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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. . .
. . . . .
.
. . . . .
.
. . . .
.
.
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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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
.
.-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
Ackerman, E. and Brill, A. S. (1965). Biochimica et Biophysica Acta 96, 357. Anthony, C. (1975). Science Progress, Oxford 62, 167. Avers, C. J. (1971). Subcellular Biochemistry 1, 25. Babel, W. and Loffhagen, N. (1979). Zeitschrqt f u r Allgemeine Mikrobiologie 19,299. Barnett, J. A., Paine, R. W. and Yarrow, D. (1979). “A Guide to Identifying and Classifying Yeasts.” Cambridge University Press, Cambridge. Beevers, H. (1969). Annals of the New York Academy of Sciences 168, 313. Betz, H. and Weiser, U. (1976a). European Journal of Biochemistry 70, 385. Betz, H. and Weiser, U. (1976b). European Journal of Biochemistry 62, 65.
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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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I. Introduction One of the topical problems of modern biochemistry is the elucidation of the mechanisms underlying the synthesis and utilization of energy-rich phosADVANCES IN MICROBIAL PHYSIOLOGY, ISBN 0-12027724-7 VOL. 24
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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
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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).
86
IGOR S. KULAEV AND VLADlMlR
M. VAGABOV
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
IGOR S. KULAEV AND VLADlMlR M. VAGABOV
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
92
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-
POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS
93
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).
94
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
POLYPHOSPHATE METABOLISM I N MICRO-ORGANISMS
97
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).
IGOR S. KULAEV AND VLADlMlR M. VAGABOV
98
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).
POLYP H OSPHATE METABOLIS M I N MIC RO - 0R GAN ISMS
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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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IGOR S. KULAEV AND VLADlMlR M. VAGABOV
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
IGOR S. KULAEV AND VLADlMlR M . VAGABOV
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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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IGOR S. KULAEV AND VLADlMlR M. VAGABOV
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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1 09
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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IGOR S. KULAEV AND VLADlMlR M. VAGABOV
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
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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.
114
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).
122
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,
POLYPHOSPHATE METABOLISM I N MICRO-ORGANISMS
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).
POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS
125
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
126
IGOR S. KULAEV AND VLADlMlR M. VAGABOV
-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
130
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.
132
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
POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS
133
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
POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS
135
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
POLYPHOSPHATE METABOLISM IN MICRO-ORGANISMS
139
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-
140
IGOR S. KULAEV AND VLADlMlR
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
141
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
142
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
151
(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
S. KULAEV
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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A. Special problems of life at low pH values . . . B. Organisms described . . . . . . . C. Physiological adaptations that meet the problems . . D. Why can’t obligate acidophiles grow at neutral pH values? 111. Alkalophilic bacteria . . . . . . . A. Special problems of life at high pH values . . . B. Organisms described . . . . . . . C. Physiological adaptations to meet the problems . . D. Why can’t obligate alkalophilesgrow at neutral pH values? IV. Concluding remarks . . . . . . . V. Acknowledgements . . . . . . . . References . . . . . . . . .
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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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TERRY A. KRULWICH AND ARTHUR A. GUFFANTI
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 +
PHYSIOLOGY OF AClDOPHlLlC AND ALKALOPHILIC BACTERIA
181
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
182
TERRY A. KRULWICH AND ARTHUR A. GUFFANTI
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
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.
264
J. G. ZEIKUS
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
268
J. G . ZEIKUS
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.
METABOLISM OF ONE-CARBON COMPOUNDS
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-)