol carbon monoxide)-' (i.e. 2 mol ATP (mol acetate produced)-I). This high cellular yield of ATP necessitates more than substrate level phosphorylation via

-

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J. G. ZEIKUS

acetate kinase alone. The carbon monoxide-dependent reduction of a formyl intermediate to the methyl level was postulated as the driving force of electron transport-mediated phosphorylation during growth on carbon monoxide (Lynd et al., 1982) because the free energy of the reaction: CO +CH20 + H20+CH30H + C02 (- 15.4 kcal; - 64.7 kJ) is sufficiently exergonic to be coupled to ATP synthesis via electron transport-mediated phosphorylation. Neglecting cell synthesis, the stoicheiometry observed for growth of B. methylotrophicumon methanol was (Zeikus et al., 1980a): 10 methanol + 2 CO2+3 butyrate. If butyrate were synthesized via transcarboxylase according to: 10 CH30H+2 C o p 6 acetyl-CoA+ 12 e12 e- 6 acetyl-CoA+butyryl-CoA 3 butyryl-CoA+ADP+P+3 butyrate+ 3 ATP

+

then the ATP gain (0.3 mol ATP (mol methanol)-') is below that needed to explain the,observed growth yield of 8.2 g cells (mol methanol)-' by solely substrate level (i.e. butyratl-acetate kinase) reactions (Stouthamer, 1979). Likewise, the growth yield on hydrogen/carbon dioxide (i.e. 1.7 g cells (mol hydrogen)-') implied that more than 1 mol ATP (mol acetate)-' is formed and electron-mediated reactions are involved. The lower cell yield on hydrogen/carbon dioxide than on carbon monoxide is consistent with the thermodynamic requirement for energy consumption in the formation of a formyl intermediate, because carbon dioxide lacks available carbon-bound electrons. V. General Metabolic Perspectives on Unicarbonotrophy

Analysis of the literature already reviewed, as a whole, provides some general metabolic perspectives on the mechanism of anaerobic CI transformation and how it differs uniquely from aerobic or phototrophic unicarbonotrophy. In the most general terms, anaerobic metabolism of one-carbon compounds is identified with maximal conservation of energy associated with transformation of C1substrates and enhanced catalytic efficiency. Both of these features are inherent to the early evolution of these bacteria and this unique mode of metabolism. In more specific biochemical terms, unicarbonotrophy of anaerobes involves direct reductive transformations of CI substrates and not indirect, oxidative CI transformations associated with aerobes and phototrophs that grow on one-carbon compounds.

METABOLISM OF ONE-CARBON COMPOUNDS

273

A . RELATION OF SUBSTRATE-PRODUCT THERMODYNAMICS

TO GROWTH EFFICIENCY

Figure 12 (p. 274) compares the energetics (i.e. based on an equivalent number of electrons per donor substrate) of one-carbon transformations associated with the metabolism of either hydrogen/carbon dioxide, methanol or carbon monoxide by M. barkeri and by B. methylotrophicum. Thermodynamic efficiency can not be compared directly to growth efficiency because it is independent of the biochemical mechanism of C I transformation, which itself is the controlling force that enables the coupling of useful energy from a chemical transformation reaction. Methanogens and homo-acetogens that transform one-carbon compounds obtain only a limited amount of potential energy content from the system compared with aerobes that utilize the same substrates. However, aerobes couple oxidation of hydrogen, carbon monoxide or methanol to reduction of oxygen; whereas anaerobes link these oxidations to the reduction of C1 substrates themselves. Therefore, the aerobes have a much higher degree of available energy (by AGwcalculations), but anaerobes are much more efficient in the conservation of energy available for cell carbon synthesis. It is worth noting that the inability to conserve energy, by aerobes, is inherent in their way of life. Oxygen, the electron acceptor that maintains the electrochemicalpotential gradient (i.e. life force) of aerobes, is itself chemically toxic and cellular energy must be expended in detoxifying both active oxygen species and their reductive products (e.g. peroxides). Thermodynamic efficiencies also are independent of kinetics, and CI transformation rates depend specifically on the properties of the biocatalysts involved. Thus, the low thermodynamic efficiencies of anaerobic C1 transformations are not necessarily associated with low reaction rates. On the contrary, the most prolific hydrogen oxidizing autotrophic species so far documented is M. thermoautotrophicum, which displays a doubling time of about 60 minutes in chemostat culture, while the rapidly growing aerobic knallgas bacterium Alcaligines eutrophus has a 192 minute doubling time (Friedrich et al., 1979). However, the aerobe is constantly faced with generation of low redox potential electrons, in the presence of oxygen, and it contains an active oxygenase (i.e. ribulose bisphosphate carboxylase) that re-oxidizes reduced cell carbon formed from net carbon dioxide fixation. Similarly, the fastest growing methylotroph is apparently not a “slow-poke’’ Methylococcus or Pseudomonas species,but rather B. methy lotrophicum which in a chemostat has a doubling time at wash-out of less than 30 minutes (T. Moench, unpublished observations). The equations shown in Fig. 12 also indicate that more favourable thermodynamic efficiencies are associated with the transformation of either

274

J. G. ZEIKUS AGo’(kcol per mol per 8 e-)

REACT1 ON

-

I. Methonosarcino borkeri A. 4 C O B. 4H,

+

2H,O

3C0,

-50.5

CH,

+

3H,O

-32.7

3 C H 4 + CO,

+

2H,O

-19. I

-

+ HCO;+

C. 4 C H 3 0 H

+

CH,

H*-

-

II.Butyribocferiurn rnefbylolrophicurn

+ 4H,O B. 4H, + 2C0, C. I O C H 3 0 H + 2HCO;A. 4 C O

CH3COO-+ 2HCO;+

-39.5

3H*

-27.6

CH3COO- H*+ 2H,O 3CH3CH,CH,C00-+

H*+6H20

-26.0

FIG. 12. Comparative energetics of C1 fermentation reaction stoicheiometries indicative of Methanosarcina barkeri and Butyribacterium methylotrophicum energy

metabolism. hydrogen/carbon dioxide, carbon monoxide or methanol by methanogens than by homo-acetogens. However, comparison of the growth yield values (g dry wt. cells (mol electron donor)-’) showed those of M . barkeri to be noticably lower than those of B. methylotrophicum (Lynd, 1981). For example, the yield value of B. methyfotrophicum during exponential growth was 8.2, while it was 3.9 for M . barkeri. This suggests that basic mechanistic differences exist between energy conservation from CI transformation reactions in methanogens and acetogens. This feature will be discussed more fully in Section VI B. Comparison of the thermodynamics of these CI transformation reactions with cell growth yields also supports and, indeed, implies the carbon flow scheme proposed in this review for unification of CI metabolism in M . barkeri and B. methylotrophicum. Notably, the thermodynamically favoured order for available free energy from CI transformations, in either methanogens or homo-acetogens, is carbon monoxide > hydrogen/carbon dioxide > methanol. However, the order of growth yields obtained with B. methylotrophicum was methanol > carbon monoxide > hydrogen/carbon dioxide (Lynd, 1981); and, methanol > hydrogen/carbon dioxide in M . barkeri (Weimer and Zeikus, 1978a). These facts suggest that carbon-bound electrons are conserved in methanol during cell carbon synthesis, and support the concept of direct assimilation of carbon at the methyl level and the synthesis of a two-carbon intermediate without the expenditure of energy in homo-acetogens and methanogens. Notably, the mechanism proposed here for acetyl-CoA synthesis in homo-acetogens shows that methylotrophic or autotrophic cell

METABOLISM OF ONE-CARBON COMPOUNDS

275

carbon synthesis reactions can actually make energy available. Furthermore, yields of anaerobic unicarbonotrophs are higher on methanol than on carbon monoxide because the substrate is more energy rich (i.e. it contains more carbon-bound electrons), and these bacteria have evolved biochemical mechanism for conserving carbon-bound electrons. Finally, one way to compare the thermodynamic efficiencies of metabolism among unicarbonotrophs is to compare ‘‘YkCa{’which Payne and Williams (1976) describe as the fraction of free energy conserved as cells divided by the energy removed from the medium. Thus, Lynd et al. (1982) used the following equation to compare aerobic and anaerobic unicarbonotrophs: thermodynamic efficiency of CImetabolism

=

(mol cells) (AGZ; cells).( 100) (mol substrate)-(AGE;of substrate) -(mol product).(AG$ product)’

By use of these calculations, the thermodynamic efficiency of carbon monoxide metabolism displayed by B. methylotrophicum (57%) approximated to that of the most efficient aerobic unicarbontrophs growing on methanol, a much more energy-rich substrate. Although comparable growth yields are not available for aerobic carbon monoxide-oxidizing bacteria, .the proposed biochemical mechanism for carbon monoxide oxidation (see Kim and Hegeman, 1982) is not consistent with the conservation of energy in carbon-bound electrons proposed for homo-acetogens. In short, anaerobes are more efficient than aerobes in C1 transformation because their metabolic machinery conserves more carbon-bound electrons. Furthermore, growth efficiencyof aerobic methylotrophs also depends directly on their ability to conserve carbon-bound electrons. Among the pathways described for aerobic methylotrophy (Quayle, 1972, 1980), more formaldehyde is assimilated via the xylulose monophosphate pathway (3 mol triose-I) than via the serine pathway (2 mol triose- I), and none via the ribulose bisphosphate pathway. Therefore, the differences in growth yield observed among methylotrophic species are directly related to the number of carbon-bound electrons, in methanol, that end up as cell carbon.

B . RELATION OF CHEMOAUTOTROPHIC ANAEROBES

TO PHOTOTROPHS AND AEROBES

In general, CItransformation reactions of chemotrophic anaerobes do not mechanistically resemble those of unicarbonotrophic aerobes or phototrophs. Notably, both phototrophic and aerobic species that metabolize one-carbon compounds are able to grow well under dark aerobic conditions, and by CI

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pathways not common to anaerobic unicarbonotrophs (Colby et al., 1979; Higgins et al., 1981; Kondratieva, 1979). The key mechanistic differences are that anaerobes utilize highly reductive C1 transformations while the aerobes and phototrophs employ more oxidative C1transformations, including those catalysed by oxygenase reactions. For example, methanol is directly oxidized to formaldehyde by a phenazine methosulphate-linked alcohol dehydrogenase, in Methylococcus species (Pate1 et al., 1973) or in Rhodopseudomonas species (Wilkinson and Hammer, 1974), whereas in M . barkeri, methanol is bound to a carbon carrier (e.g. CoM) and the methyl moiety is further transformed on other carriers (e.g. YFC) but cells lack alcohol dehydrogenase per se (Daniels and Zeikus, 1978; Weimer and Zeikus, 1978a; Shapiro and Wolfe, 1980; Kenealy and Zeikus, 1982b). Diverse aerobic methylotrophs, including Methylococcus (Whittenbury, 1981), Paracoccus (Cox and Quayle, 1975) and Rhodopseudomonas (Sahm et al., 1976) species contain ribulose bisphosphate carboxylase activity which, in itself, has an associated oxygenase activity that functions in a bioregulatory manner by oxidizing reduced cell carbon (Fritz, 1981). This feature is not consonant with a maximal conservation of energy. The phototrophic bacteria are too often generally regarded as anaerobes because their phototrophic mode of energy metabolism is inhibited by oxygen. However, Rhodospeudomonus species are facultative, and in the dark they utilize oxygen as the terminal electron acceptor and contain a complete TCA cycle (Kondratieva, 1979). Notably, in the light, the TCA cycle is used by certain phototrophs (Chlorobium species) in the reverse direction to synthesize cell carbon via coupling reductive carboxylation reactions with carbon rearrangement reactions (Evans and Buchanan, 1965; Fuchs et al., 1980; Ivanovsky et al., 1980). Further, metabolic similarities between aerobic methylotrophs and the phototrophic bacteria are also substantiated by the finding that Pseudomonus AM 1 (of single cell protein fame) displays photo-enhanced bacterial chlorophyll formation (Sato, 1978). Methanogens clearly do not display close metabolic similarities, in C1 transformation reactions, to those associated with unicarbonotrophic aerobes or phototrophs; but methanogen metabolism is mechanistically similar to homo-acetogenic metabolism in the conservation of carbon-bound electrons and in the constitutive presence of carbon monoxide dehydrogenase activity.

VI. Trends in the Significance of One-Carbon Transformations Knowledge gained from research on the anaerobic metabolism of one-carbon compounds is of tremendous importance to environmental, evolutionary and

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277

applied microbiology studies. One-carbon transformation activities are vital to understanding the biogeochemical cycling of organic matter in nature, and the biogenesis of fossil fuels (natural gas, coals and oils) in the biosphere. Anaerobic one-carbon metabolism is of great relevance to the early evolution of autotrophy and methylotrophy, and to the development of a complex biochemical apparatus for maximal conservation of energy in two phylogenetically unrelated unicarbonotrophs. Finally, the future opportunities for applications of anaerobic CI transformations in waste processing, or the synthesis of fuels, chemicals feed stocks and fine-grade biochemicals, indicate enormous potential for developing biotechnology.

A . ENVIRONMENTAL

Biomass, in the form of photosynthate, accounts for the majority of organic matter that enters the anaerobic zones of the biosphere. By and large, the sediments of aquatic ecosystems, including oceans, paddies, lakes, marshes, rivers and swamps, are the primary reservoir for organic deposition, and the niche for anaerobic CI transformation reactions in nature. Methane is by far the major CI substrate cycled in the biosphere (Vogels, 1979). In the absence of excess exogenous inorganic electron acceptors (for example, 02,N032-, NOz3-, Sod2- and S Z O S ~ -organic ), matter is decomposed by a complex microbial food chain into methane, carbon dioxide and water as the final end-products of carbon mineralization (Zeikus, 1977;Zehnder, 1978).At least four different trophic groups, which can be identified by the specificpathways they utilize for controlling catabolic carbon and electron flow, are important to the active decomposition of organic matter in anaerobic ecosystems (Zeikus, 1980a, 1982). These four groups include (i) hydrolytic bacteria, which ferment complex multicarbon compounds (e.g. saccharides, lipids and proteins) to a variety of end-products including acids (e.g. lactate, acetate and propionate), neutral compounds (e.g. ethanol and methanol) and hydrogen/ carbon dioxide, (ii) hydrogen-producing acetogens, which ferment alcohols larger than methanol, and organic acids larger than acetate, to hydrogen and acetate, (iii) homo-acetogens, which ferment multicarbon compounds, hydrogenlcarbon dioxide or one-carbon compounds into organic acids, via acetyl-CoA as an intermediate and (iv) methanogens, which ferment hydrogenlcarbon dioxide, one-carbon compounds and acetate to methane and carbon dioxide. As a whole, the function of this anaerobic food chain can be likened to a metabolic symphony of carbon and electron flow reactions. The direction of carbon and electron flow through the food chain is controlled, in part, by CI transformation reactions. In homo-acetogens, CI transformations accom-

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plish enhanced intraspecies energy conservation, since a homo-acetogen obtains more ATP for each glucose than a hydrolytic bacterium. Hydrolytic bacteria such as C. thermocellum gain more ATP via the conservation of energy associated with the utilization of phosphorylations in polymer metabolism (Ng and Zeikus, 1982). Hydrogen-producing acetogens, such as Syntrophobacter wolinii (Boone and Bryant, 1980)gain access to the anaerobic niche by utilizing substrates like propionate, in the absence of exogenous electron acceptors (e.g. Sod2-, N032-, S Z O ~ ~ but - ) , they need the activity of hydrogen-consuming species in order to function. Most importantly, the methanogens are the conductors of the metabolic symphony because their CI transforming reactions control both intraspecies and interspecies carbon and electron flow (Zeikus, 1980a, 1982). The bioregulatory roles performed by methanogens during anaerobic digestion are summarized in Fig. 13. As a consequence of C1 metabolism, methanogens influence the decomposition of organic matter by other anaerobes by removing toxic end metabolites, directing electron flow to limited reduced end-products, enhancing growth rates and yields, and by supplying essential growth factors (Zeikus, 1977, 1980a). The performance of these regulatory functions was recently demonstrated in cocultures of M . barkeri apd the hydrolytic bacterium C. butyricum grown on pectin as sole carbon and energy source (Schink and Zeikus, 1982). The coculture completely degraded pectin to methane and carbon dioxide. In addition, intermediary concentrations of hydrogen/carbon dioxide, methanol and acetate formed were simultaneously metabolized. The most important bioregulatory function of the methanogen, during pectin fermentation, was H + consumption, for the cessation of growth of C . butyricum in monoculture was due to the accumulation of H+. In coculture, the intraspecies electron flow of C . butyricum was directed by the interspecies control, elicited by M. Function perfarmed

L

PROTON REGULATION

Metabolic reaction

CH3COO-+ H-i

CH, +C02

Z ! ELECTRON REGULATICN

4H2 +C02+CH4 + 2H,O

m. NUTRIENT REGULATION

CH&OOH H2/C02 CH30H

Vitamins

Acids

Process significance Remaves a toxic metabdite

Increases thermodynamic efficiency and range of substrates metabolized

Stimulates growth of heterotrophs

FIG. 13. Bioregulatory functions performed by methanogens during anaerobic digestion of organic matter. From Zeikus (1980b).

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barkeri, to hydrogen in lieu of ethanol or butyrate. The alteration of electron flow, when coupled to methanogenesis, enabled C. butyricum to generate more energy in coculture than in the monoculture from an equivalent amount of substrate metabolized. Nutrient excretion by the methanogens was suggested because the coculture grew readily on pectin, in defined medium, but C. butyricum grew poorly in monoculture unless yeast extract was added. In aquatic surface sediment ecosystems, anaerobic digestion does not normally operate in the complete absence of exogenous electron acceptors, as interstitial waters often contain oxidized sulphur species as well as oxygen. MacGregor and Keeney (1973) demonstrated that methanogenesis in eutrophic lake sediments was inhibited by addition of excess sulphate. Martens and Berner (1974) reported that methanogenesis in marine sediments did not occur until sulphate was depleted. This information was used by Winfrey and Zeikus (1977) to test the hypothesis that the addition of excess sulphate to freshwater lake sediment altered the flow of carbon and electrons from methanogens to sulphidogens. We showed that hydrogen and acetate consumption continued in the presence of excess sulphate, but that carbon flow was altered and the [I4C]methaneproduced from [I4C]carbondioxide or [2-I4C]acetatewas decreased, whereas [I4C]carbon dioxide production from [2-I4C]acetateincreased. The major conclusions for these studies were novel at the time in that they indicated that methanogens and sulphate reducers competed for electron donors (acetate and hydrogen). These findings appear to have been misinterpreted by other investigators such as to indicate that sulphate reduction and methanogenesis are mutually exclusive in aquatic sediments (Kosnir and Warford, 1979; Smith and Klug, 1981a,b;Oremland et al., 1982a,b; K. I. Ingvorsen, personal communication). However, Winfrey and Zeikus (1977) demonstrated that, in the presence of excess hydrogen or acetate, methanogenesis continued even with excess sulphate added to the lake sediment. Our results revealed and established that the amount and rate of methanogenesis in situ depends on the amount of electron acceptors and donors available to the anaerobic digestion process. Note, Ingvorsen et al. (1981) established that under in situ conditions of sulphate concentration, both methanogenesis and sulphate reduction were very active in a eutrophic lake sediment, and both processes accounted for nearly the same amount of terminal carbon and electron removal. Methanogens were also shown to interact synergistically with sulphate reducers in sediments (Cappenberg, 1974; Cappenberg and Prins, 1974). The primary reason for non-competitive interactions between sulphate reducers and methanogens is that sulphate-reducing bacteria function as hydrogenproducing acetogens at low sulphate concentrations when their electron flow is directed by methanogens (Bryant et al., 1977). It is evident in nature that both competitive and synergistic metabolic behaviour is displayed by

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methanogens and sulphate reducers such that the population of these bacteria in situ, and the amount of acetate and hydrogen/carbon dioxide (i.e. major terminal electron and carbon donors) transformed to methane in situ ultimately depends on the environmental chemistry (available donors, acceptors, pH and salinity). However, the fact still remains that, in neutral freshwater sediments, the amount of sulphate largely regulates the flow of carbon and electrons to methane; whereas, in marine sediments the amount of electron donors is more important. When comparing carbon and electron flow patterns in different sediment ecosystems, it is imperative to document the amounts of carbon and electron donors/acceptors present in situ; failure to do this leads to unwarranted controversy since these environmental parameters vary tremendously within both freshwater or marine sediments. The importance of microbial sulphate reduction and methanogenesis in aquatic sediments is well established (Reeburg, 1969; Oremland, 1975; Strayer and Tiedje, 1978;Abram and Nedwell, 1978;Winfrey and Zeikus, 1979; King and Wiebe, 1980; Ward and Frea, 1980; Sorenson et al., 1981; Pedersen and Sayler, 1981; Zaiss, 1981; Mountfort and Asher, 1981; Barrat et al., 1981; Jones et al., 1981;Winfrey et al., 1981). The significanceof methanogensis in controlling environmental carbon and electron flow is itself influenced greatly by the sulphate concentration. Sulphate-reducing bacteria are more ubiquitous than methanogens in nature because they have a broader substrate range and derive energy via the consumption of hydrogen/carbon dioxide or acetate in the presence of excess sulphate, or via the production of hydrogen and acetate in the presence of litniting sulphate. In summary, sulphate reduction is a more biogeochemically significant process in marine sediments where sulphate is abundant (Martens and Berner, 1974), and methanogenesis is more significant in freshwater sediments where sulphate is limiting (Winfrey and Zeikus, 1977); however, neither process is mutually exclusive. Certain organic molecules are recalcitrant in anaerobic environments. These molecules accumulate and account in part for the diagenesis of fossil fuels. Methane is the most cosmopolitan fossil fuel and is generated in both marine and freshwater environments. It is worth noting here that certain carbon precursors are transformed to methane regardless of the environmental sulphate concentration. For example, methanol and methylmercaptan are transformed to methane in freshwater, marine and hypersaline sediments (Winfrey et al., 1977; Zinder and Brock, 1978; Oremland et al., 1982a; J. G. Zeikus, unpublished observations). Also, it should be noted again that acidogenic and sulphidogenic bacteria make methane as a trace gas. Lignin, the second most abundant biopolymer on earth, is not significantly biodegraded anaerobically into the precursors for C1 transformation reactions (Zeikus, 1981); thus, it accumulates and leads to the diagenesis of coals and peats.

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A review of the recent literature indicates that anaerobic methane oxidation is a conceptual phoenix. Microbiological studies reported that sulphate reducers, like methanogens (Zehnder and Brock, 1979a), produce methane (Postgate, 1979; Hatchikian et al., 1976) and transform methane (Davis and Yarbrough, 1966; Panganiban et al., 1979). Biogeochemical studies showed that methane is transformed in anoxic freshwater and marine sediments (Zehnder and Brock, 1980; Panaganiban et al., 1979; Reedburgh, 1969). However, the biospheric significance of anoxic methane oxidation is questioned here, on the basis of the biochemical transformation that accounts for methane transformation. Namely, is it a net biochemical oxidation-consumption reaction or a biochemical exchange reaction that accounts for its anoxic methane transformation? Surely, anoxic sediments, or bacterial cultures properly manipulated in a reaction chamber, will perform [14C]methane-[14C]carbon dioxide. This result is to be expected if methane-producing bacteria (methanogens, acidogens or sulphidogens) are present as catalysts because methane is formed biochemically via reduction of a methyl carrier that readily exchanges with other cell intermediates. One biochemical feature common to CI transformations of anaerobes is the active reversible exchange reactions that result in significant substrate transformation, but not total consumption. It is now clear that non-acetate fermenting methanogens (e.g. M . thermoautotrophicurn) make [I4C]methane or [I4C]carbon dioxide from [I4C]acetatevia exchange (Zeikus et al., 1975). Methanosarcina barkeri, grown on acetate, readily incorporates exogenous [I4C]carbon doxide into [I4C]methanevia exchange reaction, but it cannot make methane from carbon dioxide alone. Butyribacteriwn methylotrophicwn, during growth on carbon monoxide, can exchange 5% of [I4C]carbondioxide into [14C]carbonmonoxide but it cannot make carbon monoxide from carbon dioxide alone. Hence, the documentation of \I4C]methane+[l4C]carbon dioxide by mixed or pure bacterial systems in reaction chambers is not conclusive evidence by itself of net methane consumption as this is expected from exchange reactions. In marine sediments, where methane is depleted, the rate of [I4C]methane production in situ needs to be compared with the rate of [I4C]methane consumption in situ since exchange, and not oxidation, would be suggested if the methane consumption rate at the in situ pressure and gas concentration exceeds the production rate. In freshwater sediments, net methane oxidation was not detected in the presence or absence of excess sulphate (Winfrey and Zeikus 1977, 1979). Other studies (Panganiban et al., 1979; Zehnder and Brock, 1980), which demonstrated [I4C]methaneconversion into [14C]carbon dioxide can be mechanistically interpreted as exchange and not net consumption. In essence, the anoxic recalcitrance of methane accounts for the diagenesis of natural gas as a fossil fuel.

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B. EVOLUTIONARY

The purpose of this section is to contemplate the evolution of CI metabolism in relation to the carbon and electron flow pathways of anaerobes. An organism’s carbon and electron flow pathway is primary to the evolution of a life force because, as a whole, it represents the integration of biochemical activities which maintain an electrochemical potential gradient via the production and consumption of cellular energy. Environmental, thermodynamic and biochemical constraints imposed on methanogens and homoacetogens appear essential to the evolution of their unique mode of C1 metabolism. One question to be addressed is when did anaerobic CI metabolism originate? Certainly chemotrophic anaerobes were the first heterotrophs, methylotrophs and autotrophs on earth, as oxygen was not a significant component of the early microbial environment until it became a metabolite of photosynthesis. Recently, discussions on metabolic evolution have focused on the reducing conditions, and the significance of environmental hydrogenlcarbon dioxide and carbon monoxide on primitive earth (Hartman, 1975; Visser and Kellogg, 1978; Knoll, 1979;Walker, 1980;Hart, 1979; Halvorson and van Holde, 1980). Notably, the Oparian theory of life’s metabolic origin, from a heterotroph, was challenged by Woese et al. (1978). They suggested that autotrophs originated first because methanogens grow on hydrogen/carbon dioxide, and they are as deep phylogenetically (as ancient) as the kingdom defined as eubacteria (Woese and Fox, 1977). Hence, methanogenic archebacteria were suggested as the dominant organisms in the primeval biosphere (Woese, 1981). The validity of this concept will be addressed here by analysis of the environmental, thermodynamic and biochemical constraints of CI metabolism on homo-acetogens and methanogens. This scenario will start by comparing of hydrogen/carbon dioxide and carbon monoxide, just in case these substrates and not other complex organic goodies in the earth’s chemical soup were the substrates that early microbes consumed. Thermodynamically, carbon monoxide is a better carbon and electron donor than hydrogen/carbon dioxide for methane or acetate synthesis yet only homo-acetogens grow readily on carbon monoxide, and their growth yield is greater on carbon monoxide than hydrogen/carbon dioxide. Thermodynamically, methane is the preferred product of hydrogen/ carbon dioxide consumption, not acetate; yet homo-acetogens have a higher growth yield than methanogens at neutral pH value. Biochemically, the carbon and electron flow pathway for hydrogen/carbon dioxide metabolism is simpler in homo-acetogens because more common carriers are involved in both anabolism and catabolism (e.g. tetrahydrofolate derivations and

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corrinoids). Methanogens lose metabolic efficiency and must direct methyllevel carbon from common intermediary pools to methane via additional unique biochemical components (i.e. CoM and nickel tetrapyrrole). In methanogens, most of the CI intermediates formed are directed away from acetyl-CoA synthesis, while in homo-acetogens, acetyl-CoA is also a catabolic intermediate. In short, homo-acetogens, as members of the highly divergent eubacterial kingdom, could have preceded methanogens because of their biochemical simplicity and efficiency. Hence, homo-acetogens may have preceded methanogens and predominated on early earth. It is of interest to note again that cell synthesis reactions involving two or more carbon atoms may have a similar biochemistry in both homo-acetogens and methanogens. Nonetheless, methanogens would certainly have evolved early on earth, but not only to share in the consumption ofjuvenile hydrogen then present, or the hydrogen generated via heterotrophic fermentation of the complex organic soup, by more metabolically primitive anaerobes. Most importantly, and what points to the highly specialized aspect of methanogen metabolism, is their ability to remove and not generate toxic protons. The growth yield values reported (Taylor and Pirt, 1977) for methanogens (about 0.6 g mol hydrogen-') are much lower than for homo-acetogens (about 1.7) grown on hydrogen/carbon dioxide, which indicates more ATP equivalents generated via substrate and electron-mediated phosphorylation and acetic acid excretion mechanisms in the homo-acetogens. However, both homo-acetogens and other heterotrophic anaerobes that produce acids, alcohols or hydrogen/carbon dioxide would have ceased to exist if methanogens had not evolved to consume acetic acid and hydrogen. The methanogens lack sufficient mechanisms to couple maximal energy conservation from C1 transformation reactions. Nonetheless, the niche for methanogen metabolism was established very early because of their unique affinities for hydrogen and acetic acid and their ability to couple catabolic methyl-transfer reactions to proton consumption instead of proton production. Perhaps this physiological function is even associated with evolution of the unique membrane structural features of methanogens. An archetype methanogen can be visualized as being biochemically less sophisticated than a heterotrophic species growing on acetate with limited carbon and electron carriers. Methunosurcinu barkeri, or M . thermouutotrophicum, are too metabolically complex to serve as primitive methanogen models. Now that some common biochemical features have been identified in both homo-acetogens and methanogens, it will be of future importance to analyse protein and coenzyme structural and functional relatedness, especially those associated with C, transformation reactions (e.g. carbon monoxide dehydrogenase and corrinoid-dependent enzymes) and cell carbon synthesis (e.g. TCA cycle enzymes, pyruvate synthase and alanine dehydrogenase). This analysis

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may shed more light on the metabolic evolution of these bacteria than comparison of their basic macromolecular architecture which places them into separate kingdoms. This biochemical approach has provided many interestinginsights as to the evolutionary relatedness of other diverse bacteria (DeLey and Kerstern, 1975). Present day homo-acetogens are more complex biochemically than other simple fermentative heterotrophs that lack C1 transformation reactions and the ability to grow unicarbonotrophically. If the complex chemical soup hypothesis, and heterotrophy, was indeed the direction in which metabolic evolution proceeded, then it is likely that autotrophy and methylotrophy arose from less complicated heterotrophs via the acquisition of several key biochemical activities that transferred CI units to tetrahydrofolate and corrinoid coenzymes; namely, formyl oxidoreductases,methyltransferaseand carbon monoxide dehydrogenase. Clearly, many anaerobic species, including methanogens and sulphidogens, are not autotrophs because they require acetate as the carbon source for growth on hydrogen. Likewise, some homo-acetogens and methanogens contain carbon monoxide dehydrogenases but do not grow as autotrophs or methylotrophs. Hence, several key enzyme activities are co-ordinately required for methylotrophy and autotrophy. All anaerobic methylotrophs described to date are autotrophs. Perhaps methylotrophy arose later because of the absence of this growth substrate on primitive earth. The origins of autotrophy and methyltrophy are closely related to evolution of common CI transformation reactions in anaerobes that use carrier-bound intermediates. In addition to thermodynamic considerations, this feature eliminates toxicity problems (e.g. formaldehyde) and maximizes efficiency by eliminating the loss of free CI intermediates into the environment. Catabolic and anabolic carbon and electron flow during unicarbonotrophic growth of anaerobes is tightly coupled because CI units are assimilated and dissimilated at the methyl and formyl levels. Thus, pathways for methylotrophy and autotrophy in chemotrophic anaerobes do not bear a direct resemblance to aerobic mechanisms of unicarbonotrophy. In fact, carbon and electron flow pathways of aerobic-phototrophic autotrophy are cyclic, and not coupled by common intermediates. Lack of redox coupling between anabolic and catabolic electron flow is regulated, in part, by the oxygenase function of the ribulose bisphosphate carboxylase of the Calvin cycle, which makes this a futile but not a primitive autotrophic pathway for cell carbon synthesis. The evolutionary aspects of methylotrophy in aerobes are especially interestingwhen the ribulose monophosphate cycle of formaldehydeassimilation is considered as a template for carbon dioxide fixation by the ribulose bisphosphate pathway (Quayle and Ferenci, 1978). The formaldehyde assimilation path is more efficient because it conserves carbon-bound

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electrons in cell synthesis, but it does not display a tight redox coupling between anabolic and catabolic electron flow; this is especially evident when cells are grown on methane and free methanol and formaldehyde accumulate. Notably, the serine pathway of CI assimilation in aerobic methylotrophs shares the same mechanistic features with the glycine decarboxylase pathway of heterotrophic anaerobes. That is, serine is formed as an intermediate of one-carbon metabolism via formyl-tetrahydrofolate, but these two pathways cannot generate a two-carbon intermediate via a CI-CI condensation reaction. Methanol is a good CI substrate to compare the biodegradative strategies evolved for aerobic versus anaerobic C1 metabolism. In the presence of excess reducing potential, aerobes will continue to dissimilate carbon-bound electrons in methanol because of their more oxidative mechanism of formaldehyde and CO;!assimilation. Methylotrophic anaerobes, on the other hand, when grown mixotrophically on two electron donors, will conserve more carbon-bound electrons in methanol because of methyl and formyl assimilation. Thus, anaerobes are much less wasteful and have evolved more advanced mechanisms of energy and carbon conservation. In closing, the CImetabolism of methanogens and homo-acetogensevolved well before aerobic pathways because of the conservation of energy associated with reductive one-carbon transformations. Autotrophy and methylotrophy, in any organism so far described, is not a primitive character but it is a biochemically complex metabolic mode. In both homo-acetogens and methanogens the evolved growth mode sequence implicated by biochemical symplicity is heterotrophy-+autotrophy+methylotrophy. Unicarbonotrophic aerobes that employ reductivecarboxylations for carbon dioxide fixation via the reverse TCA cycle, and those that assimilate formaldehyde, are displaying resemblances to anaerobic pathways. Perhaps if aerobes evolve better mechanisms to eliminate the toxic effects of oxygen on cellular metabolism this will also be associated with more evolved mechanisms for conservation of carbon and electrons during growth on CI substrates. C . BIOTECHNOLOGICAL

Useful applications of anaerobic CI transformation reactions can be suggested in waste treatment, fermentation technology and applied biocatalysis. One-carbon metabolism has been used in a practical sense for over a century in the treatment of municipal, industrial and agricultural wastes by methanogenic digestion (Hughes ef al., 1982). The thermodynamic, kinetic and stoicheiometric features of one-carbon transformations displayed by homoacetogens imply a potential role for their biocatalysis in the development of a CI substrate-based fermentation industry for chemical feedstock and bio-

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I

WASTES RESIDUES CROPS-WOOD

1

I

I

COAL PEAT

I

I

PYROLYSIS CHEMICALS PRETREATMENT

Hz/CDz.

CO

cn30n

ANAEROBIC FERMENTATION TECHNOLOGY

I

I

I

FEEDSTOCKS VlTAMl N S ENZYMES

METHANE SOLVENTS GASES

ETHANOL

FOODS-FEEDS

FIG. 14. Relation of anaerobic one-carbon fermentations to developing technology for chemical and fuel production from renewable or underutilized resources. From Zeikus (1980a).

chemical production (Zeikus, 1980b). Figure 14 shows diagrammatically the role of CI metabolism in the development of chemicals and fuel production technology via anaerobic fermentation of underutilized or renewable resources. Biomass wastes and residues often need extensive pretreatments to be useful as a starting substrate for the fermentation industry. Pyrolysis of coals and peats or underutilized biomass components (i.e. lignocellulose) can be used to generate CI substrates (i.e., hydrogen/carbon dioxide, carbon monoxide and methanol). Syngas (hydrogen and carbon monoxide) has been considered as the most abundant substrate for the future chemical synthesis industry (Pruitt, 198 1). Practical applications of aerobic CI metabolism have been largely limited to single-cell protein production (Tami and Yamada, 1980).Pyrolysis chemical fermentations have not been applied in practice, but their potential in transformations for chemical feedstock, fuels or biochemicals production will be assessed below.

