Advances in
MICROBIAL PHYSIOLOGY
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Advances in
MICROBIAL PHYSIOLOGY
This Page Intentionally Left Blank
Advances in
MICROBIAL PHYSIOLOGY Edited by
A. H. ROSE School of Biological Sciences Bath University, UK
and
D. W. TEMPEST Department of Microbiology University of Shefield, UK
Volume 27 1986
ACADEMIC PRESS Harcourt Brace Jovanovich, Publishers London Orlando San Diego New York Austin Montreal Tokyo Sydney Toronto
ACADEMIC PRESS INC. (LONDON) LTD. 24-28 Oval Road London NWl 7DX US.Edition published by ACADEMIC PRESS INC. Orlando, Florida 32887
Copyright 0 1986 by ACADEMIC PRESS INC. (LONDON) LTD.
All Rights Reserved
No part of this book may be reproduced in any form by photostat, microfilm,or any other means, without written permission from the publishers
British Librory Cataloguing in Publication Data
ISBN 0-12-027727-1 ISSN 0065-291 1
Printed in Great Britain at the Alden Press, Oxford
Contributors C. Anthony Department of Biochemistry, University of Southampton, Southampton
SO9 3TU, UK
A.W. Bunch Biological Laboratory, University of Kent, Canterbury, Kent.CT2 7NJ, UK E.F. Gale Sub-department of Chemical Microbiology, Department of Biochemistry, University of Cambridge, Cambridge CB2 lQW, UK D. Kemdge Department of Biochemistry, University of Cambridge, Tennis Court Road, Cambridge CB2 IQW, UK C.J. Knowies Biological Laboratory, University of Kent, Canterbury, Kent CT2 7NJ, UK
A.J. Messenger Department of Biochemistry, University of Hull, Hull HU6 7RX, UK J.M. Turner Department of Biochemistry, University of Liverpool, Liverpool L69 3BX, UK
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Contents
V
Contributors
Mode of Action of Clinically Important Antifungal Drugs D. KERRIDGE I. 11. 111. IV. V.
Introduction The nucleus as the primary target The cell membrane as the primary target Finale Acknowledgements References
1 5 19 57 63 64
Microbial Cyanide Metabolism C.J. KNOWLES and A. W. BUNCH I. 11. 111. IV. V. VI.
Introduction Bacterial cyanide production Fungal cyanogenesis Cyanogenesis by photosynthetic micro-organisms Cyanide degradation Concluding remarks References
73 74 86 90 95 105 106
Bacterial Oxidation of Methane and Methanol C. ANTHONY I. 11. 111. IV. V.
Introduction Oxidation of methane to methanol Oxidation of methanol to formaldehyde Energy transduction during the oxidation of methane and methanol Acknowledgements References
113 116 129 179 203 203
...
CONTENTS
Vlll
Occurrence, Biochemistry and Physiology of Phenazine Pigment Production JOHN M. TURNER and ANN J. MESSENGER I. Introduction 11. Natural occurrence and some properties of phenazines 111. Biosynthesis
21 1 218 242
IV. Secondary metabolism and the physiological significance of 260 phenazine production 268 V. Acknowledgements 268 References
Nature and Development of Phenotypic Resistance to Arnphotericin B in Candida albicans ERNEST F. GALE
I. Introduction
TI. Mode of action of amphotericin 111. Assessment of amphotericin sensitivity
IV. Interactions between amphotericin, sterols and surface structures of Cundidu ulbicans V. Changes in the cell wall during the stationary phase of culture VI. The effects of oxidation and reduction VII. Nature of the cell-wall barrier and its modification by reducing agents VIII. Incorporation of glucose into the (1 -3)$-glucan fraction IX. Actions of analogues of glucose X . Conclusions and in conclusion References Note added in proof Author Index Subject Index
278 28 1 283 286 289 293 297 303 305 316 318 321 323 34 1
Mode of Action of Clinically Important Antifungal Drugs DAVID KERRIDGE Department of Biochemistry, University of Cambridge, Tennis Court Road, Cambridge, CB2 IQW, UK
I. Introduction . 11. The nucleus as the primary target . A. Griseofulvin . B. 5-Fluorocytosine 111. The cell membrane as the primary target A. Polyene macrolide antibiotics . B. Imidazole antimycotics . C. Naftifine . IV. Finale . V. Acknowledgements. . References .
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1 5 5 11 19 20
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39 56 57 63
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4
I. Introduction
The fungi comprise a large complex group of eukaryotic protista of which only a limited number are important plant and animal pathogens. Economically the fungi responsible for plant diseases and food spoilage are probably more important than those causing human and animal infections, but this article will be restricted to antimycotic drugs used in the therapy of human infections. The first description of a human mycotic infection was of oral candidosis by Hippocrates, a disease known as thrush since the time of Samuel Pepys. The causative organism, Candida albicans, was first described by Ellis (1878) and named Oodium albicans. Since that time some 100 fungal species have been found to be human pathogens and the most important of these are listed in Table 1. These organisms range from those whose habitat is soil, but yet are capable of infecting normal healthy humans, to commensal fungi which, under conditions where the hosts defense mechanisms are impaired, cause life-threatening diseases; of the latter group, C . albicans is the ADVANCES IN MICROBIAL PHYSIOLOGY, VOL. 27 ISBN 0-12-027727-1
Copyright 0 1986 by Academic Press London All rights of reproduction in any form reserved
TABLE 1. Major fungal diseases ~
Site of infection Lung
Disease Aspergillosis Blastomycosis Coccidioidomycosis Histoplasmosis
Wounds
Skin and mucous membranes
Chromomycoses Mycetomas Sporotrichosis Candidosis Dermatophytosis Pityriasis versicolor
Fungus Aspergillusfwnigatus Blastomyces dermatidis Blastomyces brasiliensis Coccidioides immitis Histoplasma capsulatum Histoplasma dubosii Cladosporium carrionii Phialophora spp. 16 species identified Sporothrix schenkii Candida albicans Epidermophyton spp. Microsporwn spp. Trichophyton spp. Pityrosporumfurfur
Normal habitat soil Soil (North America) Soil (South America) Desert areas in the U.S.A. Soil enriched with bird and bat droppings Tropics and subtropics
Soil Tropics and subtropics Commensal on mucous membranes and in the gut Usually found associated with skin or with human detritus in the soil Usually associated with skin
MODE OF ACTION OF ANTIFUNGAL DRUGS
3
most important opportunistic pathogen. In the U.K., vaginal candidosis is the most common fungal infection. It has been estimated that approximately 15% of pregnant women suffer from this condition (Odds, 1980). Candida albicans is also a predominant factor in denture stomatitis (Budtz-Jorgenson, 1974) a condition affecting a very high percentage of denture wearers. Although, in temperate climates, C. albicans and the dermatophytic fungi are the most common fungal pathogens, other fungi assume a much greater importance in tropical and subtropical regions. It has been estimated that there are about two million new systemic fungal infections each year, of which some two thousand are fatal. In the U.S.A., approximately forty million people have been infected by Histoplasma capsulatum, and in areas where histoplasmosis is endemic up to 90% of the population show a positive skin reaction to histoplasmin. Similarly, with coccidioidomycoses, a disease endemic in certain areas of the U.S.A., a high percentage of the population show a positive skin test. Success in controlling bacterial infections and changes in medical practice have resulted in a dramatic increase in the relative importance of fungal infections, in particular candidosis and aspergillosis. The problem is exacerbated by difficulties of rapid diagnosis and absence of any completely satisfactory drug for treatment of patients with systematic mycoses. Unfortunately, fungi, like their human hosts, are eukaryotic organisms and hence the number of potential targets for drug action is limited. Prior to 1950, there were no reliable or safe drugs for systemic and invasive mycoses, and only traditional empirical preparations were available for dermatomycoses (Baum, 1979; Holt, 1980a). The major breakthroughs came in the 1950swith the discovery of the polyene macrolide antibiotic nystatin (Hazen and Brown, 1950) and the finding by Gentles (1958) and Williams et al. (1958) that griseofulvin, a fungal metabolite first described by Oxford et al. (1939), could be used to cure ringworm. Developments in antifungal therapy followed rapidly thereafter. Further polyene antibiotics, including amphotericin B (Gold et al., 1956), the only polyene used to treat patients with systemic infections, were isolated and characterized (Table 2). A significant advance came as a by-product of research into antineoplastic drugs. 5-Fluorocytosine was originally developed by Duschinski et al. (1957) as a potential antitumour agent, and its antifungal activity was reported by Grunberg et al. in 1964. This compound is well tolerated even when present at high concentrations in the serum. Unfortunately, its potential as an antimycotic drug is decreased by the narrow range of sensitive fungi and by the relatively high frequency with which drug-resistant yeasts arise. The most recent clinical developments have resulted from synthesis of imidazole derivatives. The first two, clotrimazole (Plempel etal., 1969, 1970) and miconazole (Van Cutsem and Thienpont, 1972), although effective against
4
DAVID KFXRIDGE
TABLE 2. Commonly used antifungal drugs
Date introduced or discovered
Drug
Target organism Dermatophytic fungi e.g. Epidermophyton spp., Microsporum spp., Trichophyton spp.
1939/1958 1950 1953 1956
Whiffield ointment Undecenoic acid Magenta paint Aluminium chloride Potassium permanganate Potassium iodide Sulphonamides Stilbamidine 2-Hydroxystilbamidine Griseofulvin Nystatin Candicidin Amphotericin B
1957
5-Fluorocytosine
1957 1969
Pimaricin Synthetic imidazole drugs Haloprogine Naftifine
Pre- 1950
1940 1948/1952
1972 1979
Sporothrix schenkii Paracoccidioides brasiliensis Blastomyces spp.
Dermatophytic fungi Candiah albicans Candida albicans Fungi causing systemic infections Candiah albicans Cryptococcus neoformans Candiah albicans Fungi causing both topical and systemic infections Dermatophytic fungi Dermatophytic fungi
topical infections, have not proved satisfactory for systemic infections. Ketoconazole, a phenyl-imidazole-piperazine derivative, is, however, water soluble and, since high serum concentrations can be obtained after oral administration, it has obvious potential in the treatment of patients with systemic mycoses (Symoens et al., 1980). The range of compounds inhibiting fungal growth and metabolism is wide (Ryley et al., 1981; Ellames, 1982). It is neither possible nor desirable to discuss the molecular basis of action of all antimycotic compounds reported in the literature, and this chapter will be restricted to clinically important drugs. It will not be possible to discuss in detail clinical studies, and the reader is referred to the excellent book edited by Speller (1980). It is of interest to note that, unlike antibacterial drugs where the principal targets are the ribosome (protein synthesis) and the cell wall (peptidoglycan synthesis), the major targets for clinically irhportant antimycotic drugs are the nucleus and cellular membranes. Inhibitors of fungal protein and cell-wall synthesis are not used clinically. The inhibitory effects of many of the drugs and their derivatives have been studied in organisms other than fungi. Where this is relevant to our understanding of the molecular basis of action, these studies are included.
MODE OF ACTION OF ANTIFUNGAL DRUGS
5
II. The Nucleus as the Primary Target Two clinically important compounds, griseofulvin and 5-fluorocytosine, act primarily at the level of the nucleus. Griseofulvin, a drug important in the chemotherapy of dermatophytic infections, interferes with nuclear organization. 5-Fluorocytosine, which is used in the chemotherapy of systemic candidosis and cryptococcosis, interferes with synthesis of nucleic acids. A. GRISEOFULVIN
Griseofulvin (Fig. 1) was first isolated from cultures of Penicillium griseofulvin and characterized by Oxford et al. (1939). In 1946, Brian et al. isolated a substance with antifungal activity from cultures of Penicillium junczewskii and, since it caused distortions in developing hyphae, was referred to by the authors as a “curling factor”. Subsequently the “curling factor” was identified as griseofulvin (Grove and McGowan, 1947). Griseofulvin is effective against mycelial fungi, but is without effect on bacteria and most yeasts. Initial interest in this compound stemmed from its potential as a systematic fungicide for treating infected plants (Brian, 1952), but this early promise has not been realized, and it is not used in agricultural or horticultural practice. The early studies on griseofulvin have been well reviewed by Bent and Moore (1966). However, the major impetus to studies on this compound came from the findings by Gentles (1958) that griseofulvin was effectivein curing guinea pigs experimentally infected with Microsporum canis and Trichophyton mentagrophytes, and of Williams et ul. (1958) that it cured ringworm in Man. A considerable number of analogues of griseofulvin have been synthesized and tested for their efficacy in inhibiting growth of both dermatophytic fungi
kH,
FIG. 1. Structural formulae of griseofulvin (a) and 2-ethoxy-3’-benzylgriseofulvin (b).
6
DAVID KERRIDGE
and plant pathogenic fungi (Crosse et al., 1964). Although one of them, a 2-ethoxy-3’-benzyl derivative (Fig. I), was found to be more effective than griseofulvin in inhibiting growth of dermatophytes in vitro, it has not been introduced into clinical practice. Research into the mode of action of griseofulvin initially followed two routes. The first line of research stemmed from its activity as a “curling factor”, and was into its effects on fungal cell-wall synthesis. The second arose from observations by Paget and Walpole (1958, 1960) that griseofulvin impaired nuclear division in both the rat and Viciafaba, with an apparent arrest of mitosis in metaphase, and was into its interaction with cellular microtubules. It is this latter approach which has proved successful in elucidating the mode of action of this drug.
I . Molecular Basis of Antifungal Action a. Effect on Nuclear Division. Initially research was concentrated on the effects of griseofulvin on the morphology of intact cells. The effects, although diverse, had one linking feature with cellular microtubules involved in all instances. In a Pectinaria species (a marine annelid), griseofulvin and vinblastin both reversibly disrupt the mitotic spindles in the oocytes (Malewista et al., 1968). Griseofulvin inhibits flagellar regeneration in Stentor spp. (Margulis et al., 1969), and the reversible darkening associated with dispersal of melanin granules in frog-skin melanocytes (Malewista, 1971). Gull and Trinci (1973) chose Basidiobolus ranarum to study the effects of griseofulvin on nuclear metabolism. This fungus is sensitive to griseofulvin at concentrations of 10p g d - ’ and, since it has a large nucleus (25 pm long) in each hyphal compartment, division abnormalities were readily observed on staining with acridine orange. Addition of griseofulvin resulted in a decrease in radial growth rate and appearance of more than one nucleus in each hyphal compartment. The radial growth rates of fungal colonies were inversely related to the logarithm of the drug concentration; the mitotic index and drug concentration were directly related. Vinblastin, but not colchicine, had similar effects. Inhibitors of the synthesis of fungal proteins and nucleic acids lowered the radial growth rate but, unlike griseofulvin, they also decreased the mitotic index (Gull and Trinci, 1974a, b). It appeared that griseofulvin does not affect synthesis of proteins or nucleic acids, but exerts its effect specifically during the mitotic cycle by affecting microtubular structure or function. Although it was obvious at this stage that nuclear microtubules were involved, it was not clear if griseofulvin affected their assembly or their function (Grisham et al., 1973). However, development of light-scattering methods to study self-assembly of tubulin into microtubules, and of
MODE OF ACTION OF ANTIFUNGAL DRUGS
7
immunofluorescencetechniques to examine microtubules both in vivo and in vitro led to the resolution of this problem. The direct effect of griseofulvin on expression of both cytoplasmic and spindle microtubules was demonstrated very elegantly by Weber et al. (1976) using fluorescence-labelledmonospecific tubulin antibody. In mouse 3T3 cells, griseofulvin ( 5 mM) destroyed cytoplasmic microtubules. Under these conditions there was no increase in the mitotic index, and the cells were arrested in interphase. When the concentration of griseofulvin was lowered to lopi, the cells were arrested at a point close to metaphase in a manner analogous to that induced by colchicine and a variety of other spindle poisons, and did not enter anaphase. Similar effects were found with HeLa cells, where, after 24 hours exposure to griseofulvin (20 pi), the mitotic index was 75%. The appearance of nuclei shown by fluorescence staining with antitubulin antibody after exposure to griseofulvin was dependent on the concentration used. After incubation in the presence of griseofulvin (10 p ~ ) there , were well-preserved metaphase-type spindles but, at 1 0 0 p ~spindles , were not formed. Brain tubulin will, under suitable environmental conditions, polymerize into microtubules at 37" C but not at 4" C. Griseofulvin inhibited in vitro polymerization at 37" C over a concentration range of 20-200 p~ (Roobol et al., 1976; Weber et al., 1976). Addition of griseofulvin to preparations of tubulin at 0°C resulted in an immediate aggregation of protein but microtubules were not detected (Roobol et al., 1977a). The lag that precedes tubulin polymerization was markedly increased by griseofulvin, but the polymerization of tubulin that occurs in the presence of glycerol (4 M) was not affected (Roobol et al., 1977a). Although the evidence is convincing that griseofulvin inhibits microtubule formation in a concentration-dependent manner, there is some debate as to whether this results from an interaction with microtubule-associated proteins (Roobol et al., 1977b) or with tubulin dimers (Wehland et al., 1977). Roobol et al. (1977b) found that the drugprotein binding ratios determined by a non-equilibrium procedure were some 50-fold lower than those determined by equilibrium procedures, and that griseofulvin was bound to a much greater extent by microtubular-associated proteins than by tubulin dimer. The authors suggest that the high values for binding ratios found using equilibrium procedures result from a relatively non-specific interaction of the insoluble antibiotic with hydrophobic regions on the tubulin dimer. There was also a correlation between the concentrations of griseofulvin required to inhibit microtubule formation and those in which there was a sharp increase in drug binding to microtubule-associated proteins. The authors interpreted these data to support the hypothesis that binding of griseofulvin to microtubule-associated protein is responsible for inhibition of tubulin assembly. Tubulin dimers assemble into microtubules in the absence of microtubule-
8
DAVID KBRRCDGE
associated proteins if the incubation buffer contains 4 M glycerol. Wehland et al. (1977) found that this process was inhibited by griseofulvin, but only at concentrations four times greater than those inhibiting assembly in the absence of glycerol. If, however, tubulin dimers were preincubated with griseofulvin before the addition of glycerol, then the concentration of drug required to inhibit assembly was comparable to that inhibiting assembly in the presence of microtubule-associated protein, thus providing evidence for a direct interaction of griseofulvin with tubulin. More recently, Sloboda et al. (1982) studied the interaction of brain tubulin and microtubule-associated protein with radioactively labelled griseofulvin. The bound and unbound drug were separated by exclusion chromatography and it was found that 0.83 k 0.08mol of griseofulvin was bound per mol of tubulin dimer. Griseofulvin was also associated with the microtubule-associated protein fraction, but this could be accounted for by the presence of contaminating tubulin dimer. The radioactivity here corresponded to 1.11 & 0.08 mol of griseofulvin per mol of contaminating dimer. It is unlikely that the presence of griseofulvin in this fraction resulted from a non-specific binding since griseofulvin was not bound to bovine serum albumin under identical conditions. Tubulin assembly is a two-stage process, with microtubule-associated protein involved in initiation but not in subsequent tubule extension. Addition of griseofulvin to an in vitro self-assembly system after initiation resulted in an immediate cessation of elongation. Finally, depolymerization of preformed microtubules was induced by addition of griseofulvin at 37”C, and this was associated with the appearance of protein aggregates similar to those formed on addition of griseofulvin to tubulin dimer at 4°C. The evidence supports the hypothesis that griseofulvin interacts directly with tubulin dimer rather than with microtubule-associated protein. Further evidence in favour of this hypothesis has come from an analysis of strains of Aspergillus nidulans with mutations in loci coding for tubulin subunits (Morris, 1980). It would appear that, in this organism, griseofulvin inhibits growth by affecting the interaction between the a- and fl-tubulins. There is need for further detailed genetical and physical studies to elucidate the molecular mechanism of griseofulvin-induced inhibition of microtubule assembly.
b. Inhibition of Fungal Cell- Wall Synthesis. The effects of griseofulvin on the cell-wall morphology of Botrytis allii were first observed by Brian et al. (1946). At concentrations of 0.1-0.2pg of griseofulvinml-’, growing hyphae developed a distinctive regular curl, a feature that led the authors to refer to this compound as a “curling factor”. At higher concentrations, the hyphae were grossly distorted and growth was extremely slow. Later, Brian (1960) suggested that these effects could result from griseofulvin inhibiting chitin
MODE OF ACTION OF ANTIFUNGAL DRUGS
9
synthesis. Subsequent studies, reviewed by Bent and Moore (1966), did not support this hypothesis and it is likely that the hyphal abnormalities are secondary effects resulting from an impairment of microtubular function within the cell. Microtubules are present in both fungal nuclei and cytoplasm (Beckett et al., 1974) and, although the role of cytoplasmic microtubules is not well understood, it is possible that impairment of their function would affect, either directly or indirectly, synthesis and deposition of cell-wall constituents at the hyphal tip, so resulting in growth abnormalities. The curling, distortion and irregular swelling induced by griseofulvin are so marked and distinctive that they must be regarded as specific secondary effects of griseofulvin rather than a non-specific effect resulting from an inhibition of growth. c. Selectivity ofdction. The possibility that microtubules may not be the sole target responsible for growth inhibition by griseofulvin was examined by Mir et al. (1978), who synthesized derivatives of griseofulvin with different biological activities (Fig. 2). These compounds were monitored for in vivo activity by studying their effects on Physarum polycephalum and a mouse leukaemic cell line L1210, and for their in vitro activity by studying the effects on tubulin aggregation at low temperature, and inhibition of tubulin assembly at 37”C. In all cases, there was a good correlation between in vivo and in vitro effects, and the authors concluded that microtubules are the primary target responsible for the inhibitory effects of griseofulvin.
R -WH, -WH,CH,I -NH2 -OH
Derivative griseofulvin
Z’-(iodoethoxy)-griseofulvin 2’-aminogriseofulvin griseofulvic acid (enol form)
CH, lsogriseofulvin
FIG. 2. Structural formulae of certain derivatives of griseofulvin.
10
DAVID KERRIDGE
Griseofulvin interacts with tubulin from a variety of tissues and cells including sheep brain, marine annelids and myxomycetes, and it is unlikely that selectivity results from a failure to bind to specific tubulin dimers. As might be expected, bacteria, all of which lack tubulin, are insensitive, but what factors affect the variability in sensitivity among the fungi? Why, for example, is the opportunistic pathogen C. albicans resistant? El Nakeeb and Lampen (1965a, b) studied griseofulvin uptake by sensitive dermatophytic fungi, a resistant strain of Epidermophyton jloccosum, and the insensitive C. albicans and Escherichia coli. In both sensitive and resistant organisms there was an immediate small binding of the drug, independent of cultural conditions and cell viability, with values ranging from 0.04 to 0.57 pg (mg dry wt. cells)-'. In sensitive, but not resistant, organisms this was followed by a prolonged uptake extending over 24 to 48hours. This uptake was temperature dependent, required an exogenous energy source and was inhibited by dinitrophenol and sodium azide. Addition of p-fluorophenylalanine prevented uptake of griseofulvin by Trichophyton sp. and Microsporum sp., suggesting that de novo protein synthesis is required. In resistant, but not sensitive, strains all of the griseofulvin was water-extractable. In sensitive strains, some 50% of the bound griseofulvin was extractable with hot trichloroacetic acid and hot NaOH, which would suggest that griseofulvin is associated with fractions of protein and nucleic acid. Addition of purine nucleotides to the growth medium partially protected certain strains of dermatophytic fungi against the inhibitory effects of griseofulvin, but it was not possible to distinguish between an effect on uptake of griseofulvin and its interaction with cellular microtubules. Clearly a specific transport system is involved in mediating its growth inhibitory effects on sensitive cells, and resistance could result from the absence of such a system, but nothing is known of either its mechanism or specificity. It is reasonable to assume that sensitive fungi do not possess a specific transport system for this antifungal drug, and that griseofulvin, like 5-fluorocytosine, is transported illicitly into sensitive cells by a pre-existing transport system. 2. Clinical Usage
Griseofulvin is effective in curing infections caused by species of certain dermatophytic fungi, Epidermophyton, Trichophyton and Micromonosporum, that cannot be resolved by topical therapy with other antifungal drugs. It is administered orally and is usually well tolerated. The recommended dose of 1-2 g each day can be continued for up to 18 months in treating patients with onychomycosiswithout serious adverse effects (Davies, 1980; Roberts, 1980). After oral administration, most of the administered drug can be recovered from urine and faeces over the following 5 days (Lin et al., 1973). Griseofulvin
MODE OF ACTION OF ANTIFWNGAL DRUGS
11
appears rapidly in the outer layers of the stratum corneum and, depending upon the climatic conditions, values of from 5 to 45 pg of griseofulvin (mg of skin)-' may be reached in the horny layer of the skin. This association with keratinous layers of the skin is important because the drug is delivered to the site of infection, and can therefore be taken up readily by the dermatophytic fungus. Degradation of keratin by extracellular keratinases is inhibited by association of griseofulvin with the substrate, and as a result the availability of nutrients for fungal growth is diminished, so effectively lowering the growth rate. This will affect hyphal penetration into the skin and so enhance its shedding during skin growth. This could be an important factor in the treatment of patients, but is of little relevance to the molecular basis of griseofulvin action in vitro (Yu and Blank, 1973). B. 5-FLUOROCYTOSINE
5-Fluorocytosine (Fig. 3) was synthesized by Duschinski et al. (1957) as a potential anticancer drug, and, although it subsequently proved to be ineffective (Heidelberger et al., 1958), it is now used as an oral antifungal drug (Scholer, 1980). It is active against relatively few medically important fungi and, of these, the most important are C. albicans and Cryptococcus neoformans; Histoplasma capsulatum, Coccidioides immitis and the dermatophytic fungi are insensitive (Scholer, 1970). 5-Fluorocytosine has both fungistatic and fungicidal activity against C. albicans and Cryptococcus neoformans, although the latter effect requires higher drug concentrations and a more prolonged exposure (Shadomy et al., 1969, 1973; Scholer, 1970). Scholer (1974) was unable to demonstrate any fungicidal activity against Aspergillus fumigatus although a fungistatic effect was observed. The drug is usually administered orally and is rapidly absorbed giving high concentrations in serum which are well tolerated with a record dosage of 10.7kg being given over a period of 3years to a patient suffering from cryptococcosis (Zylstra, 1974). Unfortunately, its value as a chemotherapeutic agent is markedly decreased by the frequency with which resistant strains arise during therapy.