1. Methanogens

Methanogens perform a pivotal role in anaerobic digestors used for waste

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treatment because they enhance the thermodynamic efficiency and rates of organic destruction (Zeikus, 1982). The vendibility of anaerobic digestion lies in the economic treatment of wastes, and not in the production of a fuel for society’s needs. Digester designs have only changed recently to increase the catalytic efficiency of the mixed bacteria population, and to treat a broader variety of liquid wastes without the need of an aerobic pretreatment process (Hughes et al., 1982). Practical improvements in the biocatalytic activity of CI transformation reactions are needed, especially improvements in the rate of acetate transformation to methane, and enhancement of methane yields by channelling carbon and electron flow away from sulphidogenesis. Verstraete et al. (1981) have suggested an improvement in anaerobic waste treatment by process separation into acidogenic and methanogenic stages in order to accomplish greater stability and higher rates of transformation. Healy and Young (1979) have employed alkaline heat pretreatment of biomass to improve methane yield from lignin-derived soluble aromatic fermentations. Wise et al. (1978) proposed a novel process, biomethanation, to upgrade the thermal (BTU) content of coal gassification mixtures, and to remove toxic carbon monoxide via the biocatalytic application of methanogens in high pressure reactors. Vitamin B12 fermentations, as a byproduct of waste-treatment processes, or the actual development of a methanol-based fermentation process employing M . barkeri as catalyst, are of interest. The latter suggestion requires that the growth yield (g 1-I) of M . barkeri be enhanced tremendously. At present, more emphasis needs to be placed on the fundamental growthenhancement studies now in progress (Scherer et al., 1981). 2. Homo-acetogens Homo-acetogens display more potential biocatalytic applications for anaerobic C1 transformation reactions than methanogens. The potential products of homo-acetogenic metabolism are broader and more valuable. Clostridium thermoaceticum fermentations have been suggested as a novel process for the conversion of biomass-derived sugars into acetic acid (Wang et al., 1978).Two major biotechnological problems arise with normal fermentations of homoacetogens, low species tolerance to end-products, and ineffective productrecovery systems. Recently Schwartz and Keller (1982) have reported on new strains that are active at pH values below 5.0. However, economic process technology has not been demonstrated for recovery of dilute organic acids ( < 4%) from fermentation liquors. The recent discoveries of homo-acetogenic C1fermentations promise lower substrate cost and perhaps novel process design with immobilized cells and high pressure reactors. The only identified acid products of industrial use are

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acetic and butyric acids. Notably, a novel product recovery method, and a potential use for the homobutyrate fermentation of B. methylofrophicum,has been suggested (R. Datta, personal communication). Namely, to esterify chemically the fermentation liquor to yield methylbutyrate which enhances end-product recovery. Methylbutyrate represents a chemical feedstock for plastics. It is also more soluble in gasoline and has a higher octane rating than ethanol. Perhaps more exciting products can be developed from the CI metabolism of homo-acetogens by regulatory or genetic manipulation of their biochemical activities. The most notable feature to pursue is an alteration in their normal electron and carbon flow to form new products. The catabolic electron flow potential exists to engineer biochemically acetyl-CoA reduction to ethanol and butyrl-CoA reduction to butanol. Likewise, since acetyl-CoA is the direct precursor to acetic acid or cell synthesis, the metabolic potential exists to biochemically engineer carbon and electron flow away from acetate to the formation of 0x0 acids and amino acids. Unlike aerobes, homo-acetogens have evolved highly efficient mechanisms for C1 transformations and also for product excretion. Now, it will be of interest to see if genetic and biochemicaLengineering can be put to practical use in applied biocatalysis of homo-acetogens for chemical feedstock, fuels, biochemicals or enzyme production.

MI. Acknowledgements and Dedication The literature search was completed in January 1982 with access to pre-prints of 1982 references. The research was supported by the College of Agricultural and Life Sciences, University of Wisconsin-Madison and by grants from The National Science Foundation. The Department of Energy, Exxon Research Education Foundation, Shell Oil Company, the Alexander Von HumboIt Foundation and the Institute Pasteur Foundation. It is a pleasure to acknowledge the results and many ideas my students contributed to the general understanding of CI metabolism in anaerobes; these include Dr. L. Daniels, Dr. P. Weimer, Dr. D. Nelson, Dr. M. R. Winfrey, Dr. W. R. Kenealy, Dr. B. Schink, Dr. T. Moench, L. Lynd, T. Thompson, J. Krzycki, T. Phelps and R. Kerby. The excellent technical assistance of J. Lobos, P. Hegge and J. O’Brien were essential to our studies. I thank J. Krzycki, R. Kerby, T. Moench and T. Phelps for critical discussions and comments on specific aspects covered in this review. This review is dedicated to the hard work and findings of my competitive colleagues. Understanding CI transformations in methanogens and homoacetogensis an inherently difficult task because of the complexity of metabolic

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experimentation in the absence of oxygen. Equivalent experiments that can be accomplished in days with aerobes often requires weeks, months or even years of hard work with these anaerobes. REFERENCES

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The Surface Stress Theory of Microbial Morphogenesis ARTHUR L. KOCH Department of Biology. Indiana University. Bloomington. Indiana 47405. U.S.A. I. Introduction . . . . . . . . . . . . A. Caveats . . . . . . . . . . . . XI. Methods . . . . . . . . . . . . A. Soap bubbles . . . . . . . . . . . . . . . . . B. The mathematics of narrow zonal growth C. Diffusegrowth . . . . . . . . . . . D. Problems of electron microscopy . . . . . . . . E. Analysis of autoradiograms . . . . . . . . . I11. Results . . . . . . . . . . . . . A . Gram-positive rods . . . . . . . . . . B. Gram-negative rods . . . . . . . . . . IV. Discussion . . . . . . . . . . . . A. What shape ought a bacterium to have? . . . . . . B. Stress on peptidoglycan covalent bonds . . . . . . . C. Surface stress theory for cylindrical elongation . . . . . D . Pole formation . . . . . . . . . . . E. Where do the conserved and non-conserved regions join in Gram-positive ~~

rods? . . . F. Variable T mechanisms V. Summary . . . VI . Acknowledgements . . References . . .

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I Introduction

..... and ever since that day. 0 Best Beloved. all the Elephants you will ever see. besides all those that you won’t. have trunks precisely like the trunk of the ’satiable Elephant’s Child.” Rudyard Kipling ADVANCES IN MICROBIAL PHYSIOLOGY. VOL. 14 ISBN 0-12-017724-7

Copyrtghr 8 1983 Academic Press London in my/onn reserved

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The Soap Bubble Theory of bacterial morphogenesis is now old enough that the formal name of Surface Stress Theory is more appropriate. The theory provides an explanation for the shapes of several kinds of bacteria and also provides an explanation for the forces that lead to elongation of rods and to bacterial cell division, without invoking a contractile element. Such elements are clearly present in eukaryotic organisms, but, in spite of careful searching, they never have been found in prokaryotes. Bacteria exhibit three cardinal morphogenetic movements that seemingly should require either a contractile element or Divine intervention. These are division, achieving a non-spherical shape, and segregation of chromosomes. The surface stress theory offers an alternative explanation for each. The theory depends on four postulates. 1. The Cell Bridges Across Stressed Murein Before Cutting

The bacterial cell resists the hydrostatic pressure arising from the osmotic difference across the cell wall because the murein forms a covalently closed, “bag-shaped macromolecule”. In living cells, in the direction normal to the wall, the forces usually cancel. In growing regions they must precisely cancel. However, in both growing and non-growing regions the sacculus supports considerable tensile stress tending to stretch it. This tension develops even though the peptidoglycan is porous because the murein network has, within it, spaces sufficiently small to prevent evagination of the cytoplasmic membrane. Consequently, the full force of the hydrostatic pressure is transmitted from the relatively impermeable cytoplasmic membrane to the murein layer. In Gram-negative organisms, the periplasmic space contains proteins that also are osmotically active and also develop a hydrostatic pressure against the outer membrane which tends to swell up and sometimes to produce blebs and vesicles. But normally, the tension from the outer membrane is transferred to the murein and the outer membrane remains intact. The stress is transferred via lipoprotein and other bridges between the outer membrane and the murein. This is to say that the osmotic pressure of the entire cell is contained almost totally by the peptidoglycan network by transmission of those forces from the more fluid cytoplasmic (and outer) membranes. The tension developed in individual glycoside and peptide links within the wall depends on the local concentration and arrangement of stress-bearing bonds. Simple calculations, given below, show that the forces are sufficient to lower appreciably the negative free energy of hydrolysis and the energy of activation for hydrolysis. The consequence of the latter fact is that the hydrostatic forces favour spontaneous hydrolysis that could rip the wall if it becomes sufficiently weakened. Consequently, it is assumed that the biochemical mechanism for

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wall growth of bacteria operates, in general, by covalently bridging appropriate parts of the wall before making the covalent cuts that allow the sacculus to enlarge. This means that when the cuts are made, the expansion geometry will be governed by simple physical laws. Although simple from a physical point of view, these mechanisms are sophisticated biochemically: the cleavages must be made, in a co-ordinate way, of only those bonds that no longer are necessary for the integrity of the cell. Only as a consequence of previous bridging, in providing the new murein to accept the stress, do cleavages of stress-bearing wall normally take place. The mechanisms are also very sophisticated from the biological point of view in that growth is permitted to take place only in specific regions of the cell. The growth pattern is different in the various kinds of bacteria; it is the assumption of this first postulate that allows the analysis of the modes of growth of the differently shaped bacteria. 2. Hydrostatic Pressure Forces Cell Expansion A second consequence of the tension developed in the covalently closed sacculus of the living bacteria is that it provides a way for the cell to couple the formation of an enlarged wall to its success in the synthesis of new macromolecules. Thus, as the cell grows, the amount of cell materials increases because of biosynthesis and active transport. This would tend to increase the pressure, hence the tension in the wall. This, in turn, leads to a decrease in the energy of activation for the hydrolysis (or transglycosylation or transpeptidation) of bonds bearing stress relative to the same types of bonds in less stressed configuration. The action of such stress-activated enzymic activities must be related so that the only bonds that are cleaved are in positions in the wall that are protected by the unstressed murein. As mentioned, this requires either a high degree of specificity and selectivity on the part of the enzyme or the control of synthesis or its activity. A consequence of this postulate is that wall expansion normally will be coupled to growth, and the tendency for the hydrostatic pressure to increase will be countered by the expansion. Thus, a cleavage enzyme sensitive to the stress of the murein substrate is a device that tends to maintain cell pressure. The theory further supposes that momentary surges of pressure may serve as signals to the cell to initiate a new growth pattern leading to cell division. Of course, this postulate does not apply to the “square bacteria” (Stockenius, 1981) which apparently do not have a large pressure differential across the cell wall.

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3. Cell Division Results from Surface Stress Forces The effect of postulates 1 and 2 are that expansion of the wall behaves as would a soap bubble with the same geometric constraints as does the growing region of the cell. The pressurevolume work (symbolized as Pdv) is coupled to the work of extending the surface in the same way as when a soap bubble expands and acts to form new surface against the molecular forces that resist increase in the amount of surface. In the case of the living bacteria, the Pdv work functions to force the bonds in the murein to have an extended conformation. In this way the Pdv work becomes converted into an equal amount of work but now work of expansion. The latter is symbolized by TdA, where T is surface tension and dA is the increase in area. The analogy of the two cases isjust an analogy, and the surface tension, in the context of bacterial growth, while dimensionally the same, does not have the same physical basis as in inanimate surface films. Soap bubbles are quite different from bacteria. In particular, soap bubbles do not divide without an external additional source of energy. But the study of the soap bubbles as an analogue computer, and the use of digital computer simulation of various equations originallyderived with a soap bubble in mind, have been invaluable in analysing the problem of bacterial morphogenesis. It is the difference between the bacterial systems and the soap bubbles that provides the possibility of self-division.There are basically two mechanisms in prokaryotes. In one there is formation of cross walls (septa) inside the cell under conditions where the tensile forces in the surface are not relevant to their formation, followed by splitting of the septum with the variable extension of the previous septa1 murein. This means that the energy relationships are different from those with ordinary soap bubble systems and can result in spontaneous division. Ultimately, the extra energy needed for cell division derives from the biochemical energy of the wall formation during septum formation. The alternative mechanism depends on the biophysics of soap bubbles and leads to a constrictive type of division. This mechanism requires that T varies in such a way that constriction takes place spontaneously as the stress-bearing wall grows. [Biochemical changes that could affect T are to be considered elsewhere (A. L. Koch, C. L. Woldringh and I. D. J. Burdett, unpublished observations).] This requires a local decrease in T and may, by itself, lead to cell division or to at least a sufficient degree of constriction so that other mechanisms can finish the closure and carry out the cleavages of covalent bonds connecting the two daughter cells. Neither alternative requires external forces or internal scaffolding, nor macromolecules capable of exerting mechanical work on other macromolecules or cell structures. Note that

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gradations between the two are possible. If the septum is not closed when external cuts place stress on the septum, then, to varying degrees, stress will be felt on the septum's internal face where wall material is being added. 4. Cell Shape and Dimensions are Determined by the Cells' Control of Regions

of Growth and Changes in Biochemistry of Murein Formation

Several patterns of bacterial development are possible and are catalogued in Fig. 1. Heavy stippling indicates regions of wall growth. Probably most of

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FIG. 1. Patterns of growth of microbes. (a) Centripedal septal growth and reworking only at site of splitting; (b) centripedal septal growth and diffuse growth over the nascent pole; (c) constrictive growth occurring because of localized changes in biochemistry that lead to less pressure-volume work being needed for the formation of a unit of external wall area; (d) diffuse cylindrical elongation; (e) cylindrical elongation by narrow zonal growth; (f) hyphal extention resulting from apical growth; and (g) bud formation by localized wall growth.

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these occur in nature. In Fig. l(a) and l(e), growth is assumed to be narrowly zonal; and outside of those regions of the cell, growth simply does not occur. For those organisms where growth appears to be zonal, it has been sufficient to assume that within the narrow growth zone, T is constant throughout its life. In Fig. I(b) and l(d), growth has been assumed to be constant over a larger region (diffuse growth). In some of the cases of diffuse growth, it has been sufficient to assume that T is constant throughout the growth process; but for others, as depicted in Fig. l(c), (f) and (g), the surface tension must vary during the growth process. In Fig. l(f), T is always small at the apex of a growing hypha and becomes progressively smaller behind the growing tip. In Fig. l(g), T is small when the bud starts, but progressively decreases with time. In this case, because the bud remains spherical, T remains constant all over. Figure 1(c) shows the model for constrictive division of Gram-negative rods. In this type of growth, a V-shaped invagination (constriction) forms and progressively divides the cell into two. Although the profile, as viewed from the outside of the cell, may present a less invasive furrow than electron microscopic examinations of the stress-bearing peptidoglycan, the peptidoglycan is never present as a double-thick structure that is later split, as in the classical splitting of the septum in Gram-positive cells (as shown in Fig. la). The essential difference from the point of view of surface stress theory is that the peptidoglycan, during the constrictive division, supports stress very shortly after it is laid down. For the case in Fig. l(a), a septa1 wall becomes stress-bearing only during the splitting process that exposes it to the surface. Since this series is devoted to advances instead of reviews, the published work on the Surface Stress Theory will not be repeated here except to update material and make corrections. The relevant papers are: Koch et al. (1981a,b, 1982a,b) and Koch (1982a,b,c): they are marked with asterisks in the references. This article will not deal with the structure of the prokaryotic wall; for details the reader is referred to Henning (1975), Di Rienzo et al. (1978), Mirelman (1978), Sargent (1978), Rogers (1979) and Rogers et al. (1980).

A . CAVEATS

While the theory provides mechanisms for the generation of many of the morphological forms of microbes, it raises at least as many questions as it solves. These will be considered below. There are certain questions outside its scope: it does not address the shapes of spirochetes, square bacteria, mycoplasma with non-spherical shape and higher fungi, nor viruses. It provides an explanation for the stalks of Gram-negative bacteria, but not for other cellular appendages. The model, as presented here, gives no role for asymmetrical wall growth; but, on the other hand, it does not prohibit such a

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mode of growth. Likewise, no role is allotted for a precise copying process whereby an exact amount of new murein is inserted to maintain the diameter of rod-shaped bacteria. Although girdles of peptidoglycan interlinked by peptide bridges might serve as templates in the short run, in the long run it is the physicochemical properties of the cleavage enzyme, and the detailed energetics of external wall formation, that determines the width of the cell. There is an exception to this last statement: the theory assumes for Gram-positive streptococci that the wall band splits into two bands of exactly the same radius. The length of rod-shaped cells is not addressed by the model except to suggest that there may be a maximum length possible for regions of diffuse wall growth. When the cylinder length is equal to the circumference, then further growth can lead to width divergence, although kinetic factors can increase the stable length (see Thompson, 1942).

11. Methods The physical intuition of the surface stress theory is that biosyntheticenzymes can link molecules only if the chemical reaction is exergonic and the component parts at hand. What an enzyme cannot do is mechanically move or cause a molecule to achieve an energetically improbable conformation. An enzyme is also forbidden, by the second law of thermodynamics, to wait only until the molecules are in extended conformation and then to link them. Therefore, the murein polymer, as formed, must have the statistical array of conformations that depend on physicochemical situation. Usually they become extended when stressed. The extension causes more area to be covered. From model building, there could be a great increase in area. Assumptions of a two-fold increase is both modest and realistic (Oldmixon et al., 1974; Formanek et ul., 1974; Braun et af., 1973). For some of the cases considered below, a two-fold expansion fits the biological facts, but in some others the expansion factor must be much smaller. The problem of bacterial morphogenesis is the how, when, and where of the biosynthesis, as well as the cleavages that transfer the stress. Possibly the most significant advance since the first paper on surface stress theory (Koch et af., 1981a) is the incorporation into the details of the individual mechanisms for the different types of bacteria of the general principle that introduction of new units into the murein fabric usually takes place only at the cytoplasmic membrane. There is only one case (Streptococcusfuecium) that appears to require insertion not at the membrane. On the other hand, cleavages can be at more varied locations, and lead to volume expansions not arising from current biosynthesis.

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However, the problem of understanding the process of wall growth for any organism becomes one of modelling different physical possibilities and of interpreting biological experimentation. Therefore, we need to consider the methods we can use for both endeavours to advance microbial physiology.

A . SOAP BUBBLES

Soap bubbles provide an effective analogue computer for bacterial wall growth. The surface of a bubble increases when the volume increases in such a way as always to minimize the surface. This is because it takes work to break intermolecular attractive forces and, consequently, it takes work to create new surface. For the soap bubble, this work comes from the pressure-volume work. That is to say: Pdv = TdA From this can be derived the important relationship for all membranes subject to surface tension: where rl and r2 are the principal radii of curvature of the surface at any points on the surface. This equation is especially simple for a sphere or cylinder. For a sphere of radius r, du =d([4/3]nr3)= 4nr2dr,and dA = d(4nr2)= 8nrdr. Consequently, equation (1) becomes: P4nr2dr = T8nrdr

(3)

or:

P = 2T/r (4) This is also what equation (2) becomes with the specification for a sphere that r~=r2 =r. While I have chosen not to prove equation (2) here, I hope that with this check the reader will accept equation (2). The implication of this equation for a cylindrical bubble where rl = r and r2= 00 is that: P = T/r

These equations are fundamental to what follows below and were first stated by Thomas Young in 1804. But they can be empirically checked with two soap bubble pipes, wire loops, some tubing and soap solution from the toy shop, or equal mixtures of a detergent for washing dishes, glycerol and water. Actually, these relationships were popularized by C. V. Boys (1890) (see Fig. 2) and by D’Arcy Wentworth Thompson (1942) in the areas of general science and in biology, respectively.

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FIG. 2. Pressure relationship in cylindrical and spherical soap bubbles. This picture is reproduced from C. V. Boys (1890). The diagram shows the apparatus which he used to demonstrate that a cylindrical bubble must have half the radius to have the same pressure as does a spherical bubble. With this apparatus both a spherical and a tubular soap bubble were blown and then the stopcock closed. The figure shows the condition when the vertically moving element was then adjusted so that the tubular bubble becomes cylindrical. It is then observed that the radius of the spherical bubble is twice that of the cylindrical one.

Equations (2), (4), and ( 5 ) apply to wall growth because the enlargement of the covalently polymerized, bag-shaped macromolecule (or sacculus) follows equation (1) when the increase in cellular volume controls wall expansion. So the soap bubble is a suitable analogue computer for cases where there is a tight coupling of synthesis and cleavage; i.e. where Pdv work forces the expansion process directly. The soap bubble analogy is relevant to the growth of the cylindrical part of Gram-positive rods and the more spherical cocci, such as staphylococci. However, the analogy is of limited use for the other cases where Tchanges during the growth process, because Tis the same all over the soap film surface (independent of thickness of the film, because it is a surface phenomenon). For those cases where T varies, one can think about, or experiment with, molten glass, where the properties vary in special ways, depending on temperature. These analogies are useful for the cell division properties of staphylococci, the shape of Gram-negative rods, and hyphal extension. They are not easily amenable to producing shapes to be compared with microscopic observations, but I have worked with a master glass blower to check some of the conclusions from the computer studies. Equation (1) given above can lead to very sophisticated mathematics, and only for a few shapes of interest here can an analytical solution be derived (Koch et al., 1982a). Many eminent mathematicians (LaPlace, LaGrange, Euler and Maxwell among others) have worked on the calculus of variation

31 0

ARTHUR L. KOCH

problems of surface minimization, and I have no hope of doing what they could not do. Fortunately, for certain morphologies, the problem becomes tractable when it can be assumed that growth takes place in a limited region of the cell at any instant of time.

B . THE MATHEMATICS OF NARROW ZONAL GROWTH

The prime examplein this category is the streptococci where it has been known for many years (Cole and Hahn, 1962; Hughes and Stokes, 1971)that the wall is newly synthesized in the region between wall bands, and not at all in completed poles. This suggested the further simplification of assuming that shape depends on conditions where the septum splits and the wall remains rigid thereafter. There alone, it is proposed (Koch el al., 1981a), is it relevant to write the energy conservation equation (1). Because material from the septum becomes converted into external wall, the resultant equation for the slope S (=dr/dz) involves the thickness 6 of the septa1 wall relative to the external wall. A simple geometric argument leads to:

PIT = ( 2 J i i T + i i ~ ) l r

(6)

where r is the radius of the cylindrically symmetrical structure. This equation leads to a quadratic equation in S which is readily solved. Although the validity of this equation depends on the assumption of equation (l), and that the growth zone be narrow, it is possible that P, T, or 6 could change during pole formation. Consequently, if it is known, or surmised, how P/Tand 6 change during the growth process, starting from the radius of the wall band where pole formation starts (a), one can calculate, step-by-step, the course of formation of new wall. Of course, the calculation is greatly simplified if PIT and 6 are constant throughout the growth process. This, of course, can be checked experimentally from the average shape of the pole of any type of organism by using equation (6), with the substitutions that S=Tan(a). Note that equation (6) was derived with the convention that an outward slope is positive and that an invaginating wall has a negative angle and slope. If PIT is constant and 6 has certain fixed values, then equation (6) can be integrated and equations can be written out for the shapes that result. For example, a cylinder results if 6 =0 and Pr/T= 2. Because S equals 0, r does not change; thus the slope remains at zero indefinitely and r stays constant as cylindrical elongation takes place. When Pr/Tis different from 2 and 6 is 0, the shape is given by: S = +[(Pr/2T)2-

(7)

SURFACE STRESS THEORY OF MICROBIAL MORPHOGENESIS

311

The Surface Stress Theory had its first success in the agreement of the pole shape predicted for the special case where PIT is constant and 6 is 2, with the observed shape of S. faecium. The curve for this case has been designated the zonal dome. Its formula is: z =,,,z

+(2T/P) In [1 - (Pr/2T)2]

where z is the axial distance measured from the base of the pole. The axial height of the pole is given by: (9) zmax= -(2T/P) In [l -(Pu/2T)*]

In this equation, “a” is the radius at the wall band (i.e. the pole’s base). I will use the symbol a for the maximal radius of the bacterial cell of any shape. The many other shapes that could be produced by narrow growth zones can be calculated by the computer via numerical integration of the S derived from equation ( 6 ) to yield curves of r us z. Calculations for other values of 6, and for cases where PIT and/or 6 vary during the cell cycle, have been made (Koch et al., 1982a; Koch, 1982a). Some of these have been discussed in the journal papers and some more will be discussed below.

C. DIFFUSE GROWTH

If growth can occur over a significant area, then the constraints are different and sophisticated mathematics are needed; though equation (l), the conservation of energy, remains at the heart of the problem. Thus the problem involves the integral of equation (1) over all possible surfaces and the selection of that surface which minimizes differences of the Pdv work and the surface tension integrals. Although this minimization can be done formally, it does not result in general analytical equations so that a meridian can be calculated which, on revolution, describes the surface of various types of bacteria. However, it does result in an equation for the slope of the surface which, like equation (6),is in terms of the radius at that point, but as distinct for the case of a narrow growth zone, the equations for diffuse growth also depend on the slope on the adjacent region of the surface. Consequently, the only means at our disposal to calculate the surface that connects two rings of arbitrary radii “a” and “b” and separation distance “L” with an arbitrary pressure differenceand surface tension (Koch et al., 1982a)is to project tentative curves from one boundary ring of radius “a” assuming different initial slopes and only accept the curves that end up at the other boundary at the desired distance ‘‘l” and at the desired radius “b” as valid solutions.

31 2

ARTHUR L. KOCH

This involves much computing, but in principle the following equation allows generation of shapes satisfying the needed boundary conditions (Koch, 1983):

+

1 +s2 1 ASo/ro 1 +So2 To/T+PASo(l + S O ’ ) ” ~ / T In this expression, To is the value of T at the adjacent portion of the curve where the slope is SO,and A is the axial distance increment. In computation, various problems arise and various approximations must be used to get sufficiently accurate answers in different parts of the curve. But all this can be done even by a pocket programmable calculator (the Hewlett-Packard HP 41 was used to make the curves shown below) but could, of course, be done more rapidly by larger computers. Another calculation strategy that can be used was provided by Hendrick Lauwerier of the University of Amsterdam. He was readily able to show from equation (1) and the methods of the calculus of variation, that a point on the curve at (r, z) could be calculated from: r

=

[ T 2 / P2 K 2 Sin2(T)]’/2- K Cos(T)

(1 1)

and

z

=

r

rdT

(12)

where: K = ,/T2/P2-ao2T/P Cos(uo)+ao*

(13)

K is a combined constant, T is a dummy variable, and a0 is the initial angle corresponding to the initial slope, SO=Tan(u0). To summarize this section: with modern computers and with enough patience and/or computational techniques, one can describe the shapes of surfaces controlled by energy conservation. Among these there are curves that can be considered as candidates for the structures of micro-organisms.

D . P R O B L E M S OF E L E C T R O N M I C R O S C O P Y

The electron microscope provides the only way currently available to study the morphology of small, non-regularly arranged objects. As the “only game in town”, we have to deal with its problems; these can be divided into two categories-those relative to shape changes during preparation of the sample and those attributable to the fact that cells must be killed to be examined. The latter will not be treated here but can be handled by methods we have

SURFACE STRESS THEORY OF MICROBIAL MORPHOGENESIS

313

developed elsewhere (Koch and Higgins, 1982). The important point is that microbiologistscan view a particular cell only at a single instant of its life and only as a mummy. I will not discuss observational artifacts. 1. Preparation Artifacts

The cell must be fixed and dehydrated before examination in the electron microscope. Usually they are also stained with materials containing heavy atoms. In addition, when cells are sectioned there may be compression artifacts. I am not the person to discuss the myriad of detail involved in this highly technical field, although I have now worked (and argued) in the laboratories of some of the best workers in the field of microbial ultrastructure. The aspect of the problem emphasized by the Surface Stress Theory is that the peptidoglycan sacculus in the living state is under stress and must certainly change its dimension and its shape if the osmotic pressure relationships are altered. On this basis, the only acceptable preparation technique is that of very rapid freezing of microdroplets followed by freeze fracture. It clearly is not acceptable to add substances like glycerol to prevent ice crystal formation since that could affect transiently the osmotic relations. As of yet, little critical work has been done within these limitations. For the purposes needed here, the only measurement that can be made when this technique is used is the estimation of the diameter of rod-shaped organisms since fractures that happen to be transverse, as judged by their circularity, can be measured. The pictures also must bear evidence of the quickness of freezing deduced from the texture of ice crystals in the background. Only when these criteria have been satisfied am I prepared to believe the measurements as accurate estimates of the diameter in uiuo. All other available techniques must have a shrinkage artifact. If the area of the extended wall is two-fold greater than that of the unstressed wall, then the observed linear dimension would be about 71% ( = l00/42) of the living cell and the volume would be 35% ( = 100/242) as large. While more critical work is needed, the results of Woldringh et al. (1977), Trueba and Woldringh (1980), and Bayer and Remsen (1970) for the Gram-negative Escherichiu coli are clear. The diameter of the living organism as seen in phase-contrast light microscopy is larger than that of freeze-dried, critical-point dried, or median thin-sectioned material. Sacculi, or agarfiltered and dried Gram-negative bacteria, are known to be flattened and, of course, larger. They therefore incidentally approach the diameter of the living cell. The problem, albeit somewhat exaggerated, is that ofimagining the shape and size that a balloon will have after inflation from observing it empty and flattened. The other analogy which would be closer in scale would be to

314

ARTHUR L. KOCH

estimate how a leg will look in uivo from examining a nylon stocking in vitro. Critical data are not yet available for the size of living bacteria because of the limited resolution of the light microscope, although research on this point is underway (C. Woldringh and F. Brakenhoff, personal communication). For preservation of shape of whole bacteria, critical point drying seemingly would be best because surface tension artifacts are eliminated. This technique produces the smallest sized organism of any of the techniques, presumably because the flattening artifact is decreased. However, shape distortion certainly must be increased, at least in certain aspects. I found this particularly noticeable among cells that are just about to divide or have just divided and not separated. I have focused attention on such cells since they have maximum information about cell morphogenesis, because they are at a definable position within the cell cycle and because comparison measurements on different portions provide “the-subject-its-own control” type of information that tremendously decreases statistical error. As the cells shrink during critical-point drying, the distal ends of the unseparated daugher cells are not free to move toward each other as they would in liquid suspension. However, since they are affixed to a surface, stress is created in the region connecting the two, causing shape distortion in the region where they join. This artifact was observed‘ with both E. coli and Bacillus subtilis, but more so with the Gram-negative organism. So, one is currently left with selecting pictures of sectioned material that have shrunk in suspension whenever their osmotic pressure bubble burst, their wall relaxed or they became dehydrated. All one can do is hope that the resultant structure is similar in shape to the living state, although smaller. But what error, one might ask, results from errors in selection process in choosing the truly axial median sections?

2. Error in Relative Dimension Estimations from Sectioned Material The acceptance of an electron micrograph of a sectioned rod-shaped cell as an axial and median section depends on visual decisions of the worker. The first criterion is that the section must indeed be rod-shaped with rounded ends. Even so, it could be off-axis or askew. So, a second criterion is needed such as the appearance of the wall around the entire periphery of the cell (Higgins, 1976). This criterion depends on the observation that cross sections of both Gram-positive and Gram-negative walls give a crisp, tribanded structure, while in oblique sections, the appearance of the wall is diffuse. On the basis of serial sections of several organisms treated several ways, Michael Higgins and Ian Burdett believe that sections exhibiting a crisp “tripple-track” appearance are within the central 15-20% (Higgins, 1976; Burdett and Higgins, 1978) of

SURFACE STRESS THEORY OF MICROBIAL MORPHOGENESIS

315

the cell (see Figs 3, 4 and 5). As a safety factor, I will assume this second criterion to be satisfied if the section is entirely within the central 20% of the cell, but will present calculations for the limit chosen at 10,20, 30 and 40% of the width of the cell. The inclusion of off-axial or off-median sections means that the average dimensions will be underestimated and that variability would

FIG. 3. Median section of Bacillussubtilis 168s. Picture courtesy of Ian Burdett. This cell was obtained from a culture growing in glucose minimal medium with a doubling time of 45 minutes at 37°C. A statistical analysis of similar cells in the last phases of the cell cycle is given in Table 2. Note the crisp triple track outline of the wall. Compare this with the appearance of other walls in the field of view. Bar denotes 1 pm.

FIG. 4. Median section of Bacillus megaterium. Picture courtesy of Ian Burdett. Cell taken from the culture growing in broth medium with a doubling time of 19 minutes at 37°C. The section is assumed to be median because of the appearance of the wall all around the cell. Note that the clean appearance of the newly cleaved septum at the right compared with older wall. Note that the wall has a rougher appearance near poles and that the older pole is more hemispherical than the younger one. Bar denotes 1 pm.

31 6

ARTHUR

L. KOCH

FIG. 5. Median section of Escherichia coli B/rA. Picture courtesy of Ian Burdett.Cell taken from a growing culture of strain ATCC 12407with a doubling time of 45 minutes in glucose M9 medium at 37°C. The cells were fixed with glutaraldehyde,formaldehyde

and then stained with osmium tetroxide. This cell had been fixed a very short time before it would have divided. Note the more highly curved peripheral part of the nascent pole and that the completed poles are more nearly hemispherical. Bar denotes 1 pm.

be found even if there were no variability present in the objects sectioned. We must consider what error is so entailed. Figure 6 depicts the problem. The apparent diameter of the cell would range between 2a and (2aJ1 -0.22) and the apparent length between 1 and (/-2a+2uJl -0.22). So, the problem is one of calculating means and standard deviations for a very special distribution of lengths. The distribution for a circular section is shown in Fig. 7. For the width (or the length of a spherical cell), this can be done using the basic ideas of statistics with the integrals of x"dx/Jl -x2 at hand. The results are given in the second line of Table 1. This is the case of a circular section where 1= 2a. Table 1 shows the mean and coefficient of variation for the four choices of the limits for inclusion of a section in the category of median axial sections, and also for several ratios of length to width. I t is surprising how little an underestimation of length or width is made and how little a contribution to the variability of width measurements would be made with even an unstringent criterion for acceptance of a section as being axial. For elongated cells, errors in the length are smaller than in the width. Of course, the surface area or volume, which involve the products of the observed linear dimensions, have errors that are quite exactly twice or three times those of the linear dimensions.

SURFACE STRESS

THEORY OF

MICROBIAL MORPHOGENESIS

1.20

1.0.978 (2a) 1=0.994(2al I= LOO0 (2a)

sphere

L.0.20 -+-.-.-.-.-. Centre of

pole

317

._._.___._._._._._.-.

--

1= 0.9995(6al I = 0.9997(6a1 1 = 0.9992(6a)

t

.-

1=6a

FIG. 6. Error in median sections. The pole of a hypothetical cell and its axis is shown. Direction of knife cuts for the case that all of a section lies entirely within the central 20% of the cell are shown. For rod-shaped cells, knife cuts that enter the pole at this limit can range from a plane parallel to the axis, to a plane going through the centre of the cell, to a plane leaving the cell on the other side at the opposite limit point. Shown are these directions for cells with ratios of length-to-width of 1:1, 2:l and 3 : l . The computer program averages over the ranges of entry and exit points.

The bulk of the entries in the table were generated by a computer program which assumed that the point where the sectioning knife entered the cell is regularly distributed (at 41 discrete sites) over the contour of the cell within the specified criterion limits. The computer also assumed that the point where the sectioning knife leaves the cell is also regularly distributed on the other side of the cell. The computer calculated the distances by standard trigonometric formulae and computed the means and standard deviations, weighing all 412 ( = 1681) combinations as equally probable. Because the numbers of sites, though large, were not infinite and the effects for which we are searching are very small, the errors in the estimates of the errors are considerable. The greatest discrepancy should be for the first case (i.e. for a round cell or the cross section of a rod-shaped cell), and here the correct values calculated in the different way indicated above are given for comparison. The broad conclusion

318

ARTHUR L. KOCH

FIG. 7. Distribution of sections of spherical bacteria. Under the crisp triple tracks criteria, only cells sectioned entirely within the central 20% would be included. The distribution function is such that no section outside the range 97.87 to 100% of the radius would be found and most would be near the upper limit.

is that little error is involved in dealing with sectioned material meeting the criterion of a crisp triple track. Another aspect is shown in Fig. 8. This has to do with the odds of seeing the inside edge of the invaginating wall or central hole of a septum in a section. The curves are calculated on the assumption that bona fide median sections are scored for the frequency of observing holes in the septum and that the mean diameter of the holes is then calculated. On the assumption that all holes

SURFACE STRESS THEORY OF MICROBIAL MORPHOGENESIS

319

TABLE 1 . Error of cell lengths due to inclusion of non-axial sections' ~~

~

Limit for inclusion of sectionb 10%

20%

30%

40%

Cell length'

U

CV

U

CV

U

CV

U

CV

2.0d 2.0"' 3.0 4.0 6.0

0.12 0.17 0.14 0.02 0.01

0.15 0.15 0.11 0.01 0.00

0.50 0.67 0.57 0.09 0.03

0.62 0.61 0.46 0.08 0.02

1.21 1.52 1.34 0.21 0.05

1.51 1.39 1.09 0.18 0.03

2.40 2.74 2.52 1.16 0.37

3.02 2.54 2.07 0.83 0.24

' Calculated for cylindrical cells with hemispherical poles. All of the section must lie within the indicated percentage of the cell radius. U is percentage underestimation of mean length. CV is coefficient of variation due to non-axial section expressed as a percentage. Pole-to-pole cell length in units of cell width. Spherical cell. Calculated from integral expression without computational approximation. All other values obtained by numerical integration as indicated in texts.

('

are circular and centred on the cell axis, it can be shown that the ratio of the observed mean radius 6 to the actual value b is: 6/b = (z,/b2 - z2 + b2 Sin-'[z/b])/2bz where z is either the radius of the hole or the limit radius for inclusion of the section, depending on which is smaller.

3 . Curvature Measurements

For the poles of Streptococcus, there is a clear discontinuity in contour at the site of the wall band, and the wall has a symmetrical shape around the plane of the wall band. For this organism, the zonal dome theory appears to work well. The test is the fit of equations (8) and (9) to the measurements of established poles and nascent poles (Koch et al., 198 la). Another way to check this model is to measure the angle that the wall makes with the axis (or the complement of the angle with the wall band plane) at corresponding positions of the same cell or for different completed or nascent poles of corresponding portions of different cells. A particularly convenient location for measurement is at the wall band itself. We have also tried this approach (A. L. Koch and I. D. J.

320

ARTHUR L. KOCH

8FIG. 8. Probability and mean size of the hole determined by the inner edge of a septum or constriction. The mean size was calculated according to equation (14).

Burdett, unpublished experiments) with B. subtilis. However, I feel that this technique is less satisfactory in this organism because the discontinuity in slope at the presumed wall band is less marked in most cells. In fact, different observers measured somewhat different angles. This does not mean that angle measurements cannot be used successfully as a way to compare nascent poles with older poles and draw significant conclusions about morphogenesis of this organism (see below, p. 329). But for the poles of rod-shaped organisms, where the discontinuity in slope at the junction between the side wall and the pole is small, the alternative approach of measuring diameters of curvature is better. This can be done on microphotographs easily by superimposing plastic curves such as are used as templates for drafting. In particular, I have found

SURFACE STRESS THEORY OF MICROBIAL MORPHOGENESIS

321

FIG. 9. Drafting templates for estimating diameters of curvature. This commercial template was used to measure the curvatures of electron micrographs to obtain the measurements presented here.

Rotring's 842 687 Radien-Tangenten (Fig. 9) a very useful device because it contains circular arcs that are tangent to straight lines. Consequently, visual judgements can be made quite accurately of the diameter of curvature of a pole that fits best under the constraint that the pole curve be tangent to the side wall. Another advantage of this template is that it is also possible to restrict the fitting process to the 45" of arc tangent to the side walls. This device, and a variety of other available devices, can be used to make independent fits to axial curvature, i.e. the best fit of the central 90" of curvature to a completed pole. Equally subjective are curvatures calculated from digital records of photographs traced on special graphics tablets. While the computer can go directly to the radius of curvature via:

K=

d2y/dx2

(1

+[ d y / d ~ ] ~ ) ~ ' ~ '

and use statistically sophisticated ways to average the tracing, the operator must define the end points. Higgins' procedure to use the computer technique optimally and simply depends on the introduction of a quantity:

c = ( A-A')/A

(16)

where A is the contour length between a developing septum and the wall band and A' is the chord length. The values of C and A are included among the summary statistics in the computer fitting system in the laboratories in Mill

322

ARTHUR L. KOCH

Hill and Temple University so that data could be retrieved and re-expressed as diameters of curvature via:

D

(17) by using an iterative procedure to find D from A and A' or from A and C . Diameters of curvature can also be calculated using computer-stored data of the total wall surface area, the cylinder surface area, and the nascent pole surface area. These data allowed the computation of the mean surface area of completed poles. From this, an effective diameter of curvature can be calculated by assuming that the area of the sum of the surface of both poles of a cell is nD2.Similarly, a different averaging procedure can be used by taking the volume of the two poles of a cell and computing the equivalent diameter of a sphere. Measurement of many cells and use of these averaging procedures are necessary in order to draw valid conclusions since there is a fair amount of variation in the individual shapes of cells of both Gram-positive and Gram-negative species. =

A Sin(A'/D)

E . ANALYSIS OF AUTORADIOGRAMS

Bacteria are so small that ordinary autoradiography using tritiated compounds cannot locate subcellular structures. The average distance that a tritium p particle travels (1 pm) is comparable to the dimensions of the micro-organism. A very great improvement in resolution can be obtained by using a very thin film of photographic emulsion and examining the results in the electron microscope. From the distribution of silver grains, one would like to infer the distribution of tritium atoms in the cells. Most often, the samples are developed when there are only a few grains per cell or per region of a cell, and statistical procedures must be employed to estimate the mean grain count and the uniformity of distribution of isotope within the cells. In previous work it has been tacitly assumed that there are similar distributions of isotope in different cells and that the grain counts over similar positions in many cells can be accumulated, thereby the statistical problem can be overcome. Usually, what has been done is to assume that synthesis occurs in a symmetrical fashion in all cells of a common size. Thus the number of grains is measured from some reference points such as the centre of the cell or a region of constriction or septation. Then, histograms of grain counts us position in the cells have been constructed. In this way it has been established that the centre of the cell is a region of high synthesis in cells of certain class lengths (Ryter et al., 1973). Grains non-centrally located might be caused by secondary zones or diffuse

SURFACE STRESS THEORY OF MICROBIAL MORPHOGENESIS

323

growth and, in some degree, may be due to spill over of the radiation from the central zone. The latter can be accounted for, but the choice between narrow secondary zones and diffuse growth cannot easily be made. Three possibilities all lead to similar histograms: diffuse growth, many secondary narrow growth zones, or secondary growth zones that are not precisely positioned from cell to cell so that the picture is blurred. Only when the specific radioactivity is high enough, and duration of exposure of the autoradiogram is long enough so that sufficient grains are present over a cell [in addition to any intense zone grains] so that the distribution within a single cell is unequivocal, can a distinction be made between narrow, but variable, growth zones and diffuse growth. So far, this has only been done with B. megaterium (see below, p . 332). Until that has been done for the organism in question, I may argue in favour of diffuse growth and others may argue for narrow zonal growth; but autoradiographic studies, combining measurements on many cells, cannot be used as the critical test evidence. Still, we need to be able to calculate the grain distributions that would be found for various distributions of isotope within a rod-shaped bacterium. It is possible to show (Koch, 1982c) that a line source of length 2 hours, centred at position C on the X-axis of a co-ordinate system, will contribute a collective count (gl) to the strip of indefinite length perpendicular to the axis at position x of: gl = 2d2K (Tan-'[(x-C-h)/d]-Tan-'[(x-C+h)/d])

(18)

where K is a constant involving the specific radioactivity and d is the half-distance of the tritum f?-particle as measured under the experimental conditions used. This formula can be set into a computer that can then be used to calculate distributions above cells for any specified, longitudinal arbitrary distribution of radioactivity. In most cases, d and K can be treated as constant for the collection of line-segments that must be summed together to simulate the perimeter or the cross-section of the bacteria. Since the hemispherical pole has the same surface as an open-ended cylinder, equation (18) applies directly, with no further integration, for rod-shaped bacteria with hemispherical poles of uniformly labelled cells with no developingdivision site. Operationally, this is done by combining the distributions for sources with appropriate values of C and h. Of course, one must know or assume the value for d. As an example, Fig. 10 shows the curves for the distribution of silver grains along a cell of the dimensions of whole mounted Escherichia coli, such as used by Verwer and Nanninga (1980). For the shorter class of cells shown in the insert, the walls were assumed to be uniformly labelled. Two curves are shown: one assumed the entire length (cylinder plus pole) was labelled and the other assumed that the cylinder region alone was labelled. The same two choices

ARTHUR L. KOCH

324

End of pole

Plus

P Distance from centre

FIG. 10. Distribution of grains in two size classes of Escherichiu coli W7 (dup- lysA). Data from the experiments of Verwer and Nanninga (1980). The figure shows the distribution of grain counts measured from the centre (or from an observable constriction). Poles for cells of sizes between 3 . 4 3 . 8 pm and 1.8-2.2 pm (insert) are fitted by equation (18) to models where the poles are either unlabelled or labelled as highly as the side wall. For the longer cells an additional component was added to approximate the central intense zone. The same amount of radioactivity was assumed for both fits designated by the thin and thick lines: the narrower line, approximating the distribution seen in constricting cells; and the broader, approximating cells of this size class where a constriction was not visible. - . - . -, Cells showing constriction; -.-.,cells not showing constriction.

were made for the longer cells in the major part of the figure but, in addition, an extra shorter source, either quite narrow or fairly broad, was summed using suitable weights with the longer sources to produce the smooth curves shown. A quite different aspect of the analysis of autoradiograms of median sections also must be considered here. In laboratories set up to reconstruct the three-dimensional shape of the cell from median sections, this problem is taken care of by the computer program. With the systems used by M. L. Higgins at Temple University or by I. D. J. Burdett at Mill Hill, the computer calculates the volumes and the areas of specific portions of the cell via numerical summation of trapezoids of revolution. But the corresponding corrections have not been applied in published autoradiographic studies. Figure 11 shows the problem faced in the analysis of autoradiograms of

325

SURFACE STRESS THEORY OF MICROBIAL MORPHOGENESIS Surface

Section

Cylinder

21A

Complete septum

2rA

Incomplete septum

Z(r-fr

Hemispherical Dole

2arA

IA

Rotio

f

R

i-

FIG. l l . Geometric factors for the analysis of autoradiograms of median thin sections. Where 1=length of cylindrical portion, r =radius of cylindrical portion, A = thickness of section, and R=radius of inside edge of the ingrowing septum.

sections, and the factors needed to convert the counts obtained to the values that would have been obtained if whole cells could have been analysed with the same efficiency and resolution. The point is simply that a grain observed on the periphery of the rod corresponds to a great deal more radioactivity in the cell than does a grain observed near the cell axis. The considerations presented in this section were made under the tacit assumption that the efficiency of grain production was the same all over the object examined. This is not always so (see Section IV, p. 357).

111. Results

This essay is written in the standard form of a research paper to emphasize that the work and ideas arising therefrom are indeed the results of “a close and careful search”. It is also research in the arcane, but original sense of the word “of looking again”; it is not research in the modern usage of “investigation” because none of the experiments are mine. All the information presented in this section is a reassessment of work done by others. Many people have provided pictures, films and access to their notebooks and computer data banks. Other items are recalculated from published papers. In so doing, I have sometimes had to filter out ‘noise’ introduced by the draftsman, or by my ability to measure the printed graphs or computer-generated drawings. In some cases this could be done because I knew that counts had been made at precise intervals, and in others because I knew that the number of silver grains counted had to be integer. These results, recast as indicated below, provided

326

ARTHUR L. KOCH

the data to test aspects of the theory. Of course, the theory was sometimes wanting and was subsequently revised or expanded as the process of data acquisition, disputation and digestion proceeded.

A . GRAM-POSITIVE RODS

In the early phases of the development of the Surface Stress Theory (Koch et af., 1981b, 1982a), it was proposed that the Gram-positive rod had two forms of wall growth. The poles were formed by a narrow growth zone mechanism and the sides between the junctions with the pole were presumed to elongate by the process of “inside-to-outside’’ growth whereby cross-linked murein would be forced to expand as it moved more peripherally and came to bear the stress resulting from the osmotic pressure. Later cleavages interrupt the stress-bearing character of the older region wall and still later, in normal strains, the wall fragments are degraded and liberated from the cell. The early theory proposed that the poles were formed according to the zonal dome model and, once formed, the poles were rigid indefinitely. The assumption of rigidity of,pole was essential since it could be shown (Koch et al., 1982b and below, p. 347) that only if three conditions were met would stable elongation of cylindrical wall take place that would resist fluctuations to bulge catastrophically outward. These conditions are (a) rigid poles, (b) P and T remain constant, and (c) Pa/T initially 1. Adherence to these conclusions provides the simplest explanation of how an organism could achieve and maintain a rod shape with constant width without postulating scaffolding or templates. The experimental work of Pooley (1976a,b) first demonstrated with radioactive tracers that peptidoglycan was laid down near the cytoplasmic membrane and gradually moves through the wall. This has been amply confirmed by Doyle et af. (1981) and by work from J. T. M. Wouters’ laboratory (de Boer el af., 1981). At the same time, this was also demonstrated by A. R. Archibald, in Newcastle, who used nutritional shifts affecting the level of free phosphate to cause the bacilli to switch between teichoic acid and teichuronic acid production (Archibald, 1976; Archibald and Coapes, 1976). They could detect wall containing the phosphate-bearing teichoic acid because it served as a receptor of a virus. They demonstrated directly that the first appearance of new teichoic acid and hence, presumably, new murein was on the inner face of the wall. This experimental approach also showed that the poles were more highly conserved than the side walls (Archibald and Coapes, 1976; Anderson et al., 1978; Sturman and Archibald, 1978). Figures 12 and 13 show some previously unpublished pictures made as part of these studies. The inside-to-outside growth of the cylinder has been confirmed in recent

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FIG. 12. Conservation of old poles of Bacillus subtilis strain W23. Shift to phosphate excess. Photograph courtesy of Ronald Archibald. Cell sample fixed 0.8 generation after shift of chemostat to potassium limitation. By this time inside-to-outside growth has brought the phosphate-containing teichoic acid on the side walls to the surface everywhere except at the poles. Both this and Fig. 13 were made as part of the study of Archibald and Coapes (1976).

experiments done in Lausanne (Pooley et al., 1978; Schlaeppi et al., 1982)and in Louisville (H. L. T. Mobley, unpublished observations, see Koch et al., 1981b), both laboratories unequivocally demonstrating conservation of a higher degree of wall at the poles, and less conservation of wall in-between. There is a quantitative argument about just how much wall is conserved in association with the poles. The former found that a larger fraction than simply the pole is conserved; and Koch et al. (1981b), consistent with earlier results from Archibald’s laboratory, held that only the geometric pole is conserved. Figure 14 shows the conservation of the pole alone, from previously unpublished experiments from Louisville. But defining the boundary of the conserved entity is a minor skirmish; more crucial to the theory for inside-to-outside growth is the ability of the pole to resist change. In the long run, an increase in diameter of curvature of the pole would tend to bulge the

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FIG. 13. Conservation of old poles of Bacillus subtilis strain W23: shift to phosphate limitation. Photograph courtesy of Ronald Archibald. Cell sample fixed 1.7 generation after shift of the chemostat to phosphate limitation. By this time the cylinder regions have been largely replaced while the old poles still avidly absorb virus showing they retain the phosphate-containing teichoic acid formed as the poles developed.

FIG. 14. Binding offluorescein-labelled concanavalin A to Bacillussubtilis strain C33. Photograph courtesy of Ronald Doyle. This strain is wild type with rapid autolytic activity (lyt+), but is (trp-). The cell sample was fixed 3.2 generations after the temperature was raised to the non-permissive (45°C) from the permissive condition (35’C). The bar is 10 pm. Note that the fluorescence is restricted to the old pole. The photograph was part of the study of H. L. T. Mobley and colleagues (unpublished observations).

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cylindrical part, leading to more spherical (or at least wider) cells with each successive generation. In the subsections below, evidence concerning aspects of this question are presented. The pole is not formed in its final shape in the same manner as the poles of streptococci, and does turn over slowly, contrary to the original model for inside-to-outside growth of Gram-positive rods. 1. Evidence Against Fixity of the Pole During its Formation Table 2 shows the tangent angles that the pole makes with the cylindrical part of the wall of median sections of Bacillus subtilis strain 168s. Ian Burdett had prepared many sections of this organism (Burdett and Higgins, 1978; Burdett, 1980). All I did was measure, with a protractor, 13 cells in the last stages of division. While there are limitations to angle measurements, because of subjective error in locating the junctions between pole and cylinder, they have the advantage that they are independent of any magnification error that might be associated with the electron microscopy or the photographic process. The conclusion.from Table 2 is that the nascent pole makes a larger angle with the TABLE 2. Evidence for change of shape during pole formation. Data from poles of nearly dividing Bacillus subtilis 168s Tangent angles"

Diameters of curvatureb

Nascent Completed poles poles Nascent poles (degrees) (degrees) (Pm) I. Median sections' 19.1 f0.9 10.6f0.5

0.32k0.01

Completed poles (Pm) side

end

0.45k0.02 0.63f0.02

Width (Pm) 0.64kO.01

11. Glutaraldehyde-fixed whole cellsd

0.51 f O . O 1

0.47f0.02 0.58f0.02 0.533f0.004

Measured with a protractor as indicated in the text. Diameter of curvatures were measured on the 45" of arc adjacent to the side wall and on the terminal 90" of arc ending the cell. Thirteen microphotographs, selected from 292 median sections which showed cells in the final phases of the cell cycle, asjudged by nearly complete splitting of the septum, were included in the sample. AH errors are standard errors. Eleven glutaraldehyde-fixed whole cells, selected from photographs of many cells, taken from the same culture as for the median sections in Part I and showing cells in the final phase of the cell cycle as judged by the extent of the cleavage process were measured as indicated in the text.

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cylinder than does the completed pole, and therefore the pole must change its shape after formation. Further analysis on a different, more extensive, sample of data will be given elsewhere (I. D. J. Burdett and A. L. Koch, unpublished observations). The same conclusion, that the nascent poles are more curved than the completed poles, is derived by diameter-of-curvature measurements shown on the right-hand side of the table. These results are confirmed from electron micrographs of fixed whole cells from the same culture. These results exclude a narrow growth zone model for the pole of B. subtilis.

2. Evidence Against a Zonal Dome or Parabola-Shaped Pole The finding of Table 2 that the axial 90" had a larger diameter of curvature than the 45" nearest the cylinder is contradictory to all the shapes proposed in the literature for bacterial poles because the zonal dome model, the parabola (Barnett and Burdett, 1981), and the Cassinian (Oldmixon, 1974) all have the property that the curvature near the axis is smaller and increases with distance from the pale. Further confirmation of the experimental finding is given in Table 3. From the measurement of the smallest cells found in the population, it is concluded that the poles were consistently of a shape that was more highly curved near the side wall than near the axis. Thus, for a second reason, the TABLE 3. Dimensions of the ten smallest median sections of Bacillus subtilis" Diameters or diameters of curvature (pm) Left

Centre

Right

0.543 k0.021 0.529+0.040 Pole curvature, central 90" Pole curvature, 45" near cylinder 0.416 f0.016b 0.430 k0.032' Cylinder width adjacent to pole 0.506 k0.016 0.533f0.163 Cylinder width at centre 0.527 k0.015

'' Ten median sections (selected out of the 292 median sections used as the source for the cells analysed in Table 2) which on reconstruction had the smallest volumes were included in the sample. All measurements are expressed as diameters or diameters of curvature. All errors are standard errors. All measurements were made directly from the micrograph. Each photograph was oriented so that the most regular pole (rounded) was on the left. The mean width of all three positions of measurement was 0.522 pm; to be compared with the computer traced average value of 0,523 pm for all parts of all cylindrical regions. The mean radius ofcurvature of both poles of the 91 cells in the population that showed no septation was 0.51 1 k0.006 pm when calculated from the pole surface area as indicated in the text. The two measurements taken on the upper and lower sides of a cell were averaged before statistics comparing the ten cells were performed.

*

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zonal dome appears to be excluded as a model for the Bacillus subtilis pole. Postulation of a narrow growth zone, with 6 values varying in a way consistent with measurements of the septa1 thickness, overcomes this second difficulty, but not the first. So, “back to the drawing board”! Further analysis on a different, more extensive, sample of data will be given elsewhere (I. D. J. Burdett and A. L. Koch, unpublished results). 3 . Pulse Incorporation of Murein Precursor into Diflerent Parts of the Wall

Although Sturman and Archibald (1978) had noted that some slow turnover of poles takes place, I felt that further evidence was needed since cylindrical elongation seemed to require rigid poles. Antoinette Ryter and Chantal de Chastellier kindly provided me with films of their high resolution study (de Chastellier et at., 1975a,b) of peptidoglycan synthesis in Bacillus megaterium. They carried out an analysis of median sections of organisms pulse-labelled for two minutes with [3H]diaminopimelicacid. In their work, several fixation regimes were followed; from internal evidence they concluded that preparations in which the cells were boiled before autoradiography lost the bulk of their precursor pools while in several of the other treatments that they employed the cells retained precursors. So, it was the films where bacteria had been boiled that I re-analysed. My major goal was to see if the old poles acquired any radioactivity. Because B. megaterium is large and forms chains, and because the Parisian technique had extremely high resolution (half-distance, 0.083 pm), I ended up not using the computer program described above but simply classified each grain as belonging to the side wall, to a septum, to a nascent pole, to poles that had completed (but not fully separated) or to new or old poles. In order for a grain to be classified as belonging to a septum, the septum had to be visible nearby on the microphotograph. Because the septum is a zone of great biosynthetic activity, usually there was a cluster of grains there. On the other hand, the side walls usually had several grains distributed widely apart. This is strong evidence for diffuse growth, or at least multiple zones of growth for cylindrical elongation. This, of course, is consistent with the original conclusion of de Chastellier et al. (1975a, b) of diffuse cylinder-wall growth. Because B. megaterium has a habit of growing in chains it was relatively easy to decide whether a completed pole was new or if it was more than one generation old. In a few cases the distinction could not be made, but luckily most of these were cases in which no grains were associated with the poles, so classification was irrelevant. In a very few cases, an arbitrary decision was made. The grain counts on regions of the cell classified as stages in pole formation are shown in the upper part of Table 4. From the total grain counts

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TABLE 4. High resolution autographic studies of pulse-labelled boiled Bacillus megaterium

Mean Total number of Numbers grain grains cellof cells count ( fS.D.) PolesC T Y Y

I+(

new) old)

71d 29 11 27 75 105

347 105

26 37 54 50

-

4.9f3.4 3.6k3.4 2.4k1.6 1.37f1.3 0.72k0.83 0.48f0.79

Corrected total Corrected meanbnumber Geometric grain countsb of grains cell-I factor 2.83 1.57 2 2 2 2

619

Sides

105

550

98 1 165 52 74 108 100

-

6.9 2.8 2.36 2.74 1.44 0.95

1,480 5.24f2.9"'

3.24

1,728

16.45

Re-analysis of electron microscopic autoradiographs reported in de Chastellier er al. (1975b). Corrected for number of ultimate poles and for the geometric factors shown in Fig. 15. 'The shapes of nascent and age of completed poles. It was assumed that the unsplit septum had an inner radius 0.8 of the radius of the cylinder. 'The standard deviation is statistically different from the expectation of the Poisson distribution.

in the next-to-last column, it can be concludekj that about 46% of the incorporation is associated with pole structure aqd 53% with the diffuse growth of the cylinder. We will return several times to the issue, but a rod-shaped organism with hemispherical poles would need to have its pole-to-pole cell length 2.17 ( = 1/0.46) times its radius to have 46% of the wall material as pole if the thickness and density were constant in all regions (see equation 26, p. 358). This ratio is approximately what was observed through measuring the surface area of appropriate regions of the cells. This type of comparison is rough for four reasons: (1) because of the shape considerations of Fig. 8; (2) because the inside-to-outside growth process does not take exactly one doubling time for transit; (3) because unstressed wall is not immediately shed from the cell; and (4) because we need to average over a cell cycle. With respect to the critical issue of whether murein addition does or does not take place at established poles, the data are clear: radioactivity does become associated with old poles. Of course, the initial phases when the septum is developing have a much larger rate, but the incorporation in old

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poles is well above background and cannot be attributed to precursor pools. So, at this point, the Surface Stress Theory has suffered multiple defeats with respect to the inside-to-outside growth of Gram-positive rods. But please keep reading.

B . GRAM-NEGATIVE RODS

There are several fundamental differences between Gram-positive and Gram-negative organisms. The outer membrane in the latter is important in many ways. But here, the two important issues are that the peptidoglycan layer of the Gram-negative organism is very thin and that it never branches as does the wall in the Gram-positive organism as it starts septum formation (Koch, 1982a). Gram-negative organisms do not form a septum which is later split by covalent cuts. Thus they cannot lay down much murein under conditions where there is no physical stress. Consequently, shortly after covalent linkages are formed, the physical stress due to osmotic pressure must act to extend the conformation of the newly linked murein. The two critical questions that current Surface Stress Theory can approach are: How does it manage to grow in a rod shape, and how does it succeed in dividing? Both processes would seemingly be in direct conflict with the effects of the osmotic pressure that would cause the organism to becomemander. The experimental questions we can ask are: “Is the pole created in a definite shape that is then retained; and, if so, does the pole shape agree with certain special geometric curves?” and “Where does the wall enlarge?”

1 . Evidence Against Fixity of the Pole During its Formation

Table 5 gives morphometric measurements on median sections and criticalpoint dried whole cells of nearly divided Escherichiu coli. The pictures of sections were provided by Ian Burdett and the technical details of culture and preparation are extensions of those given in Burdett and Murray (1974a,b). The critical-point dried material was supplied by Conrad Woldringh. The table is organized in a similar way to Table 3. If the cells had exactly hemispherical poles, and the new poles as they developed were hemispherical, all the items in both major columns would be exactly alike. The longer of the two nascent cells were arranged on the left-hand side of the table. It can be seen that the longer is significantly more pointed (small diameter of curvature measured on the axis) than the shorter incipient daughter (right-hand side). It also can be seen that the middle of the nascent cell is slightly wider than at a completed pole or near where the new pole is forming. But the dominant

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TABLE 5. Morphometric measurement of nearly divided Escherichia coli Diameters or diameters of curvature (pm) Longest Ryter-Kellenberger fixationa length curvature, axial 90" side 45" width, end middle nascent pole curvature, nascent pole 45" width of constriction

Shortest

1.64+ 0.058 1.54f0.063 0.682f0.025 0.717+0.040 0.739 f0.032 0.679 f0.042 0.701 k0.015 0.702f0.013 0.756 f0.017 0.744f 0.012 0.732f0.014 0.732f0.014 0.468 f0.022 0.485 f 0.015 0.386f0.020

Formaldehyde-glutaraldehyde fixationb 1.96 f0.119 length 1.77 f0.100 0.621 f0.024 0.691 f0.042 curvature, axial 90" 0.794f 0.075 0.675 f0.069 side 45" width, end 0.664f0.030 0.686f0.037 0.713f0.038 0.727f0.037 middle nascent pole 0.710+0.034 0.704f0.033 curvature, nascent pole 45" 0.464f0.050 0.484f0.035 0.344f0.021 width of constriction Critical-point dried' length curvature, axial 90" side 45" width, end middle nascent pole curvature, nascent pole 45" width of constriction

1.04 f0.034 1.11 f0.032 0.434f0.011 0.422f0.011 0.461 f0.013 0.488f0.013 0.433 f0.006 0.429 f0.007 0.459 f0.008 0.461 f0.008 0.458 f0.008 0.452 f0.008 0.404f0.014 0.409 f0.01 5 0.291 f0.020

Eleven deeply constricted median sections of glucose-M9-grown Escherichiu coli, fixed by the Ryter-Kellenberger technique, were measured. All errors are standard errors. The two curvatures measured at the 45" nearest the side walls were average before statistical computations. The photographs were kindly provided by Ian Burdett. As above, except that cells from the same culture were fixed with formaldehydeglutaraldehyde, and ten deeply constricted cells were measured. As above, except that the cells were critical-point dried, and 23 deeply constricted cells were measured. The photographs were kindly supplied by Conrad Woldringh. (I

conclusion is certainly that a nascent pole has a much sharper curvature than it will have when completed. This follows from the data because the cells were taken from a steady-state balanced culture. The same conclusion can be drawn for both fixation techniques and for the critical-point dried whole cells.

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That the nascent pole is more sharply curved than expected for a hemisphere of the same diameter as the cylindrical portion is also self-evident in many pictures in the literature. On the other hand, old poles are not very far from hemispherical in average shape, but their major characteristic is that they are more variable in outline than are the newer ones. 2. Pulse Incorporation of Murein Precursors into Different Parts of the Cell

It clearly is equally important to know for the Gram-negative, as for the Gram-positive organism, where growth takes place and whether the old poles continue to incorporate peptidoglycan. For this reason the autoradiographs from the work of Verwer and Nanninga (1980) were remeasured, taking as a reference point the end of the pole. This re-analysis of their data has been published (Koch et al., 1982b), and I will simply cite the conclusion that incorporation does take place in old poles at a rate that is about one-half of that on the sides of the organism. However, this same conclusion can be drawn also from Fig. 10 (p. 324). This figure was recalculated from the computer-drawn graphs used to produce the figures in Verwer and Nanninga (1980), and kindly provided by Ronald Verwer. For each of these populations of cells, a class of small cells (insert) and two classes of large cells with the same length distribution, but differing in whether a constriction could be detected or not, are shown. The theoretical curves were fitted as described above. For each case, two alternative models were considered, (a) that the pole is labelled to the same extent as the adjoining cylinder, and (b) that it is totally unlabelled. For the moment, focus attention away from the centre of the cells. It can be seen that these two models produce curves that surround the observed distribution at the poles, implying partial labelling of old poles. The conclusion that there is murein addition to old poles can be considered definitely established from all the relevant published work (see Figs 15-18). For this work drawn from the literature, I have superimposed on the experimental curves the profiles of the shortest and longest cell measured in the size class, and have presented distances as measured from the mean position of the pole. Figures 15 and 16 are redrawn from Ryter et al. ( I 973) for E. coli growing in two mineral media. Figure 17 is redrawn from Schwarz et al. (1975) for E. coli strains growing under more favourable growth conditions. I have chosen size clases that are big enough to have completed the last cell division before the pulse of murein precursor was given. I have also chosen size classes small enough so that the cells are not likely to have started a new division site. Additionally, I have re-analysed Fig. 12 of Koppes et al. (1978) which shows the analysis of an autoradiogram of pulse-labelled wall material. I will not reproduce this figure since it contains the outline of bacterial cells of

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. ...... 1

;

I

FIG. 15. Distribution of pulse-label over sacculi of Escherichiu coli strain W7 grown in glucose medium. Culture was pulse labelled for one-eighth of a 50 minute generation time. This figure is redrawn from Fig. 1 of Ryter et al. (1973). By showing distance in pm from the mean pole position of the size classes, it can be seen that this data supports the conclusion that poles continue to incorporate label. Cell size: . . . ., 1.0-1.3 pm; - - - -, 1.4-1.7 pm; -.-, 1.8-2.1 pm; -, 2.2-2.5 pm.

the extremes of lengths contained in the size classes. I merely note that the proportions of their diagramatic cells are incorrectly drawn. Nanne Nanninga kindly provided me with films of the organism grown under the conditions used in their study; I found that the average width of PAT 84 cells, grown at 30°C, was 1.04 pm and therefore the cell outline should be about 100% larger than shown in their paper. With that change, this experiment, and those shown in Figs 15-17, all demonstrate that pulse incorporation of radioactivity labels the old poles of E. coli. Other aspects of Fig. 10, presenting data from Verwer and Nanninga (1980), are important to the argument. The two distributions shown in the major part of the figure were made from cells in the same length range class (3.4-3.8 pm): one concerns cells with an observable constriction, and the other concerns the cells where no constriction was apparent. The distribution in the former case is characterized by a more intense central peak of grains than does

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H a l f distance

1

I

I

I

0.5 Distance from pole ( p m )

I

I

0

FIG. 16. Distribution ofpulse-label over sacculi of Escherichiu coli strain W7 grown in succinate medium. For details, see Fig. 15. Doubling time is 80 minutes. Cell size: . . . ., 1.G1.3 pm; - - - -, 1.4-1.7 pm; -.-, 1.8-2.1 pm; -, 2.2-2.5 pm.

the distribution of the cells with no observable constriction. The smooth theoretical curves correspond to the same amount of radioactivity (i.e. 36% of the total in the cylinder), merely distributed differently. For the class of cells showing constriction, radioactivity was assumed to be concentrated in the central 0.2 pm of the cell length; and for non-constricting cells, the same amount of radioactivity was assumed to be distributed evenly in the central 1.2 pm. Although better fits to either curve could be made if considered individually, the agreement of the theoretical curves with the observations suggests that about the same amount of synthetic activity is involved in new pole formation in the two cases; it is just distributed more narrowly in cells where the location of the constriction is obvious. This finding is relevant to the central issue of the work of Verwer and Nanninga (1980). They were interested in whether wall growth occurred symmetrically or asymmetrically and studied this by recording the grains on each half of the cell. Because they had no way to orient the ceJJ with respect to the newer pole, they oriented the cells so that the left-hand cell had the highest grain count. They then applied a probablistic calculation to estimate the grain

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1 (a) 15

-

C In

.-

em L

:10 .In

r 0 0

0 c

c

al

E 5 n

0 0

5

10

15

20 Distance

FIG. 17. Distribution of pulse-label over the sacculi of E. coli cells grown in nutrient-rich medium. Data redrawn from Figs 2 and 3 of Schwarz et al. (1975). The original abscisa was distance, measured in mm, of a 14,000 x enlarged image. Only cells of length 2.&2.8 pm were considered. (a) Strain 103; (b) strain PAT 42; 0,30”C; 0,42°C. For details, see Fig. 15.

count to be expected if the two halves of the cell had the same probability of incorporating a tritium atom into the newly formed murein. I have repeated their derivation, but on the more general assumption that one half of the cell has a fundamental probability p of incorporating tritium, and the other half has the probability, 1 -p. This leads to the formula: R

rl = O

where R is the total number of grains on the cell and G is the number of grains on the “left-hand” cell. The grains are partitioned among the two halves according to rl + r2 =R . The formula computes the probability of a certain arrangement and then calculates the weighted average of the largest number of grains regardless of whether that is rl or r2. Whenp is assigned the value of 0.5 so that the ratio of label on the two halves is 1:1, equation (1 8a) becomes the expression of Venver and Nanninga (1 980). Figure 18 was drawn from this formula with various choices ofp and the data of Verwer and Nanninga (1980)

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15-

z L 0

=

10-

0

c

W

L

R W

In .5

e

5

-

ot

0

I

5

I

10

I

15

I

20

Number of grains cell-'

FIG. 18. Dependency of the number of autoradiographic grains on the half of the E. coli cell with more grains on the asymmetry ratio. The grains on the left-hand half of the cell were calculated from equation (1 8a); the points are taken from Table IV of Verwer and Nanninga (1980). The fit excludes the hypothesis that the tritrium atoms were uniformly distributed on both sides (asymmetry ratio= 1.1) or only on one side (asymmetry ratio= c0:l); rather the asymmetry ratio is near 2.

superimposed on it (data taken from their Table 4). Plotted in this way, some of their conclusions can be more clearly seen. First, the hypothesis that growth is completely asymmetrical ( p is 0, asymmetry ratio is 00:l) is rigorously excluded. On the other hand, the hypothesis that the distribution is exactly even ( p is 0.5, asymmetry ratio is 1: 1) is also fully excluded. No statistical tests are needed to reject either hypothesis. It is evident rather that one side is likely to get about twice as much radioactivity as the other. (The best (least squares) fit to the data gives an asymmetry ratio of 1.81.) An obvious contributing factor to this asymmetry is that the midpoint of the cell, while near the site of the future cell division, is not exactly at that site. Consequently, the potential division site, no matter on which side of the geometrical centre, will contribute a bias to the left side because of the way Verwer and Nanninga (1980) chose to collate their data. However, the magnitude of this effect cannot account for the observations as the following considerations show. If all the tritium atoms were in a finite narrow central zone that is distributed longitudinally with a 5% variation (typical of many E. cofi strains and growth conditions (Trueba, 1982)), and the half-distance of tritium autoradiography were zero, then the asymmetry ratio would be 00: 1. If, as is realistically the case, the half-distance is 0.15 pm, then the asymmetry is smaller; and if the distribution of labelled glycan was 0.2 pm wide, as is

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consistent with the data from cells where the constriction site is visible, then the theoretical asymmetry ratio based on equation (18) would be 2.07. However, when this number of grains is added to the grains uniformly distributed over the cylinder portion, the asymmetry ratio would be 1.28. Even if the zone of central incorporation were 0.4 pm, this ratio is 1.24, or if centred 10% away from the geometrical centre, it would be 1.55. Consequently, because the observed asymmetry ratio is 1.81, I confidently conclude that the anlage of the new division site is not precisely located midway at the start of the division process. I very tentatively put forth the suggestion that in the preconstriction phase the rate of wall growth increases above the rate of diffuse extension growth process at a site on the cell wall that may be quite a distance from the cell centre. However, as that site develops, it moves, or is moved, toward the centre of the cell. This would be consistent with our Renovated Replicon Model (Koch et al., 1981b) as a model both for cell division and chromosomal segregation.