H
FIG. 3. Structural formula of 5-fluorocytosine.
12
DAWJJ KERRIDGE
1. Molecular Basis of Antijiungal Action a. Inhibition of Nucleic Acid Synthesis. The molecular basis of action of 5-fluorocytosine is well understood (Polak and Scholer, 1980) and is summarized in Fig. 4. 5-Fluorocytosine is itself non-toxic, but it is metabolized by sensitive cells to give a number of derivatives that exert a growth inhibitory effect, either as a result of their incorporation into cellular macromolecules or by an indirect inhibition of macromolecular synthesis. Much of the detailed understanding of the mode of action of this compound has been derived from studies on animal tissues, and, where relevant, this information will be used to supplement the experimental data derived from fungi. Geige and Weil (1970) and Jund and Lacroute (1970) were the first to demonstrate that, after uptake into sensitive cells, 5-fluorocytosine is deaminated to 5-fluorouracil. 5-Fluorouracil is converted into the riboside triphosphate derivative and incorporated into ribonucleic acids as 5-fluorouridylate. 5-Fluorocytosine is itself not found in cellular nucleic acids. The enzymes responsible for these interconversions are all present within sensitive cells. Transport of 5-fluorocytosine into both Saccharomyces cerevisiae and C . albicuns is by a cytosine permease normally responsible for the uptake of adenine, guanine and hypoxanthine in addition to cytosine (Polak and Grenson, 1973). Uptake of these nitrogenous bases into yeasts is an energy-dependent process linked to a proton pump, with a stoicheiometry of one proton for each substrate molecule transported. The relative affinity of individual substrates for the cytosine permease can be correlated with their capacity to be protonated to the positively charged form (Foret et al., 1978). Two ionizable groups are involved in substrate binding, one (pK, 1.8) is associated with the substrate and the second (pK, 5.1) is associated with an amino-acid carboxyl group at the recognition site on the permease. Transport of a base into the cell involves production of an uncharged terniary complex comprising the carboxyl group on the permease, a proton and the substrate molecule. Uptake of a specific substrate is competively inhibited by each of the others, and the apparent K,,, values are equal to the values of Ki when used as competitive inhibitors. These values range from 6 p for ~ adenine to 60PM for 5-fluorocytosine(Polak and Grenson, 1973). Once inside the cell, the drug is deaminated by a cytosine deaminase to 5-fluorouracil. Cytosine deaminase is the essential enzyme in determining the antimicrobial spectrum of 5-fluorocytosine, and the low toxicity of the drug in the human host can be correlated with either the absence or very low levels of this enzyme in mammalian tissues. 5-Fluorouracil is then converted into 5-fluorouridylic acid, a reaction catalysed by uridine monophosphate pyrophosphorylase, an enzyme normally responsible for recycling any uracil
5-Fluorocytosine
Cytosine
pmneasc
//
Deoxyuridylate
5-Fluorocytosine
Cytosine deaminw
5-Fluorodeoxyuridylate c-
5-Fluorouracil
Uridylrtc c 5-Fluorouridydte pyrophorphorylrlc
I
S-Fluorodeoxyuridine c-- 5-Fluorouridine diphosphate diphosphate
Thymidylste synth.rDeoxythymidylate
5-Fluorouridine triphosphdte
Abnormal RNA (Aberrant protein)
FIG. 4. Metabolism of 5-fluorocytosine in yeast.
14
DAVID KERRIDGE
formed as a result of cellular catabolism. 5-Fluorouridylate then undergoes all the reactions normally associated with uridylate within the cell. The extent to which uracil is replaced by 5-fluorouracil in cellular RNA of C. albicans is considerable, up to 50% replacement can occur in both rRNA and tRNA. There was a good correlation between incorporation of radioactivity from 5-fluor0[2-’~C]cytosine into RNA in a number of strains of C. albicans and their sensitivity to the drug. The sensitive strains incorporated up to 76ng lo6 cells-’ whereas resistant strains incorporated only 2ng lo6 cells-’ (Polak and Scholer, 1975). More extensive studies on the effects of 5-fluorouracil and its derivatives on RNA synthesis have been performed with tissue-culture cells. In these cells not only is the rate of RNA synthesis decreased, but also post-transcriptional processing is affected by incorporation of 5-fluorouridylate into RNA (Tseng et al., 1978; Wilkinson et al., 1975; Glazer and Hartman, 1980). For example 5-fluorouridine at a concentration of 10 p,i not only decreased the synthesis of 45s RNA by Novikoff hepatoma cells, but completely inhibited synthesis of mature 18s and 23s rRNA. Similar findings have not been reported in C. albicans or other fungi, but it is probable that such effects occur. In bacteria, but not so far in yeast, there is evidence for synthesis of abnormal proteins on incubation in the presence of 5-fluorouracil (Mandel, 1969). Similar effects may occur in fungi since tRNA isolated from Sacch. cerevisiae after incubation in the presence of 5-fluorocytosine, although still capable of amino acylation in vitro, showed differences in labelling pattern when compared to tRNA isolated from control cells (Geige and Weil, 1970). Polak (1974) also found that incubation of Sacch. cerevisiae with the drug resulted in changes in the concentrations of amino acids in the soluble fraction and evidence for disturbance of protein synthesis. In 1957,Heidelberger et al. predicted that 5-fluorouracil would not only be incorporated into cellular RNA but also would inhibit DNA synthesis. This prediction was soon confirmed with the findings that 5-fluorouracil and its derivatives 5-fluorouridineand 5-fluoro-2’-deoxyuridine,inhibited incorporation of I4C from [14C]formate(a precursor of the methyl group of thymine) into DNA of tumour cells (Danenberg et al., 1958) and that 5-fluoro-2deoxyuridine induced thymine-less death in E. coli (Cohen et al., 1958). This inhibition of DNA synthesis results from a inhibition of thymidylate synthase by 5-fluoro-2‘-deoxyuridylate. This enzyme is responsible for de novo synthesis of thymidine within the cell and is a key enzyme in DNA synthesis. As a result, it has received considerable attention as a potential target for anticancer drugs and much of our knowledge of the mode of action of 5-fluorocytosine has been derived indirectly from such studies (Danenberg, 1977). A number of therapeutically active and widely used antimetabolites inhibit thymidylate synthase, either directly or indirectly, by inhibiting
MODE OF ACTION OF ANTIFUNGAL DRUGS
15
dihydrofolate reductase, but unfortunately the antitumour activity of such drugs is also accompanied by damage to tissues that normally proliferate rapidly. 5-Fluoro-2’-deoxyuridylic acid has played a major role in elucidating the mechanism of thymidylate synthase (reviewed by Danenberg, 1977), yet it was not until 1977 that Polak and Wain reported that addition of 5-fluorocytosine to either the yeast or mycelial form of C. albicans resulted in an immediate cessation of DNA synthesis and, in the following year, it was established that this drug is converted into 5-fluoro-2’-deoxyuridylateby C . albicans with a concomitant decrease in thymidylate synthase activity in vivo (Diasio et al., 1978). Subsequently, Wagner and Shadomy (1979) demonstrated a similar conversion in A. fmigatus. The mechanism of inhibition of thymidylate synthase by 5-fluoro-2’deoxyuridylate has been studied using purified enzymes isolated from Streptococcus faecalis, phage-infected E. coli and methotrexate-resistant Lactobacillus casei. 5-Fluoro-2’-deoxyuridylateinteracts reversibly with thymidylate synthase in the absence of 5,lO-methylene tetrahydrofolate. In the presence of this cofactor, the drug was covalently bound and the complex stable to treatment with either urea or guanidine hydrochloride; it was not dissociated on precipitation with trichloroacetic acid or by sodium dodecyl sulphate-polyacrylamide gel electrophoresis (Santi and McHenry, 1972; Langenbach et al., 1972). The inhibition results from formation of a stable terniary complex between the inhibitor, 5,lO-methylene tetrahydrofolate, and thymidylate synthase, in which the 6 position of the analogue is bound covalently to the nucleophilic catalyst of the enzyme via a thioether bond and the 5 position is linked to the one carbon of the 5,lO-methylene tetrahydrofolate (Fig. 5). This complex is analogous to the steady-state intermediate of an enzyme reaction and its formation is slowly reversible (Danenberg and Danenberg, 1978). Considerable conformational changes
OH OH
FIG. 5. The inhibitory complex at the active site of thymidylate synthase.
16
DAVID KERRIDGE
occur in the enzyme molecule as a result of complex formation; it becomes more compact with a 3.5% decrease in the Stokes radius and an increase in sedimentation coefficient. Although the enzyme is dimeric and there is evidence for two binding sites for 5-fluoro-2’-deoxyuridylate,70% of the conformational changes occur upon the binding of 1 mol of ligand to 1 mol of protein. 5-Fluoro-2’-deoxyuridine has been detected in the DNA of eukaryotic cells, but not as yet in yeasts or fungi, although by analogy one might expect small amounts to be present (Kufe et ul., 1983). These incorporated residues are excised from DNA and may contribute to the cytotoxicity of fluorinated pyrimidines. These authors were able to dissociate the effects of analogue incorporation into DNA from its incorporation into RNA and the effect on thymidylate synthase, and have provided evidence that misincorporation of 5-fluoro-2’-deoxyuridineinto DNA does, in itself, result in lethal cellular events in L1210 cells. So there is a situation where a drug, although ineffective itself, is metabolized by sensitive fungi to give metabolic intermediates, one of which is incorporated into cellular RNA, whereas the other inhibits a key enzyme in synthesis of an intermediate of DNA synthesis. 5-Fluorocytosine is not itself incorporated into cellular nucleic acids, and fluorinated pyrimidines have not been detected in yeast DNA. Obviously the fungistatic and/or fungicidal action of 5-fluorocytosinecould result from either effect. The first indication that there might be differences in the relative importance of the inhibitory effects on RNA and DNA synthesis in mediating the growth inhibitory effects came from studies on a resistant isolate of Cryptococcus neoformans, where it was found that incorporation of radioactivity from 5-fl~oro[‘~C]cytosine into the RNA fraction of the resistant isolate was similar to that into the parental strain. This was interpreted by Diasio et ul. (1978) as evidence for a resistance mechanism involving thymidylate synthase and that, for this strain at least, inhibition of DNA synthesis is more important than production of aberrant RNA in mediating the inhibitory action of the drug. A more detailed assessment of the relative importance of the effects on RNA and DNA synthesis in mediating the action of 5-fluorocytosinehas been reported by Waldorfand Polak (1983). Seventy five resistant isolates of C. albicunswere analysed and, in 75% of the strains, there was a positive correlation between their susceptibility to the drug, the incorporation of radioactivity from 5-fluorocytosineinto RNA and inhibition of macromolecular synthesis. In these strains, it was not possible to distinguish between the effects on RNA and DNA synthesis in their contribution to growth inhibition. In the remaining strains, the correlation was not so marked, with either RNA or DNA synthesis continuing at a rate comparable to the control, while synthesis of the other nucleic acid was inhibited. These
MODE OF ACTION OF ANTIFUNGAL DRUGS
17
data would suggest that the two mechanisms are not necessarily linked, and either can be responsible for growth inhibition. b. The Morphological E#ects of 5-Fluorocytosine on Fungi. Addition of the drug at the minimum growth inhibitory concentration to C. albicans does not result in an abrupt cessation of growth (Wain and Polak, 1979). There was an increase in the size of the yeast cells and continued hyphal extension in the mycelial form; this was associated with continued synthesis of RNA, protein and carbohydrate, but not DNA. In A. fumigatus, both conidial germination and hyphal extension were suppressed on addition of the drug (Wain et al., 1981). Ultrastructural changes in both nucleus and cell wall of C. albicans occurred after exposure to 5-fluorocytosine (Arai et al., 1977). After incubation for 2 hours, the nucleus was found to be enlarged, but there were no other significant morphological effects. Continued incubation for a further 12hours resulted in further enlargement of the nucleus, which became translucent with filamentous components appearing within it. The cell wall became progressively thinner, probably as a result of an increase in cell volume without a concomitant increase in cell-wall constituents. These morphological changes are exactly analogous to those observed by Cohen et al. (1958) in their studies of inhibition of DNA synthesis in both bacteria and mammalian cells, and provide further evidence in support of the hypothesis that thymidylate synthase is a primary target for 5-fluorocytosine.
c. Resistance to 5-Fluorocytosine. It is most unfortunate that clinical use of this well-tolerated drug is limited by the occurrence of resistant strains of C. albicans. Primary resistance, i.e. that characterized by growth of the majority of the cells in a population when exposed to the drug, occurs in some 8-10% of strains examined (Scholer, 1980) even though the strains had not been previously exposed to the drug and showed no cross resistance to other antifungal drugs (Schonebeck, 1973). Drouhet et al. (1974) made the very interesting observation that the frequency of isolates of C. albicans resistant to 5-fluorocytosine (25 pgml-') was much greater among strains of serotype B than serotype A. Out of a total of 455 strains examined, 5.7% were resistant to 5-fluorocytosine, of these, three resistant strains were type A out of a total of 429 type A strains, whereas 23 were type B out of a total of 27 type B strains. This imbalance between the relative frequency of drug-resistant isolates in the two serotypes has been confirmed by Auger et al. (1979), who found that, although the majority of strains they examined were serotype A, a higher proportion of resistant strains (approximately 50%) were found among the serotype B isolates. Similar results were obtained by Stiller et al. (1982) for strains of C. albicans isolated in the U.S.A. The characteristic resistance pattern for the type B serotypes is that
18
DAVID KERRIDGE
they are resistant to both 5-fluorocytosineand 5-fluorouracil but are sensitive to 5-fluorouridine, suggesting that resistance could be a result of a relative deficiency in the enzyme uridine monophosphate :pyrophosphorylase (Polak and Scholer, 1975). However, this hypothesis has not been confirmed. The efficacy of 5-fluorocytosine as an antifungal agent results from its transport into the cell and interconversion into metabolic derivatives which then interefere with synthesis of nucleic acids. Resistance to the drug can result from a loss of any of the enzymes involved in these interconversions and hence the relative frequency of resistant mutants may be high. Jund and Lacroute (1970, 1974) studied resistance mechanisms in Succh. cerevisiue and distinguished six types of resistant mutants. The evidence for these was based largely on resistance profiles to 5-fluorocytosine, 5-fluorouracil and its derivatives, and on the antagonistic effects of purines and pyrimidines. A genetic analysis was undertaken which gave the number of complementation groups involved and hence a much better assessment of the possible steps involved. The mechanisms proposed for drug resistance in Succh. cerevisiae are: (a) a deficiency in the cytosine permease (locusfcy2); (b) a deficiency in cytosine deaminase (locus fcy Z); (c) a deficiency in uridine monophosphate pyrophosphorylase (locusfurl); (d) a loss of feedback regulation of aspartate carbamoyltransferase by uridine triphosphate giving an increased de now synthesis of pyrimidines; (e) an increased de novo synthesis of pyrimidines resulting from a stimulation of orotidylate pyrophosphorylase and orotidylate decarboxylase. Where resistance results from a deficiency in either the cytosine permease or deaminase, the organisms are resistant only to the drug. In all other cases, the organisms are also resistant to 5-fluorouracil and 5-fluorouridine, except for certain strains that lack uridine monophosphate pyrophosphorylase. Resistant strains lacking this enzyme are heterogeneous and include strains both sensitive and resistant to 5-fluorouridine. In clinical isolates, the most frequently occurring resistant strains are those lacking uridine monophosphate pyrophosphorylase, strains lacking either the permease or the deaminase are found infrequently (Polak and Scholer, 1975; Kemdge and Whelan, 1984). Genetic analysis of drug resistance in C.ulbicans is hampered by the lack of a mating system. There is good evidence based on DNA content and the frequency of mutations that the organism is diploid (Okaiya and Sogin, 1979; Whelan et ul., 1980; Whelan and Magee, 1981), and it has recently been possible to demonstrate heterozygocity for both auxotrophic markers (Whelan et ul., 1980)and resistance to 5-fluorocytosine(Whelan et ul., 1981). Stiller et ul. (1982) studied factors affecting the laboratory assessment of 5-fluorocytosineresistance in C. ulbicans and reported the occurrence of three classes of strains: (a) sensitive, (b) intermediate in resistance and (c) highly
MODE OF ACTION OF ANTIFUNGAL DRUGS
19
resistant. The existence of the strains intermediate in resistance was originally reported by Normark and Schonebeck (1972) and later by Defever et al. (1982) and Whelan et al. (1981). In the latter study, it was found that three partially resistant strains were heterozygous for resistance and could give rise to both sensitive and highly resistant variants. The partially resistant strains constitute a significant fraction (3540%) of a random sample of strains in the U.S.A. (Stiller etal., 1982; Defever et al., 1982). The most common enzyme defect associated with resistance is a deficiency in the activity of uridine monophosphate pyrophosphorylase (Kerridge and Whelan, 1984) with resistant isolates possessing markedly less activity than the sensitive or partially resistant isolates. The partially resistant isolates typically possess less uridine monophosphate pyrophosphorylase than the sensitive isolates, which suggests that these strains are heterozygous at a gene that determines this activity. It is possible that heterozygosity at this locus constitutes the main obstacle to successful therapy with 5-fluorocytosine. The fact that a number of strains of C. albicans are heterozygous for drug resistance may account for the high frequency of appearance of isolates resistant to 5-fluorocytosine during therapy with this drug. It is not surprising therefore that resistant mutants can pose a serious problem when 5-fluorocytosine is used in the therapy of clinical infections. There is one further factor which may be important in the selection of resistant mutants. Peterson et al. (1983) observed that, in Chinese hamster cells, addition of 5-fluoro-2’-deoxyuridineto cultures growing in the presence of 10,UMdeoxycytidine resulted in a 6- to 90-fold increase in the Occurrence of mutants resistant to 8-azaguanineYGthioguanine and ouabain over the spontaneous mutation rate. Neither 5-fluoro-2’-deoxyuridine nor deoxycytidine alone had this effect when added to the culture. An imbalance in the concentrations of deoxyribonucleotides within the soluble pool fraction is mutagenic in cultures of mammalian cells (Meuth et al., 1979; Weinberg et al., 1981) and induction of mutations in cultured animal cells by 5-fluoro-2’-deoxyuridinein the presence of deoxycytidine is consistent with this. There is no evidence that a similar phenomenon occurs in C. albicans when incubated in the presence of 5-fluorocytosineYbut the possibility that a therapeutic agent may enhance the frequency with which mutants resistant to it may occur is somewhat alarming.
III. The Cell Membrane as the Primary Target The two most important groups of antimycotic drugs, namely polyene macrolide antibiotics and certain synthetic imidazole derivatives, have as their target the plasma membrane. Polyene antibiotics affect the structural
20
DAVID KERRIDGE
integrity of the membrane. Imidazole drugs inhibit synthesis of membrane constituents at fungistatic concentrations and interfere with the structural integrity of the membrane at fungicidal concentrations. It is perhaps surprising that, given the apparent similarity of plasma-membrane structures and function among eukaryotic organisms, this organelle should provide the most significant target for clinically important antimycotic drugs. A. POLYENE MACROLIDE ANTIBIOTICS
Polyene macrolide antibiotics comprise some 200 compounds produced usually by Streptomyces spp. (Hamilton-Miller, 1973; Ryley et al., 1981). Of these, only a few are sufficiently non-toxic to be used clinically and only one, amphotericin B, is used to treat patients with systemic fungal infections (Medoff and Kobayashi, 1980). Chemically, this group of compounds is characterized by a ring of carbon atoms closed by lactonization and containing both a system of conjugated double bonds and a hydrophilic region comprising a number of hydroxyl groups. The antibiotics in this group differ in the number of carbon atoms and double bonds in the ring; the number of hydroxyl groups, the presence or absence of a glycosidically linked carbohydrate moiety and carboxyl, aliphatic or aromatic groups attached to the ring in specific positions (Hamilton-Miller, 1973; Kobayashi and Medoff, 1977; Gale et al., 1981; Ryley et al., 1981). The structural formulae of a number of clinically important polyene macrolide antibiotics are shown in Fig. 6. The complete three-dimensional structure of amphotericin B has been elucidated from crystallographic data by Mechlinski et al. (1970). It is a rigid rodshaped molecule with opposing hydrophobic and hydrophilic faces (Fig. 7). The amino-sugar residue is present as a iarge hydrophilic group at one end of the rod, and there is a single hydroxyl group at the other end. The overall length of the rod is approximately 2.1 nm and is similar to that of a plasmamembrane phospholipid molecule, a factor of some significance in its mode of action. 1. Molecular Basis of Antifungal Action
a. Impairment of Membrane Function. Polyene macrolide antibiotics interact with plasma membranes of sensitive organisms causing an impairment of barrier function, leakage of cellular constituents and ultimately cell death. The specificity of the plasma membrane-drug interaction and extent of the resultant damage depends on both the antibiotic (in particular the nature of the charged groups and the presence of hydrophilic groups) and the lipid composition of the membrane (Norman et al,, 1976). At one extreme, filipin
MODE OF ACTION OF ANTIFUNGAL DRUGS
21
HJC
g:H
HO
NH,
Pimaricin
FIG. 6. Structural formulae of certain clinically important polyene macrolide antibiotics.
(a pentaene) causes gross disruption of plasma membranes of sensitive cells, releasing both low-molecular-weight material and small proteins (Lampen, 1966; Norman et al., 1972); at the other extreme, there are two compounds, N-succinyl perimycin (a heptaene) which induces release of K + ions only (Borowski and Cybulska, 1967) and primaricin (a tetraene) which has no effect on the permeability properties of either the plasma membrane of Acholeplasma laidlawii or egg lecithin liposomes (De Kruijff et al., 1974). However, pimaricin was subsequently shown to induce release of K+ ions from Sacch. cerevisiue (Kolter-Brajtburg et al., 1979). The inhibitory effect of polyene macrolide antibiotics is specific for eukaryotic organisms and those prokaryotic organisms, such as Acholeplasma and Mycoplasma species, containing sterols in their plasma membranes (Lampen, 1966). The sterol com-
22
DAVID KERRIDGE
FIG. 7. Space-filling model of amphotericin B.