IV. Discussion A. WHAT SHAPE OUGHT A BACTERIUM TO HAVE?

Different shaped micro-organisms occur in nature probably because those shapes are advantageous in the long term to their particular ecological strategy. We can only speculate what the factors involved might be in any particular case, but it is only a matter of design engineering to conclude what shape a cell should have if a single issue were paramount. Usually we think that microbes are small in order to increase their surface-to-volume ratio with concomitant decreases in problems of transport of nutrients and waste materials (Koch, 1971). On this basis they should be as small as possibl-ontingent on any other biological constraints that the organism must satisfy. Clearly, within a fixed volume, a spherical shape is the poorest choice because that shape minimizes the surface-to-volume ratio. A filamentous mode of growth is better and a planar habit is best. Single cells of normal bacteria do not approach a sheet-like structure, but Lampropedia has a colonial morphology of this shape and the newly recognized square bacteria approach a thin sheet. However, the filamentous mode and the planar array are only highly effective when the density of organisms is small so that there is little competition or interaction between organisms or portions of organisms. Clearly, a mycelial habit is an effective way to occupy and exploit a two- or three-dimensional habitat. On that basis, actinomycetes are to be commended. Of course, to achieve this growth habit, they must be able to grow as

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thin, long rods and also be able to branch. Most rod-shaped bacteria normally do not branch, but there are special conditions, usually deleterious, under which rudimentary branches form. But the mycelial growth pattern is in conflict with motility. (Or rather, hyphal growth is a form of motility (Koch, 1982a), albeit a slow mode.) However, for organisms that would be self-propelled in order to search for new environments, or those that would be more easily separated from their relatives or even increase the chance of being redistributed by external forces, a rod shape of fairly short length has desirable features. It combines high surface-to-volume ratio with an ability to become dispersed from siblings. Additionally, it provides the possibility that, with very little additional genetic apparatus, it may conditionally fail to permit cell separation leading to filaments; this may be a useful alternative in certain special cases (viz., during SOS repair). But to grow with a rod-shaped morphology, the organisms must be able to counteract the rounding influence of the internal hydrostatic pressure-or they must eliminate the pressure differences across the cell as I assume the square bacteria have done. Additionally they must be capable of cell division, which is also counter to the force produced by the hydrostatic pressure inside the cell. This means that work must be done and the major problem is to see how chemical work is transmuted into the necessary physical work. Lesser problems are to understand how the regulation of cell length takes place and how the regulatory system is itself regulated by the nutritional environment. There is another way to pose the problem covered in this section. This is that cellular organisms must multiply and divide by an algorithm which operates on a newly arisen organism via a series of rules producing a series of shapes that eventually result in two (or more) new organisms with essentially the same shape as the original cell. The necessity of operating under a group of rules places restraints on cell shapes, and the limitations on shapes become more severe the fewer the number, or the simpler, the rules. The rules, of course, may be precise and yet the resultant cell shapes could have high variability. As an example, Streptococcus sp. have very simple rules indeed. The cells possess immortal wall bands (Higgins and Shockman, 1976),as well as immortal genes, and at the appropriate stage of the life cycle the bands replicate semi-conservativelyand divide with the formation of a single centred septum. Soon the septum splits; and then physical laws, which we now think we are beginning to understand under the heading of Surface Stress Theory, do the rest. A more complicated case is found with Staphylococcus sp. (Giesbrecht et al., 1976) in that these organisms must create de no00 a new site of septum formation, at right angles to the previous one, since the latter is not used in the next generation, although it may possibly be used in some still later

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generation. In this case as well, we think we can understand development of two spherical cells from one spherical cell (see below, p. 351). A still more complicated case is that of the Gram-positive rod since the cylinder and the poles are formed differently. Equally more complex, but in a different way, are the additional operations that must be carried out by the Gram-negative rods. Seemingly, the complications in both cases would be much less if the wall grew under the action of narrow asymmetrical growth zones; then either kind of rod would have almost the simplicity of the Streptococcus, but unfortunately the evidence is against narrow growth zones for cylindrical extension. I believe, as will be discussed further below, that extension growth never takes place in narrow growth zones within the cylindrical regions of hydrostatically stressed cells. The spherical cocci and the rods need to have sub-algorithms to find the new division sites. For the cocci, this means that the plane for the next cell division will be chosen by different rules of different organisms. It seems that the planes of division are quite accurately 90” apart, at least as formed (Koyama et al., 1977; Tzagoloff and Novick, 1977). For the rods, this means a biochemical method is present to measure distances one hundred times the dimensions of a typical protein. There are proposals for the cell’s auto-self bisection (Gierer and Meinhardt, 1974; Koch et af.,1981b; L. Rothfield, personal communication), but they will not be discussed here. The important fact is that some bacterial species do divide in half with better precision than other kinds of bacteria, and certainly many have better precision than arguments about surface-to-volume relationships would warrant. What shape should poles have? Below I will present the shapes generated by certain simple algorithms. But it is possible that the cells may be capable of much more complex operations and “choose” their shape according to other possible goals for optimum shape. One possible goal is to have the strongest shape for resisting the internal pressure. This leads to the Cassinian equation discussed by Oldmixon (1974): (19) where n is an adjustable parameter and all other symbols have the meanings defined elsewhere in this paper. A second design criteria might be to have the external wall increase at a rate proportional to the amount of cell substance: z = I/n[a2-r2+2(a2[a2-r2])1/2]1~2

-=

dA dt

cIr

dv dt

c2v

But for a growing cell: -=

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where c2 is the specific growth rate constant; consequently: dA _ - CI/C2 dv and if growth is cylindrically symmetrical, then:

j dA/j dv = 5 2nr(l +S2)1/2dz/Jw2dz = C I / C ~

(23)

Consequently, the cell shape should be such that: J 1 + S 2 / r = constant

(24)

Figure 19 shows some of the shapes that arise for different values of the constant, assuming that the organism behaved like Streptococcus and creates new growth zones by allowing new external walls to be formed only halfway between a split wall band. Obviously, the shapes produced are, to say the least, atypical-except for the very thin rod which has the property that its volume is proportional to its surface area.

B. STRESS ON PEPTIDOGLYCAN COVALENT BONDS

The Surface Stress Theory takes as its cardinal assumption that osmotic pressure creates enough tension in covalent links in the stress-bearing regions

FIG. 19. Shapes of hypothetical cells that grow so that surface increase is proportional to volume increase. All the curves satisfy equation (24). In curve (a), the constant in equation (24) is chosen so that it is l/r. Consequently S is, and remains, zero. This corresponds to a filament of constant radius of indefinite length which may or may not be septated. Curves (b) and (c) correspond to other choices of the constant. They correspond to cylindrically symmetrical filaments of indefinite length, where each has a central bulge with a sharp discontinuity.

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of the murein to make important differences in the kinetics and thermodynamics of biosynthetic and cleavage enzyme action. It is appropriate here to detail and update the calculation given previously (Koch et at., 1981a). Assume that the volume of an average living E. coli is such that the equivalent radius is 0.8 pm and the osmotic pressure difference is four atmospheres (- 105 kPa). Then, from equation (3), the surface tension is: T = 2aP = 2 x 0.8 pm x 4 atm x (1.013- lo6 dyn ~ m atm-I)= - ~ = 650 dyn cm-' (650. lovs N cm-I).

This tension occurs in all regions of the surface. Let us consider a region where the glycan strands are twice as far apart as in the microcrystalline regions in which the glycan chains are packed 0.45 nm apart in a parallel arrangement (Labischinski et al., 1979). Then, in the direction of the chains, the stress would be (650 dyn cm)-(2*0.45nm chain-')= 5.85. dynes supported by a single chain. Of course, all covalent bonded sugar residues in a chain are stressed by this same amount. In order for a bond to be split, it must be stretched to a distance corresponding to that which puts the bonds in an activated state. This distance is typically 0.05 nm. Of course, if the bond in the wall of a normal cell becomes stretched by this amount, by whatever means, an amount of work (force x di~tance=5.83.10-~dynes.5. cm) of 2.917. 10-l3dyn cm-', or ergs, will be contributed by the stress. Multiplying this result by Avagodro's number (6.023. molecules mol-I), one obtains 1.76- 10" ergs mol-' or 17.6 kJ mol-I. Finally, dividing by the Joule (4.18 cal J-I), one obtains 4.18 kcal mol-I. This much energy would considerably displace the equilibrium point and lower the energy of activation of all chemical reactions involved in the cleavage process; it would raise the energy of activation for formation by the same amount. The consequence of the latter statement is to prohibit peptidoglycan from being formed directly into a linked stressed state. The direct chemical reaction leading to scission of the peptidoglycan to free radicals has a very high energy of activation, while the activation energy for hydrolysis at the glycoside or peptide bond, in aqueous suspension, is much lower. The exact value depends on the H+ or OH- ion concentration. The enzymic energy of activation is lower still and therefore, under the usual conditions in uiuo, is the only pathway with a significant rate. The energy of activation for the hydrolytic cleavage of typical glycoside bonds is 10 to 20 kcal mol-I (Laidler and Bunting, 1973). The decrease associated with the hydrostatic stress of 4.18 kcal mol-1 is thus quite significant because the energy of activation enters the calculation of rates in an exponential way. Now let us go over the calculation again. The actual Gram-negative organisms for which we have estimates of the osmotic pressure are rod-shaped

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instead of spherical. The tension developed in established poles is less than in the cylindrical part because “a” is a little smaller than assumed above, so T = 2Pa is smaller. For the cylindrical part we should have used T = Pa from equation (4), but the problem is a little more complex and the stress developed parallel to the axis is twice that in the direction in the cylinder normal to the axis. I leave this question for now with the note that shape may influence the energy over a two-fold range. The second assumption made above was that the pressure was four atmospheres. In the previous calculation I used five atmospheres. The latter is consistent with the scanty experimental measurements, but neglected the osmotic pressure of the usual medium which partially tends to counteract the internal pressure. The distance between glycan chains quoted in the previous calculation is that for X-ray data of walls of the Gram-positive Staphylococcus aureus, and necessarily for unstretched material. On inclusion of a factor of two to approximate this stretch, the calculated stress is considerably increased because there would be half as many chains per unit cross section supporting the tension. The tension on the peptide bonds on average should be considerablygreater than on the glycan chain. The wall does not appear to be a regular solid with a well-defined repeat distance; however, there do appear to be regions of microcrystallinity.In such regions, the stress on peptide chains would be much greater because the peptide bonds are more widely spaced than the spacing of the glycan chain. In unstressed material, the peptides occur every 2.06 nm along the glycan chain (and the glycan chains are 0.45 nm apart). Evidently there is considerable chemical variation between the wall structure in different organisms. Many peptide bonds are not involved in bridge formation because of carboxypeptidase action, because of covalent binding to lipoproteins, or because they are not properly oriented. Considering these factors, a lowering of 15 kcal mol-’ in energy of activation for hydrolysis in stress-bearing wall seems therefore quite reasonable. For the average peptide bond, a 15 kcal mol- lowering of the energy of activation would speed the cleavage process by 4 - 10l0-foldat 37°C under conditions in vivo relative to conditions in vitro. This is the basis of the postulate that the cleavage rate permitting enlargement of the wall is linked to the cell’s increase in volume via the concomitant pressure increase. A pressure increase would follow very quickly when the rate of wall expansion is not sufficient to supply adequate intracellular space for newly forming cytoplasm. The pressure would increase tremendously and very rapidly because water is nearly incompressible. This would quickly be compensated by an increased rate of hydrolysis, under normal conditions, due to increased stress. If other things remain constant, a 1% increase in pressure would increase the rate of the cleavage by 27%, and a 2% rise increases the rate by 63%. An increase in rate of 4. 10l0-foldis so large that the actual enzyme(s) that

’

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may do the cleavage step(s) important for cell-wall enlargement may never have been detected by biochemists with only unstressed murein at their disposal. The enzyme(s)may, in principle, remain undetectable in vitro until a system is developed that is sufficiently sensitive to follow the scission of a single chain. In such a system, the first break of any link in the chain results in the remaining bonds becoming virtually completely stable. Several years ago I discussed the possible existence of a class of enzymes designated as “the reflexive enzymes” (Koch, 1967). These hypothetical enzymes had the property that they bound so tightly to their products that further reaction would not occur until the product was secondarily changed. The cleavage enzyme considered here also would have the same property of being not directly detectable, and whose existence depends on either inference from results in vivo or the elaboration of a very special enzyme assay. An essential aspect of the Surface Stress Theory is that the cleavage process cleaves only stressed bonds. While the cleavage is quite easy to imagine, the selectivityconnoted by the the word ‘only’is difficult to understand. For now, we must postulate that the cleavage process acts only in ways that do not jeopardize the wall integrity and trust that future work will give an explanation of how this is accomplished. There is one case where an explanation is at hand-this is the inside-to-outside growth of cylindrical extension in Gram-positive rods, where it has been shown that the autolytic enzyme cannot normally act adjacent to the cytoplasmic side of the stressed cylinder wall because it is inhibited by the protonmotive force (Jolliffe et al., 1981).

C . SURFACE STRESS THEORY FOR CYLINDRICAL ELONGATION

One of the central problems of bacterial morphogenesisis how the rod-shaped cell maintains a cylindricalform in the face of the two-fold differential stress in the circumferential and the axial direction. If wall growth is not taking place and the sacculus is sufficiently bridged with covalent linkages, there is no problem; but, during surface extension, cleavages must be made and then the stresses would seemingly tend to round up the cell envelope. The only non-surface stress theory explanation of which I know is that there is some sort of physical template that is copied precisely as the elongation process continues. Thus if ( I ) new glycan chains of the order of 1200 disaccharide units long were arranged circumferentially around the entire perimeter of the cell, if (2) during wall growth these new glycan chains were stretched and converted into a new circularly closed structures with exactly the same number of residues as the adjacent chain, via some semiconservative replication process (that might depend on hydrogen bonding to existing

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circular structure), if (3) the new circle was attached via peptide links to only the two adjacent covalently linked circles, and if (4) only the stress-bearing bonds holding these two circles together were cleaved, then template directed cylindrical extension would be possible. But this hypothetical mechanism must be excluded for several reasons: (1) the glycan chains are believed to be far too short; (2) there would be no mechanism for the wall diameter to decrease as it does during a shift down of Gram-negative rods; (3) there would be a problem of transport and closure of the circles in all layers but the innermost one in Gram-positive rods; and (4) free energy somehow would need to be supplied to the nascent glycan chain to have the same spacing distance as the stressed template circle. Something like a template action might help in cylindrical extension, but it could not, in the long run, maintain the rod shape and the diameter generation after generation. There are three possibilities under the Surface Stress Theory. The first, which I think highly unlikely, is a narrow growth zone model. The autoradiographic evidence is sufficient to exclude a single precisely placed zone for Bacillus megaterium. It can also be excluded on thermodynamic grounds for all cases, except for the case where conditions are such that the peptide and glycan bonds form in the most extended conformation. Moreover, narrow zone cylindricalextension is inherently unstable. Equation (7), for the case in which there is no septum (8 =0):

S = ,/(Pr/2T)2- 1, (7) was quoted above. As long as PrI2T is equal to 1, the slope stays zero and cylinder extension is possible. However, a fluctuation giving a slight outward slope due to an increase in P or r or a decrease in Tis catastrophic as shown in Fig. 20. This is because the slope of the growth zone will continue to increase with time even though P and Tremain constant, or even return to their normal values. Figure 20 shows the profile of a hypothetical narrow zone cylindrical cell on further growth, if initially Pr/T was perturbed from its value of exactly 2. The only ways to prevent the narrow zonal growth catastrophe are ways that make it subordinate to another mechanism that has some other way to establish the diameter of the cylindrically growing region. The other three possibilities have been discussed in some detail elsewhere (Koch et al., 1981a,b, 1982a), so I will only outline them here. One is that the cell has previously adjusted its radius so that Pr/T= 1 and it then carries out diffuse growth, which then produces cylindricalextension. This is the case that we have suggested which may actually apply to the side walls ofGram-positive rods. As noted above, it will only function if the poles are rigid. When Pa/T is not unity, on further growth, the cell will achieve a radius so that PaIT is one. That is to say that momentary fluctuations in P, a or Tare self-correcting. This possibility depends on the assumption that P and T remain constant

w

P 01

/ ;;ar;w

!-

growth

is - - _ _ - - - - - - - - -A x-

FIG. 20. Catastrophic increase in a narrow growth zone as the results of a momentary fluctuation in P, Tor a. The profile of the cylindrical portion of the cell is shown for the hypothetical case that growth occurs at a single narrow zone. In the initial state, the cylinder has been elongating because PaIT has been exactly 2. As a result of a chance and then returned to exactly 2. This sets off a chain of events so fluctuation, momentarily Pa/T increased by that successively the cell widens at the growth zone. These increases lead to a top-shaped cell.

60

I

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under normal conditions for at least most of the cell cycle. In the simple form of the theory, P is constant because the cleavage rate tends to increase when there is an increase in pressure. T, on the other hand, is a parameter that depends on details of the chemistry of murein formation. If the biochemical mechanism stays constant, and the physicochemical environment stays constant, T will remain unmodified. The second possibility is that Pr/T is slightly greater than unity and the cell bulges a little as it elongates. If the process of cell division causes a gradual constriction, then there would be compensation during a complete cell cycle. The third possibility is that cylindrical extension takes place by apical growth. This can occur even for an organism multiplying by fission, as has been shown in the case of Schizosaccharomyces pombe (Johnson, 1965).

D . POLE FORMATION

Binary fission as executed by the schizomycetesis an energy-requiringprocess. That this is so follows from the trivial fact that soap bubbles only enlarge and do not divide (or separate from the bubble pipe) unless coupled to an energy supplying process. This implies that when cellular life first evolved, mechanical action-such as waves-was needed to cause cell division. Cells higher than the schizomycetes possess cytoskeletons and proteins that can convert high energy phosphate bond energy into the mechanical work needed for cell division. Bacteria, on the other hand, do not have these kinds of proteins and the Surface Stress Theory suggests that cell wall formation is the mechanism of energy coupling in the cell division process. There appear to be four basic patterns (but much variation occurs in each). The first is the narrow growth zone model and, in particular, the zonal dome model. The second might have been the way of fission of early wall-less creatures and may apply today to Mycoplasma sp. It depends on the properties of the exterior surface being different from the interior (septal) surface. The third is the “split-and-stretch model” that may apply to the Gram-positive organisms that do not obey the zonal dome model. The fourth is “the variable Tmodel”. This model contains the presently viable options for growth and division of Gram-negative bacteria. 1. Narrow Growth Zone and Zonal Dome Models These models have been discussed by Koch et al. (1981a, 1982a) and the relevant equations have already been presented. For comparison with the other models, it must only be pointed out that the derivation assumes that the

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septum is built of pre-stressed murein and that this material occupies the same volume when laid down, as when it later becomes externalized. This assumption is approximately correct for Streptococcus faecium where it has been found that the volume decreases 19.4%on fixation and dehydration (L. Daneo-Moore and M. Higgins, personal communication). This corresponds to a 13.4%increase in area or an increase of 6.94%in linear dimension when the wall is under tension in the living condition. Differently shaped poles arise if the ratio of the thickness of the splitting septum to the external wall (6) has different fixed values, or varies during the pole formation. Additional wall needs to be made and inserted into the wall at the site of septal splitting. The work in extending the conformation of this additional wall is derived from the pressure volume work according to equation (1). While the model seems to fit certain cases, the unresolved questions are “How is the new wall, which is intercalated at the splitting septum, inserted through the thickness of the wall?’ and “How does the septal wall form in a pre-stressed state?’ 2. Primitive Cell Division Mechanism A soap bubble without additional constraints is spherical because this

minimizes the surface for the given volume. When two spherical soap bubbles of equal volume fuse, the surface between is planar and the spherical surfaces make a 120” angle with the planar septum. After this happens, there is no tendency for the system to increase or decrease the overlap of the two spheres: it is in an equilibrium position. Similarly, if a wall-less cell could form a transverse membrane and then could initiate a splitting of that membrane, the membrane would split to the most stable position, just like the two soap bubbles, and remain in that undivided shape. Figure 2 1 shows this diagramatically. It shows the external surface, septal surface and total area. So starting either with a sphere with a septum on the left of the figure, or two touching spheres on the right, the system would spontaneously move to the configuration with the lowest total surface area where the overlap becomes 0.3817, as indicated by the arrow on the curve marked 6 = 1 . Thus, the pocket computer demonstrated what the calculus of variation or soap bubble pipes showed long ago. But, with the computer, we can now ask what would happen if 6 were different on the planar (septal) surface from that on the external surface. The figure shows that if 6 were 2 or greater, then the balance point would be shifted so that division and separation would occur spontaneously. This provides a basis for cell division: in order for the cell to divide, it must arrange circumstances so that it takes more work to make the septal material than the external wall. Thus, the work will be built in, as the septum was forming, to later favour the cell’s fission. The diagram is drawn as if that work

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requirement is satisfied by having a septum of rigid pre-stressed material S times thicker than the external covering. I did this to make an analogy with the zonal dome model, and in this form the model may explain the energetics of the formation of the spherical cocci, such as Staphylococcus sp. But it could also apply to cells without a rigid wall if the lipid bilayers making up the cytoplasmic membrane are such that the inner and outer

6= 1.0 6: 0

7---

10

I

8 6.0.5

6:l.O

6 22.0

6:lS

51 SEPTAL 6.1.0

0

0

0.2

0.4

0.6

Overlap (h)

FIG. 21. Model for division of primitive cells, mycoplasma and staphylococcal cells. Shown are the shape that two hypothetical fused soap bubbles would have if the properties of the interface was different from the properties of the external surface.

leaflets have a different composition, and that the septal membrane has a composition of the inner leaflet on both sides. In such a case, Tis different for external bilayer and septal bilayer. In this context, the thesis developed here becomes: if the T value of septal wall is equal or less than twice that of the external wall, then division will be spontaneous. This kind of mechanism may apply to a variety of spherical creatures such as L-forms and mycoplasmas.

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3. Split and Stretch Mechanism for Gram-Positive Bacteria

This type of mechanism has been briefly described as a possibility for Gram-negative bacteria (see Table 1 of Koch, 1982b) designated category V. In that paper, reasons have been presented that make it unlikely that this mechanism applies to Gram-negative bacteria; however, because of the findings already presented, that the shape of the pole changes during its formation, I now suggest that it applies to some Gram-positive rods. The Gram-positive organism forms a septum that eventually separates the cytoplasm into two compartments. This is done under physicochemical conditions that are different from those during the formation of the outside wall, whether by inside-to-outside growth of cylindrical regions, or by externalization of the previously formed septal material. The split-and-stretch mechanism assumes that there is no additional wall incorporated into the splitting septum, but there are several possible courses that the stretching can follow. One is that the septal wall is formed in some special way so it does not stretch when stressed; e.g. “split and no stretch”. This could account for the flat poles sometimes seen with Bacillus anthracis. If no stretchhg at all is possible, there is no problem in understanding how the organism maintains a fixed size. On the other hand, if the wall as laid down in the septum is extensible, poles of other shapes are possible. Several possibilities are depicted in Fig. 22 for the case where walls can eventually stretch by two-fold in area. If the split wall stretches rapidly, and the unsplit wall does not stretch at all, then a conical pole will form. If the unsplit wall stretches within its capacity to occupy the cross-sectional area of the cell, then a pole will form which, in part, is an extension of the cylinder and has a flat end. Either form could be converted into the hemispherical pole shown, with no additional material, just by the splitting of relatively few bonds that will experience an extremely large stress and result in a negligible change in surface area. Consequently, the best explanation that we have to date of the shape changes of B. subtilis presented above is that the septal disc stretches to some considerable degree as the peripheral splitting takes place, and the stress from the hydrostatic pressure acts on the unsplit septum. Therefore, the nascent pole has a highly curved shape near the cylinder and is quite flat near the axis. Subsequently, the stresses and strains work themselves out and a nearly hemispherical pole results. It would be exactly hemispherical if the total expansion were exactly two-fold. It would be a portion of a sphere of some slightly different diameter if the expansion factor were smaller or larger than two. Only if the stretching characteristics of the unsplit septum were special would a perfect hemisphere be produced as the splitting took place. Consequently, it is clearly of interest to

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353

\

I

FIG. 22. Split and Stretch Model. On the assumption that the murein fabric can

stretch two-fold in area when stressed, the shapes are shown that a pole could assume. Three cases are shown: (1) if stretch occurred as soon as stresswas applied (pill box); (2) if stretch occurred only in the split region (cone); (3) if stretch continued until the equilibrium occurred (hemisphere). The lower half of the figure shows the unsplit septum. make similar measurements of curvature and/or angles of old and forming poles of a variety of species of cocci and bacilli. There is a problem with the split and stretch models for pole formation: what keeps the septum from stretching to its full extent as soon as the septum starts to bear the stress of the external surface? Seemingly, this gives the cell end some bulbous character. Sometimes, in bacilli, a bulbous shape develops

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as spore formation starts, but not normally. Said differently, the split-andstretch mechanism creates a problem at the same time that it accounts for our observations; namely, we have assumed the cylindrical extension inside-tooutside growth was dependent on rigid poles, and here we assume that the pole shape is dependent on being attached to a rigid cylindrical portion. One way out of the dilemma is to suppose that there is a gradient of lytic action on the cylindrical portion of the cell. If near a developing pole the lytic action is slower than that on the rest of the cylindrical wall, then during the splitting and remoulding of the septum, which happens relatively quickly, the cylinder could behave as a rigid support. Subsequently, the pole would be rigid enough to support cylindrical elongation. Under normal conditions it is possible that there is a gradient of autolytic activity, being smaller in the cylindrical part near the poles and being smaller still on the poles. Since there is convincing evidence that wall material is added at a slow rate to established poles, there are two possibilities. Either there may be some slower inside-to-outside growth at the poles, and sooner or later the cells should become rounder, or the pole wall should progressively become thicker. With ways now available to identify old poles, these two possibilities can be distinguished. The critical experiment is to study the ultrastructure of those rare cells in a growing populationthat have one pole that is many generations old and the other less than one. Measurements on ten such cells would be more informative than measurements of tens of thousands of cells chosen at random.

E.

WHERE DO THE CONSERVED A N D NON-CONSERVED REGIONS JOIN I N GRAM-POSITIVE RODS?

Several types of experiments give indications that during normal growth, poles in cells, and septa in filaments, are more conserved than the side walls. The only published work inconsistent with this is that of Fan et al. (1974) who did a density isotope shift experiment. I surmise that this exception resulted from growth not being balanced. This is one limitation of the use of the density labels. Pole walls under many conditions are more resistant than cylinder walls (Doyle et al., 1981) and, as Burdett (1980) has clearly shown, under certain metabolic conditions the poles of intact cells degrade more rapidly near the cylinder. The experiment that most clearly delineates the junction of the two regions is the temperature shift experiment leading to blockade of further glucosylation of the teichoic acid. In pictures published previously (Koch et al., 1981b), as well as those shown in Fig. 19, it can be seen that the old pole retains an ability to bind fluorescent concanavalin A that side walls and new poles do not possess. The region showing fluorescence is of a thickness roughly equal to the

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radius of the cell. Almost as clear are the phosphate shift experiments of A. R. Archibald (Figs 11 and 12). They lack precision only in that the tails of the virus which is an indicator of phosphorylated teichoic acid are so long as to extend into a region of non-phosphorylated teichuronic acid. Both experiments can be criticized in that they follow the progress of a substance (glucosylated teichoic acid) that is not part of the stress-bearing part of the wall. However, the teichoic acid is covalently bound to the murein, and the work of Mauck and Glaser (1972) and Schlaeppi et al. (1982) clearly shows that teichoic acid and peptidoglycan are fragmented, during growth, in the same way. Additional evidence can be drawn from the median sections of B. megaterium and B. subtilis (Frehel et al., 1971), on effects of chloramphenicol and recovery from chloramphenicol. Pictures such as Fig. 4 (p. 3 15; supplied by Ian Burdett) show that the appearance of the wall changes with its age: new-forming pole surfaces are quite “clean” in appearance; progressively, the poles look rougher. The junction between such regions and the bulk of the cylinder, which is moderately, but uniformly, rough is evident in most pictures. Usually, the border between morphologically distinct regions occurs at the site where the cylindrical region meets the curved pole. But there are occasional exceptions to this pattern. In all of the exceptions, the border is on the cylinder not very far removed from the pole. An adherent of the Surface Theory must object strenuously to the use of autolysin-deficient mutants in studies of wall morphogenesis. Clearly, the available mutants still cleave some wall bonds and do grow at essentially normal rates, but as filaments of cells. This means that enough enzyme remains, or ordinary chemical reactions suffice, to cleave enough stress-bearing bonds to permit extension. But the situation is rather like a tug-of-war in which the weakest link, or the one that by chance has the most stress, breaks. Then even higher stress appears on a few other bonds and the result is that old cylindrical wall becomes associated with one or the other of the old poles. Therefore, I am mildly surprised that Schlaeppi et al. (1 982) did not find 100% of the label retained at septa1 sites. Rather, in minimal medium cells they found 48%, and from broth cells 24%, both numbers in excess of the contribution of the poles themselves (see below, p. 357). Since the experiments of Schlaeppi et al. (1982) can be taken as evidence against the Surface Stress Theory, I must discuss in more detail the consequence of an autolysin deficient state and return to the tug-of-war analogy. “A thread [stretched from the ends] breaks where it is weakest” (Herbert, 1740). On the contrary, if stress is developed over the length of the chain, then it is highest at the centre and the chain will break quite near its centre, as first pointed out by Hershey. Later, enough stress may develop elsewhere to give additional cleavages. For the autolysin-deficient cell, this may mean that the final resting place for old wall material can be a septum or a

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L. KOCH

pole that formed later. The reader may experiment by stretching a piece of a rubber band until it breaks, then get a friend to hold the rubber band in the middle while you pull on both ends. Alternatively, try stretching a piece of cloth. Many times the cloth will not rip if pulled from the ends, but will rip if the cloth is stretched at an edge between the fingers and thumbs of both hands with the hands in very close conjunction. This is because the stress is distributed over a larger amount of the fabric in the former than in the latter case, and it is the maximum stress felt by a bond that is higher in the latter case and starts the rip, and then the stress is relieved at most other points in the fabric and intensifed at others. The work from Archibald‘s and Doyle’s laboratories cited above has the advantage that the morphogenetic conclusion does not depend on quantitative measurements, simply on the localization of phage or fluorescence. Quantitation for the light microscope autoradiography experiments of Schlaeppi et al. (1982) is beset with a number of difficulties. There are two types of quantities to be measured: the numbers of conserved zones after a chase and the total amount of label conserved in all “clusters.” A continuously labelled culture of autolysin-sufficient dividing cells should give rise on long chase to one cluster for each extant pole and two for each nascent division site at the time’of the chase. Depending on strain and growth conditions, the number should be greater than two by an amount that depends on the number of on-going sites of septation. For the filamentous habit of growth of autolysin deficient mutants, the problem is different, largely because the concept of a cell is more nebulous; it becomes precise if we take Lb, length of a baby cell, as the length of cylinder between two adjacent septa that are observed to be just closing. Then for a length of Lb times the factor of growth increase since the chase was started, we expect two clusters plus two clusters for each intervening nascent septum present at the time of chase. In addition to this, some of the septa that develop later may give rise to two more clusters each. This is because even the partial splitting of a septum reduces the stress developed in the outermost parts of the wall so that cracks will have a larger chance of forming in the middle of cylindrical regions. In counting such units, it was the convention of Schlaeppi ef al. (1982) to consider non-end clusters overlying a septum as two clusters even though they would only become associated with separate cells if cell division could become re-instituted. To calculate the length of an average cell, the result must be multiplied by 1.4 (see Koch and Schaechter, 1962). Filamentous cell growth certainly can aid in the analysis of the wall growth of rods; however, it must be done under conditions where the filamentous habit is conditional. When labelled cells are chased under conditions where the pedigree of different parts of the structure was rigorously defined as, for example, by initiating filamentation at the initiation of achase, the counting of

SURFACE STRESS THEORY OF MICROBIAL MORPHOGENESIS

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conserved regions would be very much more significant. But this advantage is lost under conditions where filamentsgrow and fragment at random to create new chains. Then the analysis is much more difficult, both for counting the numbers of conserved units, and also the total amount of material, because the starting condition for the chase is ill-defined. Other problems arise because of the lack of normal fission mechanism. This problem is made evident by reference to the work of Fein and Rogers (1976, also see Rogers, 1979).They found that the autolysin-deficient mutants would occasionally break in the middle of filaments and, from inspection of their picture, this does not hapen at the normal sites of fission, but in such a way as to fragment and kill the cell. This leads to portions of empty cell wall attached to the end of a growing filament. How much additional conserved wall of the dead cell has remained attached to the pole of the living cells is unknown. In the photographs published by Fein and Rogers (1976), a large amount of the wall of dead cells on some living cells was evident, but the nature of the ends of most filaments was not clear. Another criticism of the quantitation aspect of autoradiography, which must be mentioned, arises in technical aspects of the autoradiography itself. When slides of whole cells are dipped, the emulsion around the cell may be thicker than on top of the cell. Therefore, disintegrations emanating from the top, and certainly the bottom, of the cell may have a lower efficiency of being counted than those arising on the side. Consequently, since a pole has more “side”, its efficiency should be higher. Note that this is a criticism of not only Schlaeppi et al. (1982), but of all experiments with whole mounted cells, including the experiments of Verwer and Nanninga (1980) that were re-examined in Fig. 10. This error could have been estimated if uniformly labelled cells had been analysed also. Let us consider the proportion of wall material in the pole part of a cell relative to the total. To start, assume that the rod has hemispherical poles, of radius r and pole-to-pole length 1. Then the pole surface area is 411r2,and the cylinder area is 2nr(l- 2r). Consequently, the percentage pole of the total wall is: 2r

%P, = 7100 Although r is nearly constant for a Gram-positive rod, 1 varies with the position of the cell in the cycle. If we deal with populations of growing and regularly dividing cells, we should choose an average. Without detailed calculation, it can be argued that this 1 is quite near the birth size. To do the sums properly, we must calculate the mean % P,between the birth size and the length when wall, for the next division site, starts to form. This, in turn, would be averaged with a value for that part of the population that has started a new

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division site before it has completed the last. Calculating the % P,in cells of this class is more difficult; the pole wall content is higher, but their length is longer. Even if we assume that the wall is of the same thickness all over the periphery, in the septum its thickness is greater and varies somewhat during development. But it is approximately twice the thickness of the peripheral wall, and of a material that may have twice the concentration of murein substance per unit volume than it does when stretched or has become stress-bearing outer wall. One more factor to be taken into account concerns details of the development of the shape of the nascent pole. How fast does the inward growth occur? How does the shape change from planar to become more hemispherical? In carrying out these averages, it is necessary to take into account that in a growing population there are, in the ideal case, four times as many cells of a fixed size range near the mean birth size than there are at the division size (Koch and Schaechter, 1962).Consequently, all in all, the various factors involved would be consistent with choosing 1 a little larger than the length of a newly formed cell l b . In many cases, 1 will be only two to three times the diameter and consequently, %P,will be in the range 30-50% of the total, unless the cells are quite long and thin. For pulse igcorporation experiments, the problem is significantly different. If poles have no turnover and the transit time for material passing through the wall is exactly one doubling time, then for each generation, new cylinder of area 4ar(lb- r ) must be made; two new poles of collective surface area 4nr2 also must be made. Consequently, if all phases of the cycle are represented:

%Pp = (r/&)100 (26) Evidently, a more elaborate calculation could be given that takes into account the population distribution and the other factors considered above. The results are not much different and I shall spare the reader. But equal time for Schlaeppi et al. (1982). First, the value of 13%or 15% for %P, given by Burdett (1974) and Burdett and Higgins (1978) is much smaller than the 30-50% suggested above. Burdett has since measured many more cells of B. subtilis 168s growing in a glucose medium with attention to including cells of the various size classes as they occur in balanced populations. So, we had his computer combine the appropriate cell wall components for 292 median sectioned cells into pole and cylinder. The calculations also include allowance for cell wall thickness, and the results were %P, = 16.51%, quite a bit less than the observsd fractions of conserved label reported by Schlaeppi et al. (1982). Our calculated value of %P,is low if septa1 wall is denser than stress-bearing wall. On the other hand, it is certainly high because rough analysis of Burdett’s electron micrographs show that the filamentous mutants studied by Schlaeppi and coworkers have a higher l/r ratio than does the strain 168s wild type, and also have thicker cross walls.

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From the analysis presented above there is every reason to believe that pole material is more resistant in vivo but does, indeed, slowly turn over. Just which portions of the pole, how much, how fast, and to what degree culture density affects the renewal are questions not presently answerable. At present, I believe we must leave open the possibilities which range from a great deal of pole turn-over at high culture densities, with autolysin-sufficient cultures, to circumstances where part of the cylinder is conserved. One possibility, pointed out by J. Mandelstam, is that the junction may occur in the cylinder at the place that spore formation normally initiates. This locus is 20%from the end of the cylinder in B. subrilis (J. Mandelstam, personal communication). Another possibility is the split and stretch mechanism proposed above. If the factor of expansion of the septal wall is greater than two-fold, then one could expect a region of cylindrical appearance that was mechanistically part of the pole. Finally, it was clearly too simplistic to imagine that at the junction site on the pole side there was no turnover, and full inside-to-outside growth in the cylinder; there must be some intergradation. So, it is not unlikely that there is some inside-to-outside growth of pole material and some slow-down of inside-to-outside growth of the cylindrical wall immediately adjacent to the poles. A critical answer to these questions of Gram-positive rod morphology will be given when studies are conducted such that at a specific time the external lytic enzyme in dilute suspensions of autolysin sufficient cells is destroyed or inactivated completely so that only enzymes made in particular regions of a cell may function where secreted from the cytoplasmic membrane and not elsewhere in the same or other cells.

F . VARIABLE T MECHANISMS

The quantity T, basic to the Surface Stress Theory, effectively summarizes the details of the energetics of the polymerization and transformation of murein to achieve the stress-bearing state. It should be a constant, if the biochemistry and physiology are constant. Some different mechanism must operate, for the Gram-negative organism where the wall is so thin [one-third of an unstretched monolayer (Braun, 1975)] and the option of forming branches in the covalent structure or thickening on the inner fold to produce septal anlage, that later split, is unlikely. We have suggested that this implies a qualitative or quantitative localized change in the polymerization process (Koch, 1982a,b; Koch et al., 1982; A. L. Koch, C. L. Woldringh and I. D. J. Burdett, unpublished observations). The theoretical considerations presented in these papers show that shapes of many kinds could be produced. Many of the possibilities can be envisaged directly from equation ( 5 )

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ARTHUR L.. KOCH

P = T/r. This equation applies to a cylindrical bubble and approximates what the equilibrium shape would form. First, notice that it is counter-intuitive: a larger pressure corresponds to a smaller radius. This is related to the phenomenon discussed above: a bubble, whether spherical or cylindrical, is metastable when connected to a supply of constant pressure. If a fluctuation takes the septum away from satisfying equation ( 5 ) , the bubble will become either infinitely small or large. (Of course, the stability changes if the ends of a cylinder have fixed radii; see p. 347.) The point of importance here is that P must be the same all over the interior of a cell, but T can be different in different regions of the growing wall. If T does vary, then the radius, at equilibrium, in different portions of the cell will be proportional to T. For example, if the cell can alter conditions so that T becomes ten-fold less in certain regions, then the cell will become ten-fold constricted in those regions. This could be exactly the situation for certain tubular appendages, such as are present in Caulobacter and Hyphomicrobium, for example. This could also be the situation in growing hyphae of mycelial organisms (Koch, 1982a): the biochemical mechanism causes T to be very low at the growing tip and decreases with distance from the apex. However, equation ( 5 ) for a cylinder can be a very bad approximation for the shapes of cylindrical bubbles. This can be seen by imagining a cylindrical bubble of radius a, satisfying P = Tc/a, connected to a spherical bubble radius a, satisfying P =2TJa. Evidently, both equilibrium situations could be simultaneously satisfied if Tc= 2Ts. Thus, a rod-shaped bacterium with hemispherical poles would be produced if the cell changed conditions so that T became halved in a central region. Note that the logic of this and the previous paragraph is quite different: a two-fold decrease in T, if done gradually, leads to a cylinder of half the radius; but an instantaneous change leads to reduction of the radius, measured from the axis of symmetry to zero, with the production of a hemispherial pole (see Fig. 23). In our recent work (A. L. Koch, C. L. Woldringh and I. D. J. Burdett, unpublished observations) we have compared the theoretical shapes and the actual shapes of nascent poles. The nascent pole shape is quite different from the finished pole (Table 4; p. 332), and the shapes observed can be fitted to the following pattern of the developing pole in E. coli: (1) wall growth takes place all over the surface; (2) it occurs to a smaller degree in old poles (presumably this is a kinetic matter); (3) at the site of future division, which is not initially very near the centre of the cell, there is a change in the conditions of polymerization so that T is decreased three- to five-fold; (4) this causes a great increase in wall synthesis locally and accounts for the well-known zones of intensive incorporation, and for the inward growth calculated from the formulas of equations (10H13). It could not account for the final cleavages that separate the cell into two daughters because that must involve some

SURFACE STRESS THEORY OF MICROBIAL MORPHOGENESIS

36 1

I A x i a l distance

FIG. 23. Shapes generated by changes in T. To the left of origin, designated by 0, cylindrical growth has and is occumng with Pa/T= 1. A shows the hemisphericalpole that results if T to the right of 0 has half the value of that on the left. The rest of the curves depict the shapes produced if Tchanges to the same degree, but more gradually. The rate of change of T is greatest for Band decreases 10-fold for each successive curve, C, D, E and F. The rate of change for curve B was such that T reached it 2-fold smaller unit value at the axial distance of 1. Note that this change in T, if very gradually approached, would eventually create a rod-shaped cylinder whose radius was 0.5.

different kind of chemistry since the sites of insertion and cleavage are differently related to each other than where the wall is simply enlarged. Lastly (5), the change in T gradually reverses itself and the pole shape gradually becomes very nearly hemispherical as Tbecomes half of that characteristic of the elongating sides. Further changes in shape are minimal either for kinetic reasons or because the equilibrium shape that is characteristic of lower values of Tare no longer possible; i.e. the cell cannot return to the original cylinder shape. The variable Tmodel, although inspired by the work of Pritchard (1974), is quite different. Pritchard’s Iog-linear model (also proposed earlier by Previc (1 970)) assumed that the pressure would progressively decrease as wall and protoplasm grow according to their own rules, and this would lead to division. Here it is assumed that the pressure is constant due to the coupling of wall synthesis to protoplasmic growth. Division might be triggered by a momentary increase in pressure, or the pressure might decrease because the change in biochemistry affecting T might temporarily alter the coupling; but basically,

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and normally for this case as those appropriate to other types of bacteria, it is the hydrostatic pressure that forces cell expansion and cell division.

V. Summary

From the physics of the situation, one might conclude that the osmotic pressure within most prokaryotes creates a sufficiently high tension in the wall that organisms are at risk of ripping themselves apart. The Surface Stress Theory holds that they avoid this, and are able to carry out certain morphogenetic processes by linking the cleavages of appropriate bonds to enzymes that are sensitive to the stress in the bonds under attack. This tends to maintain the internal pressure and couples wall growth to cytoplasmic growth. Mechanisms with widely different geometry function for different organisms, but they have in common the requirement that new murein be covalently linked, and usually in an unextended conformation. Organisms differ in the site of wall addition and site of cleavage. In the Gram-positive Streptococcus, septum formation, and septa1 splitting occurs with little stretching of the unsplit septum. In Gram-positive bacilli, the cylinder grows by the inside-to-outside mechanism, and the poles appear to be formed by a split-and-stretch mechanism. Gram-negative rods, with their much thinner wall, resist a spherical shape and are capable of cell division by altering the biochemical mechanism so that initially one-third to one-fifth of the pressure-volume work required to increase the area of the side wall is needed to increase that in a developing pole. The growth of hyphae is a separate case; it requires that muchless workis needed to force growth ofthe apex relafiveto the side wall. Some other bacterial shapes also can be explained by the theory. But at present, it is only a theory, although it is gradually becoming capable of accounting for current observations in detail. Its importance is that it prescribes many experiments that now need to be done.

VI. Acknowledgements

This work was supported by the National Science Foundation under Grant 79-1 141 and was carried out during the tenure of a Guggenheim Fellowship and written while a scholar at Centro Culturale della Fondazione Rockefeller. It was developed in Amsterdam, Bellagio, Bloomington, Edinburgh, Lausanne, Likge, London, Louisville, Oxford, Paris, Philadelphia, Tubingen, and while travelling in-between. It contains the ideas and experimental data of

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(and the results of disputations with) Ron Archibald, Ian Burdett, John Chadam, Chantal de Chastellier, Lolita Daneo-Moore, Willie Donachie, Ron Doyle, David Fan, Jean-Marie Ghuysen, Mike Higgins, Dimitri Karamata, Hendrik Lauwerier, Joel Mandelstam, Nanne Nanninga, Mary Jane Osborn, Harold Pooley, Bob Pritchard, Howard Rogers, Antoinette Ryter, Uli Schwarz, Frank Trueba, Ronald Verwer, David White, Bernard Withold, Conrad Woldringh, and Jan Wouters. The work was advanced with the technical help of Vickie Arwginski, Fred Brakenhoff, Tom Doody, John Green, Tom Lehrer, Stine Levy, John Leutscher, Nikolai Lobachevski, Hewlett Packard, Anton Philips, Cary Wang, and Jane Whalley. A11 lists in alphabetical order. All input essential to the development of the theory. Scientists are a proud lot. And as proud as they are of their data, they are even prouder of their ideas. In an article I published 12 years ago in this series (Koch,. 1971), I made a point of acknowledging the key ideas due to my students in the development of the philosophy of the feast and famine existence of bacteria. For the present paper, I have taken the ideas of others, although this time they are those of mature and successful scientists of many disciplines in whose laboratories I have worked and visited and with whom I have drunk the local beverage and argued at the bench, the pub, the garden, etc. It was Conrad Woldringh who stressed the importance of the distinction between septum formation and splitting versus constriction. He also pointed out the logical inconsistency of peptidoglycan incorporation at the splitting septum of Streptococcus and the significance of the size changes as viewed by various fixation procedure versus the living state as viewed in the phase light microscope. Jean-Marie Ghuysen pointed out the logical impossibility of wall growth at points remote from the cytoplasmic membrane. Ronald Doyle sparked my interest in the field eight years ago and developed the concept that protonomotive force could provide a mechanism for controlling wall thickness. Ian Burdett posed the question of how much error is involved in the assumption that approximately median sections are exactly median. He also provided me with electron microscope pictures of many bacteria and gave me access to his large data base of measurements on bacterial cell shape. Harold Pooley pointed out the necessity of geometrical corrections for reconstruction from median autoradiographs. Michael Higgins provided the initial clue that morphology and mechanisms of wall formation are tightly interlocked. He also provided experimental data to show that autographic emulsion can be thicker near the edges of a bacterium than elsewhere. Frank Trueba provided insight into the shapes of the nascent poles of organism and gave me a healthy distrust of the possible artifacts of electron microscopy. Nanne Nanninga made me clearly aware that it was not so easy to prove diffuse growth by autoradiography. I wish to thank Nanne Nanninga and Howard Rogers for the hospitality they extended to me during my extended visit to their

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departments. Finally, I should like to thank George Hegeman who administered my laboratory while I was in Europe and who helped clarify my writing.

REFERENCES

Anderson, A. J., Green R. S. and Archibald, A. R. (1978). Federation of European Microbiological Societies Microbiology Letters 14, 129. Archibald, A. R. (1976). Journal of Bacteriology 127, 956. Archibald, A. R. and Coapes, H. E. (1976). Journal of Bacteriology 125, 1195. Barnett, A. N. and Burdett, I. D. J. (1981). Journal of Theoretical Biology 92, 127. Bayer, M. E. and Remsen, C. C. (1970). Journal of Bacteriology 101, 304. Boys, C. V. (1890). “Soap Bubbles and the Forces that Mould Them”. Society for the Promotion of Christian Knowledge, London. Braun, V. (1975). Biochimica et Biophysico Acra 415, 335. Braun, V., Gnirke, H., Henning, U. and Rehn, K. (1973). Journal of Bacteriology 114, 1264. Burdett, I. D. J. (1974). Journal of Bacteriology 137, 1395. Burdett, I. D. J. and Murray, R. G. E. (1974a). Journal of Bacteriology 119, 303. Burdett, I. D. J. and Murray, R. G. E. (1974b). Journal of Bacteriology 119, 1039. Burdett, I. D. J. and Higgins, M. L. (1978). Journal of Bacteriology 133, 959. Burdett, I. D.=l. (1980). Journal of General Microbiology 120, 35. Cole, R. M. and Hahn, J. J. (1962). Science 135, 722. de Boer, W. M., Kryssen, F. S. and Wouters, J. T. M. (1981). Journal of Bacteriology 145.50.

de Chastellier, C., Hellio, R. and Ryter, A. (1975a). Journal of Bacteriology 123, 1184. de Chastellier, C., Frehel, C. and Ryter, A. (1975b). Journal of Bacteriology 123, 1 197. Doyle, R. J., Mobley, H. L. T., Jolliffe, L. K. and Streips, U. N. (1981). Current Microbiology 5, 19. DiRienzo, J. M., Nakamura, K. and Inouye, M. (1978). Annual Reviews of Biochemistry 47,48 1. Fan, D. D., Beckman, B. E. and Beckman, M. M. (1974). Journalof Bacteriology 117, 1330. Fein, J. E. and Rogers, H. J. (1976). Journal of Bacteriology 127, 1427. Formanek, H., Formanek, S. and Waura, H. (1974). European Journalof Biochemistry 46,279. Frehel, C., Beaufils, A.-M. and Ryter, A. (1971). Annales de I’lnstitut Pasteur 121, 139. Gierer, A. and Meinhardt, H. (1974). In “Lectures on Mathematics in Life Sciences” (S. A. Levin, ed.),vol. 7, pp. 163-180. American Mathematics Society. Giesbrecht, P., Wecke, I. and Reinicke, B. (1976). International Review of Cytology 44, 225. Herbert, G. (1740). Jocula Prudentwn. Henning, U. (1975). Annual Reviews of Microbiology 29,45. Higgins, M. L. (1976). Journal of Bacteriology 127, 1337. Higgins, M. L. and Shockman, G. D. (1976). Journal of Bacteriology 127, 1346. Hughes, R. C. and Stokes, E. (1971). Journal of Bacteriology 106,614. Jolliffe, L. K., Doyle, R. J. and Streips, U. N. (1981). Cell 25, 753.

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Johnson, B. F. (1965). Experimental Cell Research 39, 613. Koch, A. L. (1967). Journal of Theoretical Biology 15, 75. Koch, A. L. (1971). Advances in Microbial Physiology 6 , 147. *Koch, A. L. (1982a). Journal of General Microbiology 128, 947. *Koch, A. L. (1982b). Journal of General Microbiology 128,2527. *Koch, A. L. (1982~).Journal of General Microbiology 128, 2541. Koch, A. L. and Higgins, M. L. (1982). Journal of General Microbiology 128,2877. Koch, A. L. and Schaechter, M. (1962). Journal of General Microbiology 29, 435. *Koch, A. L., Higgins, M. L. and Doyle, R. J. (1981a). Journalof General Microbiology 123, 151. *Koch, A. L., Mobley, H. L. T., Doyle, R. J. and Streips, U. N. (1981b). Federation of European Microbiological Societies Microbiology Letters 12, 201. *KochiA. L., Higgins, M.L. and Doyle, R. J. (1982a). Journal of General Microbiology 128. 927. *Koch, A. L., Verwer, R. W. H. and Nanninga, N. (1982b). Journal of General Microbiology 128, 2893. Koppes, L. J. H., Overbeeke, H. and Nanninga, N. (1978). Journal of Bacteriology 133, 1053. Koyama, T., Yamada, M. and Matsuhashi, M. (1977). Journal of Bacteriology 129, 1518. Labischinski, H., Barnickel, G., Bradaczek, H. and Giesbrecht, P. (1979). European Journal of Biochemistry 95, 147. Laidler, K. J. and Bunting, P. S. (1973). “The Chemical Kinetics of Enzyme Action” pp. 196-232. Clarendon Press, Oxford. Mauck, J. and Glaser, L. (1972). Journal of Biological Chemistry 247, 1180. Mirelman, D. (1978). In “Bacterial Outer Membranes” (M. Inouye, ed.), pp. 115-166. John Wiley, New York. Oldmixon, E. H. (1974). Ph.D. Thesis, Temple University. Oldmixon, E. H., Glauser, S. and Higgins, M. L. (1974). Biopolymers 13, 2037. Pooley, H. M. (1976a). Journal of Bacteriology 125, 1127. Pooley, H. M. (1976b). Journal of Bacteriology 125, 1139. Pooley, H. M., Schlaeppi, J.-M. and Karamata, D. (1978). Nature, London 274,264. Previc, E, P. (1970). Journal of Theoretical Biology 27, 471. Pritchard, R. H. (1974). Philosophical Tramactions of the Royal Society of London Series B 267, 303. Rogers, H. J. (1979). Advances in Microbial Physiology 19, 1. Rogers, H. J., Perkins, J. R. and Ward, J. B. (1980). “Microbial Cell Walls and Membranes”. Chapman and Hall, London. Ryter, A., Hirota, Y. and Schwarz, U. (1973). Journal of Molecular Biology 78, 185. Sargent, M. G. (1978). Advances in Microbial Physiology 18, 105. Schlaeppi, J. M., Pooley, H. M. and Karamata, D. (1982). Journalof Bacteriology 149, 329. Schwarz, U., Ryter, A., Hellio, R. and Hirota, Y. (1975). Journal of Molecular Biology 9_8,749. Stockenius, W. (1981). Journal of Bacteriology 148, 352. Sturman, A. J. and Archibald, A. R. (1978). Federation of European Microbiological Societies Microbiology Letters 4, 255. Thompson, D’Arcy, W. (1942). “On Growth and Form”. Cambridge University Press, Cambridge.

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Trueba, F. J. (1982). Archives of Microbiology 131, 5 5 . Trueba, F. J. and Woldringh, C. L. (1980). Journal of Bacteriology 142, 869. Tzagoloff, H.and Novick, R. (1977). Journal of Bacteriology 129, 343. Verwer, R. W. H. and Nanninga, N. (1980). Journal ofBucteriology 144, 327. Woldringh, C. L., De Jong, M. A., Van den Berg, W. and Koppes, L. (1977). Journalof Bacteriology 131, 270.

Author Index Numbers in italics refer to the pages on which references are listed a f the end of each article.

A Abram, T. W., 260,280,289 Achatz, F., 226, 230,290 Ackerman, E., 9, 76 Adachi, O., 30, 32,82 Adamec, J., 87, 98, 132, 158, 167 Adams, A. P., 260,289 Adler, J., 200, 212 Adler, T., 220, 297 Afanas’eva, T. P., 89,90,92,93, 1 1 1, 121, 155,158,163,164 Afinogenova, A. V., 102, 107, 108, 110, 111, 112, 139, 159 Agre, N. S., 112, 170 Akiba, T., 190,208, 210 Albagnae, G., 260,295 Alger, J. R., 191, 193,213 Alking, H., 88, 158 Allan, R. A., 87, 96, 1 17, 158 Al-Rayess, H., 105, 114, 158 Altendorf, K., 175, 211 Alvarez, M., 225, 289 Anderson, A. J., 326,364 Anderson, W. B., 127, 167 Ando, A., 190, 192, 197,208 Andreesen, J. R., 219,259,260,264,265, 27 1,289,291,293 Andreev, L. V., 102, 107, 110, 111, 112, 140,167 Ankswanda, E., 232,298 Antfiony, C., 31, 76 Aoki, S., 129, 166 Apel, W. A., 181,208,209 Arakawa, H., 179,212 Archibald, A. R., 326, 327, 331,364,365 Argast, M., 136, 137, 158

Arion, W. J., 123, 145, 154, 156, 167 Armentrout, V. N., 4, 53,80 Artemenko, E.N., 114, 170 Asada, Y., 13, 78 Aseeva, J. V., 110, 112, 158, 170 Asher, R. A., 280,294 Ashworth, J. M., 105, 114, 158 Atkinson, A. W. Jr., 98, 129, 131, 158 Atsukawa, Y., 190,206,210 Augenstein, D. C., 287, 298 Augustin, J., 123, 125, 126, 170 Avers, C.J., 4, 18, 20, 76, 79 Avigad, G., 99, 101, 160 Avilova, T. V., 10, 78 Ayala, R. P., 86, 133, 138, 139, 169 Aui, A., 146,158

B Baba, M., 179,212 Babcock, K. L., 92, 114,168 Babel, W., 14, 15, 76 Babitskaya, V. G., 2, 78 Bache, R., 220,224,257,289 Bachgen, R., 230,289 Badziong, W., 225,289,297 Baere, L. de, 287,297 Balba, M. T., 280,289 Balch, W. E., 217,223,224,226,227,228, 229,230,231,236,237,257,289,291 Baltscheffsky, H., 88, 145, 146, 147, 149, 150, 152, 154, 156, 158,159,161 Baltscheffsky, M., 145,146,147,148,149, 156,158,159, 160 Bamforth, C. W., 226,289 Barany, M., 87, 99, 159 Baratti, R., 7, 8, 9, 77

367

368

AUTHOR INDEX

Baresi, L., 226, 233, 294 Barker, H. A., 217, 221, 235, 241, 259, 260,289,296 Barlow, D. J., 87, 101, 133, 159 Barnard, T., 87,98, I70 Barnett, A. N., 330, 364 Barnett, J. A., 36, 76 Barni, N., 87,161 Barnickel, G., 344,365 Barr, D. J., 55, 77 Barrat, N., 280,289 Barton, L. L.,224, 225,289 Baryshnikova, L. M., 143, 159 Bashirelashi, N., 92, 117, 159 Baskunov, V. P., 225,297 Bassham, J. A., 145,167 Batenburg-v.d. Vegte, 16,29, 78 Batt, P., 86, 98, 101, 160 Baudhuin, P., 3, 21, 22, 23, 55, 77 Bauer, F., 124, 164 Bauman, L., 144,168 Bauman, P., 144, 168 Baxter, M., 86, 87, 100, 101, 133, 138, 159,160 Bayer, M. E., 313,364 Beardsmore, A. J., 14, 15,82 Beaucherin-New-House, N., 86,101,104, 133, 134, 170 Beaufils, A.-M., 355, 364 Beck, A., 154,159 Beck, J. C., 187,208,209 Beck, J. V., 180,208 Becker, W. M., 4,81 Beckerich, J. M., 96, 117, 127, I59 Beckman, B. E., 354,364 Beckman, M. M., 354,364 Beekes, H. W., 89,91, 114, 121, 123, 170 Beevers, H., 4, 21, 76, 77 Behrens, N. H., 124,159 Beker, M. E., 115, 120, 122,164 Belikova, M. P., 89,90, 121, 163 Belkina, S.,204,205,212 Belliakova, T. N., 88, 149, 150, 156, 165 Belly, R. T., 176, 177, 179,208,209 Belozerskaya, T. A., 115,163 Belozersky, A. N., 84, 89, 99, 114, 122, 126, 154,159,163 Belyakova, T. N., 149, 154,165 Ben-Bassat, 230, 272,299

Benenson, R. A., 189, 190, 191, 192, 194, 195, 197, 198,206,210 Bennett, J., 114, 140, 159 Benziman, M., 143,159 Berezin, I. V., 10, 78 Berg, W. Van den, 313,365 Berner, R. A., 279,280,294 Best, A. N., 226,289 Best, D. J., 276, 292 Betz, H., 59, 76, 79 Bhat, J. V., 260,289 Bianchi, A. J. M., 230,294 Bianche, J. M., 280,298 Bicknell, B., 3, 77 Binder, A., 230,289 Biryuzova, V. I., 89,90,93,162 Black, F. T., 177,209 Blakemore, R., 227,291 Blaylock, B. A., 228,296 Blanchard, A. G., 194,211 Blanco, R., 189, 190, 191, 192, 194, 195, 196, 197, 198,206,210 Blaschko, H., 36, 77 Blaylock, B., 228,296 Blobel, G., 47, 55, 78 Blumenfeld, H., 194,210 Blytt, H. J., 142, 168 Bobyk, M. A., 102, 104, 105, 106, 107, 108, 110, 111, 112, 115, 120, 122, 132, 134, 135, 136, 139, 140, 153, 154,159, 160,161,163,164,166,167,171 Bock, A., 226,230,290 Bodmer, S., 230,289 Boer, P., 57,82 Boer, W. M. de, 326,364 Bogatyreva, T. G., 112, 170 Bohlool, B. B., 177, 179,208 Boller, Th., 89,94,95, 114, 117, 118, 122, 152,160,166,170 Bolun, S., 279,280,294 Bonen, L., 227,291 Bongaerts, G. P. A., 9, 77 Bongaerts, H., 233, 292 Bonner, S., 197, 198, 199,208 Booij, H. L., 122,160, 170 Boone, D. R., 278,289 Boonstra, J., 271,294 Boos, W., 136, 137,158 Bormann, C., 4,58,60,64,65,66,68,70, 74, 77 t

AUTHOR INDEX

369

Bornstein, R. F., 192, 193, 196, 197,198, Burdett, I. D. J., 314, 329,330,333, 354,

358,364 199, 200, 201, 202, 203,205, 206,207, Burlakova, E. B., 149,164,165 210,211 Burt, C. T.,87,99,159 Bos, P., 2,3, 30,31, 36,40,77,78,81 Butsch, B., 230,289 Boulle, A. L.,84,159 Butukhanov, V. D.,105, 110, 134, 135, Bourien, S., 144,167 153.159 Bowen, L., 229,298 Bowen, V. G.,227,299 Bowien, B., 219,273,289,291 C Bowyer, J. R.,204,208,209 Cabib, E., 124,159 Boyer, E.W.,190,209 Cagen, L.M.,144,159,I60 Boys, C. V., 308,309,364 Calvin, M.,154,160 Brachet, J., 84, 162 Campbell, L.L.,279,290 Bradaczek, H.,344,365 Cantino, E. C., 87,98,161 Brady, B. L., 30, 77 Cantino, M.E.,87,98,161 Brandis, A., 225, 230,232, 243,289 Cappenberg, T.E.,279,290 Branton, D.,17, 77 Caspari, D.,220,221,233,292,297 Braun, K.,259,260,267,289 Castenholz, R.W.,190,206,211 Braun, M.,223, 224,298 Cato, E. P.,259,290 Braun, V., 307, 359,364 Cedergren, R. J., 86, 101, 104, 133, 134, Bravati, B., 260,294 170 Breed, R. S., 259,264,289 Chaigneau, M., 221, 281,291 Breidenbach, R.W.,4, 77 Chambers, L. A., 86, 102, 103, 112, 139, Brentzel, H.J., 22,81 162 Brey, R.N.,187, 193,209 Chance, B., 10,80 Brice, J. M.,205,211 Chang, G. W.,221,293 Brierly, C. L.,177,209 Chapman, S. S.,220,290 Brierly, J. A., 177,209 Chastellier, C. de, 331, 332,364 Briggs, R. T.,20, 77 Chen, K.N.,227,291 Brill, A. S.,9, 76 Chen, M.,140,161 Bringer, S., 3, 7, 77 Chen, Y.T.,9, 79 Brock, K.M.,177,209 Brock, T.D.,176,177,178,179,208,209, Chernyadyev, I. I., 143, 145,160,168 223, 225,231, 242, 279, 280, 281, 292, Chernysheva, E. K.,108, 109, 115, 116, 299 118, 119, 120, 121, 122,163,164 Broda, E., 154, 155,159 Chibata, I., 106, 112,166 Chislett, M.E.,189,209 Brommer, B., 129, 130,164 Chkanikov, D.I., 114,170 Brown, A., 219,265,271,289 Chong, J., 55, 77 Brown, A. D.,187,209 Christiansen, C., 177,209 Bruschi, M.,225,292 Christensen, D., 280,296 Bruyn, J. C. de, 2, 78 Bryant, M. P.,224, 228, 238, 239, 254, Chua, N.H.,57, 77 256, 259, 261, 278, 279,289,290,291, Coapes, H.E.,326,327,364 Codd, C. A., 99, 100,169 294,297 Cohn, D.E.,197, 198,209,210 Buchanan, B. B., 143,159,276,291 Colas, J., 99,160 Buchanan, R. E.,226,259,290 Bullivant, S., 17, 77 Colby, J., 31, 77,276,290 Bu’Lock, J. D.,179, 185, 186, 209, Cole, J. A., 114,260 212 Cole, R.E.,179,213 Bunting, P. S.,344,365 Cole, R. M.,310,364

370

AUTHOR INDEX

Coleman, J. R., 86, 98, 101, 160 Colmer, A. R., 176, 213 Colvin, J. R., 227, 292, 296 Conti, S. F., 114, 161, 177, 209 Cook, G. A., 156,170 Cooney, C. L., 2, 79, 287,298 Corbridge, D. E. C., 102, 160 Cori, C. F., 145, 160 Cornell, N., 142, 164 Cortat, M., 57, 77 Couderc, F., 7, 8, 9, 77 Covo, G. A,, 145, I60 Cox, J. C., 178, 180, 183, 209, 211 Cox, R. B., 276,290,295 Crabill, M. R., 279, 290 Cramer, C. L., 89,97, 114, 117, 122, 152, 160 Cramer, W. A., 203,209 Crane, R. K., 197,209 Crang, R. E., 86, 87, 100, 101, 133, 138, 139, 160 Crofts, A. R., 204,208,209 Cross, R. J., r45, 160

D Dallam, D., 92, 117, 159 Dalton, H., 31, 77, 276, 290 Daniels, L., 223, 228, 230, 231, 232, 236, 238, 239,242, 243, 244, 245, 246, 250, 276,281,290,296,299 Darland, G., 176, 177, 209 Davidov, E., 88, 151, 168 Davidson, L. F., 179, 180, 182, 183, 187, 191,210,211 Davis, C. L., 224, 259, 261, 291 Davis, J. B., 225, 281, 290 Davis, J. J., 143, 171 Davis, R. H., 89, 97, 114, 117, 122, 152, 160 Dawes, E. A., 84, 114, 150,160 Dawson, A. G., 12, 77 Dawson, M., 225,299 Dawson, R. M. C., 221,294 Deal, P. H., 190,213 Decker, K., 218, 224, 226, 242, 259, 271, 292,297 Deinema, M. H., 85, 101, 140, 160 DeJong, M. A., 237,253,292, 313,365

DeLey, J., 284, 290 Dellweg, H., 13, 77 Denneny, J. M., 105, 132, 171 Derelanko, P., 7,80 DeRosa, M., 179, 185, 186,209,212 DeRosa, S., 185, 186,209 Dervartanian, D. V.,264, 299 DesMarais, D. J., 279, 294 Dice, J. F., 58, 78 Diekert, G. B., 224, 228, 246, 266, 290 Diel, F., 13, 77 Dierkauf, F. A., 123,160 Dijken, J. P. van, 2, 3,4, 5, 7,8,9, 10, 11, 14, 15, 16, 18,20,21,24,25,26,27,28, 29, 30, 31, 32, 33, 35, 36,40,41,42,43, 45,46,47, 51, 52, 56, 58,60,61,66, 78, 79, 81, 82 Dijkhuizen, L., 46, 77 Dilworth, G. L., 239,292 Dirheimer, G., 105, 132, 160 DiRienzo, J. M., 306,364 Ditter, B., 225, 289 Dixon, M., 9, 77 Dmitriev, A. D., 132, 136, 150, 166 Dobourguier, H. C., 260,295 Doddema, H. J., 228,242,243,250,290, 293 Doi, S., 176,213 Doman, N.G., 143, 145,160 Donini, P., 87, 93, 95, 119, 161 Doolittle, W. F., 226, 295 Doonan, B. B., 87,160 Douglas, C., 226, 230, 290 Douma, A., 70,82 Downs, A. J., 205,211 Doyle, E. K., 177,213 Doyle, K. E., 226, 296 Doyle, R. J., 306,307,309, 310,311, 319, 326, 327, 340, 342, 344, 346, 347, 349, 354, 359,364,365 Drachev, L. A., 146, 162 Draht, D. B., 20, 77 Drake, H. L., 265, 266, 267, 271, 290, 292 Drews, G., 99, 101, 132, 133, 160, 167 Drift, C. van der, 228,233,236,237,243, 253,290,292,297 Drozd, J. W., 205,211 Dubbelman,T. M. A. R., 89,91,114,121, 123, 170

371

AUTHOR INDEX

Dubinskaya, M. V., 102, 107, 108, 110, 11 1, 112, 139,159 Dudina, L. P., 10, 77 Dugan, P. R., 176, 180, 181, 208, 209, 212,228,294 Duine, J. A., 6, 77 Dukhovich, V. F., 149, 164,165 Duntze, W., 59, 77 Durr, M., 89, 94, 95, 114, 117, 118, 122, 152, 160, 170 Dutton, P. L., 148,160,204,205,209,212 Duve, C. de, 3,4,20,21,22,23,35,47,55, 57, 79 Dyakov, Yu.T., 110, 112, 114, 150, 169 Dyer, R. A., 227,291

E Ebel, J. P., 84, 89, 99, 105, 160 Ebner, E., 59, 79 Eden, G., 276,291 Edwards, C., 204,205,211,213 Efimova, T. P., 106, 110, 111, 112, 135, 171 Efremovich, N. V., 93, 130, 149, 154, 155, 156, 168 Eggeling, L., 8, 24, 35, 41, 44,45, 77, 78 Egli, Th., 35,41,42,44,45,47, 52, 66, 78 Egorov, A. M., 10, 78 Egorov, N. S., 10, 78 Egorov, S. N., 11 1,148,150,155,160,170 Egorova, L. A., 102, 140, 160 Egorova, 0. A., 10, 78 Egorova, S. A., 112, 170 Eichhoff, U., 87, 168 Eirich, L. D., 228, 236, 237,290, 291 Eisen, N., 143, 159 Elizarova, G. V., 153, 168 Ellefson, W. L., 228,237,238,290 Eloff, J. H., 87, 101, 133, 159 Emanuel, N. M., 87, 168 Engel, R. R., 220,290 Epstein, W., 175, 209, 211, 212 Erile, J. D., 242, 243, 295 Ermakova, S. A., 88, 93, 146, 149, 150, 151, 154, 156, 160, 165 Eroshin, V. K., 10, 77 Eroshina, N. V., 106, 110, 111, 112, 160 Esmon, B., 124, 168

Esposito, E., 185,209 Etcheverry, T., 124, 168 Evans, H. J., 143, 160 Evans, M. C., 276,291

F Fahlbusch, K., 221, 233, 297 Fais, D. A., 126, 163 Falkbusch, K., 259, 294 Fan, D. D., 354,364 FarkaS, V., 124, 160 Fein, J. E., 357, 364 Felicioli, R. A., 150, 161 Felle, H., 194, 209 Felter, S., 150, 160 Ferenci, T., 13, 78, 284, 295 Ferguson, S. J., 87, 101, 133, 134, 160 Ferro, S., 124, 168 Ferry, J. G., 223, 239,295 Fichte, B. A., 102, 107, 110, 11 1, 112,140, 167 Fiebig, K., 220, 233, 238, 292, 293 Fiechter, A., 35, 42, 45, 47, 52, 66, 78 Fillip, S. J., 179, 182, 183,211 Fillipovich, Yu.B., 87, 104, 164 Fischer, R. R., 146, 148, 149, 160, 161 Fishkes, H., 193,213 Fleishchaker, R. J., 287, 297 Fleron, P., 84, 142, 160 Flodgaard, H., 84, 142, 160 Flossdorf, J., 11, 81 Fogarty, W. M., 206, 211 Fogg, G. E., 190,209 Fontaine, F.E., 259,291 Formanek, H., 307,364 Formanek, S., 307,364 Fowler, S., 56, 79 Fox, G. E., 85, 171, 177, 214, 217, 223, 226, 227,228, 229,230,231, 261, 264, 282,289,291,296,298 Frank, J., 6, 77 Frea, J. I., 280, 298 Frehel, C., 331, 332, 355, 364 Freundt, E. A., 177,209 Friedberg, J., 99, 101, I60 Friedmann, H. C., 144, 159, 160 Friedrich, B., 273, 291 Friedrich, C. G., 273,291