position of the plasma membrane is important in determining sensitivity to these antibiotics, and it is the difference in relative affinities of ergosterolcontaining fungal membranes and cholesterol-containing mammalian membranes for clinically important polyene antibiotics that makes it possibIe to use these compounds to treat patients with mycotic infections (Archer and Gale, 1975). One of the earliest detectable effects of the interaction of a polyene macrolide antibiotic with sensitive cells is leakage of K+ ions. In 1966, Lampen postulated that the primary effect of nystatin and arnphotericin B is to render the plasma membrane specifically permeable to K+ ions, and that cell death results from the associated proton uptake and consequent acidification of the cell contents. This hypothesis has been modified by Palacios and Serrano (1978) as a result of their studies on inhibition of glucose and maltose fermentation in Succh. cerevisiue by nystatin and amphotericin B. Maltose fermentation was far more sensitive to inhibition by nystatin than glucose fermentation and inhibition was not reversed by addition of K + ions (Table 3). Maltose transport by Succh. cerevisiue is an active process, energized by a proton gradient (Serrano, 1977) and, unlike glucose transport, was inhibited by amphotericin B and nystatin at low concentrations. The authors proposed that the primary effect of these antibiotics is to render the cell membrane permeable to protons. Further support for this hypothesis came from direct measurements of the effect of these antibiotics on proton movement in cells depleted of their internal ATP by addition of antimycin A
MODE OF ACTION OF ANTIFUNGAL DRUGS
23
TABLE 3. Inhibition by polyene macrolide antibiotics of glucose and maltose fermentation in Succhuromyces cerevisiue. From Palacios and Serrano (1978) Fermentation rate (nmol of C0,min-'(mgdrywt.)-') Antibiotic added
Concentration (pg(mg dry wt.)-')
None Amphotericin B Nystatin Nystatin Nystatin
0.54 0.05 0.13 0.54
Glucose -KCl +KCl 53 3 44 32 3
53 30 48 48 38
Maltose -KCl +KCl
44 2 9 8 2
43 2 9 6 2
and 2-deoxyglucose. The proton gradient has a central role in functioning of the plasma membrane. It energizes uptake of amino acids and other nutrients (Foury and Goffeau, 1975) and maintenance of the internal pool of K+ ions (Pena, 1975). Rapid release of K + ions induced by addition of polyene macrolide antibiotics to sensitive cells, and used by a number of workers to monitor interaction of these compounds with the plasma membrane (Gale, 1974; Hammond and Kliger, 1974), can now be considered as a secondary effect resulting from a drug-induced increase in proton permeability of the plasma membrane. The growth inhibitory effects of polyene macrolide antibiotics are strongly modulated by the environment. Gottleib et al. (1958) first reported that addition of sterols to the growth medium protects fungi from the inhibitory action of filipin. This protection results from an in vitro interaction between drug and sterol, with a consequent lowering of the effective polyene concentration (Lampen et al., 1960). The effectiveness of different sterols in reversing the action varies from polyene to polyene; for example, ergosterol is more effective that cholesterol in antagonizing the inhibitory effects of amphotericin B on C. albicans, and the converse is true for filipin (Archer and Gale, 1975). Other hydrophobic compounds, for example fatty acids (Ianitelli and Ikawa, 1980), also protect sensitive organisms against polyene action, presumably in a similar manner. Addition of K+and M$+ ions at the relatively high concentrations of 85 mM and 45 m ~ respectively, , to cultures of Sacch. cerevisiae and C . albicans protects the organisms against the inhibitory effects of candicidin and amphotericin B methyl ester (Liras and Lampen, 1974; Kerridge et al., 1976). In this case, addition of protecting ions does not prevent the antibiotic from interacting with the membrane, but apparently maintains the internal concentration of these ions necessary for survival. With C . albicans, protection occurred only when the antibiotic was
24
DAVID KERRIDGE
present at the minimum growth inhibitory concentration; at higher concentrations the cells were killed, suggesting that, under these conditions, there is considerable disruption of the plasma membrane. The importance of the loss of internal pool of K+ ions in mediating the fungicidal effects of polyene macrolide antibiotics has been questioned. Polyene-resistant mutants, although not a clinical problem, can be readily obtained in the laboratory. In general, they have an altered membrane lipid composition, and have been useful both in the study of sterol biosynthesis (Pierce et al., 1978) and as genetic markers in following protoplast fusion in C. albicans (Pesti and Ferenczy, 1982). In one class of polyene-resistant mutant of C. albicans, release of K+ is apparently as sensitive to polyene antibiotics as the parental strain (Hsu-Chen and Feingold, 1974). Chen et al. (1978) analysed the kinetics of nystatin- and amphotericin B-induced leakage of Kf ions from sensitive strains of C. albicans, but were unable to correlate loss of K+ ions with loss of viability induced by the antibiotic. KolterBrajtburg et al. (1979) extended these studies to other polyene macrolide antibiotics and classified them into two groups according to their chemical structure and biological properties. In the first group, which includes pimaricin and filipin, concentrations required for release of K+ ions and cell death were identical, and it was not possible to separate a fungicidal from a fungistatic effect. The polyenes in the second group, which includes amphotericin B, cause considerable release of K+ ions from Sacch. cerevisiae at concentrations much lower than those required to kill cells. For antibiotics in this group, it is possible to distinguish between fungistatic and fungicidal effects. However, the dissociation of K+ leakage from the fungicidal activity of these compounds has been questioned by Malewicz et al. (1981). Interaction of the polyene with the plasma membrane is reversible, and it would be possible for cells on transfer to a drug-free medium to regain their membrane functions, even though they may have lost a considerable proportion of their intracellular potassium. If this is so, then dissociation of loss of K + ions from cell death could reflect the techniques used to measure them, and the differences between the two groups of polyene macrolide antibiotics reflect the relative ease with which the polyene-membrane interaction can be reversed. b. Molecular Models. Molecular interactions of a number of polyene macrolide antibiotics with lipid bilayers have been studied both in vivo using sensitive cells and in vitro using liposomal vesicles and black lipid films (reviewed by Gale et al., 1981 and Medoff et al., 1983). As a result of such studies, molecular models have been proposed for interaction of the antibiotic with membrane lipid and to explain the resultant change in membrane permeability (Andreoli, 1974; De Kruijff and Demel, 1974; Marty and Finkelstein, 1975; Van Hoogevest and De Kruijff, 1978). Essentially these
MODE OF ACTION OF ANTIFUNGAL DRUGS
25
FIG. 8. Molecular model for the polyene-plasma membrane interaction. From De Kruijff and Demel(l974).Reproduced with the permission of Elsevier Biomedical Press. models are similar. The hydrophobic face of the rigid polyene molecule first interacts with either a sterol molecule or an acyl side chain of a phospholipid molecule present in the membrane. These complexes then aggregate to produce an annulus comprising eight polyene molecules in which the hydrophilic face of the polyene is directed inwards producing an aqueous pore of internal diameter 0.8nm. The models are of two types. The first, proposed by van Deenen’s group, requires two opposing annuli to span the lipid bilayer (De Kruijff and Demel, 1974). In this model (Fig. 8), the mycosamine moiety is positioned at the lipid-water interface, and the solitary hydroxyl group at the other end of the polyene is embedded within the lipid bilayer. Although it is reasonable to predict that such pores may occur in artificial membranes when the antibiotic is added simultaneously to both sides of the membrane, there are obvious problems in translocating polyene molecules across a membrane when the antibiotic is added to one side only. It was suggested that a polyene-induced distortion in the membrane could aid its translocation (De Kruijff and Demel, 1974), but Aracava et al. (1981) were unable to detect any movement of amphotericin B across the membranes of multilamellar vesicles; the polyene interacted only with the outer layer. A second modified model was proposed by Marty and Finkelstein (1975) in which either a single or a double annulus could span the bilayer with, in each case, a distortion of the membrane in the vicinity of the pore (Fig. 9). Further support for a single annulus model came from studies on the effect of
26
DAVID KERRlDGE
--Polar
''70
2,znrn
-
head
-Lactone
rinq
Terminal OH group
Nvstatin malewls
-
Polar head Hydmcaftnm fails
Phcaphdipid molecule
FIG. 9. Molecular model for the polyene-plasma membrane interaction. From Marty and Finkelstein (1975). Reproduced with the permission of the Rockefeller
University Press. membrane thickness on amphotericin B-induced membrane permeability (Van Hoogevest and De Kruijff, 1978). Addition of amphotericin B to one side of egg phosphatidylcholine (containing mainly oleyl residues) vesicles induced K leakage, but for vesicles prepared from didocosanoyl phosphatidylcholineaddition of antibiotic to both sides of the membrane was necessary for leakage to occur. Addition to one side only had no effect on membrane permeability. The length of the amphotericin B molecule (2.1 nm) is somewhat shorter than the hydrophobic core of the bilayer formed from egg phosphatidylcholine(3.5 nm) and the authors proposed that flexibility of the acyl side chains allowed meniscus formation at either end of the pore, and that such a pore might shuttle up and down in the bilayer providing some mobile characteristics for the single annulus. The hydrophobic core in lipid vesicles formed from didocosanoyl phosphatidylcholine was considered too thick to allow the single annulus to span it. So there is a situation in which molecular models proposed for the interaction of an antibiotic with plasma membranes of sensitive organisms can be tested experimentally. There are a number of questions that can be asked. (1) What confirmatory evidence is there for polyene-bounded non-static aqueous pores in cell membranes? (2) Is it even necessary to postulate an aqueous pore in the plasma membrane to explain drug-induced proton permeability? (3) What membrane constituents are associated with the drug molecules? (4) How is it possible to account for the selectivity of these antibiotics? +
c. Evidence for Polyene-Bounded Aqueous Pores. If such structures exist in polyene-treated cells, their diameter will be such that it is unlikely that they would be observed by electron microscopy using either negatively stained or freeze-etched preparations. There have been a number of studies of polyenei n d u d morphological changes both in liposomes and in plasma membranes of sensitive cells (Tillack and Kinsky, 1973; Verkleij et al., 1973; De Kruijff
MODE OF ACTION OF ANTIFUNGAL DRUGS
27
and Demel, 1974; Nozawa et al., 1974; Kitajima et al., 1976; Pesti et al., 1981b; Sekiya et al., 1979, 1982). In many cases, effects were observed after prolonged incubation in the presence of the antibiotic at concentrations in excess of the minimum growth inhibitory concentration and, as a result, may not be directly related to the primary interaction of the drug with the plasma membrane. Pores were not observed in any of these studies, but it is quite clear that addition of amphotericin B and nystatin results in significant changes in membrane morphology, the most pronounced being a redistribution and clustering of intramembranous particles. This redistribution is considered by Pesti et af.(1981b) to be a specific effect of the polyene, since it did not occur on addition of nystatin to a nystatin-resistant strain of C. albicans. Other changes, including a deepening and deformation of membrane invaginations, were also observed but were considered non-specific effects. Drug-induced changes were not observed in intracellular membranes, a result cited as evidence for failure of the drug to be translocated across the plasma membrane. Similar results were obtained by Sekiya et al. (1982) in a study of the effects of amphotericin B and its methyl ester on plasma membranes of both C. albicans and erythrocytes. Differences were observed between amphotericin B and its methyl ester, the latter giving rise to elevated particle-free membrane domains towards the outside of the cell. There was also a decrease in the density of intramembranous particles. Such marked changes were not observed in erythrocyte membranes. Both groups of workers considered that changes in the distribution of intramembranous particles result from localized perturbations in the physical state of the membrane induced by a polyene-mediated dissociation of ergosterol molecules from membrane phospholipids. An interesting, but quite unrelated, observation was reported by Pugh and Cawson (1980) who found that, at the minimum growth inhibitory concentration, nystatin caused a localized collapse in the cell wall of C. albicans. Similar effects were also observed on addition of NaF and ethylenediaminetetra-acetate (EDTA) to the culture. In the absence of definitive morphological evidence for aqueous pores arising from annuli of polyene molecules within the lipid bilayer, we must look to physical methods for possible confirmation of this model. The degree of membrane damage can, to some extent, be assessed from the nature of low-molecular-weight cellular constituents released from sensitive cells or liposomes by the drug. Filipin causes gross membrane damage which, in the case of Mycoplusma spp., allows leakage of low-molecular-weight proteins, a result that is hardly consistent with formation of an aqueous pore. In contrast, amphotericin B and nystatin induce release of small molecules only, a result consistent with an aqueous pore of radius 0.4 nm (Cass et al., 1970). If pores of a finite diameter occur in membranes, then it should be possible to
28
DAVID KERRIDGE
block them. Such studies have been performed by Borisova et al. (1979) who found that organic molecules, such as tetramethyl ammonium which is approximately the sue of a hydrated K+ion, can block passage of inorganic ions and decrease the electrical conductance induced by amphotericin B in lipid bilayers. There are differences in the ion selectivity of the channel when the antibiotic is added to either one or both sides of artificial membranes. Addition to one side only results in cation-selective pores whereas, when added to both sides, the pores are anion-selective (Marty and Finkelstein, 1975). This difference is assumed to result from differences in the structures of single and double annuli. When the polyene is present on both sides of the membrane, hydroxyl groups within the double annulus will impart a positive charge to the interior of the pore relative to the bulk medium; as a result the pore will be anion selective. With a single polyene annulus, the ring of hydroxyl groups at the hydrophobic end of the annulus imparts a negative potential and hence provide a cation-selective gate at the entrance to the pore. Electrical conductance measurements can provide diagnostic evidence for the presence of antibiotic-induced aqueous pores within membranes (for references see Gale et al., 1981). Although initially it was not possible to detect discrete conductance fluctuations in polyene-treated membranes (Hladky and Haydon, 1970; Romine et al., 1977), such discrete fluctuations have now been detected in black lipid films treated with amphotericin B at concentrations ranging from 10 to 1 0 0 (Ermishkin ~ et al., 1976, 1977). Fluctuations were observed only when the polyene was added to both sides of the membrane. Addition of amphotericin ( 1 0 0 p ~to ) one side of the membrane only resulted in an increase in conductance, but discrete current fluctuations were not observed. Cholesterol was essential for polyene-induced current fluctuations and there was evidence for preferential anion selectivity of a single channel. In brain phospholipid membranes, it was found that individual polyene-induced ionic channels undergo a large number of transitions between the open and closed states during a lifetime of several minutes (Ermishkin et al., 1977). The discrepancies between these and earlier results might be related to the differences between the phospholipid preparations used (Romine et al., 1977). Discrete current fluctuations also occurred at the phase-transition temperature in lipid bilayers formed from synthetic distearoylphosphorylcholine, and it was suggested that such ionic channels result from lipid domains interacting within the bilayer (Antonov et al., 1980). If this is so, then it is likely that “aqueous pores” are an indirect effect of an interaction of the polyene molecules with membrane sterols resulting in localized changes in membrane fluidity and are not necessarily formed from an annulus of eight drug molecules. d. Role of Membrane Constituents in Polyene Action. Early studies on the
MODE OF ACTION OF ANTIFUNGAL DRUGS
29
interaction of polyene macrolide antibiotics both in vivo with prokaryotic and eukaryotic organisms, and in v i m with liposomal vesicles and black lipid films, emphasized the importance of membrane sterols in determining both the selectivity of these antibiotics and sensitivity of the organisms to these compounds (Hamilton-Miller, 1973; Norman et al., 1976; Gale et al., 1981). Membrane lipids of Acholeplasma laidlawii can be altered by growth in the presence of different sterols, and this has proved invaluable in studying the structural requirements of the membrane sterol necessary to confer polyene sensitivity (De Kruijff et al., 1974). For cells to be sensitive to polyene antibiotics, the membrane sterol must have a 3-8 hydroxyl group, a planar ring structure and a hydrophobic side chain at the C-17 position. Using polyene-induced leakage of K+ ions to measure polyene sensitivity, Gale (1973, 1974) found that ergosterol-containing C. albicans was more sensitive to amphotericin B methyl ester than cholesterol-containingmouse LS cells; the converse is true for filipin. Similar results were obtained by Chen et al. (1977), who found that C. albicans was more sensitive than human erythrocytes to nystatin, amphotericin B and its methyl ester. Cells or liposomal vesicles containing ergosterol are more susceptible to disruption by amphotericin B than those containing cholesterol (De Kruijff et al., 1974; Archer and Gale, 1975; Archer, 1976; Teerlink et al., 1980). It would appear that the clinical effectiveness of amphotericin B results from its greater affinity for ergosterol-containing fungal membranes than cholesterolcontaining mammalian membranes. Simultaneous addition of exogenous sterols protects sensitive cells against the growth inhibitory effects of polyene macrolide antibiotics and, again, the selectivity correiates well in that ergosterol is more effective than cholesterol as an antagonist of amphotericin B methyl ester when tested against C. albicans (Archer and Gale, 1975).Here, of course, protection results from a physical interaction between sterol and polyene with a decrease in the effective drug concentration. Mutant fungi resistant to amphotericin B, and other polyene macrolide antibiotics, are not a particular clinical problem (Athar and Winner, 1971; Hamilton-Miller, 1973), but such resistant mutants can be readily produced in the laboratory. In the majority of cases, the sterol constituents of the resistant mutant differ from those of the parental strain, but there are inconsistenciesin the results. Polyene-resistant mutants of C. albicanr selected by continuous culture in the presence of antibiotic were found by Athar and Winner (1971) to have a decreased ergosterol content, whereas strains selected after mutagenesis had an enhanced ergosterol content (Hamilton-Miller, 1972a). Detailed analysis of the lipid content of polyene-resistant mutants of C. albicans have been camed out by Subden et al. (1977) and Pierce et al. (1977), but these results have been more important in helping to elucidate the
30
DAVID KERRIDGE
pathway of sterol biosythesis in this organism than understanding the mode of action of polyene macrolide antibiotics. It is clear from the data of Pierce et al. (1978) (Table 4) that, although membrane sterols are important in the disruptive interaction of polyenes with cellular membranes, strains with apparently similar sterol compositions can have markedly different sensitivities, so that other factors must also be involved. The first evidence that the presence of sterols within the membrane was not an essential factor in determining sensitivity to these antibiotics came from studies on polyene-induced glucose release from liposomal vesicles (HsuChen and Feingold, 1973). Under certain conditions, sterols far from enhancing the disruptive action of polyenes have the opposite effect. Liposomes prepared from lecithin with dipalmitoyl or distearoyl side chains are sensitive to polyenes but, if a sterol is incorporated into the bilayer, they are resistant. It was suggested that these results might be explained by osmotic swelling of the liposomes and subsequent breakage (De Kruijff et al., 1974), but similar results were obtained by Archer (1976) using Mycoplasma mycoides subspecies Capri in which the membrane composition had been modified by growth in media supplemented with different sterols. Since this organism is not osmotically fragile, nor does it leak K+ ions at 2"C, it is unlikely that these results can be explained in this way. It would appear that both membrane sterol and the physical state of the membrane determine the sensitivity or resistance to polyenes, and a disruptive interaction demands that membrane lipids are in an ordered state. Application of a variety of spectroscopic techniques to analysis of polyene-lipid interactions has led to considerable advances in our understanding of the mode of action of these drugs (Medoff et al., 1983). These studies have provided information on two important aspects of polyene action: (1) the stoicheiometry and specificity of polyene-lipid interaction, which is of considerable importance in understanding the selectivity of these drugs, and (2) conformational changes in the polyene molecule that occur on association with membranes, from which it is possible to gain information on possible stages in the drug-membrane interaction. Sterol-polyene complexes have been demonstrated in both water and water-ethanol mixes (reviewed by Medoff et al., 1983). This does not prove conclusively that such complexes exist in lipid bilayers, but it does provide circumstantial evidence for their existence. Interaction between drug and lipid is strongly influenced by the environment (Gruda et al., 1980) and, as a result, there are considerable problems in interpreting data from such studies. Amphotencin B molecules aggregate in aqueous media at concen~ ~ et ul., 1982)and, in forming polyene-sterol trations as low as 0 . 5 (Mazerski aggregates, the sterol molecules must compete with these aggregation forces. Addition of ethanol prevents not only polyene aggregation but also
TABLE 4. Sterol composition of polyene-resistant strains of Candida albicans. From Pierce et al. (1978)
Strain Parent c7 E4
Minimum growth inhibitory concentration ( p g d - ' ) Amphotericin B Nystatin 0.23 1.36 6.9
c4
500
D10
500
8 25 102 250 lo00
Total sterol (Yo dry wt.) (YOesterified) 0.15 0.189 0.367 0.722 0.499
27 19 34 26 26
Major sterol (YOof total sterol) Ergosterol (62.5) Ergosterol (70.4) Ergosta-5,8,22-trienol (46.5) 24-Methyl-24,25-dihydrolanosterol
24-Methyl-24,25-dihydrolanosterol
32
DAVID KERRIDGE
formation of the sterol-polyene complex. There is a narrow range of ethanol concentrations where the interaction can be successfully monitored. Gruda et af. (1980), using these optimum conditions, demonstrated a differential affinity towards ergosterol and cholesterol. Readio and Bittman (1982) overcame the problem of polyene aggregation in aqueous media by studying interaction of amphotericin B and its derivatives with sterols incorporated into egg phosphatidylcholine vesicles. In all cases, one polyene molecule interacted with one membrane sterol molecule. There was a significant difference between the binding constants of amphotericin B to the membraneassociated ergosterol and cholesterol, with amphotericin B being bound an order of magnitude more firmly to ergosterol than to cholesterol, a result consistent with the selectivity of this drug. Other studies on the stoicheiometry of interaction of polyene molecules with membrane sterols have been performed using either spectrophotometric techniques or permeability changes in membrane vesicles to monitor the interaction (reviewed by Medoff et af., 1983)and the values obtained range from less than unity to greater than three for the sterol/drug ratio. Circular dichroism is the spectrophotometric method most suitable for following changes in molecular conformation of polyene molecules that occur on interaction with membrane constituents (Bolard et af., 1980). Amphotericin B exists in a number of conformational states when associated with phospholipid vesicles. These states depend not only on the cholesterol content and physical state of the membrane (Boudet and Bolard, 1979), but also on the time elapsed after addition of the drug to vesicle preparations (Bolard and Cheron, 1982). These differences in circular dichroism spectra reflect not only the final conformation of the drug within the lipid bilayer but also intermediate stages occurring after the first adsorption on to the membrane surface. The importance of the physical state of membrane lipids in determining their affinity for amphotericin B has been further emphasized by Bolard et af. (1981) in a study of the transfer of amphotericin B between vesicles in different physical states. The exchange between vesicles was rapid with a half-time of about 30 seconds, and cannot be accounted for by vesicle fusion which is a much slower process. There was a difference in the association constant for the drug with membranes in the gel and the liquid-crystalline state, with the binding of amphotericin B to phospholipids in the gel state being some 200 times higher than to phospholipids in the liquid-crystalline state. Interaction of amphotericin B with phospholipid vesicles in the gel state induced fusion or aggregation of vesicles, which did not ocur when the vesicles were in the liquid-crystalline state. Proton efflux from vesicles is very dependent on amphotericin B content; at low concentrations not only is the efflux slower but it is also incomplete,
MODE OF ACTION OF ANTINNGAL DRUGS
33
a result consistent with a pore model where a minimum number of drug molecules per vesicle is required for leakage (Bolard et al., 1981). Similar studies have been carried out by Vertut-Croquin et al. (1983) to assess the relative efficiency of interaction of amphotericin B with multilamellar vesicles containing ergosterol or cholesterol. Again, numerous amphotericin B conformations were demonstrated by circular dichroism, and it is likely that one or at the most two conformers of amphotericin B are responsible for druginduced permeability. The active conformers were the same when either cholesterol or ergosterol was present in the bilayer. The concentration of amphotericin B necessary to attain these conformational states was greater when cholesterol was present in the lipid bilayer than when ergosterol was the membrane sterol, a result to be expected from the selective action of this antibiotic. Sterol-free vesicles were also disrupted by amphotericin B but the sensitivity was lower than when sterols were present. Conformational changes occurred in the absence of sterols, but these have been interpreted as involving formation of an amphotericin B-phospholipid mixed vesicle (Bolard et al., 1980). Not all polyene macrolide antibiotics behave as amphotericin B. The aromatic heptaene macrolide antibiotics, candicidin and vacidin A, are interesting in that they are biologically more active than the non-aromatic heptaenes and, unlike these compounds, give rise to only two types of lipid-polyene complex in unilamellar vesicles (Mazerski et al., 1983). The first was observed in the absence of sterols or in vesicles containing less than 10mol YOof cholesterol. Under these conditions, there was no effect on membrane permeability, and it was assumed that polyene molecules were adsorbed in monomeric form on the surface of the bilayer. The second type of interaction was apparently associated with formation of a polyenecholesterol complex, and was responsible for changes in permeability. The physical state of the membrane lipids and sterol content af€ected only the relative proportion of the two conformational states.
f. Inhibition of Membrane-Associated Enzymes by Polyene Macrolide Antibiotics. So far, we have assumed that interaction of polyene antibiotics with membrane lipids and the resulting impairment of structural integrity is solely responsible for their growth inhibitory effects. The lipid environment is important in controlling the activity of a number of membrane enzymes (Singh et al., 1979) and, if amphotericin B and other polyenes modify this environment, then the activity of these enzymes may be affected, at least indirectly. The first indication that this might occur came from observations by Solov’eva et al. (1976) that, at concentrations well in excess of the minimum growth inhibitory concentration, polyenes inhibited lactate dehydrogenase and adenosine triphosphatase (ATPase) of C. albicans.
34
DAVID KERRIDGE
Dipple and Houslay (1979) examined the effects of amphotericin B on the adenylate cyclase of rat-liver plasma membranes. At low antibiotic concentrations, there was a marked effect on Arrhenius plots for this enzyme, presumably resulting from lateral redistribution of membrane lipids associated with formation of a complex between antibiotic and membrane cholesterol. The chitosomal chitin synthase of Mucor rouxii is inhibited by both amphotericin B and nystatin at concentrations in excess of the minimum growth inhibitory concentration (Rast and Bartnicki-Garcia, 1981). Inhibition by amphotericin B appeared to be non-competitive with a K,value of 0.13 KIM. The effect of nystatin was complex in that, although the enzyme was inhibited at concentrations in excess of 0.1 mM, there was a marked stimulation of activity at 0.05 mM. Filipin had a slight inhibitory effect and pimaricin was without effect on the activity of this enzyme. The 16s chitin synthase produced by digitonin treatment of chitosomes from Aspergillus bisporus is also inhibited by amphotericin B methyl ester. A Dixon plot gave a 4 value of 0 . 0 9 m ~and, unlike data for Sacch. cerevisiae, indicated a competitive type of inhibition (Hanseler et al., 1983). Earlier studies (Keiler and Cabib, 1971; Kinsky, 1972; Lyr and Seyd, 1978) had failed to show any inhibition of chitin synthase by polyenes, but Rast and Bartnicki-Garcia (1981) considered that this resulted from an excess of sterol in the crude membrane fractions used by these workers as a source of chitin synthase. Although there is evidence from these results for a sterol or a sterol-binding site in chitosomal chitin synthase from these organisms, it is unlikely that chitin synthase is a primary target in a polyene-induced growth inhibition. Chitin synthase activity in polyene-resistant mutants of C. albicans was found by Pesti et al. (1981a) to be significantly higher than that in the parental strain, and the zymogen component from the membrane fractions of the mutant strains was more susceptible to trypsin digestion than those from the parental strains. This effect was assumed by the authors to result from changes in the lipid environment of the membrane in the vicinity of the enzyme, and the inhibitory effects of polyene antibiotics on chitin synthase observed in vitro may result from similar changes. g . Resistance to Polyene Macrolide Antibiotics. When considering resistance to clinically important antimycotic agents, it is essential to relate this to the drug concentrations at the site of infection. Obviously for superficial infections this is not a particular problem, but for systemic infections it is an important consideration since many of the drugs used are themselves toxic, and the concentration that can be tolerated in the serum during therapy is low. Hamilton-Miller (1972b) determined the resistance profiles of a number of clinical isolates of C. albicans; an appreciable proportion of these strains were not inhibited in vitro by these drugs at concentrations equivalent to those attained in serum.
MODE OF ACTION OF ANTIFUNGAL DRUGS
35
The occurrence of mutants of C. albicans and other pathogenic fungi resistant to polyene macrolide antibiotics is not a clinical problem, and patients are not normally monitored for the presence of drug-resistant organisms. However, Dick et al. (1980) examined isolates of C. ulbicuns from patients undergoing extensive therapy for acute leukaemia and bone-marrow transplantation, and observed a significant incidence of polyene-resistant yeasts. They suggested that, for patients at risk and undergoing prolonged therapy, there was a need to monitor for the presence of drug-resistant fungi. The production and selection of polyene-resistant strains of fungi in the laboratory is relatively easy, and in general resistant organisms are distinguished from parental strains by an alteration in the lipid composition of cell membranes. These studies have already been considered in relationshipto the role of membrane lipids in determining the sensitivity of the organism to these antibiotics. Many authors have emphasized the importance of sterols in mediating drug-induced permeability changes and the role of these mutants in determining the pathways of sterol biosynthesis, but there have been few detailed genetic analyses of polyene resistance in fungi. In one study, two phenotypically distinct sets of polyene-resistant mutants of Sacch. cerevisiue were isolated, the first being respiratory competent and having no lipid requirements for growth and the second being respiratory deficient and requiring either an unsaturated fatty acid or a sterol for growth. The latter groups of mutants were more resistant. Genetic analysis demonstrated that these strains fell into six distinct groups and that, in this instance, nystatin resistance was a recessive character (Molzahn and Woods, 1972). Drug resistance in bacteria is often associated with enzymicinactivation of an antibiotic, but there have been no reports of a similar phenomenon being important in fungi resistant to polyene macrolide antibiotics. Nystatinresistant mutants of certain dermatophytic fungi have been isolated, and Capek et al. (1970) reported that Trichophytonmentagrophytes,Trichophyton rubrum and Microsporum gypseum produce an inducible enzyme that degrades nystatin. Intrinsic resistance was also found in the strains of dermatophyte and this was associated with an altered sterol content (Capek and Simek, 1972). One characteristic property of micro-organisms is their ability to adapt to environmental changes. Candida albicans and other pathogenic fungi are no exception, and environmentally induced changes in the cell envelope of these organisms may affect either the composition of the target organelle or penetration of the drug through the cell wall. We have already seen how modification of the plasma membrane of Mycoplasma and Acholeplasma species has a dramatic effect on their sensitivity to polyene antibiotics, and similar results have been obtained for yeasts. Koh et al. (1977) isolated a strain of C. albicans requiring an unsaturated fatty acid for growth. By
36
DAVID KERRIDGE
growing this strain in the presence of different unsaturated long-chain fatty acids, it was possible to modify the membrane composition and, as a result, sensitivity to amphotericin B methyl ester. Sensitivity to amphotericin B methyl ester (measured by release of K+ ions) ranged from 0.08 f 0.02 pg ml-' for organisms grown in the presence of palmitoleic acid ((&) to 1.2 f 0.3 pgml-' for organismsgrown in the presence of oleic acid (Clk.). Membrane sterols were little affected by the nature of the fatty-acyl supplement. Stationary-phase cultures of the mutant strain, like those from the parent, were less sensitive to amphotericin B methyl ester and, for cells grown in the presence of oleic acid, protoplasts were also resistant. Resistance in this instance was associated with the plasma membrane rather than the cell wall. Similarly, with a strain of C. albicans deficient in membrane ergosterol and hence resistant to nystatin, addition of ergosterol to the growth medium resulted in organisms regaining their sensitivity to nystatin (Mas and Pina, 1980). This change was readily reversible and, after growth in the absence of sterol, the organism was once more resistant. Changes in susceptibility to polyene-induced release of K+ ions occur during growth of C. albicans in batch culture where, after cessation of growth, the organisms become progressively more resistant to amphotericin B (Gale, 1974; Hammond and Kliger, 1974; Hammond et al., 1974). In batch culture, the environment is continuously changing and, as a result, it is difficult to determine which factors are responsible for changes in polyene sensitivity of the organisms. Johnson er al. (1978) overcame this problem by studying polyene-induced release of K+ ions from C. albicans grown in continuous culture, and monitored the effects on polyene sensitivity of a variety of factors ranging from the nature of the limiting substrate to the specific growth rate. Considerable differences in the susceptibility of these cells were observed; carbon-limited cultures showed highest susceptibility. In general, the susceptibilityincreased with growth rate and with growth at low temperatures and at low pH values. There were also differences in the concentration of K + ions within the pool. Unfortunately, the authors did not correlate these differences with changes in composition and organization of the cell envelope. Far more extensive studies have been carried out by Gale and his colleagues on the biochemical basis of the polyene resistance in C. albicans which develops after the cessation of growth. Here the variation in sensitivity to polyene-induced release of K+ ions results from changes in the composition of the cell wall since protoplasts derived from phenotypically resistant cells are sensitive (Gale er al., 1975). There are changes in ultrastructure, with walls of stationary-phasecells being thicker (211 f 58 nm) than those of exponentially growing cells (143 f 22 nm) and lacking the characteristic layered appearance of exponentiallygrowing cells (Cassone et af., 1979). This
MODE OF ACTION OF ANTIFUNGAL DRUGS
37
phenotypic change was readily reversible with the walls regaining their characteristic layered appearance, and the cells becoming sensitive to amphotericin B methyl ester on transfer to fresh growth medium. Although there were obvious changes in cell-wall ultrastructure after cessation of growth, no comparable changes were detected in the porosity of the wall to polyethylene glycols (Cope, 1980b). Polyene molecules must penetrate the fungal cell wall to reach the plasma membrane, and the organization of such a complex structure will be important in passage of molecules as large as amphotericin B. Scherrer et al. (1974) determined the porosity of walls of Sacch. cerevisiae to polyethylene glycols, and found that molecules with an M,value greater than 600 and with an Einstein stokes radius of 0.81 nm are excluded. Clearly, if the wall of C. albicans is similar in structure to that of Sacch. cerevisiue, then amphotericin B with an M, value of 960 will have difficulties in crossing it. This is borne out by the fact that release of K+ions from protoplasts of C. albicans occurs immediately on addition of amphotericin B whereas, with intact cells, there is an appreciable time lag before it can be detected. Chemical analyses of walls of C. albicans have shown that changes in the lipid composition occur after cessation of growth, but there is no evidence that these are responsible for the development of phenotypic drug resistance (Gale et al., 1975). The resistance of stationary-phase cells to amphotericin B methyl ester can be modified in a number of ways, namely (1) by treatment with thiol agents such as mercaptoethanol and N-ethylmaleimide (Gale et al., 1975), (ii) by manipulating environmental conditions, e.g. O2tension and pH value (Gale et al., 1977, 1978) and (iii) by treating cells with exogenous glucanases (Gale et al., 1980a). Although factors that affect the phenotypic resistance of C. albicans are many and various, there is an underlying pattern in that all treatments that decreased the activity of endogenous /?-glucanases increased resistance to polyenes, and all of those that increased the activity of these enzymes decreased drug resistance (Notario et al., 1982; Table 5). Both (1-3)- and (1 +6)-/?-glucans are major structural components of the wall of C. albicans, and the synthesis and deposition of these molecules will involve not only membrane-associated glucan synthases but also endogenous B-glucanases within the wall (Johnson, 1968; Farkas, 1979; Notario, 1982). The cell wall is not a static structure, and its molecular organization results from a balance between synthesis and partial breakdown of its structural components. It is not surprising that changes in wall organization resulting from an imbalance between synthetic and degradative pathways of B-glucan metabolism can have such a pronounced effect on the sensitivity of C. albicans to polyene macrolide antibiotics. However, the question that must be asked is: what, if any, is the clinical significance of this phenotypic resistance in C. albicans? Fungal lesions will
38
DAVID KERRDGE
TABLE 5. Beta-Glucanase activity and polyene resistance in Candida albicans. From Notario et al. (1982) Condition or treatment
/3-Glucanase activity
Stationary-phase cultures Growth medium supplemented with glutamate (stationary-phase cultures) pH Value of the medium (a) maintained at 7 (b) maintained at 8 Oxygen content of the growth medium (a) increased (b) decreased Addition of (a) trichodermin (b) mercaptoethanol (c) N-ethylmaleimide Incubation of the culture with (a) chitinase (b) /3-glucanase
Resistance to amphotericin B methyl ester
Decrease Increase
Increase Decrease
Increase Inactivation
Decrease Irreversible increase
Decrease Increase
Increase Decrease
Decrease Increase Inhibition
Increase Decrease Irreversible increase
Increase
Decrease Decrease
?
contain an heterogeneous population of C. albicans with both growing and non-growing cells present and, if the non-growing cells have a resistance greater than the amphotericin B concentration in the serum (which owing to its toxicity will be low), then there may be a reservoir of infection which can develop on cessation of therapy. 2. Clinical Usage
Amphotericin B is the only polyene macrolide antibiotic sufficiently nontoxic to be used in the treatment of patients with systemic fungal infections. It is administered by intravenous infusion and this is usually accompanied by nausea, vomiting, fever and local thrombophlebitis. These conditions normally abate on prolonged administration and can be controlled by other drugs (Medoff and Kobayashi, 1980). Kidney damage is invariable, and during polyene therapy renal function must be monitored. The effects of polyene antibiotics on the renal medulla have been examined by Brevis et al. (1984). These authors consider that cellular damage results from anoxia associated with the increased O2demand required to maintain the electrolyte balance in cells whose membrane permeability has been increased by addition
MODE OF ACTION OF ANTIFUNGAL DRUGS
39
FIG. 10. Structural formula of chlormidazole.
of drug. No one has suggested that a similar effect can be responsible for the fungicidal effects of these antibiotics. Apart from the nephrotoxicity, all toxic effects of amphotericin B disappear once therapy is discontinued. Similar problems do not occur when these antibiotics are used to treat patients with topical and gastrointestinal infections (Medoff and Kobayashi, 1980). The role of amphotericin B in the therapy of systemic infections is almost certainly complex and not restricted to a disruptive interaction with the fungal plasma membrane. There have been a number of reports of a stimulatory effect of this drug, at low concentrations, on macrophages. It also has potent humoral immunostimulant effects in mice, and augments cell-mediated immune responses. A discussion of these effects is outside the scope of this article, but clearly they are of great importance in understanding the therapeutic effect of this compound in vivo (Medoff et al., 1983). B. IMIDAZOLE ANTIMYCOTICS
The most recent major advance in the control of both human and plant mycoses has come with the introduction of synthetic benzimidazole, imidazole, and triazole drugs. The original impetus to these studies came from observations by Woolley (1944) that benzimidazole inhibited growth of a number of organisms, and that these inhibitory effects were reversed by addition of adenine or guanine. These findings were extended by a number of groups of workers and, in 1958, chlormidazole (Fig. 10) was introduced into clinical practice (Herrling et af., 1959). This compound has now been replaced by other synthetic imidazole drugs. The first of these was clotrimazole (Plempel et al., 1969),and this has been followed by miconazole (Van Cutsem and Thienpont, 1972), econazole (Thienpont et al., 1975), tioconazole (Jevons et al., 1979), isonconazole (Godefroi et al., 1969) and ketoconazole (Heeres et al., 1979). Structural formulae of these compounds are shown in Fig. 11. Many other imidazole derivatives have been synthesized and tested for their antimycotic activity but as yet have not been introduced into clinical practice. Imidazole antimycotics are effective against a wide range of pathogenic fungi and Gram-positive bacteria. In general, they are fungistatic at the
X-t-Z I Y
Miconazole
H
H
Econazole
H
H
o-" Cl
lsoconazole
H
H
CI
H -CSH I+
CI
$Flfi Cl H
H
Tioconazole
H
1
CI
a CI
Ketoconazole
H
H
FIG. 11. Structural formulae of certain clinically important imidazole antimycc drugs.
MODE OF ACTION OF ANTIFUNGAL DRUGS
41
minimum growth inhibitory concentration and fungicidal at higher concentrations. These drugs, except for ketoconazole, are poorly soluble in water, and their interaction with sensitive fungi is markedly affected by environmental conditions, which may account for the reported differences in the sensitivity of fungi to these compounds and the difficultiesin reproducibly determining the minimum growth inhibitory concentrations (Holt, 1980b).
I . Molecular Basis of Antifungal Action Clinically important imidazole drugs are characterized by an unsubstituted imidazole ring and tetrahedral symmetry at the atom to which this is joined. The molecule is also predominantly hydrophobic (Tolkmith et al., 1967).The imidazole moiety is the only reactive functional group in these molecules and is important in the interaction with their target(s) within the cell. Structure-activity relationships in the azole drugs have been examined by a number of workers. It is clear that the lipophicity of these compounds is related to their activity in vitro, and this may reflect the ability of the drug to penetrate biological membranes and to inhibit membrane-bound enzymes (Bawden etal., 1983). Heeres (1983) found no clear correlation between the activity of the mole drugs in particular the 1,3-dioxolan-2-y1 methyl derivativesin inhibiting the growth of C. albicans in vitro and in vivo in animal model systems. These compounds have a variety of inhibitory effects on membrane-associated functions in sensitive organisms, and the relative importance of these effects in mediating growth inhibition may be influenced by the nature of the imidazole and the fungal pathogen, as well as the environment. There is a concensus of opinion that fungistatic effects result from an inhibition of membrane sterol synthesis, and the fungicidal effects from an interaction with, and consequent impairment of, the barrier function of the plasma membrane. However, many other effects have been reported which may be important in the inhibitory action of these compounds during therapy and these will be discussed later. a. Inhibition of Sterol Biosynthesis. Inhibition of cholesterol biosynthesis in rat liver by 1-alkyl imidazoles was first reported by Baggaley et al. (1979, but it was not until 1978 that Van den Bossche and his colleagues (Van den Bossche et al., 1978) observed a time- and concentration-dependent inhibition by miconazole of incorporation of radioactivity from ['4C]acetateinto the sterol fraction of C. albicans. The most interesting observation was that, associated with the growth inhibitory effects and decrease in ergosterol biosynthesis, there was an accumulation of 14a-methyl sterols (Table 6). These changes were interpreted as resulting from a specificinhibition of sterol demethylase by miconazole, and an associated accumulation of intermediates
42
DAVID KERRIDGE
TABLE 6. Miconazole-induced changes in the sterol composition of Candida albicans. From Van den Bossche et al. (1978) Incubation time (hours)
Miconazole concentration (M)
Membrane sterols present
4 4
0 10-~
4 16
10-9
16
10-7
24
10-7
Ergosterol, lanosterol Ergosterol, obtusifoliol, 4,14-dimethylzymosterol, lanosterol, 24-methylene dihydrolanosterol 1CMethylfecosterol Ergosterol, obtusifoliol, 24-methyldihydrolanosterol, lanosterol 14-methylfecosterol,obtusifoliol, 4,lCdimethyllanosterol, lanosterol 1CMethylfecosterol
in ergosterol biosynthesis (Fig. 12). Biosynthesisof sterols is affected not only in vivo but also in vitro where an inhibition of sterol synthesis from mevalonate by cell-free systems has been reported (Marriott, 1980; Pye and Marriott, 1982; Gadher et al., 1983; Van den Bossche et al., 1984). In 24-Methylene dihydrolanosterol
Obtusifoliol
Ergosterol
14Methylfezosterol
FIG. 12. Site of inhibition by imidazole drugs in biosynthesis of ergosterol.
MODE OF ACTION OF ANl"GAL
43
DRUGS
Ustilugo muydis (maize loose smut) both miconazole and dodecyl imidazole inhibit sterol demethylation at the minimum fungitoxic concentration; at higher concentrations, dodecyl imidazole also inhibits 2,3-oxidosqualene cyclization and subsequent transmethylation. The diversity of these effects was assumed to be due to binding of the drug to sterol-carrier proteins (Henry and Sisler, 1979). Addition of clotrimazole to Succh. cerevisiue also inhibits 3-hydroxy-3-methylglutaryl-CoAreductase, but this results from feedback inhibition by accumulated intermediates of sterol biosynthesis (Berg et ul., 1981). At concentrations higher than those required to inhibit sterol demethylase there are changes in the relative proportion of saturated fatty-acyl residues (Van den Bossche et ul., 1981). This change in lipid composition could result from a direct effect on the fatty acid desaturase system or an indirect effect resulting from changes in the sterol composition of the plasma membranes. Much of our knowledge of sterol demethylation in yeasts stems from studies on Succh. cerevisiue, where it has been demonstrated, in both anaerobically and aerobically grown cells, that 14a-demethylation of lanosterol, an initial stage in its conversion into zymosterol and hence ergosterol, is catalysed by a cytochrome P-450 NADPH-cytochrome P-450 reductase system (Aoyama and Yoshida, 1978a, b; Aoyama et ul., 1981). There are three oxygenation steps involved in this reaction with the methyl carbon ultimately being removed as formic acid (Fig. 13). The initial oxidation is inhibited by CO, whereas the oxidation of the alcohol and aldehyde derivatives is unaffected by CO. It is characteristic of cytochrome P-450 that it binds CO and that the CO complex has a major absorption band at 450 nm. The fact that 14a-methyl sterols, rather than the alcohol or aldehyde derivatives, accumulate would suggest that the first reaction is inhibited. Microsomal mono-oxygenases are involved not only in sterol biosynthesis but also in a variety of other reactions, including inactivation of certain toxic
CH,
CH~OH
NADPH HCOOH
CHO
FIG. 13. Reactions involved in sterol demethylation.
44
DAVID KERRIDGE
and carcinogenic agents. Clotrimazole is one of the most effective inhibitors of hepatic aryl hydrocarbon hydrolase, with 50% inhibition of this enzyme occurring at a concentration of 70- (Kahl et al., 1980). Spectral analysis of the interaction of clotrimazole with the reduced form of the hepatic cytochrome P-450 gave a Type I1 spectrum (Schenkman et al., 1967) and a double-banded Soret region with peaks at 427 and 446 nm characteristic of an interaction between heterocyclic compounds with a nitrogen atom in the ring and the haem iron in cytochrome P-450. The effects of a number of agriculturalfungicides on sterol demethylationby cell-free preparationsfrom Sacch. cerevisiae and rat liver were examined by Gadher et al. (1983) who compared their relative efficiencies as inhibitors with a series of triazole derivatives. The inhibitory effects were considered to result from binding of the heterocyclic nitrogen atom of the fungicide to the protohaem iron thus excluding 02,the normal sixth ligand. Differences in relative efficiencies of these heterocyclic fungicides as inhibitors of sterol demethylation were assumed to reside in the remainder of the molecule, with the non-heterocyclic part of the molecule binding to the lipophilic binding site on cytochrome P-450 normally occupied by the 14a-methyl sterol. Similar effects on cytochrome P-450 of C.albicans have been reported by Van den Bossche and his collaborators (Van den Bossche et al., 1984). The next question to be asked is: how does inhibition of sterol demethylation and accumulation of sterols possessing a C-14 a-methyl group cause cessation of growth? Saccharomyces cerevisiae is capable of synthesizingall of its sterolsde novo under aerobic conditions, but has an absolute requirement for exogenous sterols when grown under strict anaerobic conditions (Andreasen and Stier, 1954). This finding has been exploited by Nes et al. (1978) to examine the relationship between sterol structure and its ability to support anaerobic growth of this organism. Growth occurred normally on addition of ergosterol; it was decreased when ergosterol was replaced by cholesterol and, under these anaerobic conditions, was not supported by lanosterol. Lanosterol can be incorporated into phospholipid vesicles, but the presence of a methyl group on C-14 of the sterol molecule weakens the interaction of the sterol a face with nearby phospholipid acyl side chains by abolishing contact with C-12 and weakening interactions at C-5 and C-7. This weakening of the interaction between sterol and phospholipid enhances membrane fluidity with an associated impairment of membrane function and a resultant inhibitory effect on growth (Bloch, 1983). What is the evidence that interaction of imidazole drugs with sterol demethylase is responsible for their fungistatic effects? Pye and Mamott (1982) compared the activities of a number of imidazole derivatives against both growth of C. albicans and sterol demethylase activity in virro (Table 7), and found no obvious correlation between the two effects. But as they point
45
MODE OF ACTION OF ANTIFUNGAL DRUGS
TABLE 7. Inhibition of sterol demethylase in vitro and correlationwith inhibition of growth of Candida albicans. From Pye and Marriott (1982) Imidazole drug
Tioconazole Ketoconazole Parconazole Butaconazole Econazole Clotrimazole Miconazole
Sterol demethylase IC, (nM) 50 f 10 50 f 10 60 f 20 60 f 20 90 f 20 120 & 10 20 200
*
Minimum growth inhibitory concentration
(w) 8.4
46
34 120 33 33 30
out, not only are environmental factors important in determining the mimimum growth inhibitory complex, but also the relative ease of penetration to target sites within the cell may vary from one imidazole derivative to another. Direct evidence for the involvement of sterol demethylase in the inhibitory action of imidazole drugs has been obtained by Sud and Feingold (1981). Saccharomyces cerevisiae grows under strict anaerobic conditions, provided that ergosterol and an unsaturated fatty acid are added to the growth medium, and under these conditions the mimimum growth inhibitory concentrations for miconazole and clotrirnazolewere 13p~ and 36 p~ respectively. These values were lowered to 0.2 p~ and 1.1 p~ under aerobic conditions. The fungicidal concentration, under both aerobic and anaerobic conditions, was 10p ~ at; this concentration the plasma membrane was disrupted, allowing entry of methylene blue into the cells. Further support for the hypothesis that sterol demethylase is the primary target has come from studies of a strain of Ustilago maydis deficient in sterol C-14 demethylase (Walsh and Sisler, 1982). Growth of this strain was considerably slower than that of the parental strain and, surprisingly, not increased by addition of ergosterol. This may result from the fact that ergosterol is poorly taken up by yeasts. This strain had essentially the same characteristicsas those of the drug-treated wild-type strain, and was insensitive to miconazole and other inhibitors of sterol demethylation except at high concentrations. The role of sterol demethylase as a primary target for imidazole antimycotics has been questioned by Taylor et al. (1983). In a study of the effects of imidazole drugs and 15-azasterol on growth of two strains of Sacch. cerevisiae, one blocked in lanosterol C-14 demethylation and A5-desaturation, and the other in 2,3-oxidosqualene cyclization (this latter strain having an absolute requirement for both a sterol and an unsaturated fatty acid), the authors observed that growth was inhibited by clotrimazole and miconazole
46
DAVID KERRIDGE
at concentrations similar to those required to inhibit growth of the parental strain. Growth of the parent strain was unaffected by ergosterol and the authors interpreted these data as eliminating the possibility that sterol biosynthesis is a primary target for these compounds. Further supporting evidence for the importance of sterol demethylase as the primary target for the imidazole antimycotics both in vivo and in vitro has come from studies by Van den Bossche et al. (1980). Oral administration of ketoconazole to rats infected with C. albicans resulted in decreased incorporation of radioactivity into ergosterol and an increased incorporation into methylated sterols by the yeast. Higher concentrations of drug were required for a similar degree of inhibition of cholesterol synthesis in rat liver. The results in vivo correlated well with those obtained for inhibition of sterol biosynthesis in cell-free systems from both yeasts and rat liver, and with studies of binding of imidazole derivatives to cytochrome P-450 in microsomal preparations from both yeast and rat liver. In addition to the effects of imidazole drugs on fungal metabolism discussed in the preceding sections, it has also been reported that ketoconazole, at concentrations attainable during therapy, will displace corticosteroids from the corticosteroid-bindingprotein present in C. albicans (Stover et al., 1983). There is, however, no evidence that this phenomenon has any relevance to the antifungal activity of this or related imidazole drugs. The imidazole drugs cause hormonal perturbationsin the host during therapy (Pont er al., 1982a, b) resulting from their effect on cytochrome P-450-mediated reactions (Loose et al., 1983). Not only do these drugs inhibit sterol biosynthesis but they also displace steroid hormones from serum-carrier proteins (Gross0 et al., 1983) and these may be responsible for the side effects of these drugs during therapy. Ergosterol biosynthesis inhibitors are important not only in medicine but also in agriculture, and Schwinn (1983) has reviewed their history and contribution to both medicine and agriculture. b. Impairment of Membrane Function ( i ) Barrier function. A direct interaction of the hydrophobic moiety of an imidazole drug with lipid bilayers can affect both membrane structure and its barrier function, resulting in loss of cellular constituents, and the fungicidal effects of these drugs may result from such a physical interaction (Sud and Feingold, 1981).There have been innumerable studies during the past decade that support this contention. Iwata et al. (1973a) reported an impairment of membrane function and loss of intracellular constituents as one the primary effects of clotrimazole on C. albicans. Subsequent studies by Swamy et al. (1974) and Cope (1980a) confumed and extended these findings. In general, effects on membrane permeability are induced at concentrations higher than
MODE OF ACTION OF ANTIFUNGAL DRUGS
47
TABLE 8. Reversal by lipids of imidazole action on Candida albicans. From Yamaguchi (1977)
Fraction added (0.4mg ml-') Control Total lipid extract Phospholipids Phospholipids (hydrogenated) Trig1ycerides Triglycerides (hydrogenated) Sterols Sterol esters
Minimum growth inhibitory concentration ( p ~ ) Clotrimazole Miconazole 1.4 5.6 11.2
4.2 15.8 13.4
0.7 5.6
2.1 33.6
1.4 1.4 1.4
4.2 4.2 4.2
those required to inhibit growth, but the importance of these permeability effects is often difficult to assess since the minimum growth inhibitory concentration is markedly affected by the environment. The membrane disruptive effects of miconazole are not restricted to sensitive fungi, and haemolysis of mammalian erythrocytes is induced by the drug at concenet al., 1976). Haemolysis was prevented by trations of 1 8 8 (Swamy ~ ~ addition of serum to the suspending medium when the drugs bind to serum constituents in preference to the erythrocyte membrane. Radioactive miconazole binds to the erythrocyte membrane and, clearly, a direct interaction of the drug with the membrane is responsible for the disruptive effects. Both the growth inhibitory effects and the disruptive effects of imidazole drugs on protoplast membranes of C. albicans were antagonized by CaZ+and M$+ ions (Swamy etal., 1974). This antagonism may result from competition between the divalent ions and miconazole in its positively charged form (pK,value for the imidazole group is 6.65) for negatively charged groups within the cell. The interaction of both clotrimazole and miconazole with C. albicans was also antagonized by addition of lipids to the growth medium (Table 8). In contrast to polyene macrolide antibiotics, the inhibitory effects of the imidazole drugs were not antagonized by sterols and sterol esters (Yamaguchi 1977, 1978). The antagonistic effects are limited to phospholipids and acyl glycerides containing unsaturated fatty-acyl residues and sterols activated by ultraviolet radiation and fatty acids possessing a trans configuration. In view of the affinity of these compounds for imidazole antimycotics, the author suggested that they may be involved in interaction of these drugs with the plasma membrane. The presence of free fatty acids in
48
DAVID KERRIDGE
phospholipid liposomes sensitized these structures to the disruptive action of the imidazole drugs (Yamaguchi and Iwata, 1979; Sud et al., 1979). There was little loss of internal glucose from phospholipid vesicles at concentrations but, when oleic acid (30mol%) was below 25pg of miconazole d-' incorporated into the vesicles, there was a significant leakage of glucose at concentrations as low as 3 pg of miconazole ml-'. Growth of Gram-positive, but not Gram-negative, bacteria is inhibited by imidazole antimycotics and the inhibition is also prevented by unsaturated and unesterified fatty acids (Van den Bossche et al., 1982). In Staphylococcus aureus, the inhibitory effects were reversed when the fatty acid was added up to 2 hours after addition of drug. Although oleic acid reversed the inhibitory effects of miconazole on Staphylococcus aureus, it did not affect uptake of radioactively labelled drug measured after 24 hours incubation. In this instance, it would appear that antagonism does not result from a lowering of the effective drug concentration by an in vitro association of lipid and drug. This differs from the interpretation given by Yamaguchi (1977) that the spectral changes which occur on mixing miconazole and egg lecithin result from formation of a hydrophobic complex, with a consequent lowering of the effective drug concentration. Oleic acid was not as effective an antagonist of miconazole action against Staphylococcus aureus as it was for C. albicans, and it did not reverse the inhibitory effects of ketoconazole on C. albicans. Binding of miconazole to C. albicans is very dependent on the conditions under which the yeast has been grown, with more drug binding to cells harvested after cessation of growth than to those harvested in the exponential phase of growth (Cope,198Oc). Exponential-phase cells bind approximately 1 pg of drug (mg dry wt of yeast)-', a value comparable to that reported for binding to Staphylococcus aureus (Van den Bossche et al., 1982). Interaction between imidazole drugs and artificial lipid bilayers has also been studied by measuring drug-induced changes in electrical conductivity (Arndt et al., 1982), and by differential scanning calorimetry (Van den Bossche et al., 1982). The electrical conductivity in lipid bilayers, produced from oxidized cholesterol was increased by a factor of 1.65 & 0.35 at 1 O p ~ miconazole and by 3.57 & 1.98 at 2 0 p miconazole, ~ and similar effects were seen in bilayers formed from lipids extracted from baker's yeast. The effect was reversible and clearly supports the existence of a direct disruptive interaction of the drug with lipid bilayers at concentrations similar to those required to inhibit growth of C. albicans (Amdt et al., 1982). Analysis by differential scanning calorimetry of the interaction of miconazole and ketoconazole with multilamellar vesicles, formed from dipalmitoyl phosphatidylcholine,showed no significant effect on the enthalpy of melting, suggesting that neither compound bound to the lipid constituents. Miconazole changed the organization of the lipid bilayer when present in
MODE OF ACTION OF ANTIFUNGAL DRUGS
49
high concentrations (10% molar ratio), and this change in membrane organization could be responsible for its fungicidal action. Ketoconazole was located in the bilayer, but had little effect on lipid organization, a feature consistent with a lack of fungicidal activity (Van den Bossche et al., 1982). Insertion of miconazole and ketoconazole into lipid bilayers has been examined by computer modelling (Brasseur et al., 1983), and the findings are consistent with those obtained by differential scanning calorimetry (Van den Bossche et al., 1982). The model predicts that the two dichlorophenyl groups of miconazole are inserted in the hydrophobic phase of the lipid bilayer with the imidazole moiety remaining in the hydrophilic phase. Each miconazole molecule occupies a mean area of 0.9nm2 and is surrounded by seven dipalmitoyl phosphatidylcholinemolecules. There is no suggestion of drug molecules associating with each other within the membrane (as with polyene macrolide antiobiotics). Ketoconazole, a watersoluble imidazole derivative, is different in that, when inserted into the lipid bilayer, it is the piperazine moiety that is oriented towards the hydrophobic region, with the dichlorophenyl groups remaining in the hydrophilic phase. The area occupied by the drug molecule is 0.3 nm2 and such an orientation would not appear to disrupt the bilayer organization. However, the deacylated derivative has an inverted orientation in the lipid monolayer with the piperazine moiety oriented towards the hydrophilic phase. This results in an increase in the area occupied by the drug to 0.9 nmz and this compound is fungicidal (Fig. 14). This disorganization of the lipid bilayer may well explain the impairment of barrier function when C. albicans is treated with miconazole at fungicidal concentrations. Failure of ketoconazole to disrupt the lipid bilayer is consistent with it acting primarily as a fungistatic drug. Imidazoles apparently have two basic effects, namely an inhibition of sterol demethylase, responsible for growth inhibition, and at higher concentrations a physical disruption of the plasma membrane responsible for the fungicidal effects. This physical disruption may be responsible for the therapeutic action of these drugs when used to treat topical infections, where high local concentrations can be achieved. Imidazole drugs also affect a variety of other metabolic systems at concentrations equivalent to those inhibiting growth. Certain of these effects may be a consequence of either changes in membrane sterols resulting from an inhibition of the sterol demethylase, or physical disruption of cellular membranes. ( i i ) Inhibition of membrane transport. In 1974, Van den Bosche reported that at 1 0 . 4 1 a~ concentration well below the minimum growth inhibitory concentration, miconazole had selective effects on the uptake of specific nutrients into C. albicans.There was a decrease in uptake of adenine, guanine and hypoxanthine, no effect on uptake of glucose and leucine, and an
50
DAVID KERRIDGE
FIG. 14. Molecular models for the insertion of imidazole derivatives into lipid bilayers: (a) miconazole, (b) a deacylated derivative of ketoconazole and (c) ketoconazole. From Brasseur et al. (1983). Reproduced with the permission of Pergamon Press Ltd.