AUTHOR INDEX

372

Fritz, G. J., 276, 291 Frolov, V. N., 146, 162 Fuchs, G., 218, 219, 223, 224, 226, 228, 230,232,238,239,243,244, 245,248, 250,276,290,291,294,296,297,299 Fuhs, G. W., 140,161 Fujii, T., 7, 8, 13, 78, 190, 192, 208 Fukui, S.,3,4,5, 16, 18,20,21,24,29,47, 47, 50, 51,65, 78,80,81,82, 190, 192, 208 Fukuzumi, F., 4,47,80 Fuller, R. C., 114, 161 Fulton, G., 242, 243, 290,296 Funayama, S., 59, 78 Furuya, T., 177,209 Fyodorov, V. D., 145, 146,168

G Gadian, D. G., 87, 101, 133, 134, 160 Galtseva, G. G., 114, 170 Gambacorta;A., 179, 185, 186, 209,212 Gancedo, C., 59, 78 Gancedo, J. M., 59, 78 Gandman, I. M., 110,158 Gardener, S., 280, 292 Gargus, J. J., 200,212 Garland, P. B., 174, 174, 175, 182, 209 Gaugas, J. M., 40, 78 Gee, J. M., 190,209 Genthner, B. R., 224, 259,261,291 Gerhardt, B., 55, 78 Gertrude, W., 230,296 Gest, H., 147, 162 Gezelius, K.,97, 105, 107, 110, 114, 126, 127,161 Ghambeer, R. K.,265,291,296 Gibbons, N. E., 226, 259,290 Gibson, J., 227,291 Gibson, Q. H., 9, 78 Gibson, T., 189,209 Gierer, A., 342, 364 Giesbrecht, P., 341, 344,364,365 Gilbert, W., 87,166 Gilles, H. H., 228, 246, 290,292 Gilula, N. B., 17, 77 Glaser, L., 355,365 Glass, T. L., 226,294 Glauser, S., 307, 365

Glick, B. R., 225,291 Glonek, T., 87, 99, 159, 161 Glynn, R., 87, 170 Gnirke, H., 307, 364 Gokhlerner, G. B., 154, 161 Goldberg, A. L., 58, 78 Goldman, B. M., 47, 55, 78 Goldman, D. J., 200, 212 Golobov, A. D., 88, 151, 168 Golovlev, E. L., 106, 110, 111, 112, I60 Goncharova, I. A., 2, 78 Gonina, S. A., 11 1, 136, 137, 166 Goodman, E. M., 92, 114,161 Gorham, P. R., 190,212 Gororov, N. N., 10, 78 Gorts, C. P. M., 59, 78 Gottschalk, G., 144, 220, 221, 223, 224, 233, 238, 259, 260, 267, 289, 292, 293, 294,297,298 Gottwald, M., 264, 291 Gould, J. M., 203, 209 Grant, W. D., 190, 191,210,213 Grauer, A., 150, 167 Graves, L. B. Jr., 53,80 Green, D. E., 145, 160 Green, R. S., 326, 364 Greville, G. D., 174, 201, 210 Griffin, J. B., 92, 117, 167 Grigor’eva, S. P., 110, 164 Grime, D. W., 221,294 Gruber, P. J., 4, 81 Guerrini, A. M., 87, 93, 95, 119, 161 Guffanti, A. A., 179, 180, 182, 183, 187, 189, 190, 191, 192, 193, 194, 195, 196, 197, 198, 199, 200, 201, 202, 203, 205, 206,207,208,210,211,212 Guillory, R. J., 146, 148, 149, 160, 161 Gunnarsson, L. A. H., 230,295 Gunning, B. E. S., 98, 129, 131, 158 Gunsalus, R. P., 228,236,237,238,291 Gupta, R., 227, 291 Gusev, M. V., 154, 161 Guthrie, J. D., 144,168 Guynn, R. W., 142, 156,161,164

H Habets, L. H. A., 85, 101, 140,160 HBgele, E., 59, 80

AUTHOR INDEX

Hahn, J. J., 310, 364 Hale, E. M., 190, 210 Hall, B. G., 256, 291 Hall, J. B., 153, 161 Halling, P.J., 180, 211 Halvorson, H. O., 282,291 Hamdan, I. Y., 2,80 Hamilton, W. A., 174,210 Hammer, G., 276,298 Hammes, W., 227,292 Hammond, R. C., 276,292 Han, Y. W., 261,291 Haney, M. E., 187,210 Hanselmann, K., 230,289 Hansen, B., 124, 168 Hansler, G., 4, 80 Hanson, R. S.,225,281,294 Harder, W., 2,3,4, 5,8,9, 10, 11, 14, 15, 16, 18, 20,21,24,25, 26,27,28,29,32, 33, 35, 37, 38,41,42,45,46,41, 50, 51, 52,53, 54, 56, 58, 60,61,62,64,66,68, 70, 12, 73, 77, 78, 79,81 Harold, F. M., 84, 85, 86, 99, 104, 111, 112, 114, 126, 132, 133, 134, 136, 161, 174, 187, 193, 196,210 Harold, R.L., 86, 104,161 Hart, M. H., 282,291 Hart, W., 225, 281,294 Hartlieb, R.,226,296 Hartman, H., 282,291 Hass, V., 259,289 Hatchikian, E. C., 220,221,225,281,291, 292,299 Haug, A., 178, 182, 183, 184, 186, 210, 212,213,214 Haywood, G. W., 9, 32, 79 Hayward, H. R., 221,292 Hazeu, W., 2, 16, 29, 78 Healy, J. B., 287,292 Heck, U., 89,94, 166 Heckmann, G., 264,293 Heefner, D. L., 193,210 Heer, U.,59,80 Hegeman, G. D., 219,275,293 Hegge, P.W., 223,224,230,257,261,264, 267,272,299 Held, A. A., 55, 78, 87, 97, 114, 125, 171 Held, W., 13, 77 Heldt, H. W., 145, 146, 154, 156, 159

373

Hellio, R., 331, 335, 338, 364, 365 Hemmings, B. A., 59, 78 Henning, D. H., 231,299 Henning, U., 306,307,364 Herbert, G., 355,364 Heslot, M.,96, 117, 127, 159 Hespell, R. B., 227,291 Hess, A., 205,210 Hess, W., 53, 79 Hesse, J., 175,209 Hesse, S.,22,81 Higgins, I. J., 221,226,237,242,276,292, 298 Higgins, M. L., 306, 307, 309, 310, 31 1, 313, 314, 319, 326, 329, 341, 344, 341, 349, 358, 359,364,365 Hildebrandt, A., 92, 114, 115, 116, 161 Hilpert, R., 227, 292 Hinkelman, W., 4,5,16, 18,20,24,47,80, 81 Hippe, H., 220,221,227,233,292,297 Hirota, N., 200,210 Hirota, Y., 322, 335,336,338,365 Hoare, D. S., 276,295 Hoch, H. C., 4,80 Hoddinott, M.H., 192, 194,210 Hoehn, M.M.,187,210 Hoffmann, H. P.,18, 79 Hoffman-Ostenhof, O., 84,104,114,142, 161 Hoffstein, J., 192, 193, 197, 198,200,205, 207,211 Hogg, R. W., 200,212 Holde, K. E. van, 282,291 Holdeman, L. V., 259,290 Hollander, R.,179, 205,210 Hollanes, F., 260,294 Holltta, E., 40,79 Holmes, R.,3,4, 5,47, 55, 79 Holzen, G., 226,297 Holzer, H., 58, 59,60, 66,70, 77, 79,80, 82 Hori, T., 14, 79 Horig, J. A., 264,293 Horikoshi, K., 190, 197, 206, 208, 210, 211 Horio, T., 145, 146, 147, 161, 167 Horiuti, Y., 147, 161 Hosoi, K., 147, 167 Hostalek, Z., 112, 132, 135, 161, 164

374

AUTHOR INDEX

Hou, C. T., 7,80 Hougland, A. E., 187,213 Howard, B. H., 225,292 Hsu, D. S., 143, 168 Hsung, J. C., 178, 182, 183,210 Hu, S. H., 266, 267, 271, 292 Hu, S. I., 265, 266, 290 Hubbard, A. L., 56, 79 Huber, H., 230,296 Huber-Walchli, V., 94, 161 Hughes, D. E., 99,114,132,160,161,285, 287,292 Hughes, R. C., 310,364 Hungate, R. E., 225, 260, 292, 294 Huser, B. A., 231,299 Hutchinson, T. E., 87, 98, 161 Hutten, T: J., 228,233,236,237,243,253, 290,292,297

I Ibragimov, A. R., 154, 156, 165 Igamnasarov, R. P., 110, 112,161 Imae, Y., 194, 200,210,213 Imaizumi, F., 5, 16, 29, 78 Imhoff, D., 257,259,260,292 Indge, K. J., 89,94, 117, 152, 161 Ingelman, B., 108, 161 Ingenito, E., 136, 171 Ingle, M. B., 190,209 Ingledew, W. J., 178, 180, 183, 192, 194, 209,210,211 Ingvorsen, K., 225, 279, 292, 299 Inouye, M., 306,364 Invorsen, K., 226,299 Ipata, F. L., 150, 161 Irie, M., 148,161 Irion, E., 265, 293 Isaev, P. L., 146, 148, 161, 162 Isherwood, F. A., 9, 79 Itoh, T., 177, 209 Ivanov, M. V., 225,297 Ivanovsky, R. N., 276,292 Iwamura, T., 114, 162 J

Jacobus, W. E., 153, 168 Jaenchen, R., 228,246,290,292

Jaffe, J. S., 176, 213 Jagow, G. von, 13, 79 Janecovic, D., 230, 296 Janssen, F. W., 3, 7, 79 Jarrell, K. F., 232, 242, 292, 296 Jarrell, K. G., 227, 292 Jaspers, H. T. A., 91, 123, 162 Jeener, R. R., 84, 162 Jensen, Th., 86, 100, 101, 133, 138, 159 Jensen, T. E., 86, 87, 99, 100, 101, 133, 134, 138, 139,160, 162,164 Jewell, K. J., 184, 212 Jewell, M. J., 184, 212 John, P., 226,298 John, P. C. L., 98, 129, 131, 158 Johnson, B. F., 349,364 Johnson, E., 259,291 Johnson, G. S., 127, 167 Jolliffe, L. K., 326, 346, 354, 364 Jones, C. W., 205,211 Jones, H. E., 86, 102, 103, 112, 139,162 Jones, J. B., 223, 231, 239, 292 Jones, J. G., 280, 292 Jones, 0. T. G., 145, 162 Jost, M., 99, 162 Jorgenson, B. B., 280,296 Jungermann, K.A., 218, 219, 224, 226, 242,259,271,292,297 Jungermann, K. A., 218,297

K Kaback, E., 192, 199,200,205,212 Kaback, H. R., 191, 192, 194, 196, 197, 198,209,210,211,212,213 Kaczorowski, G. J., 198,211 Kadomstseva, V. M., 138, 167 Kalebina, T. S., 88, 150, 151, 160 Kallas, T., 190, 206, 211 Kamen, M. D., 259,289 Kanai, R., 129, 162, 166 Kandler, O., 226, 227,261,292,293,298 Kaneko, H., 177,209 Kano, M., 11, 41, 79 Karamata, D., 327, 355, 356, 357, 358, 365 Karnovsky, M. J., 17,20, 77 Karnovsky, M. L., 20, 77 Karzanov, N. N., 10, 78

AUTHOR INDEX

Kashket, E. R., 194,211, 212 Kasper, Ch. B., 145, 164 Kato, J., 106, 112, 166 Kato, N., 4,5,7, 11, 12, 14, 15, 25,41, 79, 80, 81, 82 Katsumata, M., 147, 161 Kaufer, B., 224,297 Kaulen, A. D., 146, 162 Kawamoto, S., 5,16,18,20,21,24,29,47, 50, 51, 65, 78, 81, 82 Keck, K., 99, 162 Keeney, D. R., 279,294 Keister, D. L., 146, 148, 149, 162, 167 Keizer, I., 3,4,5, 16,21,24,47,50,52,58, 82 Keizer-Gunnink, I., 38, 53, 54, 72, 82 Kell, D. B., 87, 101, 133, 134, 160, 203, 211,242,243,250,293 Keller, F. A., 287, 296 Kellogg, R. M., 282, 297 Kelly, C. T., 206, 211 Kelly, D. P., 243, 297 Keltjens, J. T., 228, 236, 293, 297 Kemp, M. B., 13, 15, 79 Kenealy, W. R., 228, 232, 238, 239, 241, 243, 244, 245, 246, 247,250,251, 252, 253,255,276,293,299 Kerby, R., 220, 223, 254, 259, 267, 268, 270,271,272,275,294 Kersten, K., 284,290 Kerwin, R. M., 3, 7, 79 Kessel, M., 87, 101, 133, 162 Khan, S., 200, 211 Kheinaru, E. Kh., 10,81 Khokhlov, A. A., 149,165 Kholodenko, V. P., 138, 167 Kiebig, K., 221, 233,297 Kihara, M., 194,214 Kim, K. E., 221,293 Kim, Y. M., 219,275,293 King, G. M., 280,293 King, M. T., 156, 170 King, T. E., 145, 162 Kirchniawy, F. H., 218,292, 297 Kirk, M., 145, 167 Kitada, M., 197, 200, 206, 210, 211 Kiyomiya, A., 192, 197, 200,211, 212 Klebanova, L. M., 126,163 Klee, B., 228,290 Klein, J., 287, 295

375

Klemme, J. H., 147, 162 Kleppe, K., 9, 77 Klevicikis, S. C., 280, 298 Kline, E. S., 92, 117, 167 Klingenberg, M., 13, 79, 145, 146, 154, 156, 159 Klotz, L. C., 226,294 Klug, M. J., 279, 296 Kluge, M., 287,295 Klungsoyr, L., 145, 162 Kluyver, A. J., 217, 223, 239, 293 Knaff, D., 204,205,212 Knobloch, K., 145, 162 Knoll, A. H., 282,293 Kobayashi, H., 193,210 Kobylansky, A. G., 87, 104, 164 Koch, A. L., 307,309,310,311,312,313, 319, 323, 326, 327, 333, 335, 340, 341, 342, 344, 346, 347, 349, 352, 354, 356, 358, 359,360, 363,364,365 Koike, M., 132, 166 Koivusalo, M., 11,81 Kokurina, N. K., 88, 89, 90, 121;163 Kolot, M. N., 111, 136, 165 Komagata, K., 2,40, 79 Kondrashin, A. A., 146, 148, 162 Kondratieva, E. N., 276,293 Kondratuva, E. N., 276,292 Konheiser, U., 228, 290 Konig, H., 227, 230,292, 293, 296 Konings, W. M., 271, 294 Konoshenko, G. I., 88, 89, 90, 93, 108, 109, 110, 111, 112, 114, 115, 119, 120, 121, 132, 144, 146, 150, 151, 155, 162, 163, 164,165, 168 Konovalova, S. V., 108, 1 12, 132, 164 Kopp, F., 57, 77 Koppes, L., 313, 335, 365 Kornberg, A., 84, 104, 142, 162 Kornberg, S., 84, 104, 162 Koshiya, K., 202, 211 Kosnir, B. R., 279, 293 Kostlan, N. B., 140, 169 Kowalska, H., 106, 107, 135, 169 Koyama, N., 179,192,197,200,202,21 I, 212,214 Koyama, T.,342,365 Kozlova, T. M., 49, 64, 70, 80 Krasheninnikov, I. A., 88, 89, 90,92,93, 115, 117, 121, 132, 155, 162, 163,164

376

AUTHOR INDEX

Langeworthy, T. A., 173, 175, 176, 177, 179, 184, 185, 186, 187, 211, 212, 213, 226, 227, 297 Lanyi, J. K., 197, 212 Large, P.J., 9, 31, 32, 79 Larsen, S. H., 200,212 La Rue, T. A., 30,40, 79 Laskin, A. J., 7,80 Lawry, N. H., 86, 100, 133,164 Lawson, J. M. R., 142,164 205,206,207,208,2J0,211,212 Lawson, J. W., 156,161 Layvenieks, M. G., 115, 120, 122, 5, Kryssen, F. S., 326, 364 Kryzcki, J. A., 223, 224, 257, 261, 264, Lazarow, P. B., 21, 55, 56, 57, 79,80 267,299 Lee. J. D.. 2.40. 79 Krzycki, J. D., 228, 231, 233, 239, 240, Ledall, J.; 221,’225, 228, 238, 264, 281, 291,292,293,298 241,253,254,293 Lehle, L., 124, 164 Kuhl, A., 84, 86, 89, 99, 129, 163 Leigh, J. A., 260,293 ICuhn, W., 238,293 Lentz, K., 264,293 Kula, M. R., 11, 12, 80, 81 Levin, G. V., 85, 140, 164 Kulaev, I. S.,84,85,86,87,88,89,90,91, 92,93,94,96,97,99,102,104,105,106, Levine, D. W., 2, 79 107, 108, 109, 110, 111, 112, 114, 115, Lewis, B., 229,298 116, 117, 118, 119, 120, 121, 122, 123, Lewis, B. J., 227,291 124, 125, 126, 127, 129, 130, 131, 132, Lewis, R. J., 192, 199, 200,204, 205,212 133, 134, 135, 136, 137, 138, 139, 140, Li, L. F., 265,293 143, 144, 145, 146, 147, 148, 149, 150, Liberman, E. A., 146, 148, 161 151, 152, 153, 154, 155, 156, 157, 158, Lichko, L. P., 89, 96, 117, 138, 152, 165, 167 159, 160, 161, 162, 163, 164, 165, 166, Liebermann, L., 84, 114, 164 167,168,169, 170,171 Lin, E. C. C., 154, 171 Kumagai, H., 32, 82 Lindley, N. D., 15, 79,82 Kupriyanov, V. V., 153,168 Lindmark, D. G., 219,293 Kurahashi, K., 143, 170 Lindstrom, E. B., 280,289 Kuratomir, M., 265,295 Linnemans, W. A. M., 57,82 Kusaka, I., 190, 197,208 Lipmann, F., 154,165 Kushner, D. J., 187,189,209,211 Liss, E., 85,88,89,95, 114, 122, 164, 165 Kuwashima, S.,114, 162 Lisson, T. A., 189,211 Liu, Chi Li, 225,293 Ljungdahl, L. G., 217,219,257,259,264, L 265, 266, 267,271,289,291,293,295, 296,298,299 Labischinski, H., 344,365 Lobakova, E. S., 88, 150, 151,160 Laidler, K. J., 344, 365 Lobanok, A. G., 2, 78 Laimins, L. A., 175,211 Lochmiiller, H., 143, I71 Lam., K. S.,145, 164 Loeblich, A. R. 111, 226, 294 Lambert, M., 96, 117, 127, 159 Lambina, V. A., 102, 107, 108, 110, 111, Loeblich, L. A., 226,294 Lomagen, N., 14, 15, 76 112, 139,159 Loginova, L. G., 102, 140,160 Lamm, L., 264,293 Langen, P., 84, 85, 88, 89, 95, 114, 122, Loginova, N. B., 143,159 Lohmann, K., 84, 114, 122,164,165 164,165 Krasil’nikov, N. A., 112, 155, 163 Kreger, N. J. W. van Rij, 4, 5, 16, 81 Kritsky, M. S.,91,92, 108, 109, 114, 115, 116, 118, 119, 120, 121, 122, 126, 127, 163,164 Krochta, J. M., 261,291 Kroger, A., 224,293 Krulwich, T. A., 179, 180, 182, 183, 187, 189, 190, 191, 192, 193, 194, 195, 196, 197, 198, 199, 200, 201,202,203,204,

AUTHOR INDEX

Lopilato, J., 197,212 Lord, J. M., 55, 56, 79 Lowe, E. J., 102, 160 Ludwig, 11, J. R., 87, 114, 119, 165 Luehrsen, K. R., 227,291 Lugtenberg, B., 136, 137, 170 Lund, B. M., 190,209 Lunde, M., 87,161 Lundgren, D. G., 176,209 Lusby, E. W. Jr., 87,95, 114, 119, I65 Lusta, K. A., 110, 111, 168 Lynd, L. H., 220,223,224,254,259,261, 264, 267,268,270,271,272, 274,275, 293,294,299 Lyngdahl, H. G., 144,165 Lyulina, N. V., 153, 168

M Macario, A. J. C., 230,294 Macario, C. D. E., 230,294 McBride, B. C., 228,236,294 McCarthy, B. J., 226,294 McCoy, E., 259,291 MacDonald, R. I., 197,212 MacGregor, A. N., 279,294 McInnerey, M. J., 254, 256,294 Mack, H. M., 190,213 McKellar, R. C., 227,296 McKenna, W. R., 22,81 McLachlan, J., 190, 212 McLaughlin, C. S., 87,95, 114, 119, I65 MacLeod, R. A., 197,213 MacNab, R. M., 191, 193, 194,200,211, 213,214 Macy, J., 144, I65 Madsen, F. M., 220,290 Magrum, L. J., 177, 214, 217, 223, 226, 227, 228, 229, 230, 231, 282, 289, 291, 298 Mah, R. A,, 226,230,231,233,234,241, 255,294,296,299 Mahadevan, S., 242,243,295 Maiorova, I. P., 126, 163 Malamy, M. H., 137, 171 Malmgren, H., 108, 122,161,165 Maloney, P. C., 194,212 Mandel, IC51G., 187, 191, 192, 193, 196, 198, 199,206,211,212

377

Mandl, I., 150, 167 Mandy, W. J., 276,295 Maniloff, J., 227,291 Mann, M., 197, 198, 199,208 Mann, T. M., 180, 187, 191,210 Mannheim, W., 179,205,210 Mansurova, S.E., 88,89,90,92,93, 11 1, 117, 121, 122, 123, 134, 143, 144, 145, 146, 147, 148, 149, 150, 151, 152, 153, 154, 155, 156, 158, 160, 162, 163, 164, 165, 167,168,170 Mao, M. W. H., 180,212 Mapson, L. W., 9, 79 Maraeva,O. B., 110, 111, 136, 137, 165, 166 Margulis, L., 153, 155, 165 Marsh, C., 279, 294 Marsh, L. M., 279, 280,294 Martens, C. S., 279, 280, 294 Marti, T.,226,299 Martin, P. A. W., 180,212 Martin, S. M., 225, 291 Martin, W. G., 225, 291 Martinez, R. J., 86, 165 Martinoia, E., 89, 94, 166 Marty, D. G., 230,294 Marty, 0. G., 280, 298 Massey, V., 9, 78 Masters, C., 3, 4, 5, 47, 55, 79 Matile, Ph., 57,74, 77,80,89,94,96, 114, 117, 118, 124, 152,166 Matin, A., 180,212, 214 Matin, M., 180,212 Matrosov, A, G., 225,297 Matsuhashi, M., 342,365 Matsuura, S.,194,213 Mattenheimer, H., 108, 122, 150, 166 Matteuzzi, D., 260,294 Mauck, J., 355,365 Maxam, A. M., 87,166 Maxwell, D. P., 4, 53, 80 Maxwell, M. D., 4,80 Mayberry, W. R., 179,184,186,187,211, 212,213 Mayberry-Carson, K. J., 184,212 Mayer, F., 25, 27, 28,82,260, 289, 293 Mecke, D., 59,80 Medvedeva, G. A., 49,64,70,80 Mehta, R. J., 10,80 Meinhardt, H., 342,364

378

AUTHOR INDEX

Meisel, M. N., 49, 64, 70, 80 Melgunov, V. I., 89, 115, 121, 163, 166 Menzies, R. A., 143, 168 Metcalf, G., 190, 209 Metzger, W. C., 194, 211 Michaels,G., 143,145,150, 154, 155,166, 171 Michels, J. B., 271, 294 Michels, P. A., 271, 294, Mihara, S . , 129, 166 Mikelsaar, S. K., 10, 81 Miller, E., 259, 294 Miller, J. J., 87, 96, 117, 158 Miller, S. M., 154, 166 Miller, T. L. 219, 294 Millonig, G., 185, 212 Mills, A. A., 190, 191, 210, 213 Milner, Y., 145, 166 Minin, A. A., 144, 162 Mink, R. W., 228,294 Minnale, L., 186, 209 Minton, N. J., 146, 148, 149, 162 Mirelman, D, 306, 365 Mitchell, P., 148, 155, 166, 174, 196,212, 213,242,294 Mitchell, R., 181, 213 Mitchell, R. M., 226, 294 Miwa, N., 47,80 Miya, T., 7, 81 Miyachi, S., 129, 166 Mobley, H. L. T., 306,326, 327,340, 342, 347, 354,364,365 Moll, J., 232, 295 Monti, L., 189, 191, 194, 196, 210 Moor, H., 17, 77 Moore, A. L., 13, 80 Moore, M. R., 264,293 Moore, R., 226, 294 Moore, W. E., 259, 290 Morris, J. G., 242, 243, 250, 293 Moskovitz, B. R., 145, 166 Mountford, D. O., 242, 280, 294 Moura, I., 228, 238, 298 Moura, J. J. G., 228, 238, 298 Mudd, S . , 132, 166, 168 Mudgett, M., 87, 161 Muhammed, A., 99, 132,161 Muhlethaler, K., 17, 77 Muller, D., 59, 80 Muller, S., 99, 160

Murata, K., 106, 112, 166 Murray, E. G. P., 259, 264,289 Murray, G. M., 4,80 Murray, R. G. E., 333,364 Mushak, P. A., 140,169 Myery, T., 87, 161

N Nagumo, T., 177,209 Nagy, M., 89, 94, 95, 114, 117, 118, 122, 152, 160 Nakamura, K., 306,364 Nanninga, N., 306, 323, 324, 326, 335, 336, 337, 338, 339,357,365 Navon, G., 87,99,166 Neat, C. E., 21,80 Neben, I., 12,80 Nedwell, D. B., 280, 289 Neeff, J., 59, 80 Negi, T., 148, 161 Neill, A. R., 221, 294 Nelson, D. R., 230, 231, 232, 280, 281, 298,299 Nesmeyanova, M. A., 110,111,132,136, 137, 150, 165, 166, 168 Neuberg, C., 150, 167 Neuhaus, J., 59, 80 Neumann, D., 59, 77 Neupert, W., 57, 80 Neurauter, C., 219, 265, 271,289 Newcomb, F. H., 4,81 Ng, T. K., 278,294 Nicholls, D. G., 178, 180, 183, 209 Nicolaus, B., 185, 186, 209 Nieuwdorp, P. J., 16, 29, 78 Nikitin, D. I., 102, 107,110,111, 112, 140, 167 Niklowitz, W., 99, 167 Nikolaev, N. N., 104, 108, 112, 164 Nilshammar-Holmvall, M., 98, 131, 169 Nilsson, J. R., 86, 98, 101, 160 Nishikawa, H., 2, 80 Nishikawa, K., 147, 161 Nishikawa, N., 7, 81 Nishikawa, R., 147, 167 Nishizawa, T., 12, 15, 79 Nisman, Y., 220,298 Nivochkova, A. T., 70,80

AUTHOR INDEX

Nordlie, R. G., 123, 145, 154, 156, 167 Northcote, D. H., 17, 77 Norum, K. R., 21,80 Nosoh, Y., 179, 192, 197, 200, 202, 211, 212,214 Novichkova, A. T., 49, 64, 70, 80 Novick, R., 342, 365 Novik, P., 124, 168 Nurse, P., 151, I67

0 Oberlies, G., 245, 294 O’Brien, W. E., 143, 144, 145, 150, 154, 155, 167, 171 O’Connor, M. L., 14, 15,80 Odom, J. M., 225,294 Offenbacher, S., 92, 117, 167 Ogata, K., 2, 7, 11, 14, 25, 30, 32, 41, 79, 80,81, 82 Ogawa, S., 87, 99, 166, 168 Ohashi, H., 14, 79 Ohashi, S., 84, 167 Ohnishi, T., 13, 80, 204, 213 Ohshino, N., 10, 80 Ohshino, R., 10, 80 Ohsuji, M., 2, 80 Ohta, K., 192, 200, 212 Ohwaki, K., 260,294 Okorokov, L. A., 89, 96, 110, 112, 114, 117, 138, 150, 165, 167 Okorokov, V. A., 117, 152, 167 Oldmixon, E. H., 307,330, 342,365 Oliver, S . G., 87, 114, 119, 165 Omori, Y., 7, 25, 79 Oostra-Demkes, G. J., 11, 82 Oparin, A. I., 153, 154, 167 Ordal, G. W., 200, 212 Oremland, R. S., 279,280,294 Orgel, L. E., 154, 159 Orme-Johnson, W. H., 242,243,290,296 Oro, T., 226,297 Ortigoza, R. O., 99, 101, 170 Orton, W. L., 89, 121, 171 Oshima, T., 179,212 Osmundsen, H., 21,80 Ostrovsky, D. N., 87, 89, 98, 101, 167 Ostroumov, S. A., 146, 167 Ostrowski, W., 106, 132, 169

379

Osumi, M., 4, 5, 16, 18,20,21,24,25,28, 29, 47, 50, 51, 55, 65, 7480, 81,82 Otto, R., 8, 9, 10, 11, 81, 82 Overbeeke, H., 335,365 Owens, J. D., 3, 77 Ozick, D., 189, 191, 194, 196, 210

P Packer, L., 17, 77 Padan, E., 191, 193,200,212,213,214 Paine, R. W., 36, 76 Pal, H. S., 2, 80 Palji, A., 143, I59 Palm, P., 230, 296 Panganiban, A. T., 225,281,294 Paoleila, P., 219, 293 Papineau, D., 187, 210 Parish, R. W., 4, 80 Parker, D., 257,296 Parker, D. J., 264, 265,294 Parris, M., 154, 166 Parthasarathy, M. V., 87, 98, 132, 158, 167 Pastan, I. H., 127, 167 Pastuszak, I., 106, 107, 135, 167 Patel, R. N., 7, 80, 276, 295 Patt, T. E., 225, 281,294 Payne, W. J., 275,295 Peck, H. D., 225,294 Peck, H. P., 225, 293 Peckman, K., 229,298 Pedersen, D., 280,295 Pedersen, T. A., 145, 167 Peel, J. L., 190, 209 Peeters, B. P. H., 237, 253, 292 Penniall, R., 92, 117, 167 Perkins, J: R., 306, 365 Perski, H. J., 232,295 Peterson, W. H., 259, 291 Peverly, J. H., 87, 98, 132, 167 Pfennig, N., 220, 224, 226,289,295, 296 Pidhorskyi, V. S., 5, 81 Piechulla, K., 228, 290 Pieterse, A. J. H., 87, 101, 133, 159 Pilon, S. A. F., 16, 26, 47, 51, 52, 53, 58, 60, 61, 62, 66, 82 Pirt, S. J., 231, 232, 243, 283, 295, 297 Plack, R. H. Jr., 193,212

380

AUTHOR INDEX

Platonenkova, L. S., 10, 78 Polyakov, V. Yu., 89, 90, 93, 115, 158, 164 Pomoshchnokova, N. A., 49,64,70,80 Pooley, H. M., 326, 327, 355, 356, 357, 358,365 Porter, J. S., 194,209 Postan, I., 259, 260,296 Postgate, J. R., 221, 225, 281, 293,295 Poston, J. M., 265,295 Powell, M. J., 55,80 Previc, E. P., 361, 365 Privot, A. R., 226,229,295 Prince, R. C., 204,209 Prince, R. J., 204,205,212 Prins, R. A., 279, 290 Pritchard, R. H., 361,365 Pruitt, R. L., 286, 295 Pullaiach, T., 147, 167 Putnins, R. F., 88, 167

Richter, C., 129, 168 Rijn, H. J. H. van, 57,82 Ris, H., 99, 168 Rits, S., 116, 132, 170 Ritter, M. T., 259, 291 Robbi, R., 55,80 Roberts, L. M., 55, 56, 79 Robertson, D. E., 196, 198,211 Rogers, H. J., 306, 357, 364, 365 Roggenkamp, R., 4, 5, 8, 16, 18, 20, 24, 47,80,81 Romanenko, V. M., 5,81 Romesser, J. A., 236, 237, 238, 291, 295 Ronnow, P. H., 230,295 Rose, C. A., 231,295 Rose, I. A., 143, 170 Roseman, S., 197,213 Rosen, B. P., 187,191,193,208,209,222, 213 Rosenberg, H., 102, 168 Rosing, J., 188, 203, 212 Q Rottenberg, H., 87, 170, 191, 203, 212 Quayle, J. R:, 2, 13,14, 15, 78, 79,80,82, Rowley, J. R., 87,98,170 Rozhanets, V. V., 87, 104, 156, 163,164 226,275,276,284,289,290,295 Rozzi, A., 287,297 Rubtsov, P. M., 89,90,92, 93, 114, 115, R 129, 130, 131, 149, 154, 155, 156, 164, 168 Ramos, S., 191, 196, 211,212 Ruelins, H. W., 3, 6, 79 Rao, P. Y.,146, 167 Ratner, E. N., 102, 107, 110, 111, 112, Rupprech, E., 218,219,292,297 Rusch, H. P., 92, 114,161, 168 140,167 Ruwart, M. J., 183, 184, 212,213 Raveed, N. J., 149, 162 Ryter, A., 322, 331, 332, 335, 336, 338, Reddy, C. A., 279,290 355,364,365 Reeburg, W. S., 280,281,295 Reeves, R. E.,88,142,143,144,145,154, 167,168 5 Rehn, K., 307,364 Reid, G. A., 192, 194, 210 Sagan, L., 153, 155, 168 Reinicke, B., 341, 364 Sagdullaev, S. N., 93, 1 10, 120, 130, 170 Remennikov, V. G., 146, 148,162 Sahm, H., 3,4,5,7,8, 10, 11, 12, 13, 16, Remsen, C. C., 3 13,364 18,20,24,35,41,42,43,44,45,47,58, Rensburg, W. L. J. van, 87, 101, 133,159 60,64,65,66,68,70,74, 77, 78, 79,80, Renthal, R., 197,212 81,228,233,234,235,253,276,295 Renz, P., 264,293 Sakai, T., 24, 25, 28, 80 Reshetnyak, V. I., 87, 89, 98, 101, 167 Sakazawa, C., 12, 15, 79 Reverly, J. H., 87, 98, 132, 158 Saks, V. A., 153,168 Rhoads, D. B., 175,211,212 Salhany, J. M., 87, 99, 168 Rhodin, H., 3,80 Sall, T., 132, 168 Rich, P. R., 13, 80

AUTHOR INDEX

381

Samin, E., 260,295 Schwarz, U., 322,335,336, 338,365 Samsonoff, W., 177,209 Schwencke, J., 89, 94, 95, 104, 109, 114, Samuilov, V. D., 146, 148, 161, 162, 167 117, 118, 120, 122, 150, 152, 160, 168, Santos, M. H. I., 228,238,298 I70 Sapienza, C., 226, 295 Schwermann, K., 59, 78 Sargent, M. G., 306,365 Scott, D., 276, 292 Satir, B., 17, 77 Scott, K. J., 114, 140, 159 Satir, P., 17, 77 Scrutton, M. C., 143, 168 Sato, K., 276,295 Searcy, D. G., 177,178,182,213,226,296 Sato, M., 24, 25, 28, 55,80 Segal, H. L., 58, 81 Sauer, F. D., 242,243,295 Segal, W., 220,296 Sauer, H. W., 92,114, 115, 116,161,168 Seliverstova, L. A., 70,80 Sauer, L., 92, 114,161 Selyach, T. O., 88, 150, 151,160 Saunders, V. A., 145,162 Semenov, A. Yu, 146,162 Sawyer, M. H., 144, 168 Senior, P. Y., 84, 114, 150,160 Sayler, G. S., 280,295 Sepetov, N. F., 87,89,98, 101,167,168 Schaechter, M., 356, 358,365 Seppet, E. K., 153,168 Schatz, G., 57, 80,81 Sergeev, N. S., 104, 108, 112, 164 Schauer, N. L., 223,239,295 Serrano, R., 144, 168 Schaupp, A., 219,265,271,289,295 Severin, A. I., 110, 111, 136, 166,168 Schekman, R., 124, 168 Shabalin, Yu, A., 89, 91, 97, 104, 118, Schellenberg, M., 59,70,82, 89,95, 171 121, 123, 124, 125, 127, 131, 152, 164, Scherer, P., 218, 219, 228, 233, 234, 235, 168, 170 253,287,295,297 Shady, A., 134, 143, 145, 152, 153, 164, Schimke, R. T., 58,81 168 Schink, B., 220, 254, 278,295 Shakhov, Yu,A., 88, 147, 149, 150, 151, Schlaeppi, J. M., 327, 355, 356, 357, 358, 154, 156,165,167,168 365 Shama,A. M.,92,117,146,149,156,165 Schlanderer, G., 13, 77 Shapiro, J., 85, 140, 164 Schlegel, H. G., 154,168,219,260,289 Shapiro, S., 228, 236, 237, 238, 276,291, Schleifer, K. H., 261,292,298 296 Schmidt, G. W., 57, 77 Shaposhnikov, V. N., 145, 146,168 Schmidt, W., 218,292 Sheldelin, V. H., 145, 162 Schnellen, Ch. G. T. P., 217, 223, 239, Sherod, D. W., 264,293 293,295 Shimzu, S., 7,82 Schoberth, S., 223,224,257,267,289,296 Shin, K. C., 7,82 Schnitker, U., 224,291, 297 Shioi, J. I., 194, 213 Schofield, A. K., 191,210 Shively, J. M., 99, 168 Scholten, J., 85, 101, 140, 160 Shnyukova, E. U., 140,169 Schon, G., 219,292 Shockman, G. D., 341,364 Shulman, R. G., 87, 99, 166, 168,170 Schonheit, P., 232, 246,295 Schubert, K. R., 228,232,244,247,250, Sibel’dina, L. A., 87,89,98,101,167,168 255,293 Sicko, L. M., 86, 100, 133, 162 Schuldiner, S., 191, 193, 194, 200, 212, Sicko-Goad, L. M., 86, 100, 101, 133, 213,214 138, 139, 169 Schulman, M., 257,265,291,296 Siefert, E., 226, 296 Schiitte, H., 11,81 Sievert, R., 221, 233, 297 Simisker, Y.A., 10, 81 Schwartz, J. S., 62,81 Schwartz, R. D., 287,296 Simkiss, K., 102, 169 Schwartz, S., 220, 290 Simms, E., 84, 104, 162

382

AUTHOR INDEX

Simon, B. M., 280,292 Simon, R.D., 119,169 Simonis, W., 129,162,170 Singh, R.N.,99,168 Sintsov, N.V., 276,292 Siu, P.M.L., 142,169 Skryabin, K.G., 89,90,92, 115,148,150,

Stahl, D. A., 227,291 Stanley, P.E.,202,213 Starkey, R.,220,296 Steblyak, A. G., 1 1 1, 123,170 Steckenbrandt, E., 85,169 Stedingk, L. V. von, 88, 145,146,147,

152, 154,156,158, 159,161 154,163,164,165,169 Steere, R. L., 17,77 Skulachev, V. P., 146,148,154,156,161, Sterkenburg, A., 88, 158 Stetten, M. R., 123,145, 156,169 162,167,169, 194,213 226,230,232,243,289,296 Slabova, 0.I., 102,107,110, 111, 112, Stetter, K.O., Steveninck, J. van, 89,91,114,121,122, 140,167 123,162,170 Slater, E. C., 188, 203,212 Stewart, W.P., 99,100, 169 Slayman, C. L., 194,209 Stich, H., 99,162 Slechta, L., 142,161 Stichler, W., 248,250,291 Slonczewski, J. L.,191,193,213 Stock, J., 197,213 Smith, G. G., 184,213 Stoecknius, W., 303,365 Smith, L. D., 259,260,296 Stokes, E., 310,364 Smith, M.R.,177,213,226,233,234,241, Stokes, J. L.,189,213 255,294,296 Stouthamer, A. H.,203,213,271,272, Smith, N. R., 259,264,289 Smith, P.F., 177,179,184,186,187,211, 296 212,213 Strayer, R. F., 280,296 Streips, U. N.,306,326,327,340,342, Smith, R. L., 279,296 Snozzi, M., 230,289 346,347,354,364,365 Sodano, S., 186,209 Strom, T., 13,78 Sokolovsky,V.Yu,91,92, 114,115,117, Strong, K., 185,213 Stroobant, P.,196,211 127,146,149,156,165,169 Solimene, R.,87,93,95,119,169 Stupperich, E.,230,243,245,250,276, 291,296 Sorensen, E. N.,193,209 Sorenson, J., 280,296 Sturman, A. J., 326,331,365 Sorokin, Y. I., 225,296 Sudyina, E. G., 140,169 Sugiyama, T., 184,213 South, D. J., 142,144, 168 Souza, K.A., 190,213 Sukovatova, L.V., 10,77 Sundberg, I., 98,131,169 Souzu, H., 89,121,169 Susman, P., 189,191,195,197,210 Spencer, J. F. T., 30,40,79 Suzuki, J., 147,167 Spencer, R.W., 242,243,290,296 Suzuki, M., 24,25,28,80 Speth, V., 17,77 Switzer, R.L.,58,81 Sprey, B.,3,7, 77 Swoboda, B. E. P., 9,78 Sprott, G. D., 227,232,242,292,296 Sysuev, V. A., 110,112,114,150,169 St. John, A. G., 58, 78 Szabo, A., 18, 79 Stachelin, L.A., 17,77 Stackebrandt, E.,227,261,264,291,296Szumilo, T., 107,169 Stadtman, E. R.,259,260,295,296 Szymona, M., 106, 107, 132,135, 150, Stadtman, T.C., 221,223,228,231,236, 167,169 Szymona, O., 106,107,108,112,115,135, 239,241,260,292,296 154,163,169 Stadtman, T. K., 259,260,296 Stafford, D., 285,287,292 T Stahl, A. J. C., 150,160 Stahl, D., 229,298 Tafft, H. L., 145,156,169

AUTHOR INDEX

383

Tokuda, H., 196, 197,211,213 Taggart, J. V., 145, 160 Tolbert, N. E., 21,81 Takagi, A., 132,168 Tomita, K., 148, 161 Talpasayi, E. R. S., 99, 169 Tommassen, J., 136, 137, 170 Tami, Y.,286,296 Tanaka, A., 3,4, 5, 16, 18,20,21,24, 29, Tonomura, K., 7, 8, 13, 49, 78, 81 Tornabene, T. G., 226,227,297 47, 50, 51, 65, 78,80, 81,82 Torriani, A., 136, 171 Tanaka, K., 49,81 Towel, J. P., 260, 295 Tani, K., 106, 112, 166 Tani, N., 7,80 Trelease, R. N., 4, 81 Trilisenko, L. V., 89,91,92, 11 1,114,121, Tani, T., 4, 5, 81 123, 127, 170 Tani, Y., 7, 11, 12, 14, 15, 25,41, 79,81, 82 Trotsenko, Y. A., 13,81 Tanner, R. S., 223, 224, 227, 257, 261, Trueba, F. J., 313, 339,365 Trumpower, B., 204,213 264,267,289,291,296 Tsai, L., 239, 298 Tanner, W., 124,164 Tsiomenko, A. B., 89, 91, 104, 118, 121, Taylor, C. D., 228, 236, 238,297 123, 124, 125, 126, 131, 152, 164, 168, Taylor, G. T., 232,243, 283,297 170 Tempest, D. W., 88, 158 Tempole, K. L., 176, 213 Tsofina, L. M., 146, 148, 161 Teranishi, Y., 4, 18,20,21,24,29,47,78, Tsubouchi, J., 49, 81 80 Tsuchiya, T., 197,212 Terekhova, V. A., 110,112,114,150,169 Tsuji, K., 225, 297 Tereshin, I. M., 106, 110, 111, 112, 135, Tsydendambaev, V. D., 134, 152, 153, 171 168 Termkhitarova, N. G., 106, 110, 111, 112, Tsyrenov, V. Zh., 105, 110, 134, 153, 159 160 Thauer, R. K., 218, 219, 223, 224, 225, Tumerman, L., 116, 132, 170 226, 228, 230, 231, 232, 236, 239, 242, Tuovinen, 0. H., 180,212 Tupik, N. D., 140, 169 243,244,245,246,248,250,259,265, 266, 271,289,290,291,292,294,295, Turkstra, E., 85, 101, 140, 160 297,299 Turnbill, C. E., 190,213 Thilo, E., 85, 87, I69 Turtle, J. H., 181, 208 Thomas, J. A., 179,213 Tyrsin,Yu. A., 115, 117, 132, 164 Thomm, M., 230,296 Tzagolaff, H., 342, 365 Thompson, D’Arcy W., 307, 308,365 Tzeng, S. F., 228, 238, 239,297 Thompson, J., 197,213 Thompson, T. E., 223,224,225,228,232, U 244,247,250,255,257,261,264,267, 293,299 Uchida, T., 106, 112, 166, 229,298 Thomson, R. H., 186,209 Thore, A., 146, 158 Uchino, F., 176,213 Thurman, R.G., 22,81 Uffen, R. L., 219,226,239,297 Tiedje, J. M., 280, 296 Ugurbil, K., 87, 170 Ullrich, W., 129, 170 Tierney, G. V., 204,208 Tijssen, J. P. F., 89,91, 114, 121, 123,170 Umnov, A. M., 111, 112, 114, 115, 121, 123, 148, 150, 155, 162, 164, 170 Tillberg, J. E., 87, 98, 170 Tindall, B. J., 190, 213 Umnova, N. S., 111, 123, 170 Uotila, L., 11, 81 Tobek, I., 112, 132, 135, 161, 164 Urech, K., 59,70,82,89,94,95,114, 117, Todd, M. M., 18,81 118, 122, 152,160, 170,171 Toews, M. L., 220,297

AUTHOR INDEX

384

Uryson, S. O., 89,93, 104, 106, 108, 11 1, 112, 132, 155, 158, 163, 164, 169, 170 Uspenskaya, V. E., 143, 145, 160 Utter, M. F., 143, 170 Uwajima, T., 32, 82

v Vagabov, V. M., 89,91,92,97, 104, 11 1, 114, 118, 121, 123, 124, 125, 126, 127, 129, 131, 152, 163, 164, 168, 170 Vaillancourt, S., 86, 101, 104, 133, 134, 170 Vainio, H., 146, 158 Vainshtein, M. G., 225,297 Valikhanov, M. N., 93, 110, 112, 120, 130, 161, 170 Vaughn,L.E.,89,97, 114, 117, 122, 152, 160 Vedder, A., 189, 190,213 Veech, R. L. 142, 156, 161, 164, 170 Veenhuis, M., 3,4,5,16,18,20,21,24,25, 26,27,28,29, 32,33,35,37,38,41,42, 45,46,47,50,51,52, 53, 54, 56, 58,60, 61,62,64,66,68,70,72,73, 78,81,82, 243,290 Veloso, D., 156, 161 Velzeboer, C. T. M., 260,289 Vermeulen, C. A., 18,82 Verstraete, W., 287, 297 Verteletskaya, N. L., 89,90,92, 115, 148, 164 Verwer, R. W. H., 306,323,324,326,335, 336, 337,338, 339, 357,365 Vierstra, R., 184, 213 Vigil, E. L., 18,81 Visser, C. M., 282, 297 Voelz, H., 99, 101, 170 Voelz, V., 99, 101, 170 Vogels, G. D., 228, 233, 236, 237, 242, 243,250,253,277,290,292,293,297 Volfova, O., 11,82 Volkova, M. V., 88, 151, 168 Volloch, V. Z., 116, 132, 170 Vorob’eva, E. I., 108, 110, 112, 132, 164

W Wagner, F., 4, 5, 7, 8, 10, 11, 13, 16, 18, 20, 24, 47, 78, 79,80,81

Waites, M. J., 15, 79, 82 Waksman, S. A., 176,213 Walker, J. C. G., 282,297 Walsh, F., 181, 213 Walt, J. P. van der, 30, 81 Walter, R., 238, 293 Walther, R., 221, 233, 259, 294,297 Wang, D. I. C., 287,297 Wang, G. Y., 287,297 Ward, D. M., 280,298 Ward, J. B., 306,365 Ward, P. M., 226,294 Ward, T. E., 280,298 Warford, A. L., 279,293 Warner, R. R., 86,98, 101,160 Warren, L. G., 142, 168 Watt, G. D., 264,299 Waura, H., 307,364 Wazer, J. R. van, 84, 170 Weber, D. J., 53, 79 Weber, L. J., 257,298 Webers, H. A. A. M., 85, 101, 140, 160 Wecke, J., 341,364 Weigert, W., 84, 161 Weimberg, R., 89, 114, 121,171 Weimer, P. J., 221, 223, 224, 228, 230, 231, 232, 234, 238, 241, 246, 247, 253, 255, 257, 261, 264, 267, 274, 276, 281, 298,299 Weinstein, R. S., 17, 77 Weiser, U., 59, 76 Weiss, N., 261, 298 Weiss, R. L., 177, 186, 209, 213 Weller, H., 184, 213 Welty, F. K., 265,298 Weringa, K. T., 217,260,298 West, I. C., 196, 213 White, F. H., 260,296 Whitelaw, V., 175, 209 Whitman, W. B., 228,232,298 Whittenbury, R., 31, 77, 276, 290, 298 Wiame, J. M., 84, 89, 171 Wiebe, W. J., 280, 293 Widdel, F., 224,295 Wiegel, J., 223, 224, 260, 298 Wieker, W., 87, 169 Wiemken, A., 59, 70,82, 89, 94,95, 114, 117, 118, 122, 151, 152, 160, 161, 166, 167, 170, 171 Wiley, W. R., 189, 213

AUTHOR INDEX

Wilkinson, T. G., 276,298 Williams, M. C., 275, 295 Williams, R. J. P., 203, 213 Williams, S. G., 202, 213 Willison, J. C., 226, 298 Willsky, G. R., 137, 171 Wilson, B., 180,212,214 Wilson, T. H., 154, 171, 194, 197,212 Winder, F. G., 105, 132, 171 Winfrey, M. R., 279, 280, 281,298 Winter, J., 221, 226, 227, 230, 233, 254, 256,292,296,298 Wise, D. L., 287,298 Wisendanger, S., 220,298 Woese, C. R., 85,171, 177,214,217,223, 226,227,228,229,230,231,261,264, 282,289,291,296,298 Woldringh, C. L., 313,365 Wolf, D. H., 58, 59, 82 Wolf, G., 179, 210 Wolfe, R. S., 217,221,223,224,226,227, 228, 229, 230, 231, 232, 233, 236, 237, 238, 239, 241, 242,243, 254, 256, 257, 260, 264, 267, 276,289, 290,291, 293, 294,295,296,297,298,299 Wolin, M. J., 230, 294 Wolkin, R., 231, 233,239, 240,241, 253, 254,255,293 Wood, H. G., 88,142,143,144,145,150, 154, 155, 160, 166, 167, 171, 217, 257, 259, 264, 265, 266, 267, 271, 290, 291, 292,293,294,296,298 Wood, J. M., 228,238, 241,298 Wood, N. P., 219,293 Wool, S. H., 87, 97, 114, 125, 171 Worthington, R. D., 186, 209 Wouters, J. T. M., 326, 364 Wu, T. F., 264, 265,294 Wuhrmann, K., 231,299 Wunderl, S., 230, 296

X Xavier, A. V., 228, 238, 298

Y Yabuki, M., 190, 192, 197,208 Yabute, A., 148, 161

385

Yagi, T., 225, 297, 298 Yagil, E., 136, 171 Yakovleva, L. A., 89, 121, 163 Yamada, E. W., 88, 167 Yamada, G., 30,82 Yamada, H., 4,5,7, 12, 15,32, 79,81,82, 286,296 Yamada, M., 342,365 Yamane, T., 87,99,166, 168 Yamashita, J., 147, 161 Yamazaki, R. R., 21,81 Yamazaki, S., 239,298 Yamazaki, Y., 179,214 Yang, L. L., 184, 185,214 Yang, S. S., 264,298,299 Yarbrough, H. F., 225, 281,290 Yarrow, D., 36, 76 Yasuhara, S., 5, 16, 18,20,21,24,29,47, 50, 51, 65, 78,81,82 Yazykov, A. I., 129, 130, 164 Yike, N. J., 146, 162 Yoshida, A., 84, 104, 114, 171 Yoshida, H., 145, 171 Yoshida, S.A., 132, 166 Yoshimura, S., 147, 167 Young, L. Y., 287,292 Young, M. R., 143,168 Younis, M. S., 105, 114, 158

z Zaichkin, E. I., 110, 11 1, 166 Zablen, L., 229,298 Zablen, L. B., 227, 291 Zaiss, U., 280, 299 Zajac, J., 150, 169 Zaritsky, A,, 194,214 Zavarzin, G. A., 223, 230, 299 Zbarsky, I. B., 92, 171 Zehnder, A. J. B.,223,225,226,231,242, 277.281.299 Zeikus, J. G., 220,221,223,224,225,226, 227, 228, 229, 230, 231, 232, 233, 234, 236, 238,239,240,241,243,244,245, 246, 247, 249, 250, 251, 252, 253, 254, 255, 257,260,261,262,264,267,268, 270, 271, 272, 274, 275, 276, 277, 278, 279, 280,281, 286, 287, 289,290, 292, 293,294,295,298,299

386

AUTHOR INDEX

Zeller, E. A., 36,82 Zemlyanukhina, 0. A., 104, 118, 131, 152,168 Zhang, H., 261,291 Zhilina, T. H., 223, 230, 233,299 Ziegler, H., 248, 250, 291 Zilberstein, D., 191, 193, 200, 212, 214 Zillig, W., 230, 296 Zinder, S. H., 226, 230, 280,296,299 Zuckerman, R. S., 179, 182, 183,211 Zuckier, G., 136, 171

Zurrer, H., 230, 289 Zuzel, T., 256, 291 Zuzina, M. L., 106, 110, 111, 112, 135, 171 Zwart, K. B., 3, 4, 24, 29, 32, 33, 35, 37, 38,41,46,47, 52, 53,58,60,64,66,68, 70, 72, 73, 82 Zvyagilskaya,R. A., 88,93,146,149,154, 156,165 Zyakun, A. M., 225,297 Zychlinsky, E., 180,212,214

Subject Index A

pyrophosphate synthesis in, 145 pyrophosphate utilization in, 143 Acetabularia sp., polyphosphate metabpyruvate phosphate dikinase in, 144 olism in, 129 Acetobacterium woodii Acetabularia crenuiata carbon monoxide consumption, 224 polyphosphates in, development and, formate consumption, 224 130 hydrogen/carbon dioxide consumppolyphosphate metabolism in, 129 tion, 223 Acetabularia mediterranea metabolism, 267-270 adenosine triphosphate :polyphosmethanol consumption, 224 phate phosphotransferase in, 130 methanol production by, 220 chloroplasts, polyphosphates in, 93 one-carbon transformations in, 257 polyphosphate metabolism in, 129 Acetogenium kiuui, taxonomy, 260 enzymes, 114 Acetyl-coenzymeA Acetate kinase synthesis, in homo-acetogens, 257,269 in Clostridium thermoaceticum, 265 in methanogens, 248 regulation, in Methanosarcina barkeri, Achrobacter sp., polyphosphate (meta225 phosphate)-dependent NAD+ Acetates kinase in, 106 fermentation by Methanosarcina sp., Acidophilic bacteria, 175-187 233 physiology, 173-214 metabolism, by Methanosarcina barsurface properties, 184-186 keri, 246 Acinetobacter sp. in Methanobacterium thermoautohigh molecular-weight polyphosphates trophicum, 243 in, 140 regulation, in Methanosarcina sp., inorganic polyphosphates in, 85 255 Actinomycetes, shape, 340 methanogenesis from, 239 Active transport, cell expansion and, synthesis in Clostridium thermoaceti303 cum, 265 Adenosine triphosphatase synthesis in in methanogens, 248 homo-acetogens, 27 1-272 Acetic acid, production by homo- Adenosine triphosphate acetogens, 288 synthesis, in Saccharomyces carlsberAcetobacter sp., polyphosphate (metagensis, 1 18 phosphate)-dependent NAD+ methanogenesis and, 242-243 kinase in, 106 oxidative phosphorylation and, Acetobacter suboxydans 201-203 387

388

SUBJECT INDEX

Adenosine triphosphate:polyphosphate phosphotransferase in Acetabularia mediterranea, 130 Aerobacter aerogenes polyphosphate kinase in, 104 polyphosphate metabolism in, 132 enzymes, 1 12 polyphosphates in, 86, 134 tripolyphosphatase in, 150 Aerobes, relation to chemoautotrophic anaerobes, 275-276 Agaricus bisporus, polyphosphate hydrolysis in, 126 Alanine metabolism by Methanosarcina barkeri, 247 synthesis, in Methanobacterium thermoautotrophicum, 244, 245, 246 Alanine dehydrogenase, regulation, in Methanobacterium thermoautotrophicum, 255 Alcaligenes sp., pyrophosphate-dependent phpsphofructokinase in, 144 Alcaligines eutrophus, growth efficiency, 273 Alcohol oxidase degradative inactivation, 66 in Candida boidinii, 64, 65, 66 in Hansenula polymorpha, 64,65,66, 67 inactivation after exposure to excess methanol, 62-64 at low growth rates, 60-62 during bud formation, 60 in crystalline peroxisomes from Kloeckera sp., 25 in methanol metabolism, 6 in microbodies, 16 in peroxisomes, molecular substructure and, 24 kinetics, oxygen concentration and, 8 K, values for oxygen of, 9 localization, cerous ions in, 20 peroxisomal, regulation of synthesis, 41 productivity, in Hansenula polymorpha, 45 in Kloeckera sp., 45 properties, 7 recrystallization, 29

Alcohol peroxidase, in crystalline peroxisomes, 28 Algae, polyphosphate metabolism, 129- 132 Alkalophilic bacteria, 187-207 extracellular enzymes, 206 physiology, 173-214 Allomyces sp., polyphosphates in, localization, 125 Amine oxidase degradative inactivation, 66 in microbodies, 35 K, values for oxygen of, 9 methylamine metabolism and, 32 peroxisomal matrix, 29 synthesis, regulation, 46 Amino acids, polyphosphates and, in yeast, 119 D-Amino acid oxidase degradative inactivation, 65 in peroxisomal matrix, 29 in peroxisomes, 21 K,,, values for oxygen of, 9 synthesis, regulation, 46 Aminopeptidases I, in Saccharomyces cerevisiae, location, 59 Aminopeptidases 11, in Saccharomyces cerevisiae, location, 59 Ammonia, assimilation by Methanobacterium thermoautotrophicum, 244 Anabolism in Methanobacterium thermoautotrophicum, 243-246 in Methanosarcina barkeri, 246248 Anacystis nidulans polyphosphate granules, 100 polyphosphate kinase in, 104 polyphosphates in, 86, 134 localization, 134 Anaerobes consumption of one-carbon compounds by, 221-226 one-carbon metabolites, transformation by, 2 18-226 Ankistrodesmus braunii, polyphosphate metabolism in, 129 Arginine binding to polyphosphates, in yeasts, 117 polyphosphate complex, in yeast, 119

SUBJECT INDEX

Arthrobacter sp., polyphosphate phosphohydrolases, 110 Aspartate metabolism by Methanosarcina barkeri, 247 synthesis, in Methanobacterium thermoautotrophicum, 244, 245 Aspersillus sp.. polyphosphate phospho-

hydrolases, 110 Aspergillus niger amine oxidase from, 32 polyphosphate metabolism in, 114 Aspergillus oryzae, tripolyphosphatase in, 150 Aspergillus wentii, polyphosphate phosphohydrolases, 110 Autolysin-deficient mutants, wall morphogenesis and, 355 Autoradiograms, analysis, 322-325 Autoradiography, conserved and nonconserved regions and, 356 Autotrophic pathways in Methanosarcina barkeri, 248-251 in Methanobacterium thermoautotrophicum, 248-25 1 Autotrophy in evolution, 284

B Bacillus subtilis, median section, 3 15 Bacillus sp. polyphosphate phosphohydrolases, I10 transmembrane electrical potential, 194 tripolyphosphatase in, 150 ubiquinone in, 205 Bacillus A-007 alkaline tolerant, 189 sodium ion transport, 197 Bacillus No. 8-1, sodium ion transport, 197 Bacillus acidocaldarius, 176 cell wall, characterization, 186 external pH, 183 internal pH, 179, 180 neutral pH, 187 transmembrane electrical charge gradient, 182

389

Bacillus alcalophilus alkaline tolerant, 189 ATP synthesis, 203 oxidative phosphorylation and, 201 cytochrome levels, 205 cytoplasmic pH regulation, 191, 192 membrane colour, 204 membrane lipids, 206

membrane vesicles, cytochromes, 204 respiratory chain, 204 Rieske protein, 205 sodium ion transport, 193, 197, 198, 199 solute transport in, 196, 198 transmembrane electrical potential, 195 Bacillus anthracis, pole formation, 352 Bacillus cereus, alkaline tolerant, 189 Bacillus circulans alkaline tolerant, 189 cytoplasmic pH regulation, 191 solute transport in, 196 transmembrane electrical potential, 194 Bacillus coagulans, 176 Bacillus firmus alkaline tolerant, 189 ATP synthesis, 202 oxidative phosphorylation and, 201 cytoplasmic pH regulation, 191, 192 membrane colour, 204 membrane lipids, 206 pH and, 207 respiratory chain, 204 sodium ion transport, 193, 197, 199 solute transport, 197, 198 transmembrane electrical potential, 194, 195 Bacillus megaterium autoradiograms, analysis, 323 median section, 3 15 chloramphenicol and, 355 peptidoglycan synthesis in, 331 pulse-labelled boiled, high resolution autographic studies, 332 Bacillus pasteurii alkaline tolerant, 189 cytoplasmic pH regulation, 192 transmembrane electrical potential, 194, 195

390

SUBJECT INDEX

Bacillus subtilis binding of fluorescein-labelled concanavalin A to, 328 cell curvature, electron microscopy and, 320 median sections, chloramphenicol and, 355 dimensions, 330 poles, conservation, 327, 328 zonal dome model, 33 1 shape, changes, 352 pole formation and, 329 solute motility, 200 Bacteria phosphorylation in, pyrophosphates and, 142-145 polyphosphate utilization in, mechanism, 134 preservation of shape, 3 14 shape, 302 Bacteriaceae, taxonomy, 229 Bacteroides fragilis, pyrophosphatedependent phosphofructokinase in, 144 Bacteroides symbiosus pyrophosphate utilization in, 143 pyruvate phosphate dikinase in, 143 Basidiomycetous moulds, methanol metabolism by, 3 Bdellovibrio bacteriovorus 1,3-diphosphoglycerate:polyphosphate phosphotransferase in, 108 polyphosphate glucose kinase in, 107 polyphosphate hexokinase in, 112 polyphosphates, localization, 102 metabolism, 139 polyphosphate phosphohydrolases, 110 tripolyphosphate hydrolase in, 11 1 Benzidine, diamino-, catalase in Saccharomyces cerevisiae and, 18 Biocatalysis, anaerobic one-carbon transformations and, 285 Biomethanation, 287 Biosynthesis cell expansion and, 303 polyphosphates, enzymes in, 103-1 14 Biotechnology, one-carbon transformations and, 285-288

Blastocladiella emersonii, vacuoles, polyphosphate granules in, 98 Brevibacterium sp. polyphosphate(metaph0sphate)dependent NAD+ kinase in, 106 polyphosphate phosphohydrolases, 110 Brevibacterium ammoniagenes, phosphorylation in, pyrophosphates and, 143 Bud formation, alcohol oxidase in, inactivation, 60 Butyribacterium methylotrophicum adenosine triphosphatase synthesis, 27 1 carbon monoxide consumption, 223 carbon monoxide formation by, 220 carbon monoxide metabolism, thermodynamic efficiency, 275 catabolism and anabolism, unification of carbon flow pathways, 270 electron photomicrographs, 263 formate consumption, 224 growth efficiency, 273 substrate-product thermodynamics, 273 unicarbonotrophy and, 274 growth yields, 268 values, 274 hydrogen/carbon dioxide consumption, 223 metabolism, 267-270 methanol consumption, 224 methylotrophy in, 259 morphology, 262 one-carbon metabolism, 269 spore morphology, 260 Butyribacterium rettgeri carbon monoxide consumption, 224 methanol consumption, 224 Butyric acid, production by homoacetogens, 288

C Calcium, in polyphosphate granules, 139 Caldariella sp. lipid structure, 185 membrane lipids, 186

SUBJECT INDEX

Caldariella acidophila caldariellaquinone, 186 membrane lipids, 185 Caldariellaquinone, 186 Candida sp. alcohol oxidase from, properties, 7 methanol metabolism by, 2 Candida boidinii (see also Kloeckera sp.) alcohol oxidase from, properties, 7 alcohol oxidase inactivation in, 64,65, 66 amine oxidase from, 32 catalase inactivation in, 64, 65, 66 crystalline peroxisomes, molecular substructure, 28 formate dehydrogenase from, 12 fructose bisphosphatase in, methanol metabolism and, 15 methanol metabolism in peroxisomal enzymes, regulation of synthesis, 41 peroxisomal matrix, 29 peroxisomes, 17 development in, 49 protease inactivation in, 68 thiokinase in, methanol metabolism and, 15 Candida guilliermondii polyphosphate phosphohydrolases, 110 polyphosphate synthesis in, 151 Candida utilis methylamine metabolism, 32, 33 peroxisomes in, 33 ultrastructure, 34 Carbon dioxide consumption by anaerobes, 225 formation from formaldehyde, 11-12 /hydrogen, consumption by anaerobes, 224-225 methanogenesis from, 236238 methanogenesis from, 239 metabolism, by Methanosarcina barkeri, 246 in Methanobacterium thermoautotrophicum, 243 Carbon monoxide Clostridium thermoaceticum growth on, 224

391

consumption by Desulfouibriodesu[furicans,'225 microbial metabolism, 219 Carbon monoxide dehydrogenase in Methanobacterium Ihermoautotrophicum, 239 in Methanosarcina barkeri, 254 regulation, in Methanosarcina barkeri, 255 Carboxypeptidase S, in Saccharomyces cerevisiae, location, 59 Carboxypeptidase Y, in Saccharomyces cerevisiae vacuoles, 59 Carotenoids, conformation, 146 Catabolism, methanogens, 235-243 Catabolite inactivation, peroxisomal enzymes, regulation by, 59 Catalase assemblage in peroxisomes, 55 degradative inactivation, 66 in Candida boidinii, 64, 65, 66 in Hansenula polymorpha, 64,65,66, 67 in Kloeckera sp., 65 hydrogen peroxide removal by, 8 inactivation after exposure to excess methanol, 62-64 in microbodies, 16, 35 in peroxisomes, molecular substructure and, 24 in Saccharomyces cerwisiae, diaminobenzidine and, 18 methylamine metabolism and, 32 peroxisomal, regulation of synthesis, 41 productivity, in Hansenula polymorpha, 45 in Kloeckera sp., 45 synthesis in yeasts, regulation, 46 Cations, accumulation, in acidophilic bacteria, 175 Caulobacter sp., variable T mechanisms, 360 Cell bridges across stressed murein before cutting, 302-303 Cell carbon synthesis in methanogens, 243251 Cell curvature, electron microscopy _ .and, 3 19-322

392

SUBJECT INDEX

Cell diameter, electron microscopy and, 316 Cell dimensions electron microscopy and, 314-319 growth and, 305-307 Cell division bacterial, 302 constrictive, 304 primitive, mechanism, 350-351 surface stress forces and, 304305 Cell expansion, hydrostatic pressure and, 303 Cell poles formation, 349-354 shape, 342 Cell shape, growth and, 305-307 Cell wall expansion, growth and, 303 growth, Gram-positive rods, 326 growth, symmetry, 337 pulse incorporation of murein precursor, 331 tension, incell expansion, 303 Cerous ions, alcohol oxidase localization by, 20 Chemiosmotic hypothesis acidophilic bacteria, 174 alkalophilic bacteria, 174 Chemotrophic anaerobes, one-carbon compound metabolism by, 215-299 Chemoautotrophic anaerobes, relation to phototrophs and aerobes, 275-276 Chlorella sp. pyrophosphate synthesis in, 145 vacuoles, polyphosphate granules in, 98 Chlorella eIlQsoidea, polyphosphate metabolism in, 129 Chlorella pyrenoidosa, polyphosphate granules, potassium ions and, I32 Chlorobiwn thiosulphatophilum polyphosphate metabolism, enzymes, 114 pyrophosphate synthesis in, 145 Chromophores, membrane-associated, alkalophilic bacteria, 200 Chromosomes, segregation, 302

Cladosporium herbarum, polyphosphate phosphohydrolases, 110 Clostridium sp. alkalophilic, 190 formate production by, 218 methanol formation by, 220 Clostridium aceticum growth on carbon dioxide/hydrogen, 217 taxonomy, 260 Clostridium barkeri phylogenesis, 261 taxonomy, 259-260 Clostridium butyricum, formate production by, 218 Clostridiumformicoaceticum electron carriers in, 264 formate production by, 219 taxonomy, 260 Closlridium kluyeri, formate production by, 218 Clostridium lactoacetophilum, as homoacetogen, 260 Clostridiumpasteurianum carbon monoxide consumption, 224 formate production by, 219 methane production by, 221 phylogenesis, 261 Clostridium ramosum, phylogenesis, 26 1 Clostridium sporogenes, methanet hiol production by, 220 Clostridium sticklandii, as homoacetogen, 260 Clostridium thermoaceticum adenosine triphosphatase synthesis, 72 1 corrinoid enzyme from, 265 electron carriers in, 264 fermentation, applications, 287 formate production by, 219 homo-acetogenic pathway, 266 hydrogen/carbon dioxide consumption, 224 metabolism, 259 one-carbon metabolism, 264-267 taxonomy, 260 Clostridium thermoautotrophicum carbon monoxide consumption, 224 hydrogen/carbon dioxide consumption, 223

SUBJECT INDEX

Clostridium thermoautotrophicum (cont.) methanol consumption, 224 taxonomy, 260 Clostridium thermosaccharolyticum, phylogenesis, 261 Coal, Thermoplasma acidophilum growth on, 177 Cobalamins in methanogens, 228 methyl reduction and, 235 Coenzyme M in methanogens, 228 methanogenesis and, 236 from hydrogen/carbon dioxide or methanol, 236 Coenzyme YFC, in methanogens, 228 Corrinoids in acetate synthesis by Clostridium thermoaceticum, 265 in homo-acetogens, 264 in Methanosarcina barkeri, 228 Corynebacterium spp. polyphosphate: adenosine monophosphate phosphotransferase in, 105 polyphosphate kinase in, 105 polyphosphate(metaphosphate)dependent NAD+ kinase in, 106 polyphosphate phosphohydrolases, 110 polyphosphate utilization in, ATP synthesis and, 134 Cotton plant, polyphosphatase in, 110 Critical-point drying, cell shape and, 3 14 Cutting across stressed murein, cell bridges and, 302-303 Cylindrical elongation, surface stress theory for, 346349 Cyanobacteria alkalophilic, 190 high molecular-weight polyphosphates in, 100 polyphosphate granules, metal concentration, 138 polyphosphate metabolism, nucleic acids and, 133 polyphosphates in, intracellular localization, 134 Cyanophycin, 119 Cytochrome a in Bacillus alcalophilus membrane vesicles, 204

393

Cytochrome a3 in Bacillus alcalophilus membrane vesicles, 204 Cytochrome b in Bacillus alcalophilus membrane vesicles, 204 in homoacetogens, 264 in Mycoplasmas, 179 Cytochrome c in Bacillus alcalophilus membrane vesicles, 204 in Mycoplasmas, 179 Cytochrome d oxidase in Mycoplasmas, 179 Cytoplasmic malate dehydrogenase, peroxisomal, regulation by inactivation. 59

D Degradation, polyphosphates, enzymes in, 103-114 Degradative inactivation peroxisomal enzymes, 6 6 9 regulation by, 58 Dehydrogenases in methanol metabolism, 6 Desulfotomaculum sp., sulphate reduction by, 225 Desulfovibrio sp. carbon monoxide consumption, 225 methane production by, 221 methylamine production by, 22 1 sulphate reduction by, 225 Desulfovibrio desulfuricans carbon monoxide consumption, 225 hydrogen/carbon dioxide consumption, 225 methane consumption, 225 sulfate reduction, methanol in, 225 Desulfovibrio gigas hydrogen/carbon dioxide consumption, 225 low molecular-weight polyphosphates, cation binding, 139 magnesium tripolyphosphate granules, tripolyphosphatase and, 112 polyphosphate granules, 102 Desulfovibrio vulgaris carbon monoxide consumption, 225