acceleration of uptake of adenosine, deoxyadenosine and guanosine. These results were originally interpreted as a direct and selective effect on membrane transport. Nutrient uptake was determined after prolonged exposure to the drug, and an alternative explanation would be that drug-induced changes in the composition of the plasma membrane are responsible. These results were not confirmed by Yamaguchi and Iwata (1979) who observed that, at the minimum growth inhibitory concentration, miconazole and clotrimazole lowered both the rates of uptake, and the final pool concentrations of leucine, lysine and other amino acids in starved suspensions of C. albicans. This inhibition was partially relieved by addition of glucose to the incubation medium. The drugs also induced an efflux of pool amino acids. The apparent discrepancy between these findings and those of Van den Bossche (1974) might be attributable to differences in experimental conditions used, with one group studying drug action under starvation conditions and the other under conditions of growth. Yamaguchi and Iwata (1979) considered that inhibition of amino-acid uptake is a secondary nonspecific effect resulting from a disruptive interaction of imidazole drugs with
MODE OF ACTION OF ANTIFUNGAL DRUGS
51
the plasma membrane. There may, however, be a more direct effect on nutrient transport since miconazole competitively inhibits the plasma membrane-bound ATPase of Schizosaccharomyces pombe and also the lipid reconstituted form of the enzyme (Dufour et al., 1980). The Ki value of miconazole for the plasma-membrane enzyme was 5.3 PM and for the lipidreconstituted form 47 PM,the former value being similar to the concentration required to inhibit growth of the organism. At these concentrations, miconazole also induced a rapid stoicheiometricexchange of protons and K+ ions when added to intact cells incubated at pH 4.5 in the presence of glucose. Miconazole-induced loss of K + ions from Sacch. cerevisiae was found by Borst-Pauwels et al. (1983) to be accompanied by shrinkage of cells. This response was heterogeneous in that certain of the shrunken cells still retained barrier properties in their plasma membranes. Bacillus megaterium is also sensitive to miconazole and, at the minimum growth inhibitory concentration, although there was little loss of cellular K+ ions the plasma-membrane ATPase was significantly inhibited (Patricia Skeggs, unpublished observations). Clearly, the effects of imidazole drugs on nutrient transport must be re-examined in the light of these findings to distinguish between a specific effect on the transport proteins, an effect on the proton-pumping plasmamembrane ATPase, and a non-specific impairment of membrane barrier function. (iii) Mitochondria1 function. One further membrane-associated function that has been implicated as a target for imidazole antimycotics is mitochondrial oxidative phosphorylation. In an early study, Dickinson (1977) observed that, at low concentrations, miconazole had an uncoupling effect on rat liver mitochondria; at higher concentrations, more extensive damage occurred with an associated loss of matrix proteins and an inhibition of O2uptake. Inhibition of O2uptake could have resulted from either gross membrane damage, or from a direct interaction with, and inhibition of, respiratory-chain components. Delhez et al. (1977) also reported that the mitochondrial ATPase of Schizosaccharomyces pombe is inhibited by ~ . miconazole with a 50% inhibition of activity (IC,) occurring at 3 0 ~ De Nollin et al. (1977) reported a decrease in the specificactivities of cytochrome c oxidase and peroxidase and an increase in the specific activity of catalase after prolonged incubation of both C. albicans and Sacch. cerevisiae in the presence of miconazole at the minimum fungistatic concentration. When miconazole was present at a fungicidal concentration (10 p),there was complete loss of activity. These authors proposed a possible mode of action involving these enzymes, with cell death resulting from an increased internal concentration of H202. The possibility that changes observed after incubation for 5 hours in the presence of the drug were secondary effects
52
DAMD ICERRIDGE
resulting from cessation of growth was not excluded. Further evidence for involvement of the respiratory chain as a possible primary target for ketoconazole has come from the work of Uno et al. (1982). This drug at a concentration of 9 4 completely ~ ~ inhibited growth of C. albicans strain 7N, whereas at lower concentrations (0.4 to 47 PM) inhibition was incomplete. The growth inhibitory effect was not reversed by addition of ergosterol, and under anaerobic conditions ketoconazole was ineffective as a growth inhibitor. At concentrations as low as 1. O ~ M ,ketoconazole immediately inhibited both endogenous and exogenous (in the presence of glucose) respiration by 20 to 30%. Oxidation of NADH by intact mitochondria isolated from C. albicans was also inhibited by ketoconazole at concentrations as low as 1 . 0 (13%) ~ ~ and 70% inhibition at 0 . 1 ~ In . a subsequent paper, Shigematsu et al. (1982) extended these studies and provided spectrophotometric evidence for a specific and direct interaction of ketoconazole with cytochrome c oxidase and considered that this might be the primary site of action. This result is analogous to that of De Nollin et al. (1977), although these authors considered that miconazole inhibited synthesis rather than functioning of mitochondrial enzymes. Additional evidence for a specific effect on yeast mitochondria has come from studies by Wilm and Stahl(l983) who observed a preferential inhibition by econazole of synthesis of the mitochondrial-membrane enzymes, cytochrome c oxidase and succinate dehydrogenase, but not phenylalanyltRNA synthetase. After prolonged incubation in the presence of radioactive econazole, the highest specific activity .(c.p.m. (mg of protein)-') was found in the mitochondrial fraction. The results were related to the protein contents of the cellular fractions and, given the cellular disorganization that would have occurred during the incubation period, the validity of these findings can be questioned. Kawai et al. (1983) compared the relative effects of a number of imidazole drugs on mitochondrial oxidative phosphorylation, stimulation of latent ATPase and mitochondrial swelling functions, and the differences they observed between clotrimazole, miconazole and econazole they considered resulted from the different degrees of chlorination of the molecules. There may well be other explanations for the differences since, clearly, clotrimazole has quite a different structure from the other two drugs. c. Metabolism of Nucleic Acids. There have been a number of reports on the effects of imidazole drugs on lymphocytes, but two recent ones concerning thymidine uptake are of particular interest (Alford and Cartwright, 1983; Buttke and Chapman, 1983). In human lymphocytes, ketoconazole decreased the uptake of thymidine into cells at concentrations lower than those required to damage the cell membrane. This effect was detected after 10 minutes exposure and Alford and Cartwright (1983) proposed that this
MODE OF ACTION OF ANTIFUNGAL DRUGS
53
decrease in uptake resulted from an effect on the plasma membrane-associated thymidine kinase thereby preventing phosphorylation of thymidine via the thymidine rescue pathway. Similar results were reported by Buttke and Chapman (1983) from their study of the effects of ketoconazole on mitogeninduced DNA synthesis and cholesterol biosynthesis in both human and mouse lymphocytes. The decrease in thymidine incorporation was detected when ketoconazole was added either at the same time as the mitogen or up to 55hours later. Unlike Alford and Cartwright (1983), these authors assumed that thymidine incorporation gave a direct measure of DNA synthesis and did not consider the more likely possibility that, after prolonged exposure to a drug known to interfere with membrane synthesis and function, an earlier stage in thymidine uptake and metabolism was affected. Until this possibility has been excluded, it cannot be assumed that ketoconazole has a direct effect on DNA synthesis in lymphocytes.
d. Morphological Eflects. There have been a number of studies of the effects of imidazole drugs on the cellular morphology of sensitive organisms (see, e.g. Iwata et al., 1973b; De Nollin and Borgers, 1975; Preusser, 1976). Apart from the inhibition of yeast-mycelium transformation in C. albicans, the most significant changes that occur on prolonged incubation in the presence of these drugs at growth inhibitory concentrations are associated with modification to cellular membranes. However, it is not possible from such studies to distinguish between direct effects resulting from interaction of the drugs with membrane lipids or inhibition of sterol demethylase, or indirect effects resulting from autolytic changes occurring after cessation of growth. One effect of imidazole drugs, with clear clinical implications, is that at low concentrations they inhibit the yeast-mycelium transformation in C. albicans (Borgers et al., 1979). Addition of ketoconazole (10 mi) or miconazole (1 p ~ ) to cultures of C. albicans growing in a pseudomycelium-promotingmedium completely inhibits outgrowth of hyphae from the yeast inoculum. The relative efficiency of these two drugs in inhibiting pseudomycelium development in C. albicans is at variance with data obtained with normal growth medium, where miconazole is far more effective in inhibiting growth of the yeast form of the organism. Ketoconazole is effective in inhibiting growth of C. albicans in mixed culture with human fibroblasts and leucocytes (De Brabander et al., 1980; Borgers and Waldron, 1981). Ketoconazole (0.01 pgml-') suppressed growth of C. albicans and completely inhibited mycelial development, but was toxic to mammalian cells only at a concentration of lOOpgml-'. When C. albicans was grown in mixed culture with polymorphonuclear leucocytes and macrophages, the yeasts were rapidly engulfed but not eliminated, largely due to the fact that the engulfed cells produced mycelia which grew out of the cell and which are more resistant to
54
DAMD KERRIDGE
the lytic action of leucocytes. In the presence of ketoconazole, mycelium formation was suppressed and the fungi eliminated. Ketoconazole does not inhibit initiation of hyphal development in C. albicans but affects their subsequent elongation (Aerts et al., 1980; Johnson et al., 1982). Imidazole drugs differ in their effectiveness in decreasing hyphal growth with ketoconazole being more effective than either miconazole or tioconazole (Johnson et al., 1983). Inhibition was time-dependent in that the rate of hyphal extension in the presence of the drug decreased progressively during the course of the incubation. The progressive decrease in hyphal growth rate is consistent with an inhibition of ergosterol biosynthesis with the ergosterol “pool” being exhausted before these drugs exert their effect on hyphal growth. Suppression of mycelium development by ketoconazole may account for the efficiency of this drug in curing patients suffering from candidosis and for the large differences in minimum growth inhibitory concentrations in vivo and in vitro. Minagawa et al. (1983) proposed a simple alternative explanation for the relative efficiences of ketoconazole in vivo and in vitro. It has been known for some time that the minimum growth inhibitory concentrations of imidazole drugs are dependent on the pH value of the growth medium (Holt, 1975; Cope, 1980a) and the values for ketoconazole ranged from 40 pg ml-’ at p H 3 to 0.02pgml-’ at pH 7 (Minagawa et al., 1983). The pH value of mammalian tissues susceptible to C. albicans infections will normally exceed 5, whereas that for many media used to determine drug susceptibilitieswill fall below this value during growth of the yeast, and this may well be the most important factor in determining relative efficiences in vivo and in vitro. The relative efficiences of ketoconazole in vivo and in vitro can be correlated with protonation of the piperazine moiety of the molecule (pK,2.94) where a decrease in protonation is associated with a lowering of the minimum growth inhibitory concentration. The pK, value of the imidazole moiety is 6.51, and it would appear that this group is not important in determining the efficiency of this drug in vivo. Support for the clinical significance of imidazole inhibition of yeastmycelium transformation comes from studies of a strain of C. albicans isolated from a treatment failure. Germ-tube development in serum was equivalent to that in the control strain but, in the presence of ketoconazole (0.5 pg ml-’), development of germ tubes in the control strain was inhibited by approximately 70% over a 6-hour incubation period, whereas that of the resistant strain was inhibited by 10% (Warnock et al., 1983). The molecular basis of this inhibition of the yeast-mycelium transformation is not understood, but it may be a direct effect on synthesis and deposition of structural components of the cell wall or an indirect effect reflecting changes in composition of plasma-membrane lipids induced by inhibition of sterol demethylase. The latter explanation is supported by studies of the effects of
MODE OF ACTION OF ANl"GAL
DRUGS
55
fenpropimorph and imazalil (both inhibitors of sterol biosynthesis) on Penicillium italicurn (Kerkenaar et al., 1984). These compounds at low concentrations induced the production of distorted germ tubes and irregular deposition of (1+3)- and (1+4)-8 polysaccharides and the authors considered this to be a secondary effect of the drugs resulting from interference with sterol biosynthesis. e. Conclusions. Inhibition of sterol demethylase appears to be responsible for the fungistatic action of these compounds, and' direct impairment of membrane integrity responsible for fungicidal effects. The relative importance of these and all of the other observable effects in inhibiting fungal growth, both in the laboratory and in infected animals, will ultimately depend upon the organism, the environment in which it is growing and the specificdrug used. Resolution of this problem requires isolation and characterization of mutant strains of yeast resistant to these drugs. Fortunately for the clinician, the occurrence of strains resistant to these antimycotic drugs is not a problem, nor is it easy for the scientist to obtain drug-resistant strains of C. albicans in the laboratory. Ryley et al. (1984) examined two resistant strains of C. albicans isolated from cases of treatment failure. Resistance for these strains resulted from failure of the drug to penetrate to the sterol demethylase within the cell. The authors assumed that this resulted from failure of the drug to cross the plasma membrane. However, their results did not exclude the possibility that the drug failed to penetrate the cell wall. The mechanism by which imidazole drugs are transported across the plasma membrane is completely unknown, but if mutants blocked in drug transport are available, then this problem should be rapidly resolved. There are considerable phenotypic variations in the sensitivity of C. albicans to imidazoleinduced release of K+ ions (Gale et al., 1980b;Cope, 1980a). As with polyene macrolide antibiotics this resistance develops after cessation of growth, and since protoplasts were sensitive is apparently associated with changes in the cell wall. A similar phenomenon has also been observed by Beggs (1984) who found that the fungicidal, but not the fungistatic, effects of miconazole and ketoconazole were affected by the growth phase, with resistance developing after cessation of growth. In contrast to the earlier studies, it would appear that changes in the plasma membrane rather than the cell wall may be responsible for the development of drug resistance. The involvement of the plasma membrane in resistance to imidazole drugs has also been observed in cultured rat cells (Ikesaki et al., 1984) where it would appear that the cellular level of unsaturated fatty acids is correlated with the sensitivity of these cells to clotrimazole. However, the authors did not exclude the possibility that other membrane constituents may also be involved. Selectivity appears to result, at least in part, from the different affinities of
56
DAVID KERRIDGE
HCI
FIG. 15. Structural formula of naftifine (a) and the metabolic reaction inhibited by it (b).
the host and parasite cytochrome P-450sterol demethylase (Van den Bossche et al., 1984), and isolation and characterization of strains with a modified
demethylase would greatly aid our understanding of the molecular basis of action of these drugs. If inhibition of hyphal extension by these drugs is a direct, rather than an indirect, effect, then it would appear that the primary target in vivo is different from the primary target in vitro. The inhibition of hyphal development could be of prime importance in enabling the host's normal defence mechanism to eliminate the fungus. C. NAFTIFINE
Naftifine is an allylamine derivative and represents a new chemical group of synthetic antifungal compounds (Fig. 15a). It is highly active, both in vitro and in vivo against dermatophytic fungi, with minimum growth inhibitory concentrations between 0.01 and 0.2pgml-', but less active against other pathogenic fungi (Georgopoulos et al., 1981;Petranyi et al., 1981). It has been used topically to treat patients suffering from dermatophytic infections. I . Molecular Basis of Ant&fiungalAction
Like the imidazole antimycotics, naftifine inhibits sterol synthesis in sensitive fungi (Paltauf et al., 1982). Incubation of Candidaparapsilosisin the presence of the drug resulted in accumulation of squalene. There were changes in the lipid composition of the organism including a diminution in cellular ergosterol by 75% after exposure of C. parapsilosis to naftifine at concentrations in excess of the minimum growth inhibitory concentrations for
MODE OF ACTION OF ANTFUNGAL DRUGS
57
15hours. However, these may be secondary changes resulting from inhibition of sterol biosynthesis and cessation of growth. In both intact cells and cell-free systems, inhibition of cellular biosynthesis of membrane sterols resulted from an inhibition of squalene epoxidase (Fig. 15b). Similar results have been obtained by Ryder et af.(1984) for the effect of naftifine on growth and sterol biosynthesis in C. albicans. Morphological changes observed in C. parapsifosisafter exposure to naftifine include accumulation of lipid particles within the cytoplasm, alterations in the plasma membrane and a thickening of the cell wall. These are consistent with an inhibition of membrane sterol synthesis (Meingassner et al., 198I; Meingassner and Sleytr, 1982).
IV. Finale Considerable progress has been made during the past two decades both in the treatment of patients suffering from fungal infections and in our understanding of the modes of action of the clinically important antimycotic drugs. It is, however, quite clear that antifungal therapy lags far behind antibacterial therapy and, given the increase in relative importance of fungal infections, there is need for new and more effective antifungal drugs. There is a need in particular for drugs that can be administered orally to patients suffering from either topical or systematic infections. Development costs are high; Ryley et al. (198 1) have estimated that to put a new antifungal drug on the market costs up to f25,000,000. Since this cost is ultimately recovered from profits, the market for the drug must be a substantial one. The empirical search for antimycotic drugs has been successful in the past and clearly should be continued. In addition, modification of existing drugs have proved particularly successful with the imidazole drugs where we have progressed from the insoluble clotrimazole, suitable only for treating patients suffering from topical infections, to the water-soluble ketoconazole, which can be administered orally and is effective in the therapy of both topical and systemic infections. There is still considerable interest in the development of imidazole and triazole inhibitors of sterol demethylase as therapeutic drugs. Selectivityof these compounds resides, at least in part, in the relative affinities of these compounds for the active site of the cytochrome P-450 sterol demethylases in the host and the fungus, and this might provide a rapid laboratory assessment of the potential clinical value of new derivatives (Van den Bossche et al., 1984). Ketoconazole is a prime example of a compound that is far more effective in vivo than would have been predicted from its action against pathogenic fungi in culture in the laboratory, and it is obvious that, in screening for antifungal drugs, testing in vivo with infected animals must be performed in parallel with studies in vitro.
58
DAVID KERRIDGE
Modification of the other antifungal drugs has not proved SO successful. A large number of griseofulvin derivatives were prepared and their activity was tested against both plant and animal fungal pathogens (Crosse et al., 1964), but so far not one has replaced griseofulvin as the drug of choice in treating patients with recalcitrant dermatophytic infections. Presumably the limited range of sensitive fungi and the lack of toxicity of this drug have not provided the impetus for further development. The polyenes have been extensively modified since the first derivative was synthesized by Lechevalier et al. (1960) and by now a considerable number of derivatives have been synthesized and tested for their therapeutic value (Schaf€ner and Borowski, 1961; Schaffner and Mechlinski, 1972; Bruzzese et al., 1975; Falkowski et al., 1975, 1979; Jarzebski et al., 1982) but, as yet, none has replaced amophotericin B in control of systemic fungal infections. The selectivity and lack of toxicity of 5-fluorocytosine depend upon it being transported by the cytosine permease into sensitive fungi and subsequently deaminated to 5-fluorouracil, and there is little scope here for improvement by chemical modification. Combination therapy with two or more drugs has proved effective in the control of bacterial infections, and a combination of amphotericin B and 5-fluorocytosine has been used successfully to treat patients suffering from systemic fungal infections and is recommended in cases of cryptococcosis (Holt, 1980b). Medoff et al. (1972) provided evidence that potentiation of the effects of 5-fluorocytosine by amphotericin B resulted from polyeneenhanced uptake of the other drug. Conflicting results have been reported by Beggs and Sarosi (1982b), who observed an inhibition by amphotericin B of 5-fluorocytosine uptake into both 5-fluorocytosine-sensitive and -resistant strains of C. albicans, and suggested that these drugs have a sequential effect with amphotericin B acting first to kill the majority of cells and 5-fluorocytosine preventing growth of survivors. The latter explanation for the efficiency of combined therapy seems most probable, although it has been reported that, at concentrations that fail to inhibit growth, amphotericin B accelerated leucine uptake into mouse 3T3 fibroblast cells (Foresti and Amati, 1983). These authors considered that polyene-induced changes in membrane fluidity were responsible for activation of the leucine transport system. This seems to be a reasonable explanation since it is difficult to see how a drug whose primary effect is to render the plasma membrane permeable to protons could enhance uptake of a compound whose transport is energized by a proton gradient. There is also evidence that ketoconazole and 5-fluorocytosine have additive effects in C . albicans, but the biochemical basis of this is not understood (Beggs and Sarosi, 1982a; Odds, 1982). Not all combinations of antifungal drugs show additive or synergistic effects; polyenes and imidazole derivatives are antagonistic both in vivo (Schacter et al., 1976) and in vitro (Dupont and Drouhet, 1979), the latter effect being strongly influenced by the environ-
MODE OF ACTION OF ANTIFUNGAL DRUGS
59
ment. One explanation for this antagonism is that, in the presence of the imidazole antimycotic drug, there is an alteration in the sterol constituents of the plasma membrane and this renders the cells resistant to polyene macrolide antibiotics (Sud and Feingold, 1983). The efficacy of any chemotherapeutic drug depends upon differences between the parasite and its host, and basic research into the comparative biochemistry of the two organisms may provide a guide for development of the next generation of antifungal compounds. Metabolic differences may not be enough and it must be remembered that, in treating patients with fungal infections, we are dealing with a complex system, not merely a culture of a fungus within a test-tube. Factors such as the distribution of the drug within the host’s tissue, and its effect on the normal defense system, may be as important as the specific inhibitory effect on the fungal pathogen. An approach with obvious potential for the future is the development of drugs effective in inhibiting synthesis of the fungal cell wall. The two major structural polysaccharides, chitin and /3-glucan, are absent from man and, by analogy with the bacterial cell wall, might be expected to provide a target for clinically important drugs. There are a number of antifungal antibiotics that inhibit synthesis of these polysaccharides and mannoproteins yet none is used clinically. Two classes of compound are of particular importance as inhibitors of chitin synthesis in fungi; they are the polyoxins and the nikkomycins or neopolyoxins (Isono et al., 1969; Uramoto et al., 1978). These compounds are produced by strains of Streptomyces and their structures are shown in Fig. 16. They are of no clinical importance although C. albicans is inhibited by polyoxin D at millimolar concentrations (Becker et al., 1983). However, they are important in agriculture in Japan where polyoxin D is used in the control of peach black spot (caused by Alternaria kikuchiana). These compounds are potent competitive inhibitors of chitin synthase in vitro with 4 values in many cases lower than the K,,, value for the normal substrate namely UDP-GlcNAc. There are frequently wide discrepancies between the minimum growth inhibitory concentrations for inhibition of fungal growth and the K,values for the inhibition of chitin synthase in vitro (Gooday, 1977), and this is usually associated with failure of the antibiotic to reach its target within the cell. These compounds can be considered as di- or tri-peptide analogues, and are transported into both filamentous fungi (Mitani and Inoue, 1968; Hori et al., 1977) and C. albicans (Mehta et al., 1984) by a peptide transport system. This is another example of the importance of illicit transport of an antifungal drug in mediating its growth-inhibitory effects, and where resistance usually results from failure of the drug to penetrate to its enzyme within the cell rather than a modification of the target. Three tripeptidyl polyoxins have been synthesized by Naider et al. (1983). These
H,NCH o=7HNc$Y I HCR: I HOCH
OH
OH
I
CH,OCNHI I
0
R Polyoxin A
CH20H
Polyoxin D
COOH
H k HO Me
o
R,
R2 OH
HO
OH
HO OH
HO
UDP-N-acetylglucosamine
R2#zf+R
OH
Nikkomycin X
NH,
HO OH
zjCHo floH Me
0
I
I
N
Nikkomycin 2
FIG. 16. Structural formulae of certain polyoxin and neopolyoxin antibiotics.