394

SUBJECT INDEX

Desulfovibrio vulgaris (cont.) hydrogen/carbon dioxide consumption, 223, 225 Dictyostelium discoideum polyphosphate :adenosine monophosphate phosphotransferase in, 105 polyphosphate glucose kinase in, 107 polyphosphate granules in, 97 polyphosphate kinase in, 105 polyphosphate metabolism in, 114,126 polyphosphate phosphohydrolases, 110 Dihydroxyacetone synthase in methanol metabolism by yeasts, 14

1,3-Diphosphoglycerate:polyphosphate phosphotranferase, 107- 108 in polyphosphate metabolism, occurence, 115

E Ectothiorhodosporasp., alkalophilic, 191 Electrochemical proton gradient, alkalophilic bacteria, 194 Electron flow in methanogenesis, 238 Electron microscopy cell curvature and, 319-322 cell shape and, 312-322 polyphosphate granules, localization and, 100 preparation artifacts, 3 13-3 14 relative dimension estimates from sectioned material, 314319 Elongation, rods, 302 Endomyces magnusii high molecular-weight polyphosphates in, localization, 121 polymerized polyphosphates, localization, 121 polyphosphates in, 90, 98 intracellular, 92 metabolism, 89 polyphosphate phosphohydrolases in, 111 pyrophosphate synthesis in, 146 Energy in formaldehydeoxidation, 12-1 3 Entamoeba histolytica phosphorylation in, pyrophosphates and, 143

polypyrophosphate hydrolysis in, 142 pyrophosphate-dependent acetyl kinase in, 144 pyrophosphate-dependent phosphofructokinase in, 144 pyrophosphate utilization in, 143 pyruvate phosphate dikinase in, 143 Environment, one-carbon transformations and, 277-28 1 Enzymes extracellular, alkaline stability and, 206 in biosynthesis of polyphosphates, 103-1 14 in degradation of polyphosphates, 103-1 14 peroxisomal, degradative inactivation, 64-69 inactivation, 58-76 regulation by inactivation, 58-60, 60-64 regulation of synthesis, 41-47 variations, in polyphosphate metabolism in micro-organisms, 112-1 14 Escherichia coli autoradiograms, analysis, 323, 324 cell diameter, electron microscopy and, 313 cytoplasmic pH regulation, 191 intracellular orthophosphate in, polyphosphates in regulation of, 136 intracellular polyphosphates, exogenous orthophosphate and, 137 median section, 3 16 methanol production by, 220 morphometric measurements, 334 phosphohydrolases in, 137 pole formation, fixity and, 333 poles, shape, 360 polyphosphate metabolism in, 1 16,132 polyphosphate phosphohydrolases in, 111 potassium transport in, 175 pyrophosphate synthesis in, 145 sacculi, distribution of pulse-label, 336, 337 sodium ion transport, 193 solute motility, 200 tripolyphosphatase in, 150 tripolyphosphate hydrolase in, 11 1

395

SUBJECT fNfJEX

Ethanesulphonic acid, 2-mercapto-, in methanogens, 228 Ethylamine, metabolism by yeasts, 36 Eubacterium limosum carbon monoxide consumption, 224 methanol consumption, 224 taxonomy, 259 Eukaryotes high molecular-weight polyphosphates, intracellular localization, 89-99 polyphosphatemetabolism in, 114-132 Eutrophic lake sediments, methanogenesis in, 279 Evolution one-carbon transformations and, 282-285 phosphorus metabolism, polyphosphates in, 1 5 3 157

F Factor F4m, in methanogens, 228,238 Factor F ~ H ) in methanogens, 228 synthesis, in Methanobacterium thermoautotrophicum, 246 Fermentation technology, anaerobic one-carbon transformations and, 285 Ferredoxin in homo-acetogens, 264 Ferrous ion, Thiobacillus ferrooxidans growth and, 181 Filamentous cell growth, wall growth and, 356 Flavobacterium sp., alkalophilic, 190 Fluidity, membranes, in Thermoplasma acidophilwn, 184 Fluorescence microscopy in polyphosphate detection, 87 Formaldehyde alcohol oxidase synthesis and, 41 consumption, by anaerobes, 223 by sulphidogenic bacteria, 225 fixation, xylulose phosphate pathway, 14 formation from methanol, 6-1 1 metabolism in Methanobacterium thermoautotrophicum, 243

oxidation, energy from, 12-13 to carbon dioxide, 11-12 Formaldehyde assimilation path in evolution, 284-285 Formaldehyde dehydrogenases in peroxisomes, 2 1 in yeasts, 11 methylamine metabolism and, 32 Formate alcohol oxidase synthesis and, 41 consumption, by anaerobes, 223 by homo-acetogens, 224 metabolism, by Clostridium thermoaceticum, 265 in Methanobacterium thermoautotrophicum, 243 methanogenesis from, 239 production by anaerobes, 218 Formate dehydrogenase in Clostridium thermoaceticum, 265 in methanogenesis, 239 in peroxisomes, 21 in yeasts, 1 1 methylamine metabolism and, 32 S-Formylglutathione hydrolase, in Hansenula polymorpha, 12 Freezing in elec,tron microscopy, 3 13 Fructose bisphosphatase in Candida boidinii, methanol metabolism and, 15 in Hansenula polymorpha, methanol metabolism and, 15 in methanol metabolism by yeasts, 14 in peroxisomes, 2 1 peroxisomal, regulation by inactivation, 59 Fungi intracellular localization of high molecular-weight polyphosphates in, 89 polyphosphate metabolism in, 114-128 1

ti

Glucose oxidase, K, values for oxygen of, 9 Glutamate metabolism by Methanosarcina barkeri, 247

SUBJECT INDEX

396

Glutamate (cont.) synthesis, in Methanobacterium thermoautotrophicum, 244, 245 Glutamine: 2-oxoglutarate aminotransferase, regulation, in Methanobacteriwn thermoautophicum, 255 Glycerol alcohol oxidase synthesis and, 42 catalase synthesis and, 42 Glycine decarboxylase in homoacetogens, 257 Glycoside, hydrolytic cleavage, energy of activation, 344 Glyoxysomes, germinating oil seedlings, 4 Gram-negative rods cell division, 362 morphogenesis, surface stress theory, 333-340 Gram-positive bacteria growth, 362 split and stretch mechanism for pole formatian, 352-354 Gram-positive rods conserved and non-conserved regions, 354-359 shape, 342 surface stress theory, 326333 Growth cell bridges and, 302 cell dimensions and, 305-307 cell shape and, 305-307 cell wall expansion and, 303 diffuse, 3 1 1-3 12 efficiency, unicarbonotrophy and, substrate-product thermodynamics, 273-275 microbes, patterns, 305 Gulunolactone oxidase, K,,,values for oxygen of, 9

H Halobacterium sp. alkalophilic, 190 membrane lipids, 185 Hansenula sp., methanol metabolism by, 2 Hansenula polymorpha

alcohol oxidase in, 8 properties, 7 inactivation, 64, 65, 66, 67 after exposure to excess methanol, 62 synthesis, regulation, 42 asci, peroxisomas in, 54 ascospores, peroxisomes in, 54 bud formation, inactivation of alcohol oxidase and, 60 catalase activity in, localization, 18 catalase inactivation in, 64, 65, 66, 67 after exposure to excess methanol, 62 cell morphology, methanol metabolism and, 5 concurrent metabolism of C- and N-sources, peroxisomes and, 37-40 crystalline peroxisomes, 27 catalase removal from, 25 molecular substructure, 24 substructure, 26 formate dehydrogenase from, 11 fructose bisphosphatase in, methanol metabolism and, 15 generative reproduction, peroxisome development in, 53 methanol oxidation by, 10 methylamine metabolism, 32 peroxisomes in, 33 organelles, 16 peroxisomal breakdown in, generative reproduction and, 72 mechanism, 70 peroxisomal division in, 5 1 peroxisomal enzymes, regulation of synthesis, 41 peroxisomes, 17 crystalline alcohol oxidase in, 26 development in, 47, 48, 51 localization with cerium chloride, 22

staining, 19 recrystallized alcohol oxidase, 29 rough endoplasmic reticulum, 56 sphaeroplast, ribosomes and, 56 thiokinase in, methanol metabolism and, 15 He1i.r aspersa, polyphosphate granules in, 103

SUBJECT INDEX

Homo-acetogens in biotechnology, 287-288 in evolution, 283 one-carbon transformation, 257-272 spore morphological properties, 260 Hydrogen /carbon dioxide, consumption by anaerobes, 224-225 methanogenesis from, 236-238 consumption by anaerobes, 225 Hydrogen ions, transport, alkalophilic bacteria, 193 Hydrogen peroxide, removal, 8 Hydrostatic pressure, cell expansion and, 303 L-a-Hydroxy acid oxidase degradative inactivation, 65 in peroxisomal matrix, 29 in peroxisomes, 21 Hyphae, growth, 362 Hyphomicrobium sp., variable T mechanisms, 360 Hyphomicrobium X , methanol oxidation by, 10

I Inactivation alcohol oxidase, after exposure to excess methanol, 62-64 at low growth rates, 60-62 during bud formation, 60 catalase, after exposure to excess methanol, 62-64 peroxisomal enzymes, regulation by, 58-60,60-64 Isocitrate lyase, assemblage in peroxisomes, 55 a-Isopropylmalate synthase, peroxisomal, regulation by inactivation, 59

K Kloeckera sp. alcohol oxidase, properties, 7 synthesis, regulation, 42 catalase inhibition in, 65 crystalline peroxisomes, molecular substructure, 24, 25, 26

397

formate dehydrogenase from, 12 microbodies, 18

L Lampropedia, shape, 340 Levorin, polyphosphate utilization during synthesis of, 135 Light, polyphosphate metabolism in Chlorella ellipsoidea and, 129 Lignin, biodegradation, methanol production and, 220 Lipopolysaccharides, membrane-associated, in Thermoplasma acidophilum, 184 Luxury uptake, phosphates, 140 Lysine, binding to polyphosphates, in yeasts, 117

M Magnesium, binding to polyphosphates, in yeast vacuoles, 1 17 Magnesium tripolyphosphate in bacterial cells, 102 Manganese, binding to polyphosphates, in yeast vacuoles, 117 Mannoproteins, biosynthesis, localization, 124 Membranes fluidity, in Thermoplasma acidophilum, 184 in bioenergetic processes, 155 Membrane lipids, alkalophilic bacteria, 206 Menaquinones in homo-acetogens, 264 in mycoplasmas, 179 Merrulina lacrimans, pyrophosphate synthesis in, 145 Metabolic traps in micro-organisms, polyphosphates and, 141 Metabolism in methanogens, regulation, 251-256 unification, 25 1-256 of polyphosphates in eukaryotes, 114132 one-carbon compounds, by chemotrophic anaerobes, 2 15-299

SUBJECT INDEX

398

Metabolism (cont.) in homo-acetogens, 264-272 in yeasts, peroxisomes in, 1-82 phosphorus, evolution, polyphosphates in, 153 pyrophosphates in, 153-1 56 polyphosphates, in micro-organisms, 83171 pyrophosphates and, 150-153 variations in enzymes of, 1 12-1 14 in prokaryotes, 132-141 Metals, in cyanobacteria polyphosphate granules, 138 Metallogonium sp., external pH, 181 Methane consumption by Desulfovibrio desulfuricans, 225 environmental significance, 277 fossil fuel, 280 production by anaerobes, 221 Methanethiol consumption, by anaerobes, 223 by sulphidogenic bacteria, 225 production by anaerobes, 220 Methanobacteriaceae, taxonomy, 229 Methanobacteriales, taxonomy, 229 Methanobacterium sp. Methanobacterium sp., membrane lipids, 185

methanogenesis, electron flow, 238 physiology, 227 taxonomy, 229 Methanobacterium bryantii protoplasts, 227 taxonomy, 230 Methanobacterium formicicum formate consumption, 223 formate dehydrogenase in, 239 Methanobacterium ruminantium, formate dehydrogenase in, 239 Methanobacterium hungatii, sphaeroplasts, 227 Methanobacterium thermoautotrophicum anabolism, 243-246 autotrophic pathways, 248 carbon monoxide consumption, 223 carbon monoxide dehydrogenase in, 239 cell carbon synthesis, 243 growth efficiency, 273

internal pH, 242 methanogenesis, 237 electron flow, 238 from hydrogen/carbon, 236 methyl-coenzyme M reductase in, 237 physiology, 232 taxonomy, 229,230 Methanobacterium vanneili, formate dehydrogenase in, 239 Methanobrevibacter sp., taxonomy, 229 Methanobrevibacter arboriphilus, 23 1 Methanococcaceae, taxonomy, 229 Methanococcales, taxonomy, 229 Methanococcus sp. membrane lipids, 185 physiology, 227 taxonomy, 229 Methanococcus vanneili culture, 23 1 formate consumption, 223 Methanogens bioregulatory roles, 278 catabolism, 235-243 cell carbon synthesis, 243-251 cell-wall composition, 227 culture, 231 in biotechnology, 286287 in evolution, 283 interaction with sulphate reducers in sediments, 279 one-carbon compound transformation in, 226256 physiology, 227-235 sulphate in fresh water lake sediment and, 279 taxonomy, 228 Methanogenesis adenosine triphosphate synthesis and, 242 from hydrogen/carbon dioxide, 236238 from methanol, 236238 Methanogenium sp., taxonomy, 229 Methanol assimilation, 13-16 consumption by anaerobes, 223 dissimilation, 6-13 in sulphate reduction by Desulfovibrio desulfuricans, 225 metabolism, by Candida boidinii, 2

SUBJECT INDEX

Methanol (cont.) by Methanosarcina barkeri, 246 peroxisomes and, 5 3 0 methanogenesis from, 236238 oxidation to formaldehyde, 6 1 1 peroxisomal enzymes in oxidation of, regulation of synthesis, 41 production by anaerobes, 220 Methanomicrobiaceae, taxonomy, 229 Methanomicrobiales, taxonomy, 229 Methanomicrobium sp., taxonomy, 229 Methanoplasma elizabethii, taxonomy, 23 1 Methanopoieticum sp., taxonomy, 229 Methanopterin, carboxy-7,8-dehydro-, in methanogenesis from hydrogen/ carbon dioxide or methanol, 236 Methanopterin, carboxydihydro-, in metabolism in methanogens, 253 Methanosarcina sp. acetate fermentation, 233 consumption of one-carbon compounds by, 223 culture, 234 membrane lipids, 185 physiology, 227 taxonomy, 229,230 Methanosarcina barkeri acetate consumption, 221 acetate fermentation, 233 anabolism, 246248 ATP synthesis, 242 ATPase, 243 autotrophic pathways, 248 batch growth curve, 233 carbon monoxide consumption, 223 carbon monoxide dehydrogenase in, 239 carbon monoxide metabolism, 240 cell carbon synthesis, 243 consumption of one-carbon compounds by, 223 corrinoids in, 228 cultural adaptation, 256 culture, 231 growth efficiency, substrate-product thermodynamics, 273 unicarbonotrophy and, 274 growth yield values, 274 metabolic versatility, 233

399

metabolism, regulation, 253 unification, 25 1 methanogenesis, 237, 240 electron flow, 238 from hydrogen/carbon, 236 methanol fermentation by, 217 methanol metabolism, 276 methylamine production by, 220 methylotrophy, 253 one-carbon metabolism, 269 physiology, 232 taxonomy, 230-23 1 Methanosarcina biotype 2, acetate metabolism by, 233 Methanosarcina mazei consumption of one-carbon compounds by, 223 taxonomy, 231 Methanosarcina methanica, taxonomy, 23 1 Methanosarcina vacuolata consumption of one-carbon compounds by, 223 taxonomy, 231 Methanosarcina voltae, culture, 232 Methanosarcinaceae, taxonomy, 229 Methanospirillum sp. membrane lipids, 185 physiology, 227 taxonomy, 229 Methanothermus feridus, taxonomy, 230 Methanothrix soehngenii, taxonomy, 23 1 Methionine, S-adenosyl-, binding to polyphosphates, in yeasts, 117 Methyl-coenzyme M in methanogenesis, from hydrogen/ carbon dioxide or methanol, 236 from methanol, in Methanosarcina barkeri, 237 Methyl coenzyme M reductase in Methanobacterium thermoautotrophicum, 237 Methylamine as nitrogen source, peroxisomes and, 30-3 1 consumption, by anaerobes, 223 by sulphidogenic bacteria, 225 metabolism, peroxisome role in, 3040 by yeasts, 3

400

SUBJECT INDEX

Methylamine (cont.) methanogenesis from, 239 production by anaerobes, 220-221 Methylococcus sp. methanol oxidation in, 276 ribulose bisphosphate carboxylase activity, 276 Methylotrophy in homo-acetogens, 259 in Methanosarcina barkeri, 253 in evolution, 284 Microbodies 1-carbon compound metabolism and, 3 enzymic composition, cytochemistry and, 18 methanol metabolism and, 5 methylamine metabolism and, 35 enzymic composition, 35 peroxisomes and, 16 Micrococcaceae, taxonomy, 229 Micrococcus sp. alkalophilic, 190

polyphosphate(metaph0sphate)dependent NAD+ kinase in, 106 polyphosphate phosphohydrolases, 110 Microcystic aeruginosa, alkalophilic, 190 Micro-organisms polyphosphate, distribution in, 85 metabolism in, 83-171 pyrophosphates and, 15&153 variations in enzymes of, 112-1 14 Morphogenesis bacterial, biosynthesis in, 307 microbial, surface stress theory, 301365 Motility, bacterium shape and, 341 Moulds, alcohol oxidase from, properties, 7 Murein addition to cell poles, 335 extension, 307 formation, cell shape and, 305-307 precursor, pulse incorporation, into cell walls, 331 into Gram-negative rods, 335-340 stressed, cell bridges before cutting, 302-303

Mycobacteria, polyphosphate metabohsm in, 132 Mycobacterium smegmatis, polyphosphate localization in, 101 Mycobacterium tuberculosis, polyphosphate glucose kinase in, 107 Mycoplasma sp., 177 fission, 349

N Narrow growth zone, pole formation and, 349 Narrow zonal growth, mathematics, 310-311 Neurospora crassa 1,3-diphosphoglycerate:polyphosphate phosphotransferase in, 108 high molecular-weight polyphosphates in, localization, 121 phosphorus metabolism in, 91 polyphosphate depolymerase in, intracellular localization, 120 polyphosphate kinase in, 104 polyphosphates in, 92, 127 growth and, 128 intracellular, 92 localization, 97 metabolism, 89, 114 localization, 121 polyphosphate polyphosphohydrolases in, 109 polyphosphate phosphohydrolases in, 111 tripolyphosphatase in, 150 intracellular localization, 1 12 tripolyphosphate hydrolase, 11 1 Nickel tetrapyrrole, methanogenesis and, 236 Nicotinamide adenine dinucleotide dehydrogenases in mitochondria, 13 Nicotinamide adenine dinucleotidedependent dehydrogenase in methanol oxidation in yeasts, 10 Nicotinamide adenine dinucleotide in methanol metabolism by yeasts, 12 Nicotinamide adenine dinucleotide, formation, methanogenesis, 238

SUBJECT INDEX

Nocardia sp., glucose phosphorylation, 135 Nocardia erythropolus, polyphosphate phosphohydrolases, 110 Nocardia erythropolus, tripolyphosphate hydrolase in, 1 11 Nocardia minima, polyphosphate glucose kinase in, 106107 Nostoc pruniforme, polyphosphate granules, 100 Nuclear magnetic resonance, 31P, polyphosphate localization by, 101 Nucleic acids, metabolism, polyphosphate metabolism and, 132

0 One-carbon compounds consumption by anaerobes, 22 1-226 metabolism, by chemotrophic anaerobes, 2 15-299 in homo-acetogens, 264-272 in yeasts, peroxisomes in, 1-82 metabolites, transformation by anaerobes, 218-226 production by anaerobes, 2 18-221 transformation, in homo-acetogens, 257-272 in methanogens, 226256 significance, 276288 Orthophosphates, regulation, 138 Oxidases, hydrogen peroxide-producing, K, values for oxygen of, 9 formaldehyde, energy from, 12-1 3 to carbon dioxide, 11-12 methanol, to formaldehyde, 6-1 1 2-Oxoglutarate synthase, Methanobacterium thermoautotrophicum, 244, 245 Oxygen in Earth’s atmosphere, 154

P Parabola-shaped pole, Gram-positive rods, 330-331 Paracoccus sp., ribulose bisphosphate carboxylase activity, 276 Paracoccus denitrijicans one-carbon compound consumption, 226

401

polyphosphate synthesis, mechanism, 134 Pectin, metabolism by yeasts, 2 Penicillum sp., polyphosphate phosphohydrolases, 1 10 Penicillium chrysogenum orthophosphate regulation in, 138 polyphosphate metabolism in, 114 Peptidoglycan covalent bonds, stress on, 343-346 in cell membranes, 326 synthesis in Bacillus megaterium, 33 1 Peroxisomes assemblage, 55-58 biogenesis, 40-58 crystalline, molecular substructure, 2430 degradation, 58-76 subcellular events in, 69-76 development, in spore formation, 5355 in vegetative growth, 47-53 enzymes, degradative inactivation, 6 4 69 inactivation, 58-76 regulation, by inactivation, 60-64 of synthesis, 4 1 4 7 function, 16-23 adaptation, 4 in spore formation, 53-55 in animal cells, 21 in concurrent metabolism of C- and N-sources, 37-40 in metabolism of one-carbon compounds in yeasts, 1-82 in methanol-grown yeasts, metabolism and, 21 in methanol metabolism, 5-30 by yeasts, 23 in methylamine metabolism, role, 3040 microbodies and, 3 structure, 16-23 pH, cytoplasmic, regulation, 191-193 Phosphates, metabolism in microorganisms, pyrophosphates and, 150-153 Phosphate shift experiments, Grampositive rods, conserved and nonconserved regions and, 355

402

SUBJECT INDEX

Phosphoenolpyruvate carboxykinase, peroxisomal, regulation by inactivation, 59 Phosphoenolpyruvate carboxylase, Methanobacterium thermoautotrophicum, 244 Phosphohydrolases, in Escherichia coli, 137 Phosphorus metabolism, evolution, polyphosphates in, 153-1 57 pyrophosphates in, 153-157 Phosphorylation in bacteria, pyrophosphates in, 142145 oxidative, ATP synthesis and, 201203 pyrophosphates and, 1 4 5 150 Phototrophs, relation to chemoautotrophic anaerobes, 275-276 Physarum sp., polyphosphate metabolism in, 116 Physarum pobcephalum phosphorus metabolism in, 91 polyphosphates in, 127 localization, 92 metabolism in, 114 Physiology acidophilic bacteria, 173-214 alkalophilic bacteria, 173-214 Phytophthora infestans polyphosphate phosphohydrolases in, 110

tripolyphosphatase in, 150 tripolyphosphate hydrolase in, 1 12 Pichia sp. alcohol oxidase from, properties, 7 formaldehyde fixation in, 16 methanol metabolism by, 2 Pichia pastoris, alcohol oxidase from, properties, 7 Plectonema boryanum polyphosphate granules, 100, 101 polyphosphates in, localization, 133 strontium in, 138 Poles formation, fixity and, 329-330 Gram-negative rods, 333-335 shape and, 329 nascent, shapes, 360

Polymethylamines, metabolism, by methanogens, 233 Pol yphosphatase competition with polyphosphate kinase, 135 in Aerobacter aerogenes, 112 Pol yphosphates biosynthesis, enzymes in, 103-1 14 degradation, enzymes in, 103-1 14 granules, cytological, 86 high molecular-weight, 89-142 in evolution of phosphorus metabolism, 153-1 57 in microbial metabolism, physiological role, 141-142 in primordial organisms, 154 intracellular localization, 89-103 in eukaryotes, 89-99 in prokaryotes, 99-103 inorganic, 8 4 8 5 detection, 86-88 distribution in micro-organisms, 85 extraction, 88 fractionation, 86-88 identification, 86-88 metabolism, in eukaryotes, 114-132 in micro-organisms, 83-1 7 1 variation in enzymes of, 112 in Neurospora crassa, 121 in prokaryotes, 132-141 nucleic acid metabolism and, 132 Polyphosphate :ADP phosphotransferase, 103-105 Polyphosphate depolymerase in Neurospora crassa, 119 intracellular localization, 120 in polyphosphate metabolism, 118 Polyphosphate glucokinase, in Streptomyces levoris, 136 Polyphosphate :D-glucose 6-phosphate phosphotransferase, 106-107 Polyphosphate hexokinase in evolution, 155 occurrence, 112 Polyphosphate kinase competition with polyphosphatase, 135 in Aerobacter aerogenes, 1 12 in algal vacuoles, 131 in polyphosphate metabolism, 118

SUBJECT INDEX

Polyphosphate kinase (cont.) in Saccharomyces carlsbergensis, 1 18 reverse activity, in Saccharornyces carlsbergensis, 1 18 Polyphosphate (metaphosphate)-dependent NAD+ kinase, 105-106 Polyphosphate overplus, yeasts and, 85 Polyphosphate phosphohydrolases, 109112

Polyphosphate polyphosphohydrolases, 108-109 Polyporus sp., alcohol oxidase from, properties, 7 Polysaccharides in Saccharomyces carlsbergensis, polyphosphates and, 123 Poria contigua alcohol oxidase in, 3 properties, 7 Potassium accumulation, in acidophilic bacteria, 175 polyphosphate granules in Chlorella pyrenoidosa and, 132 Prokaryotes high molecular-weight polyphosphates, intracellular localization, 99- 103 polyphosphate metabolism in, 132-141 Propionibacteriwn reudenreichii, serine phosphorylation in, 144 Propionibacterium shermanii phosphorylation, pyrophosphates and, 142 polyphosphate glucose kinase in, 107 pyrophosphate-dependent phosphofructokinase in, 144 serine phosphorylation in, 144 Propionibacterium technicum, serine phosphorylation in, 144 Propionic-acid bacteria polyphosphate metabolism in, 132 pyrophosphate utilization in, 143 Protease, degradative inactivation, in Candida boidinii, 68 Protein 'e' in Escherichia coli, 137 Proteinase A, in Saccharomyces cerevisiae vacuoles, 59 Proteinase B, in Saccharomyces cerevisiae vacuoles, 59

403

Proteolysis in spore formation in yeasts, 59 Proteus vulgaris, methanethiol production by, 220 Proton-motive force, 174 Pseudomonas sp., alkalophilic, 190 Pseudomonas marina, pyrophosphatedependent phosphofructokinase in, 144

Pteridines in homo-acetogens, 264 Pulse incorporation experiments, conserved and non-conserved regions and, 358 Pyrophosphates high molecular-weight, in evolution of phosphorus metabolism, 153-1 57 in phosphorylation in bacteria, 142145

in primordial organisms, 154 inorganic, metabolic and physiological role, 142-1 53 phosphorylation and, 145-1 50 polyphosphate metabolism in microorganisms and, 150-153 Pyrophosphate :ADP phosphotransferase, in Corynebacterium sp., 135 Pyrophosphate-dependent acetyl kinase, 144

Pyrophosphate-dependent phosphoenolpyruvate carboxykinase, 143 Pyrophosphate-dependent phosphofructokinase, 144 Pyruvate phosphate dikinase, 143 Pyruvate synthase, Methanobacterium thermoautotrophicum, 244, 245

Q Quinoproteins in methanol metabolism, 6

R Regulation cytoplasmic pH, 191-193 metabolism, in methanogens, 25 1-256 peroxisomal enzymes, 41-47 by inactivation, 58-60 Renobacter sp., polyphosphates, localization, 102

404

SUBJECT INDEX

Renobacter vacuofatum polyphosphate and ATP in, 140 polyphosphate glucose kinase in, 107 polyphosphate hexokinase in, 112 polyphosphate phosphohydrolases, I10 tripolyphosphate hydrolase in, 1 11 Renovated Replicon Model, 340 Respiratory chain, alkalophilic bacteria, 203-205 Rhodopseudomonas sp. methanol oxidation in, 276 ribulose bisphosphate carboxylase activity, 276 Rhodopseudomonas acidophilia, one-carbon compound consumption, 226 Rhodopseudomonas capsulata, one-carbon compound consumption, 226 Rhodopseudomonas palustris phosphorylation in, pyrophosphates and, 143 pyrophosphate synthesis in, 145 Rhodopseudohonas rubrum phosphorylation in, pyrophosphates and, 143 pyrophosphate synthesis in, 145, 152 Rhodopseudomonas viridis, pyrophosphate synthesis in, 145 Rhodospir;illum sp., formate production by, 218 Rhodospirillum rubrum polyphosphates, localization, 134 synthesis, 147 pyrophosphatases in, 147 Ribose alcohol oxidase synthesis and, 42 catalase synthesis and, 42 Rieske protein, Bacillus alcalophilus, 205 Ristellaceae, taxonomy, 229 Rods elongation, 302 shape, 341 Rough endoplasmic reticulum in Hansenula polymorpha, 56 Rozella allomycis, polyphosphates in, localization, 125 polyphosphate granules in, 97 Rubredoxin in homo-acetogens, 264

S

Saccharomyces sp., pyrophosphate synthesis in, 151 Saccharomyces carlsbergensis mannan enzymes, 126 polyphosphates in, 90, 91 biosynthesis, 125 enzymes, 126 polyphosphate kinase ~ . . in, 1 18 polymerized polyphosphates, localization, 121 polysaccharides in, polyphosphates and, 123 reverse polyphosphate kinase activity, 1 I8 vacuole pool, polyphosphates in, 96 Saccharomyces cerevisiae catalase in, diaminobenzidine and, 18 orthophosphates in, 122 polyphosphates, 122 synthesis, 151 polymerized polyphosphates, localization, 121 proteases in, 59 pyrophosphate synthesis in, 146 tripolyphosphate in, 95 Saccharomycopsis lipolytica low molecular-weight polyphosphates in, 96 polyphosphates in, 127 Scenedesmus sp., vacuoles, polyphosphate granules in, 98 Sediments, sulphate reducers in, methanogens and, 279 Septum formation, 304 Gram-negative rods, 333 Shape bacteria, 302 ideal, 34&343 during pole formation, 329 variable T mechanisms and, 361 Soap bubbles theory, 308-310 Sodium ions symport, alkalophilic bacteria, 197 transport, alkalophilic bacteria, 193 Soil, polyphosphatases in, 110 Solute motility in alkalophilic bacteria, 196201

405

SUBJECT INDEX

Solute transport in alkalophilic bacteria, 196-20 1 Sorbitol alcohol oxidase synthesis and, 42 catalase synthesis and, 42 C-Sources, concurrent metabolism with N-sources, peroxisomes in, 3 7 4 0 N-Sources, concurrent metabolism with C-sources, peroxisomes and, 37-40 Spirillaceae, taxonomy, 229 Spirillum volutans, volutin granules, polyphosphates in, 99 Split and stretch model, 353 Spore formation, peroxisome development and function in, 53-55 Square bacteria, shape, 340 Staphylococcus sp., shape, 341 Streptococci, wall growth, 310 Streptococcus sp. curvature measurements, electron microscopy and, 3 19 formate production by, 218 septum formation, 362 shape, 341 Streptococcus faecalis, sodium ion transport, 193 Streptococcus faecium cell wall growth, 307 pole formation, 350 shape, surface stress theory, 3 1 1 Streptococcus mitis, carbon monoxide formation by, 220 Streptomyces levoris polyphosphate glucokinase in, 136 polyphosphate metabolism in, 135 polyphosphate phosphohydrolases, 110 tripolyphosphate hydrolase in, 11 1 Streptomycetes, polyphosphate metabolism in, 132 Stress on peptidoglycan covalent bonds, 343-346 on murein before cutting, cell bridges, 302-303 Strontium in cyanobacterium polyphosphate granules, 138 Sugars, transport through cytoplasmic membrane, polymerized polyphosphates in, 122

Sulfolobus sp., internal pH, 180 lipids, 186 membrane lipids, 186 pili, 186 Sulfolobus acidocaldarius, 177 glycerol diethers, structure, 185 internal pH, 180 Sulphates in fresh water lake sediment, sulphidogens and, 279 reducers in sediments, methanogens and, 279 reduction, anaerobes and, 225 Sulphidogens, sulphate in fresh water lake sediment and, 279 Surface properties, acidophilic bacteria, 184186 Surface stress forces, cell division and, 304305 Surface stress theory for cylindrical elongation, 346349 Gram-positive rods, 326 microbial morphogenesis, 301-365 Surface structures, alkaline stability and, 206 Surface-to-volume ratio, bacterium shape and, 340 Synechococcus sp. alkalophilic, 190 pH and, 206

T Teichoic acid in cell walls, 326 Temperature shift experiments, Grampositive rods, conserved and nonconserved regions and, 354 Tetrahymena pyriformis granules, polyphosphates in, 98 magnesium tripolyphosphate in, 102 Thermodesulfotobacterium commune, hydrogen/carbon dioxide consumption, 225 Thermoplasma sp. lipids, 186 membrane lipids, 186 Thermoplasma acidophilum, 117 internal pH, 178 membrane-associated lipopolysaccharides in, 184

406

SUBJECT INDEX

U membrane-bound ATPase, 185 membrane fluidity in, 184 pH and, 187 transmembrane electrical charge gradient, 182 ThermusJavus polyphosphates, ATP and, 140 localization, 102 Thin-layer chromatography in polyphosphate detection, 87 Thiobacillus acidophilus, internal pH, 180 Thiobacillusferrooxidans, 176 ATP synthesis in membrane vesicles, 181 external pH, 181 internal pH, 180 transmembrane electrical charge gradient, 183 Thiobacillus thiooxidans, 176 Torulopsis sp., methanol metabolism by, 2 Transmembrane electrical charge gradient in acidophilic bacteria, 182-183 Transmembrane electrical potential, alkalophilic bacteria, 194 Transmembrane pH gradient, acidophilic bacteria, 178-1 82 Tricarboxylic acid cycle Methanobacterium thermoautotrophicum, 244 Methanosarcina barkeri, 247 Trichosporon sp., methylamine metabolism in, 32 Thiokinase in Candida boidinii, methanol metabolism and, 15 in Hansenula polymorpha, methanol metabolism and, 15 Tripolyphosphatase, 150 Tripolyphosphate hydrolase in bacteria, 111

Tuberoidobacter sp., polyphosphates, localization, 102 Tuberoidobacter mutans polyphosphate and ATP in, 140 polyphosphate phosphohydrolases, 110 tripolyphosphate hydrolase in, 11 1

Ubiquinone in Bacillus sp., 205 Unicarbonotrophy, metabolic perspectives, 272-276 Urate oxidase in peroxisomes, 21 K , values for oxygen of, 9 synthesis, regulation, 46 Uricase assemblage in peroxisomes, 55 in peroxisomal matrix, 29

v Vacuoles peroxisomal breakdown and, 70 proteolysis in yeasts and, 59 yeast, high molecular-weight polyphosphates in, 94 polyphosphates in, chain length, 94 Variable T mechanisms, 359-362 Vegetative growth, yeasts, peroxisomes development in, 47-53 Vibrio succinogenes, formate consumption, 224

W Waste treatment anaerobic one-carbon transformations and, 285 methanogens in, 286287 Wolinella succinogenes, formate consumption, 224

X Xanthine oxidase, K,,,values for oxygen of, 9 X-ray energy-dispersion microanalysis with electron microscopy, polyphosphate granules and, 101 Xylose alcohol oxidase synthesis and, 42 catalase synthesis and, 42 Xylulose phosphate pathway in formaldehyde fixation, 14

SUBJECT INDEX

Y Yeasts alcohol.oxidase from, properties, 7 intracellular localization of high molecular-weight polyphosphates in, 89 metabolism of one-carbon compounds in, peroxisomes in, 1-82 peroxisome function adaptation in, 4 polyphosphates in, 85 metabolism in, 114128 spore formation, peroxisome development and function in, 53-55 tripolyphosphatase in, 150 vacuoles, high molecular-weight poly-

407

phosphates in, 94 polyphosphates in, chain length, 94 vegetative growth, peroxisomes development in, 47-53 Yellow fluorescent compound in methanogenesis from hydrogen/carbon dioxide or methanol, 236

2 Zeikusella sp., taxonomy, 229 Zonal dome Gram-positive rods, 33&33 1 models, pole formation and, 349

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