MODE OF ACTION OF ANTIFUNGAL DRUGS
HO,
OH'
61
,OH
OH
FIG. 17. Structural formulae of echinocandin (a) and papulacandin (b).
compounds inhibited growth of C. albicans but were inactive against chitin synthase in vitro. It would appear that these inactive prodrugs are transported into cells where they are converted by cellular enzymes into an active form. This system may well provide a model for the development of novel antifungal drugs. Glucan synthesis is inhibited by two groups of antibiotics, namely the echinocandins (and related aculeacins; Fig. 17a) and the papulacandins (Fig. 17b). The former group are cyclic peptides and are produced by strains of Aspergillus (Benz et al., 1974; Satoi et al., 1977) and the latter group by Papularia sphaerosperma (Traxler et al., 1977). These antibiotics inhibit /I-glucan synthesis both in vivo and in vitro. Incubation in the presence of these
62
DAVID KERRIDGE
0 II
n = 8.9.10.11.
-
CH,OH
NH - C H I
fi0
FIG. 18. Structural formula of tunicamycin (n can be 8, 9, 10 or 11).
compounds results in rapid lysis of growing cultures of fungi (Baguley et af., 1979; Cassone et al., 1981; Schroder, 1981; Schroder and Kerridge, 1981; Perez et al., 1981, 1983; Varona et al., 1983; Yamaguchi et al., 1982). Cell lysis, which can be prevented by addition of an osmotic stabilizer, results from an imbalance between the /I-glucan synthase and endogeneous glucanase activity, with these antibiotics inhibiting glucan synthesis rather than stimulating the cell wall-associated glucanases. The structure of cell-wall mannoproteins and the sequence of reactions involved in synthesis of the polysaccharide moiety of the molecule are well characterized (Ballou, 1976). Tunicamycin (Fig. 18), an antibiotic produced by Streptomyces liposuperijicans(Takatsuki et al., 1971), is a potent inhibitor of fungal glycoprotein synthesis and it does this by inhibiting formation of dolichol pyrophosphate GlcNAc from UDPGlcNAc and dolichol phosphate (Lehle and Tanner, 1976). Unfortunately is also inhibits similar reactions in both bacteria and mammalian cells and has little potential use as a selective antimycotic antibiotic. Bacitracin A (Fig. 19), one of a number of cyclic peptide antibiotics produced by Bacillus licheniformis and Bacillus subtilis (Johnson et al., 1945), is also a potent inhibitor of glycoprotein synthesis. In Sacch. cerevisiae, it inhibits formation of the dolichol diphosphate chitobiose unit, so resulting in accumulation of the dolichol diphosphate GlcNAc derivative (Reuvers et al., 1978). In bacteria, this antibiotic binds to a Css polyprenol diphosphate inhibiting its hydrolysis to the monophosphate, thus preventing its acting as a carrier in synthesis of cell-wall peptidoglycan (Stone and Strominger, 1971). Unfortunately, none of the antibiotics inhibiting synthesis of the fungal wall polysaccharides is effective in vivo. These
MODE OF ACTION OF A N " G A L
(L-CYS) 2
63
(~-1Ie) I
C=O
D-Phe 10
1
11 L-lle
L-His 9
1
12 D-Om
t
L-Aso8
3 L-Leu
4 D-Glu
-
~-1le 5 +
DRUGS
signifies a C
+
l
D Am
L-LYS
7
6-
N bond
FIG. 19. Structural formula of bacitracin.
+
signifies a C-rN bond.
compounds have been far more useful as molecular probes in studies of cell-wall biosynthesis than as therapeutic drugs. Selective toxicity can be considered as one specific topic in comparative biochemistry. It is unfortunate that so far studies on the molecular basis of action of antimycotic drugs have been far more important in enhancing our knowledge of comparative biochemistry than studies on comparative biochemistry have been in enabling us to design new and effective drugs. Studies on the mode of action of antimycotic compounds will not in themselves enable us to develop clinically useful drugs. Basic research must be coordinated with an investigation of the interaction of the parasite with its host, the effect of the drug on the hosts metabolism and the distribution of the drug within the hosts tissues. The research reviewed in this article, has, I hope, outlined current knowledge of the mode of action of the clinically important compounds and indicated the areas where future developments might be expected to occur.
V. Acknowledgements I would like to thank the Medical Research Council for financial support and my colleagues W.L. Whelan, K.G. Simpkin, R.O. Nicholas and R.D. Cannon for many helpful discussions during the preparation of this article and for critically reading the manuscript.
64
DAVID KERRIDGE
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Microbial Cyanide Metabolism CHRISTOPHER J. KNOWLES and ALAN W.BUNCH Biological Laboratory, University of Kent, Canterbury, Kent CT2 7NJ,UK
I. Introduction
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11. Bacterial cyanide production
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Cyanide-producing species . . Conditions of cyanide production. . Pathways of cyanide formation . . Cyanide degradation by Chromobacteriwn violaceum . . . E. Relationship of cyanogenesis to primary metabolism III. Fungal cyanogenesis . . A. Species producing cyanide . . . B. Cyanide-linked plant diseases . . . C. Cyanogenesis by pure cultures of fungi . . . D. Cyanide degradation by cyanogenic fungi . . IV. Cyanogenesis by photosynthetic micro-organisms. A. Species that form cyanide . . . . . B. Pathways for the production of cyanide. . C. The role of cyanide production in photosynthetic micro-organisms V. Cyanide degradation . . . A. Cyanide degradation by phytopathogenic fungi . B. Cyanide resistance and degradation by bacteria . VI. Concluding remarks . References . . . . . . . .
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I. Introduction
It is the aim of this article to review cyanide metabolism in micro-organisms. This is a surprisingly diverse subject and it is necessary to limit the topics covered. In particular the metabolism of organic cyanides (nitriles, RCN) will not be discussed except where relevant to the metabolism of inorganic cyanide (HCN). Nitrile metabolism has been reviewed recently by Chamberlain and Mackenzie (1981) and Jallageas et al. (1980). Cyanide can be detoxified by conversion into thiocyanate by rhodanese. There is little ADVANCES IN MICROBIAL PHYSIOLOGY, VOL27 ISBN 0-12427727-1
Copyright 0 1986 by Academic Press London All rights of reproduction in MY form reserved
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evidence, at least in micro-organisms, that this is the intracellular role of rhodanese, whose primary function appears to involve sulphane sulphur transformations. Cyanide detoxification by rhodanese is thus an accessory function of the enzyme (Volini and Alexander, 1981). Rhodanese has been reviewed by Westley (1980, 1981) and will not be further discussed here. The term “cyanide” will be used loosely and refers to both the cyanide anion (CN-) and undissociated hydrogen cyanide (HCN). The pKa of cyanide is 9.3; it is therefore present largely as HCN at physiological pH values. HCN is volatile (boiling point 26°C) and is less dense than air. Hence cyanide formed by microbial cultures will be rapidly lost to the environment. In addition, cyanide is highly reactive and complexes tightly to metals such as nickel, copper, zinc, iron and gold (Towill et al., 1978). In this article it will be assumed that cyanide is supplied to microbial cultures as KCN or NaCN unless otherwise stated. Cyanide also reacts reversibly with keto groups to form cyanohydrin derivatives. As a result of its reactivity, many metallo and other enzymes are strongly inhibited by cyanide. It is therefore often used as a metabolic inhibitor (Solomonson, 198l), especially of cytochrome oxidases (Henry, 1981; Palmer, 1981). The role of cyanide as an inhibitor will not be covered in this review. This is because in inhibitor studies cyanide does not function as a metabolite except in certain instances where it is formed by cyanogenic organisms, for example where it inhibits respiration (Niven et al., 1975) or nitrate reductase (Solomonson and ‘Spehar, 1981; see below). Microbial cyanide metabolism has been reviewed previously be Knowles (1976) and summarized by Vennesland et al. (1982). Algal and bacterial cyanide formation (cyanogenesis) have been reviewed by Vennesland et al. (1981b) and Castric (198 l), respectively. The book “Cyanide in Biology” (Vennesland et al., 1981b) is an invaluable source of information on other aspects of cyanide metabolism.
II. Bacterial Cyanide Production A.
CYANIDE-PRODUCING SPECIES
Cyanide formation (cyanogenesis) by bacteria was first noted before World War I (Emerson et al., 1913; Clawson and Young, 1913). Subsequently there have been a series of surveys to detect cyanogenic species (Patty, 1921; Lorck, 1948; Sneath, 1956,1960; Bugel and Muller, 1963;Michaels and Corpe, 1965; Goldfarb and Margraf, 1967; Castric, 1975; Freeman et al., 1975, 1976; Askeland and Morrison, 1983). These surveys have shown that Chromobacteriwn violacewn, many but not all strains of Pseudomonas aeruginosa and Pseudomonas Jluorescens, and a few strains of some other Pseudomonas
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species ( P . chloraphis, P. aureofaciens) are cyanogenic, whereas a wide range of other bacteria do not form cyanide. Early tests for cyanide, often by the picrate method, were insensitive and it is probable that many cyanogenic organisms were missed. Recently, more sensitive and specific assay systems have been developed (Freeman et al., 1975, 1976; Castric and Castric, 1983), as has a rapid detection system (Castric and Castric, 1983). It has been suggested that cyanide production by P. ueruginosa,in patients suffering massive septaecemia from infections of severe burn wounds, may be intimately related to the high mortality caused by this bacterium (Contreras et al., 1963; Goldfarb and Margraf, 1967). B. CONDITIONS OF CYANIDE PRODUCTION
Lorck (1948) was the first to show that glycine stimulates cyanide formation by growing bacteria. This was confirmed by Wissing (1968) for an unidentified Pseudornonas species. Michaels and Corpe (1965) showed that optimal cyanogenesis by C. violaceurn growing on glutamate as the source of carbon and nitrogen required the addition of both glycine and methionine to the growth medium. Similar results were obtained by Castric (1977) for glutamate-grown P. aeruginosa. When glycine was provided as the sole source of nitrogen for C. violacewn, growing with glucose as the carbon source and in the presence of methionine (which was unable to act as a nitrogen source), there was little formation of cyanide. In the presence of ammonia, cyanogenesis was enhanced (Collins et al., 1980). Glycine can be replaced by threonine, but not by serine, for optimal cyanide production by cultures of P. ueruginosu,or by cultures or suspensions of C. violaceurn (Collins et d.,1980). This is probably because threonine is converted into glycine without formation of a C-1 unit, whereas serine is converted, via the action of serine hydroxymethyltransferase, into glycine plus a C-1 unit (methylene tetrahydrofolate, THF), the latter being a precursor of methionine. Radiolabellingstudies with P.ueruginosahave shown that cyanide is formed from glycine, serine or threonine (in the presence of methionine) but that cyanide is formed preferentially from glycine when both glycine and threonine or serine are present (Castric, 1977). In the presence of glycine, L-methionine can be replaced by its metabolites S-adenosyl-L-methionine or cystathionhe, or by D-methionine or DLmethionine sulphoxide for optimal cyanogenesis by cultures of C. violaceurn (Collins et al., 1980). Cyanide is produced by P . ueruginosa for a brief period at the end of the exponential growth phase and the start of the stationary phase (Castric, 1975; Askeland and Morrison, 1983). The ability of suspensions of bacteria, harvested at various times after inoculation, to form cyanide from glycine has
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CHRISTOPHER J. KNOWLES AND ALAN W. BUNCH
been termed the cyanogenic capacity by Castric et al. (1979). Cyanogenic capacity of bacteria is low in the early and mid-exponential growth phases, and is izlduced in the late-exponentialphase before the appearance of cyanide. It is then rapidly degraded by the bacteria, to almost zero at the time of maximal cyanide content of the cultures. Measurements of the ability of cell-free extracts to convert glycine into cyanide by the cyanide synthase enzyme system showed a corresponding temporal pattern of induction and loss of activity (Castric et al., 1979). It seems likely that the loss of cyanide synthase activity is due to autodestruction of the enzyme rather than the appearance of a specific metabolite or an inducible protease (P. A. Castric et al., 1981). A similar sequence of events is seen for C. violaceum (Rodgers and Knowles, 1978; Bunch and Knowles, 1982) but, in addition, the cyanide, once produced, is rapidly degraded by the bacteria. During the period that cyanide is lost, fi-cyanoalanine accumulates in the medium (Rodgers, 1981, 1982; Macadam and Knowles, 1984). Beta-Cyanoalanine is formed from cyanide and cysteine or 0-acetylserine by fi-cyanoalaninesynthase, which is present at low activity during the exponential growth phase and which is induced to a maximum value at the time the cyanide content of the cultures is greatest (Macadam and Knowles, 1984). The formation of cyanide for a brief period at the end of the exponential growth phase is typical of secondary metabolism (Drew and Demain, 1977; Demain et al., 1979; Malik, 1980; Hopwood, 1981), as is the requirement for a primary metabolic precursor (glycine) plus an activator (methionine). Cyanogenesis is also typical of secondary metabolism in that the conditions of synthesis are narrower than those required for growth (Weinberg, 1971, 1978). Cyanide formation by P. aeruginosa is optimal at 34°C to 37"C, whereas the growth yield is unaffected by growth at 25°C to 40°C (Castric, 1975). Cyanogenesis by C. violaceum is less sensitiveto temperature (Rodgers and Knowles, 1978). Variations in the contents of iron (FeSO, or FeC1,) and phosphate in the medium, at concentrations that do not alter the growth rate or growth yield, affect the level of cyanogenesis by P. aeruginosa (Castric, 1975; Meganathan and Castric, 1977), C. violaceurn (Rodgers and Knowles, 1978), P. aeruginosa and P.fIuarescens (Askeland and Morrison, 1983). In each case, increases in concentrations of iron or phosphate stimulate cyanide formation, although phosphate concentrations above 1 0 m also inhibit cyanogenesis by P.aerugmosa (Castric, 1975). Formation of cyanide synthase in P. aeruginosa is repressed by growth in media containing only low concentrations of iron and phosphate (Castric et al., 1979). Growth of either C. violaceum or P. aeruginosa on glutamate alone (conditions of low cyanogenesis), as well as on glutamate in the presence of glycine and methionine (high cyanogenesis), results in the formation of
MICROBIAL CYANIDE METABOLISM
77
cyanide synthase activity (Castric et al., 1979; Bunch and Knowles, 1982). Indeed there is slightly more synthase activity formed under the former growth conditions, indicating that exogenous glycine and methionine are not required for induction of the enzyme. However, the intracellular concentration of glycine could be the trigger, since P. A. Castric et al. (1981) have shown that its intracellular concentration increases during growth and reaches a peak at the time cyanide synthase is most active. The point at which cyanide production is maximal in batch cultures of P. aeruginosa is coincident with the time of minimal O2content of the medium. Once cultures enter the stationary phase, O2 content of the medium rises rapidly, presumably due to a decrease in the rate of respiration (P. A. Castric et al., 1981; Castric, 1983). This is coincident with termination of cyanogenesis and disappearance of cyanide synthase. Interestingly, imposition of anaerobic conditions at this time prevents loss of cyanide synthase activity. K. F. Castric et al. (1981) have shown that the yield of cyanide is related to the O2concentration of the medium: too high or too low a concentration of O2at the time of induction of cyanogenesis results in lower yields of cyanide. There is not an obligate requirement for O2 for cyanogenesis by C. violaceum (Nazly et al., 1981). Anaerobic growth in a medium containing glucose, fumarate (as an electron acceptor to permit growth to occur) and NH,Cl, supplemented with glycine and methionine, resulted in significant formation of cyanide. Wissing (1974) showed that, for cyanogenesis by suspensions of a Pseudomonas species, O2could be replaced by the electron acceptors phenazine methosulphate (PMS), methylene blue or 2,6-dichlorophenolindophenol (DCIP). Similarly, low levels of cyanogenesis occurred with suspensions of aerobically-grown C. violaceum incubated anaerobically in the presence of glycine and PMS, DCIP or fumarate (Nazly et al., 1981). C. PATHWAYS OF CYANIDE FORMATION
Studies using [ l-'4C]glycine, [2-14C]glycine,["Nlglycine and [methyl-14C]methionine have shown that cyanide and CO, are formed by oxidative decarboxylationof glycine in C. violaceum (Michaels et al., 1965; Brysk et al., 1969; Bunch and Knowles, 1982):
o v NH,CH,COOH
--
o
HCN
v
+ CO, + 4[H]
Cyanide was also shown to be derived from the methylene carbon of glycine by P. fluorescens strain S and P. aeruginosa strain 1-73 (Askeland and Morrison, 1983). Castric (1977) reported that the cyanide carbon was derived from both the methylene (63%) and carboxyl(36%) groups of glycine, by P. aeruginosa strain 9-DZ. This could be due to randomization of the C-1 and C-2 carbons of glycine either via formation of a symmetrical intermediate in
78
CHRISTOPHER J. KNOWLES AND ALAN
W. BUNCH
the cyanogenic pathway or by reversible utilization of glycine in primary metabolism, for example by incorporation into a purine and degradation to glyoxylate followed by transamination (Castric, 1981) or by formation of a symmetrical metabolite such as oxalate (Bunch and Knowles, 1981). It is not known what intermediates are involved in the conversion of glycine into cyanide and whether the “cyanide synthase” system consists of one or more enzymes. A start has been made on characterization of the enzyme(s)involved, but this work has been hampered by the sensitivity of the synthase system to 02. Wissing (1975, 1983) and Wissing and Andersen (1981) have shown that extracts of a Pseudomonas species can form cyanide from glycine using PMS as an electron acceptor. The activity is present in the membrane fraction but can be solubilized by Triton X-100. It is activated by dithiothreitol and is sensitive to 0,. P. A. Castric et al. (1981) have obtained active cell-free extracts of P. aeuroginosa that produce cyanide from glycine when incubated with glycine either aerobically, or anaerobically in the presence of PMS, DCIP or ferricyanide. The enzyme is inactivated by 02,but is stabilized against O2 toxicity by glycine and partially by dithiothreitol. Bunch and Knowles (1982) found that cyanide synthase of C. violacewn is located primarily in the particulate fraction but can be solubilized by detergents. Only a low level of cyanide formation from glycine occurs with extracts incubated aerobically, but there is considerable activity in the presence of PMS, DCIP or phenazine ethosulphate when incubated either aerobically or anaerobically. NAD+, NADP+, FMN, FAD and a range of other redox dyes are ineffective as electron acceptors. Oxygen is toxic to the synthase but glycine provides partial protection. Extracts are active only if they are isolated and maintained in the presence of dithiothreitol and stored under an atmosphere of N,. It is possible to conclude tentatively that in vivo the cyanide synthase system is similar in each of these bacteria and is membrane-bound. The natural electron acceptor for the process is probably a component of the respiratory system rather than O2itself. In vitro, the natural acceptor can be replaced by PMS and DCIP. K. F. Castric et al. (1981) suggested that the decline in O2content of the medium and the rise in intracellular glycine concentration at the end of the exponential-growth phase in batch cultures could be a signal to the cell to synthesize the synthase and begin cyanide production. The requirement to remove glycine is aggravated by provision of exogeneous glycine and methionine, resulting in greater formation of cyanide. The subsequent rise in the 0,concentration of the medium as respiration decreases, possibly due to partial inhibition by cyanide, and the decrease in glycine concentration (via conversion into cyanide) in the early stationary phase destabilizes the
79
MICROBIAL CYANIDE METABOLISM
synthase resulting in its selective destruction. In addition, the potentially toxic cyanide is further metabolized by C . violaceum to 8-cyanoalanine (see Section 1I.D). Several proposals have been made for possible intermediates involved in conversion of glycine into cyanide (cf. Knowles, 1976; Wissing and Anderson, 1981; Castric, 1981). The first of these potential pathways relates to the intermediates found in the well established plant cyanogenicglycoside system (Vennesland et al., 1982; Conn, 1981). Depending on the plant species, a range of amino acids are converted into cyanogenic glycosidesby the following pathway: Amino acid
-
N-Hydroxyamino acid
Cyanogenic glycoside *
+ducose
-I -
co2
a-Hydroxynitrile
Aldoxime
Nitrile
On injury or death, cyanogenic glycosides are exposed to a glycosidase and an hydroxynitrilase, which in healthy plants are in a different cellular compartment or tissue, resulting in release of cyanide: Cyanogenic glycoside
-Blucosc+
-
a-Hydroxynitrile
Cyanide + aldehyde
Some plants, as well as C. violacewn, are able to form p-cyanoalanine from cyanide and cysteine (see Section 1I.D). By analogy, it is possible that glycine is converted into cyanide by bacteria as shown in Fig. l(a). Alternatively, decarboxylationcould occur after oxime formation (Fig. lb). Wissing (1974) has suggested that an unstable imine could be formed. Cyanoformic acid also could be an intermediate (Knowles, 1976). This route is shown in Fig. l(c). This proposal is given weight in that amino acid oxidases probably function via formation of imines (Hafner and Wellner, 1971). Michaels et al. (1965) suggested that oxamic acid could be an intermediate (Fig. Id). As yet there is no evidence for any of these routes. Formation of cyanide from oxamic acid, N-hydroxyglycine, formaldoxime or glyoxylic acid oxime did not occur with extracts of C . violaceum, but N-hydroxyglycine was an inhibitor of cyanogenesis from glycine (Bunch and Knowles, 1982). Wissing and Andersen (1981) did not find any cyanogenesis from oxamic acid or formaldoxime by extracts of a Pseudomonas species. D. CYANLDE DEGRADATION BY
Chromobacterium violacewn
Chromobaterium violaceum is known to synthesize at least three enzymes capable of metabolizing cyanide.
80
W. BUNCH
CHRISTOPHER J. KNOWLBS AND ALAN
(a)
H,C
CH,
-H2C
\
\
NH,
* HCN 7R.OH
NH.OH
CO,
N-Hydroxyglycine
H,C
\
-H,C
-
CH
\
NH,
Formaldoxime
1 I
NH.OH
N.OH
1-[
N-Hydroxyglycine
(C)
"O 'H .[H,C
\
? H / COOH NH,
H,C
COOH ]-HCN
NH
+ CO,
N
/COOH \
-0c
a,
Glyoxylic acid oxime
Iminoacetic acid
(4
--rHCN
\ NH,
Cyanoformic acid
-[
(c*H]-HCN
+ CO,
NH2 Oxamic acid
Cyanoformic acid
FIG. 1. Potential pathways for bacterial cyanogenesis from glycine.
(a) By the action of rhodanese (Rodgers and Knowles, 1978): S,Oi-
+ CN-
-
SO:-
+ SCN-
(b) By the action of y-cyano-a-aminobutyric acid synthase (Brysk and Ressler, 1970; Ressler et af., 1973). The overall process is: Homocystine
+
2CN-
-
y-Cyanoaminobutyrate
+ SCN- + homocysteine
This probably takes place via a non-enzymic step: Homocysthe + C N -
Homocysteine
followed by an enzymic step: y-Thiocyanoaminobutyrate + CN-
-
+ y-thiocyanoaminobutyrate
SCN-
+ ycyanoaminobutyrate
81
MICROBIAL CYANIDE METABOLISM
(c) By the action of 8-cyanoalanine synthase (Brysk et al., 1969; Rodgers and Knowles, 1978; Rodgers 1981, 1982; Macadam and Knowles, 1984): CN-
-
+ cysteine (or o-acetylserine)
&Cyanoalanine
+ H2S(or acetate)
All three enzymes have been detected in C. violaceum (Rodgers and Knowles, 1978), and their concentrations increase in the post-cyanogenic period. Incubation of washed, resuspended, early exponential phase C. violaceum cells with cyanide and glycine under conditions of low aeration results in formation of y-cyanoaminobutyrate and p-cyanoalanine (Brysk and Ressler, 1970). However, rhodanese and y-cyanoaminobutyrate synthase probably play only a minor role, if any, in the further metabolism of cyanide in growing cultures of C. violaceum. The reactions catalysed by both enzymes result in formation of thiocyanate. In cultures of C. violaceum, no build-up of thiocyanate has been detected (Rodgers, 1981, 1982; Macadam and Knowles, 1984). In cultures of C. violaceum, j?-cyanoalanine accumulates in appreciable quantities in the medium (Table 1) in the period when the cyanide concentration is decreasing (Rodgers, 1981; Macadam and Knowles, 1984). The buildup of jkyanoalanine is particularly noteworthy in bacteria grown under conditions of high cyanogenesis. Suspensions of harvested C. violaceum cells are also able to form B-cyanoalanine, when incubated with cyanide and serine (Brysk et al., 1969; Rodgers, 1981, 1982). In these experiments, some of the /3-cyanoalanine was metabolized to aspartate. Beta-Cyanoalaninesynthase from C. violaceum has been purified to homogeneity (Macadam and Knowles, 1984). The physicochemical properties and TABLE 1. Formation of fl-cyanoalanine by cultures of Chromobacteriwn violacewn
Additions to the growth
Maximum cyanide content
01M)
Maximum fl-cyanoalanine content 01M)
-
20
60
GIycine Methionine Glycine + methionine
80
170 140 370
70 230
Chromobacteriwn violaceum was grown in a minimal salts medium with 10 mM glutamateas the sole carbon, nitrogen and energy source (Macadam and Knowles, 1984). The medium was supplemented with 2 m glycine ~ and/or 0.5 m~ methionine, as indicated above. Growth terminated 10 to 12
hours after inoculation due to glutamate depletion. The cyanide content of the medium was maximal 12 to 13 hours after inoculation and then declined. The fl-cyanoalanine content was maximal 18 to 24 hours after inoculation.
82
CHRISTOPHER J. KNOWLES AND ALAN W. BUNCH
substrate specificities of enzymes having 8-cyanoalanine synthase activity are shown in Tables 2 and 3. The properties of 8-cyanoalanine synthase from C. violaceurn are very similar to those of cysteine synthase or serine sulphydrylase from other bacteria. However, 8-cyanoalanine synthase of C. violaceurn does not exhibit these activities. It is thus a true 8-cyanoalanine synthase, like the mitochondria1 enzyme of Blue Lupine. Plant and C. violaceurn 8-cyanoalanine synthases have different physicochemical properties. Beta-Cyanoalanine synthase from C. violaceurn catalyses 8-cyanoalanine formation from cysteine or 0-acetylserine, but has little activity with serine or cystine (Macadam and Knowles, 1984). Cyanide can be replaced as the co-substrate by a range of thiols. Methionine is a non-competitive inhibitor. E. RELATIONSHIP OF CYANOGENESIS TO PRIMARY METABOLISM
The reasons for production of secondary metabolites by micro-organisms have not been resolved (Drew and Demain, 1977; Demain et al., 1979; Malik, 1980; Hopwood, 1981). It is generally supposed that it is the process of secondary metabolism rather than the products that is important. At times of metabolic stress, when growth slows down, the demands for protein synthesis, RNA synthesis and turnover, DNA replication and energy metabolism, for example, decrease at different rates. There is thus a potential build-up of primary metabolites, particularly at metabolic branch points where efficient regulation to satisfy varying demands for different metabolites is difficult to attain. Secondary metabolism could be a method for removal of such primary metabolites. If we are to comprehend fully secondary metabolism, we must focus considerable attention on regulation of the preceding primary metabolic pathways. We know little so far about the primary metabolic pathways leading to bacterial synthesis of cyanide and 8-cyanoalanine. As mentioned earlier, K. F. Castric et af. (1981) suggested that cyanogenesis is a response to a build-up of the intracellular glycine concentration. This process is clearly aggravated by addition of glycine and methionine to the growth medium. Figure 2 shows the relevant primary and secondary metabolic pathways. The key primary metabolic enzymes are serine hydroxymethyltransferase and glycine cleavage enzyme which catalyse conversion of serine into glycine and glycine into C 0 2 and ammonia, respectively. In each case the extra C-1 unit is conserved as methylene tetrahydrofolate, and is used for synthesis of purines, histidine, thymidine and, especially, methionine. Serine and glycine are incorporated into proteins. In addition, glycine is used for purine synthesis. Dev and Harvey (1982) have elegantly shown, in Escherichia coli growing exponentially, that the demand for C-1 units is greater than the requirement
TABLE 2. Physicochemical properties of enzymes having /3-cyanoalanine synthase activity from plants and bacteria Properties
Species
Plants Blue lupine Blue lupine White lupine Lotus tenub BaEteria
Optimum pH Molecular References value weight 1 2,3 4 5
9.5 8.8 9.5 9.0
Salmonella typhimuriunf
6.7
Escherichia colt Bacillus m e g a t e r i d
11 8
9.0 -
9 10
Enterobucter species 10-lb Chromobacterium violaceum
Subunit molecular weight
53,000 52,000 50,420 -
(no subunits) -
68,000
34,000 x 2
9.1510.0
60,000 to 70,000 68,000
34,000 x 2
9.15
71,067
35,057 x 2
Pyridoxal phosphate Spectral Ratio of A,, :A,,, content Isoelectric peak (mol-I) point (PI) (nm) (A,,: A4,,) 4.7 4.57 -
Effect of acetylserine on spectrum
405
410 -
2.1
3.5
412nm peak shifts to 470 nm
412nm peak shifts to 470 nm 410nm peak decreases
-
412
1.5
5.0
412
3.3
1.7
4.94
410
3.6
405
"No jcyanoalanine activity detected; it has only cysteine synthase activity. bPrincipaUy cysteine synthase activity. References: 1, Hendrickson and Conn (1969); 2, Akopyan et al. (1975); 3, Akopyan and Goryachenkova (1976); 4, Galoyan et ul. (1981); 5, Floss et al. (1965); 6, Kredich and Tomkins (1966); 7, Becker et ul. (1969); 8, Castric and Conn (1971); 9, Yanese el al. (1982a, b); 10, Macadam and Knowles (1984); 11, Dunnill and Fowden (1965).
TABLE 3. Substrate specifities for enzymes having p-cyanoalanine synthase and cysteine synthase activities Activity (%) p-Cyanoalanine synthase (+ KCN) Cysteine synthase (+H2S) References
Plants Blue lupine (mitochondria1extract) (soluble fraction) Blue lupine (mitochondria1 extract) White lupine Lotus tenuis Vicia sativa
1
2,3 4 5 12
Cysteine 100 0.51 100 100
0-Acetyl-L-serine 5.4
10.2 0
-
Serine
Serine
0
4.1
-
100
-
0
-
4 100
100 0
Bacteria Escherichia colf Escherichia coli Bacillus megateriwn" Enterobacter species 10-1 Salmonella typhimuriwn Chromobacteriwn vwlaceum
11 6 8 13 6,7 10
50 -
0.04 0
-
100
0-Acetyl-L-serine
0.05 0.4
-
84
30 -
100
-
0
-
-
0
100 100 100
5.2
0
100 0
0.002 0
'Recalculated from data provided in the papers. References: As given in Table 2; 12, Nigam and Ressler (1964); 13, Yanese et al. (1982~).
-
-
85
MICROBIAL CYANIDE METABOLISM
0-Acetylserine
Intermediary metabolism
-- t
I serine
Cysteine
WyanoPIPninc
-
I
Glycine
HCN
+ CO,
Histidine Thymidine Methionine
FIG. 2. Primary metabolicpathways related to the secondary metabolic pathways for cyanide and &cyanoalanine formation.
for glycine. Therefore extra glycine has to be synthesized such that it can be utilized by the glycine cleavage enzyme to provide the extra C-1 units. Serine hydroxymethyltransferase is synthesized constitutively by C. violuceum,but glycine cleavage enzyme has been detected at only an extremely low concentration (Beechey and Knowles, 1984). It is possible that excess glycine has to be formed to satisfy the requirement for C-1 units during growth. Serine hydroxymethyltransferase is constitutive and competitivelyinhibited by glycine and cysteine. If the supposition that the function of secondary metabolism is to remove one or more primary metabolites that have built up during conditions of metabolic stress is correct, then it might be expected that maintenance of bacteria under conditions that enhance production of secondary metabolites would prolong the period of their survival. In some instances, this appears to be the case (Weinberg and Goodnight, 1970; Gentry et ul., 1971; Smith et ul., 1974). In the case of cyanogenesis by C. violuceum,the effect of cyanogenesis on the length of viability is more difficult to categorize as advantageous or detrimental (Macadam and Knowles, 1983). Cultures grown on glutamate alone (low cyanogenesis) survived much longer than those grown on glutamate plus glycine and methionine (high cyanogenesis). However, since cyanogenesis is likely to be a response to the need to remove glycine and/or methionine, the addition of these amino acids to the growth medium probably aggravates this situation and, despite enhanced cyanogenesis, accelerated death occurs. Cyanogenesis also can be repressed by lowering the medium phosphate and iron content. Under these conditions survival was, unexpectedly, also shortened (Macadam and Knowles, 1983).
86
CHRISTOPHER J. KNOWLES AND ALAN
W. BUNCH
III. Fungal Cyanogenesis A. SPECIES PRODUCING CYANIDE
Cyanide production by the fungus Marusmius oreades was the first report of microbial cyanogenesis (Von Losecke, 1871). Since then it has been claimed that cyanogenesis is a widespread phenomenon within the class Basidiomycetes where members of at least 52 genera produce cyanide (Locquin, 1944; Bach, 1956).These, and more recent, studies have also shown members of the Ascomycetes and Zygomycetes to be cyanogenic (Bach, 1956; Singer, 1975; Hutchinson, 1973; Saupe, 1981). However, there are some doubts about the accuracy of some of the assay systems used to detect cyanide production (Bach, 1956). This is particularly true when samples are incubated for several hours and the cyanide produced is detected by a sensitive assay system. Under these conditions, positive results can be due to contamination of samples by cyanogenic bacteria (Saupe, 1981). As a result, there are only a few examples where cyanogenesis has been used as a taxonomic aid (e.g. Singer, 1975). This is in contrast to higher plant cyanogenesis which has successfullybeen employed as a useful chemical taxonomic characteristic (Hegnauer, 1977; Saupe, 1981). B. CYANIDE-LINKED PLANT DISEASES
It has been shown that several plant diseases involving fungi progress with the liberation of cyanide in the host plant tissues (Vennesland et ul., 1982; Conn, 1981). Winter crown rot or snow mould disease is the best studied cyanide-linked disease. The fungus involved is a psychrophilic basidiomycete which can attack a wide range of forage plants including Medicago sutivu (Cormack, 1948).Ward et ul. (1961) obtained three isolates of this fungus. Isolate A was highly pathogenic to Medicago sutivu, whereas isolate B exhibited only moderate pathogenicity. Most of the cyanide produced during the invasion of Medicago sutivu originates from the host plant cyanogenic glycosides (Colotelo and Ward, 1961). Cyanide is released, after conversion of these compounds into a-hydroxynitriles, by a B-glucosidase produced by the fungus. Isolate B of the fungus has been shown to possess an oxynitrilaseenzyme (Stevens and Strobel, 1968), although cyanide release from a-hydroxynitriles can proceed non-enzymically. Type B isolates also synthesize cyanide from their own cyanogenic compounds (see Section 1II.C). Cyanide made in this way does not appear to be crucial for development of the disease in Medicago sutivu, but may be important when other plants are attacked by this fungus (Ward et al., 1961).
MICROBIAL CYANIDE METABOLISM
87
Isolate C of the psychrophilic fungus was not pathogenic to any of the plants tested, nor did it synthesize cyanide. Other cyanide-linked diseases also develop in a similar way. For example, copperspot disease of Lotus corniculatus is caused by the fungus Stemphylium loti. Cyanide is released following the action of a fungal 8-glucosidase on the cyanogenic glumsides of the host plant (Millar and Higgins, 1970). This fungus has not been reported to synthesize cyanide from its own cyanogenic compounds. The fungus that causes Fairy Ring disease of grasslands and parks, M. oreades, can itself form cyanide (Lebeau and Hawn, 1963). Although a fungal 8-glucosidase has also been implicated in the development of Fairy Ring disease, it is likely that cyanide originating from the fungus is important (Filer, 1965, 1966). C. CYANOGENESIS BY PURE CULTURES OF FUNGI
Many fungi are able to synthesize cyanide. As discussed above, this ability may be important for development of some cyanide-linked diseases. However, much less is known about the mechanism and physiology of cyanogenesis in fungi compared with bacterial or algal cyanogenesis. This is undoubtedly due to the difficulty of growing and handling the fungi. Two fungi, isolate B of the snow mould fungus and M. oreades Fr., have provided most of the information available on fungal cyanogenesis. Thus, for the time being, it can be only assumed that other fungal cyanogenic systems are similar. 1. Possible Intermediates for the Production of Cyanide
Ward and Thorn (1966) have shown that cyanide production by cultures of isolate B of the snow mould fungus is stimulated by addition of glycine to the culture medium. Using radiolabelled glycine it was demonstrated that the nitrogen and C-2 atoms are converted into cyanide without breaking the carbon-nitrogen bond (Ward et al., 1977). The C-1 atom of glycine is mainly converted into CO, (Bunch and Knowles, 1980). Serine and aspartic acid also can act as precursors of cyanide, although much less efficiently (Ward et al., 1977). Production of cyanide from serine presumably occurs after its conversion into glycine by serine hydroxymethyltransferase. It is possible that aspartic acid is also metabolized to cyanide via glycine by a more complicated route. However, Ward et al. (1977) proposed that aspartate may be converted into cyanide via glyoxylic acid cyanohydrin (see below). The small amounts of cyanide originating from aspartate, under the experimental conditions employed, would indicate that this pathway, if it exists, is of little significance compared to cyanide production from glycine.
88
CHRISTOPHER J. KNOWLFS AND ALAN W. BUNCH
Ward (1964) reported that isolate B of the snow mould fungus produces as unstable cyanogen in cultures grown in the presence of high glucose concentrations. A similar substance also was detected in cultures of M. oreades grown under the same conditions (Ward, 1964; Ward et al., 1971). This compound became labelled when [2-'4C]glycinewas given to isolate B of the snow mould fungus. Although the same amount of the cyanogen was produced when [1-'4C]glycinewas used, it was not labelled (Ward and Thorn, 1966). Tapper and MacDonald (1974) have shown that the compound is probably glyoxylic acid cyanohydrin, although they also detected smaller amounts of a second cyanogenic compound, possibly pyruvic acid cyanohydrin: OH
1
H-C-COOH
I
CN Glyoxylic acid cyanohydrin
OH
I
H, C - C - C O O H
I
CN m v i c acid cyanohydrin
Bunch and Knowles (1 980) found only traces of these compounds when lower concentrations of glucose were used in culture media. It is possible therefore that these compounds are formed by the reaction of cyanide with fermentation products which would be less abundant under the conditions used by the latter workers. This remains to be confirmed, but it is hard to envisage the cyanohydrins as intermediates in conversion of glycine into cyanide. Very little is known about the metabolic route for production of cyanide in fungi. Ward et al. (1977) have shown that, if N-hydro~y[2-'~C]glycine or [2-'4C]glyoxylicacid oxime are given to isolate B of the snow mould fungus, very little label is recovered in cyanide. These compounds would be intermediates if glycine is converted into cyanide by a system similar to that found in higher plants (see Section 1I.C). However, as intact fungi were used in this experiment, it is possible that the compounds were not transported into the organism. It has been claimed that cyanogens similar to those produced by higher plants occur in the snow mould fungus (Stevens and Strobel, 1968), but this has been shown to be erroneous by Ward et al. (1971). In addition, Tapper and MacDonald (1974) found no evidence that plant-like cyanogens are produced by the snow mould fungus, or by M.oreades. The biosynthesis of cyanide by fungi may therefore be similar to that Seen in bacteria (see Section 11) although, as yet, there are no reports about the properties of the fungal cyanide synthase system.
MICROBIAL CYANIDE METABOLISM
89
2. Physiology of Cyanide Production The stage of growth of the snow mould fungus, in batch cultures, at which cyanide is first detected in the medium depends on the nutrients present (Bunch and Knowles, 1980, 1981). If it is grown in a basal salts medium with glucose as the carbon source, cyanide is first observed at the beginning of the stationary phase. Cyanide is not synthesized until the glucose has been completely depleted from the culture medium (Bunch and Knowles, 1980). In contrast, cyanide is produced continuously throughout growth and in the early stationary phase when acetate replaces glucose in the medium (Bunch and Knowles, 1980). Cultures grown on media containing both glucose and acetate produce cyanide only after exhaustion of glucose. This is typical of the manner in which many secondary metabolites are produced by microorganisms during batch cultivation (Drew and Demain, 1977). Although carbon catabolite repression of cyanogenesis is implicated, addition of high concentrations of cyclic AMP (adenosine 3’,5’-phosphate) to cultures have no effect on the pattern of cyanide production by glucose-grown cultures (Bunch and Knowles, 1980). Cyanogenesis is greatly stimulated if glycine is present in culture media. Radiotracer experiments have shown that the C- 1 atom of glycine, in glucosegrown cultures, is catabolized to CO, during and after growth (Bunch and Knowles, 1980). There is little conversion of the C-1 atom into cyanide. The C-2 atom is metabolized to cyanide only at the beginning of the stationary phase, coincidental with maximal cyanide production by the cultures. Very little CO, is produced from the C-2 atom. This indicates a switch in the metabolic fate of glycine during batch culture. Glycine, present in acetate-grown cultures, is metabolized quite differently. In these cultures, cyanide may be derived from either of the carbon atoms of glycine. Little I4CO2is recovered from either [1-14C]-or [2-14C]glycineunder these conditions. It is probable that the carbon atoms of glycine are scrambled during primary metabolism of acetate (Bunch and Knowles, 1981). Unlike bacterial cyanogenesis, methionine has little affect on the amount of cyanide produced (Bunch and Knowles, 1980). In addition, cyanide production by the snow mould fungus is not sensitive to the concentration of phosphate or Fez+in the culture medium. Ward and Thorn (1966) reported that betaine, NN-dimethyl glycine or formaldehyde inhibited cyanide production by the snow mould fungus. It is interesting to note that if betaine or NN’-dimethyl glycine are added to the medium during growth of the fungus on glucose plus glycine, cyanide is produced continuously (Bunch and Knowles, 1981).
90
CHRISTOPHLIR J. KNOWLES AND ALAN
W. BUNCH
D. CYANIDE DEGRADATION BY CYANOCENIC FUNGI
Cyanogenic fungi might be expected to develop systems to protect themselves from the cyanide that they have produced. So far this possibility has been investigated only in the snow mould fungus. Strobe1 (1964) showed that the snow mould fungus is able to assimilate KI4CN with formation of radiolabelled alanine and glutamate. Formation of both amino acids probably involves an initial chemical reaction of cyanide and ammonia with the appropriate aldehyde, which is then converted enzymically into the amino acid (Strobel, 1966, 1967): N H ,HCN
-
RCHO LRCHCN RCHCOOH
The level of formation of the amino acids was low under the conditions used in the experiments. Formation of the amino acids may be a way of preventing build-up of nitriles resulting from the non-enzymic reaction of cyanide with cellular constituents. Bunch and Knowles (1980) reported that cyanide, added to stationaryphase cultures of the snow mould fungus, was metabolized principally to CO,. Very little assimilation of cyanide into alanine and glutamate occurred. The ability to degrade cyanide to COz was optimal at the time of maximal cyanogenesis, regardless of the composition of the culture medium; the pathway involved is unknown. Formation of B-cyanoalanine from cyanide has not been observed by this fungus.
IV. Cyanogenesis by Photosynthetic Micro-organisms A. SPECIES THAT FORM CYANIDE
As will be seen, there are several possible routes for the formation of cyanide by photosynthetic micro-organisms (Vennesland et al., 198la, 1982; Solomonson and Spehar, 1981). Unlike fungi and non-photosynthetic bacteria, the possibility that some photosynthetic micro-organisms synthesize cyanide by one or more of these pathways was only appreciated relatively recently (Gewitz et al., 1974). This, and the fact that only small quantities of cyanide are usually produced, has resulted in relatively few reports of cyanogenesis by photosynthetic micro-organisms (Vennesland et al., 1981a). To date the alga Chlorella vulgaris, and the blue-green bacteria Anacystis nidulans, Plectonema borganum and Nosroc muscorum, are the only photosynthetic micro-organisms definitely known to be cyanogenic (Vennesland et
MICROBIAL CYANIDE METABOLISM
91
al., 1981a). However, it is likely, given the nature of the cyanogenic pathways involved, that many more of these micro-organisms will be shown to be cyanide producers. B. PATHWAYS FOR THE PRODUCTION OF CYANIDE
Photosynthetic micro-organisms synthesize cyanide from a wide range of metabolites by at least two distinct systems (Vennesland et al., 1982). 1. The Amino Acid Oxidase-Peroxidase System Production of cyanide by an alga was first demonstrated by Gewitz et al. (1976a) using Chorella vulgaris. They showed that cyanide is formed in small amounts when extracts are illuminated in the presence of O2 and supplemented with Mn2+ions and peroxidase. A large number of amino acids and related compounds were tested as possible precursors of cyanide. D-Histidine was found to be the best promotor of cyanogenesis. Other aromatic amino acids, but not glycine, could also promote cyanide formation (Gewitz et al., 1976b). These experiments also showed that although extracts of Chlorella vulgaris can release cyanide from amygdalin, a plant cyanogen, the cyanide produced under oxidative conditions was not made in this way (Gewitz et al., 1976a). It is interesting that the New Zealand spinach plant has a system for producing cyanide similar to that found in extracts of Chlorella vulgaris (Gewitz et al., 1976b), as well as forming cyanogenic glucosides in the grana. Further studies revealed details of the mechanism by which histidine is converted into cyanide by extracts of Chlorella vulgaris. Pistorius et al. (1977) showed that a soluble protein, plus a component of the particulate fraction of extracts, are necessary for cyanogenesis. The soluble protein was found to be a D-amino acid oxidase. This was partially purified by Pistorius and Voss (1977) and characterized as a flavoprotein. A wide range of amino acids, including glycine, could be oxidized by this protein with the liberation of ammonia from the a-amino group, and the reduction of O2 to Hz02.The particulate component of extracts has to be present in reaction mixtures for cyanide production to occur. It is not certain what the identity of the active component is, but it can be replaced with horseradish peroxidase or certain redox metals such as manganese and bound iron (Pistorius et al., 1977). Pistorius et al. (1977) revealed that production of cyanide from histidine and other aromatic amino acids is a general reaction catalysed by D-amino acid oxidoreductases (deaminating, EC 1.4.3.3.) and L-amino acid oxidoreductases (deaminating, EC 1.4.3.2.) when they are supplemented with peroxidase or suitable metal ions. The stoicheiometry of this process was investigated by Gewitz et al. (1980) using snake venom L-amino acid oxidase and horseradish
92
CHRISTOPHER J. KNOWLES AND ALAN W. BUNCH
peroxidase. Under optimal conditions, this system converted 72% of the added histidine into cyanide. Other products of the reaction were imidazoleCaldehyde, imidazole4carboxylic acid, CO, ,ammonia, water and imidazole acetic acid. The amount of CO, produced equalled the quantity of histidine oxidized and the sum of the ammonia plus cyanide formed. Oxidation of histidine to cyanide required more 0, than histidine oxidation by L-amino acid oxidase in the absence of peroxidase. Cytochrome c, heme or ferricyanide could substitute for peroxidase, but were less effective. Other amino acids were converted into cyanide by this system, but overall yields were less than for histidine. Catalase inhibited the process, whereas superoxide dismutase caused an increase in 0,consumption and cyanide production, and there was a larger accumulation of H,O,. Hydroxylamine can interact with many oxidants to yield, amongst other products, nitrite. When hydroxylamine was added to histidine and L-amino acid oxidase, little nitrite was produced but, in the presence of peroxidase, hydroxylamine was converted into nitrite. This reaction was inhibited by superoxide dismutase (Gewitz et al., 1980). Vennesland et al. (1 98 la) have proposed a plausible reaction sequence to explain these observations:
NH
(3)
20;
+ 2Ht
0 2
+COz
+ HZOz
+ 2H
+ 2 0 ; + HzO
The first step has been well characterized (Bright and Porter, 1975). If no peroxidase is present, the imino acid is rapidly hydrolysed non-enzymically to the keto acid and ammonia. The product, in the case of histidine, is imidazole pyruvic acid which can then react with HzOzto give imidazole acetic acid and CO,. If peroxidase is present then reactions (2) and (3) could take place. The presence of 0;)or some other activated form of oxygen, is implicated by the conversion of hydroxylamine into nitrite. Cells of Chlorella vulgaris do not excrete cyanide into the culture medium in more than trace amounts (Gewitz et al., 1976a). In contrast, in vivo production of cyanide by whole cells of the blue-green bacterium A. nidulans could easily be detected (Pistorius et al., 1979). Cyanide production by A.
MICROBIAL CYANIDE METABOLISM
93
nidulans could be stimulated by histidine. Larger quantities of cyanide were produced if peroxidase, or certain redox metals, were also present. This suggests that either the amino acid oxidase is located in the outer part of the cells, or the imino acid intermediate is excreted (Vennesland et al., 1981a). Not surprisingly, an L-amino acid oxidase was demonstrated in the cells. This enzyme has been purified to homogeneity by Pistorius and Voss (1980). It has two subunits, each of 49,000 molecular weight, and contains one molecule of FAD per molecule of enzyme. Unusually it acts only on basic amino acids. Histidine is oxidized at a much slower rate. It is inhibited by divalent cations and orthophenanthroline. This latter observation implied a requirement for a metal ion which has been shown to be zinc (Vennesland et al., 1981a). Recently the L-amino acid oxidase of A. nidulans has been reported to be associated with photosystem I1 (Pistorius and Voss, 1982). Intact cells of P . borganum and Nostoc muscorum can also synthesize cyanide from histidine. The presence of peroxidase in the medium stimulates cyanide production (Vennesland et al., 1981b). It would therefore seem that an amino acid oxidase is involved, although this needs to be confirmed.
2. The Glyoxylic Oxime System Chlorella vulgaris has been shown to possess a second system for generating cyanide. Solomonson and Vennesland (1972) reported that extracts cause cyanide formation from hydroxylamine and glyoxylate. Later studies (Solomonson and Spehar, 1981) indicated that the first step in this pathway, the reaction of hydroxylamine with glyoxylic acid to form glyoxylic acid oxime, may proceed non-enzymically. The enzyme system that catalyses cyanide production is soluble and stimulated by Mn2+ ions and ADP (Solomonson and Spehar, 1979; Solomonson and Vennesland, 1972). Using dialysed extracts, Solomonson and Spehar (1979) revealed that, for cyanide production to occur, there is an absolute requirement for ADP. Some combinations of ATP and ADP caused additional stimulation of the system, inferring a relationship between cyanide production and energy charge. Analogues of ADP decreased the response to ADP. High-performance liquid chromatography of the assay system has shown that ADP is metabolized to AMP, but there is no correlation between this and cyanide production. The extracts produced cyanide from glyoxylic acid oxime, a reaction that is also stimulated by ADP. To date the overall metabolic pathway involved is unknown. Recent work has shown that there is a similar system in spinach extracts, but radiotracer studies did not reveal much information on the steps between glyoxylic acid and cyanide (Solomonson and Spehar, 1981).
94
CHRISTOPHER J . KNOWLES AND ALAN W. BUNCH
C. THE ROLE OF CYANIDE PRODUCTION IN PHOTOSYNTHETIC MICRO-ORGANISMS
Cyanide can inhibit a wide range of metabolic processes (Solomonson, 1981), but the most pertinent effect in photosynthetic micro-organisms seems to be inhibition of the reduced form of nitrate reductase (Lorimer et al., 1974).This enzyme is essential for assimilation of nitrate. It catalyses the reduction of nitrate to nitrite by NADH. Chlorella vulgaris can form a cyanide-inactivated enzyme in vivo in the absence of added cyanide (Lorimer et al., 1974; Gewitz et al., 1978). Whether this is a common phenomenon in photosynthetic micro-organisms is not known. Inactivation is particularly noticeable in Chlorella vulgaris when cells are transferred from a medium containing nitrate to a medium containing ammonia. Under these conditions the concentration of the active form of the enzyme declines to a low level faster than the inactive form (Vennesland et al., 1981a). It has been proposed that cyanide production acts as a signal to stop a variety of metabolic processes, including assimilation of nitrogen from nitrate (Solomonson and Spehar, 1977; Solomonson, 1978). The physiological importance of cyanide production from glyoxylate and hydroxylamine or histidine in such a system has yet to be assessed. It is interesting to note, however, that the glyoxylate system may account for production of glycolate by Chlorella vulgaris grown under conditions that are optimal for cyanide production (Warburg and Krippahl, 1960). However, hydroxylamine is no longer considered to be intermediate in the reduction of nitrate to ammonia (Hewitt, 1975; Vennesland and Guerrero, 1979).The scheme for involvement of the glyoxylic oxime system in regulation of nitrate reductase is given below, where NR represents nitrate reductase (Vennesland et ul., 1982): COOH
I HC=O
+
NHZOH
f3JATPIADPf8
HC=NOH
-
HCN
NR (active)
HCN
NR-CN
oxidant
It should be noted that all nitrate reductases are inhibited by cyanide, but the strength of binding varies from one source to another. Moreover, the level of synthesis of nitrate reductase in photosynthetic micro-organisms can fluctuate greatly. Many of these fluctuations cannot be ascribed to the effects of cyanide (Hewitt et al., 1979; Vennesland et ul., 198la).
MICROBIAL CYANIDE METABOLISM
95
V. Cyanide Degradation Cyanide degradation by cyanogenic bacteria and fungi has been dealt with in earlier sections of this review. The mechanisms involved include formation of B-cyanoalanine, y-cyano-a-aminobutyrate and then a-amino acids, and, in the case of the snow mould, CO, production by an unknown pathway. This section will concentrate on cyanide degradation by non-cyanogenic species. A. CYANIDE DEGRADATION BY PHYTOPATHOGENIC FUNGI
I . Relationship to Pathogenicity
Over 2000 species of plants produce cyanogenic glycosides and other cyanogenic compounds (Conn, 1980). The reasons for production of these compounds are not known. Injury to the plant cells by water stress, harvesting, disease and infection, causes release of cyanide (Conn, 1981; Vennesland, et al., 1982). A function for the cyanogenic glycosides could be to protect the plant from predation by micro-organisms. If this is the case, then the degree of protection afforded would depend on the relative damage caused by the cyanide released to the plant tissues and to the phytopathogenic microorganism (Fry and Myers, 1981). Successfulinvasion of the host plant could occur if the invading pathogenic micro-organism was able to stimulate breakdown of the cyanogenic glycosides of the host plant, and was itself cyanide resistant. This appears to be the case with the snow mould basidiomycete and, possibly, Marasmius oreudes (for references see Section 1II.B). The fungi appear to produce an extracellular glucosidase which promotes release of cyanide from the cyanogenic glycoside of the host plant. Since the fungi are themselves cyanogenic, this process may be supplemented by cyanide formation. The fungi, at least in pure culture, can tolerate cyanide produced by their own cyanogenic apparatus. In addition, the snow mould can detoxify cyanide by converting it into C02. It thus seems that damage to the plant tissues caused by cyanide formation, as a result of fungal invasion, must be greater than the damage to the invading micro-organisms, permitting phytoparasitism to occur. Many cyanogenic plants are successfully invaded by non-cyanogenic phytopathogenic fungi (Fry and Myers, 1981). In these cases, the fungi degrade cyanide released by the plant via conversion into formamide by the action of cyanide hydratase (EC 4.2.1.66; formamide hydro-lyase): HCN
+ HZO
-
HCONHZ
Fry and Evans (1977) showed that cyanide hydratase was produced by 11
96
CHRISTOPHER J. KNOWLES AND ALAN W. BUNCH
pathogens of cyanogenic plants, 9 out of 14 pathogens of non-cyanogenic plants, and only one out of six non-pathogenic fungi. Stemphylium loti causes copper spot disease of birdsfoot trefoil (Lotus corniculatus). Millar and Higgins (1970) showed that S. loti excretes a Jl-glucosidase that releases cyanide from linamarin and lotustralin, the cyanogenic glycosides of L . corniculatus. Two types of plants were studied; both formed cyanogenic glycosides, but only one released cyanide (HCN+). The other type (HCN- ) did not form a j?-glucosidase.Since S. loti was pathogenic and caused cyanide release from both plant types, it is clear that the /I-glucosidase promoting cyanide release must have been of fungal origin. Gloeocercospora sorghi causes zonate leaf spot disease of cyanogenic sorghum species. Infected plant tissues contain less cyanogenic glycoside (dhurrin) than healthy tissues (Fry and Munch, 1974). When primary leaves of 8-10-day-old sorghum plants were inoculated by G. sorghi, there was progressive development of target leaf spot disease, and complete necrosis and tissue colonization of the leaves within 3.5 days. During this period the cyanogenic glycoside content of the leaves decreased by 85% (Myers and Fry, 1978a, b, c). The plant hydroxynitrilase activity was constant during this period, but Jl-glucosidase activity increased about 20-fold; this activity very probably stemmed from the fungus rather than the plant. Cyanide hydratase activity could not be detected in healthy plants, or before 18 hours after inoculation, but increased 200-fold within the next 24 hours. Other potential cyanide-degrading enzymes, rhodanese and Jl-cyanoalanine synthase, were present only in low concentrations. In addition to degrading cyanide, pathogens of cyanogenic plants must be able to form a cyanide-resistantrespiratory system. Both S. loti and G . sorghi are known to have inducible cyanide-resistant respiratory pathways (Fry and Millar, 1971a, b; Fry and Munch, 1975). 2. Properties of Cyanide Hydratase Cyanide is quantitatively and irreversibly converted into formamide by spores or mycelia of S. loti, or mycelia of G. sorghi induced for cyanide hydratase activity (Fry and Millar, 1972; Fry and Munch, 1975; Nazly and Knowles, 1981; Nazly et al., 1983). Cyanide hydratase from homogenates of spores of S. loti was purified 5-fold on DEAE-cellulose, and a further 3-fold on Sephadex G-200. The enzyme eluted in the void volume of Sephadex G-200, suggesting that it has a molecular weight of at least 6 lo5(Fry and Millar, 1972). Cyanide hydratase from mycelia of G. sorghi was purified 3- to 6-fold on Sephadex G-100 (eluting in the void volume) and a further 1.5- to 5-fold on DEAE-Sephadex. The enzyme eluted slightly behind the void volume of Bio-Gel A-15M,
-
97
MICROBIAL CYANIDE METABOLISM
-
suggesting that it has a molecular weight of at least 2 lo6(Fry and Munch, 1975). The enzyme was not sedimented by centrifugation for 20 minutes at 2O,OOOg, or at 100,OOOg for 90 minutes, but was sedimented at 100,000g for 6 hours (Fry and Myers, 1981). Thus, the cyanide hydratases of S. loti and G. sorghi are large enzymes. This contrasts with two enzymes from Fusarium solani which convert cyanide into ammonia, and have molecular weights of 17,000 and 40,000 (Shimizu et al., 1968; Shimizu and Taguchi, 1969). The cyanide hydratases from G. sorghi and S. loti have broad pH optima with maximal activity at pH 7.0 to 9.0 (the pK, value of cyanide is 9.3; Fry value for cyanide of the S. and Millar, 1972; Fry and Munch, 1975). The K,,, loti enzyme is in the range 15 to 2 7 m ~ (Fry and Millar, 1972; Nazly and Knowles, 1981; Nazly et al., 1983), whereas the G.sorghi enzyme has a &, value of 25 to 2 7 m ~(Fry and Munch, 1975; Nazly et al., 1983). Both enzymes are stable to dialysis and 10mM EDTA (ethylenediaminetetraacetate), NH,Cl, KC1, MgC12or CaCl, (Fry and Millar, 1972; N. Nazly and C.J. Knowles, unpublished observations). Cyanide hydratase of S. loti, G.sorghi and several other fungi pathogenic to cyanogenic plants is found at low or undetectable concentrations when they are grown in pure culture, but are dramatically induced by the addition of 0.1 to 5 mM cyanide to the cultures 12 to 24 hours before harvesting (Fry and Evans, 1977; Fry and Myers, 1981; Nazly et al., 1983). The product of the reaction (formamide) can also induce cyanide hydratase, apparently synergistically with cyanide (Fry and Myers, 1981). 3. Industrial Potential
Large quantities of cyanide are formed due to natural synthesis (Knowles, 1976; Conn, 1981; Vennesland et al., 1982) and industrial production (Anonymous, 1976a, b). Industrial cyanide effluents pose a major hazard, and many processes have been proposed for detoxification. These include treatment by alkaline chlorination, hypochlorite, bleach, peroxide or ozone, ion exchange or electrolysis(Howe, 1963,1965; Green and Smith, 1972; Scott and Ingles, 1980; Zabbon and Helwick, 1980). In addition, a range of biological treatments have been proposed using sewage systems, but these have not found widespread acceptance (Pettet and Mills, 1954; Ludzack and Schaffer, 1960; Nesbitt et al., 1960; Winter, 1963;Mikami and Misono, 1968; Shimizu et al., 1968; Howe, 1969; Fuji and Oshimi, 1973; Raef et al., 1975; Furuki et al., 1975). Discovery of cyanide hydratase (Fry and Myers, 1981) for conversion of toxic cyanide into the relatively non-toxic formamide (Brinkham and Kuhn, 1978), and the ability of the snow mould to degrade cyanide to C02(Bunch and Knowles, 1981), suggests that cyanide-degrading fungi could be used to detoxify industrial cyanide wastes.
98
CHRISTOPHER J. KNOWLES AND ALAN W. BUNCH
Spores of S. loti lost 50% of their cyanide hydratase activity within 2 days at 25°C (Fry and Millar, 1972), whereas mycelia of S. loti lost 50% of their cyanide hydratase activity within 15 hours at 22-24°C. Cyanide hydratase from G. sorghi lost half of its activity in 3-5 days (Nazly et al., 1983). For industrial use, the enzymes must be considerably more stable. This has been achieved by immobilization (Nazly and Knowles, 1981; Beardsmore and Powell, 1981; Nazly et al., 1983), using polyelectrolyte flocculating agents (Lee and Long, 1974). Immobilized S. loti, G. sorghi and Fusarium moniliforme have been tested in continuous packed-bed column reactors (Nazly et al., 1983). Continuous detoxificationof at least 70 mM cyanide has been achieved, with 100% conversion into formamide, for 45 hours using S. loti, and 30 days using G. sorghi. Inclusion of 1m~ glucose in the feed enhanced stability up to 55 hours and 40 days, respectively. Heat-treatment wastes containing NaCN and KCN also usually contain barium salts and carbonates (Anonymous, 1976a, b). Cyanide degradation by the immobilized fungi was unaffected by trace concentrations of a range of metals, and higher concentrations of barium salts and Na,CO, (Nazly and Knowles, 1981). However, metal finishing wastes contain a range of heavy metals that severely inhibit cyanide degradation by the immobilized fungi. B. CYANIDE RESISTANCE AND DEGRADATION BY BACTERIA
I . Bacteria Pathogenic to Cyanogenic Plants As indicated above, fungi that are phytopathogenic to cyanogenic plants form cyanide hydratase to detoxify cyanide by conversion into formamide (see also Fry and Myers, 1981). It might, therefore, be expected that bacteria pathogenic to cyanogenic plants would also form cyanide hydratase or other cyanide-degrading enzyme systems. Rust et al. (1980) have tested a range of bacteria for their sensitivity to cyanide by inclusion of cyanide in a glucose plus nutrient broth medium. The bacteria tested included several strains of P. syringae (pathogenic to sorghum) and Xanthomonas manihotis (pathogenic to cassava), as well as a range of bacteria pathogenic to non-cyanogenic plants. In every case growth was 50% inhibited by 0.06 to 0.17 m~ cyanide. These concentrations of cyanide are considerably lower than the local concentration of cyanide that would form in the leaves of cassava or sorghum if even a small proportion of their cyanogenic glycosides were degraded on infection. These results are, at first sight, both puzzling and unexpected. However, they should be accepted with caution. It is possible that the cyanogenic glycosides are not degraded on bacterial infection. Alternatively, in vivo the
MICROBIAL CYANIDE METABOLISM
99
bacteria might degrade cyanide released from the plant cyanogenic glycosides; possibly a co-nutrient for cyanide detoxication is supplied by the plant or there is a trigger substance present for induction of a bacterial cyanide degrading system.
2. Cyanide Resistance as a Taxonomic Test Inhibition of growth by cyanide is often used as a taxonomic test to differentiate betweeen species of the Enterobacteroaceae (Cowen, 1974). For example, most Salmonella species and Escherichia coli are cyanide-sensitive bacteria, whereas Citrobacterfreundii and Enterobacter species are often cyanideresistant. The medium used is a broth containing peptone, NaCl, phosphate and 1 to 4 m KCN (Msller, 1954). This medium can be made more stable and sensitive by inclusion of an indicator and/or making it semi-solid with agar (Gershman, 1960; Munsen, 1974). Porter and Knowles (1 979) have investigated the factors responsible for cyanide resistance by some species of Enterobacteriaceae.They showed that Citrobacterfreundii and Enterobacter aerogenes were able to grow in proteose peptone containing 1mM KCN, whereas E. coli failed to grow in peptone containing only 100 p~ KCN. In contrast, none of the organisms grew in a lactate-minimal salts medium containing 100 p~ KCN. The nutrient responsible for conferring cyanide resistance in the peptone medium was L-cysteine. Citrobacterfreundii or Enterobacter aerogenes grew on lactate-minimal salts medium containing 1m~ KCN if 2 m Lcysteine ~ was present, but not if a mixture of 21 other amino acids, or each of the 21 amino acids alone, were present. L-Cysteine could be replaced by L-cystine, D-cystine or thio-acetic acid. D-Cysteine and N-acetyl-L-cysteine had some protective effect but cysteamine, thioglycolate, homocysteine and mercaptoethanol were ineffective. It might be expected that cyanide detoxification occurs in the resistant species by the action of B-cyanoalanine formation, possibly by a cysteine synthase with secondary /?-cyanoalaninesynthase activity (see Macadam and Knowles, 1984). This has not been tested, but it seems that cyanide is not metabolized by the bacteria since cyanide is lost from uninoculated medium and cultures of cyanide-sensitive or cyanide-resistant bacteria at a similar rate. Another possibility is that the biosynthetic pathway for cysteine formation is cyanide-resistant in some species, whereas in other species it is cyanide-sensitive; this would not explain the protective effect of some other sulphur-containingcompounds. Cystine and homocystine react with cyanide (Ressler et al., 1973), so it is possible that cysteine in the growth medium
100
CHRISTOPHER J. KNOWLES AND ALAN
W. BUNCH
could be progressively oxidized to cystine by some bacteria, thus detoxifying it. 3. Cyanide Degradation
There have been several reports that bacteria or sewage systems degrade cyanide with release of COz or ammonia (Howe, 1963, 1965; Ludzack and Schaffer, 1960; Nesbitt et al., 1960; Pettet and Mills, 1954; Skowronski and Strobel, 1969; Winter, 1963). Skowronski and Strobe1 (1969) isolated a strain of Bacillus pumilus that grew rapidly in a trypticase soy/yeast extract/glucose medium containing 0.1 M KCN. The cyanide was degraded with release of C 0 2 and ammonia. They claimed that some growth could occur due to utilization of cyanide as the source of carbon plus nitrogen. With cyanide in the growth medium, Oz uptake was greatly stimulated. A strain of Bacillus cereus was isolated by McFeters et al. (1970) that could grow in nutrient medium containing 1mM KCN. Growth in the presence of cyanide resulted in development of a cyanide-resistant respiratory system and enhancement of formation of cytochrome b and cytochrome oxidase (proba& ably cytochrome a Sakai et al. (1981) isolated a range of bacteria that were able to grow in a rich medium containing 2 m~ cyanide. From these, a secondary screen was performed to isolate bacteria that formed ninhydrin-positive material (amino acids) when they were incubated in media containing cyanide. Three types of bacteria were obtained; those that formed B-cyanoalanine, valine, or alanine and aspartate. One of the organisms that formed B-cyanoalanine,Enterobacter strain 10-1, was studied further. This bacterium, when grown in the presence of 1 to 5 m cyanide, ~ accumulated almost stoicheiometric amounts of 8cyanoalaninein the medium. Resting cell suspensions formed B-cyanoalanine from cyanide when incubated in the presence of 0-acetyl-L-serine, L-serine, m-alanine or L-cysteine. Beta-Cyanoalanine synthase activity was found to be 98.5% in the soluble fraction of the cell. 0-Acetyl-L-serine was the most active substrate; L-cysteine and L-cystine gave only about 15% of the activity. The small amount of 8-cyanoalanine synthase activity in the particulate fraction, or in a detergent-solubilized fraction from it, was active only with 0-acetyl-L-serine as the substrate. The enzyme activity from the soluble fraction of the cell has been purified to homogeneity (see Tables 2 and 3). The purified enzyme catalysesformation of cysteine from 0-acetyl-L-serine and Na,S at 245-fold the rate of formation of B-cyanoalanine from 0-acetyl-L-serine and NaCN (Yanese et al., 1982a, b). The affinity for NazS (K,0.5 m ~ is)10-fold greater than the affinity for NaCN (K, 5 . 2 m ~ ) indicating , that it is an 0-acetyl-L-serine sulphydrylase
+
MICROBIAL CYANIDE METABOLISM
101
(cysteine synthase) rather than a true B-cyanoalanine synthase, as found for C. violaceum (Macadam and Knowles, 1984). The enzyme is constitutive: inclusion of cyanide in the growth medium does not cause it to be induced (Yanese et al., 1982a). The enzyme from Enterobacter strain 10-1 thus resembles the enzyme studied earlier by Castric and Conn (1971) in Bacillus megaterium. The latter enzyme is also principally a cysteine synthase, with a small amount of /I-cyanoalanine synthase activity, using cysteine, 0-acetyl-L-serine or serine as the substrate. Moreover, this enzyme was inducible when grown in media containing sulphate or L-djenkolic acid, rather than L-cysteine, as the source of sulphur, but was not induced by addition of cyanide to the medium. Escherichia coli (Dunnill and Fowden, 1965; Lauinger and Ressler, 1970), C. violaceum (Brysk et al., 1969) and B. megaterium (Castric and Strobel, 1969; Castric and Conn, 1971) can convert fl-cyanoalanineinto asparagine or aspartate. Beta-Cyanoalanine hydratase, which catalyses asparagine formation, has been partially purified and characterized from the plant, blue lupine (Castric et al., 1972). Little is known about the enzyme involved in B-cyanoalanine assimilation by the bacteria mentioned above. The conversion of fl-cyanoalanine into aspartate by E. coli is probably a function of asparaginase (Lauinger and Ressler, 1970). Yanese et al. (1982~)have isolated several bacteria that utilize fl-cyanoalanine as a source of nitrogen. Of the isolates, two were shown to release .asparaghe and aspartate into the culture medium, whereas a fluorescent Pseudomonas species (strain 13) rapidly degraded fl-cyanoalanine without accumulation of asparagine and aspartate. With resting cell suspensions of strain 13, disappearanceof fl-cyanoalanine was accompanied by formation of aspartate. Cell-free extracts catalysed formation of aspartate from fl-cyanoalaninewithout formation of asparagine, but also exhibited asparaginaseactivity. The fl-cyanoalanine-degrading enzyme was partially purified and shown to be separate from the asparaginase activity. This enzyme had no asparaginase activity, but catalysed formation of both asparagine and aspartate from fl-cyanoalanine. BetaCyanoalanine-degradingactivity was induced by growth on fl-cyanoalanine, but not by growth on asparagine and a range of nitriles and amides as sources of nitrogen, whereas asparaginase activity was found for growth on asparagine, b-cyanoalanine or several other nitriles are amides. The fl-cyanoalanine-degradingenzyme has been purified to homogeneity (Yanese et al., 1983). It has a molecular weight of about lo6 and has 30 identical subunits. The enzyme is totally specific for B-cyanoalanine, and does not degrade asparagine. The ratio of asparagine to aspartate formed is 2.2: 1.
102
CHRISTOPHER J. KNOWLES AND ALAN W. BUNCH
4. Cyanide Utilization
As already discussed in this article, it is known that (a) some plant pathogenic fungi form cyanide hydratase to convert cyanide into formamide, (b) the snow mould can convert cyanide into CO,, and (c) CO, can be released from cyanide by cyanide-acclimatized sewage systems. These data suggest that it should be possible to isolate micro-organisms able to grow on cyanide as a source of carbon and/or nitrogen, provided they are able to form a cyanideresistant respiratory system and other cyanide-resistant metabolic processes (Solomonson, 1981), as well as the enzymes involved in cyanide assimilation. Possible pathways for assimilation of cyanide have been discussed by Knowles (1976). The discovery of cyanide hydratase (Fry and Myers, 1981) suggests that a possible route for evolution of CO, is via the activities of cyanide hydratase, an amidase and formate dehydrogenase, with formamide and formate as intermediates: HCN
7
HCONH2 + H20 H C O O H n C 0 2 NAD+
NADH+H*
If this route occurs, then methylotrophic bacteria (Large, 1983) able to utilize cyanide as a source of carbon (and nitrogen) should exist. Evidence for such a pathway also comes from studies on the plant, loquat (Eriobotryajaponica), which detoxifiescyanide to formate by a three-enzyme sequence (Shirai et al., 1977; Shirai, 1977). Formaldoxime was proposed as an extra intermediate: HCN
- +H20
HCONH,
(H2C = NOH)
+ H20
HCOOH
Other possible routes of cyanide assimilation (Knowles, 1976) include formation of /3-cyanoalanine followed by conversion into aspartate,and formation of a-hydroxy- or a-amino-nitriles followed by metabolism to the corresponding a-hydroxy or a-amino acids. There have been several reports of the isolation of cyanide-utilizing microorganisms. Pettet and Ware (1955), and Ware and Painter (1955), isolated a cyanide-utilizing bacterium from a percolating filter that had been seeded with cyanide-acclimatized sewage sludge. The organism was strictly autotrophic and grew on silica gel but not on agar plates. Ammonia was released from cyanide. The organism was Gram-positive and filamentous. It was probably an actinomycete. Using acclimatized sludge obtained from Pettet and Ware, Winter (1963) reported the isolation of two bacteria that grew on cyanide as a source of carbon and nitrogen. In contrast to the organism of Ware and Painter (1955), these bacteria were facultative autotrophs that grew rapidly and profusely on
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cyanide as well as on several organic compounds. They were Gram-positive rods with sparse aerial hyphae, and were probably actinomycetes. Furuki et al. (1972) isolated a bacterium that utilized cyanide as a source of nitrogen from soils that had been polluted with cyanide-containingeffluents from an electroplating plant. The organism was alkaliphillic and grew best on a medium containing up to 2 0 m cyanide. ~ Glucose, fructose, mannose and galactose were sources of carbon and energy, and ammonia or urea as well as cyanide could be used as sources of nitrogen. Growth was relatively slow; with cyanide as the nitrogen source there was a lag phase of 5 days, and the stationary phase was attained 11 to 12 days after inoculation. Raynaud and Bizzini (1959) isolated bacteria from pond mud that were able to grow on 1 to 10mM NaCN. It was claimed that three types of bacteria were obtained: (a) slow-growing Gram-positive cocci, (b) a sporulating Gram-positive rod that grew a little more rapidly, and (c) a rapidly growing Pseudomonm species. The last organism exhibited a lag phase of 1 to 5 hours during which the cyanide concentration in the medium decreased; growth commenced when no cyanide remained. Unfortunately, the bacterium isolated by Ware and Painter (1955) has not been retained (H. A. Painter, personal communication). Cultures of the bacteria isolated by Winter (1963) were kindly provided by J. A. Winter to P. A. Collins and C. J. Knowles, but appear to have lost their ability to grow on cyanide (unpublished observations). The other cyanide-utilizing organisms mentioned above are not generally available. Extensive efforts by N. Nazly, R. H. Harris and C. J. Knowles (unpublished observations) to isolate bacteria that grow on cyanide as a carbon source have been unsuccessful. The toxicity of cyanide presents problems in isolating bacteria that are able to utilize it; a concentration high enough to support reasonable growth might in fact prove to be too toxic to permit growth to occur. Since the amount of nitrogenous source needed for growth is lower than the requirement for carbon, it might be easier to isolate bacteria that use cyanide as a nitrogen source, in the presence of a carbon source such as glucose. Moreover, under these conditions, cyanide is not required to act as the source of energy. The toxicity problems also can be decreased by providing cyanide (since it is volatile) as a vapour for enrichment cultures to isolate bacteria. Cyanide-limited fed-batch or continuous culture could be used to obtain growth in liquid culture. Harris and Knowles (1983a) have isolated bacteria that grow on cyanide as a source of nitrogen. The bacteria were isolated from samples of river mud from a single site. Using glucose as the carbon source, and with cyanide supplied as a vapour, a series of Gram-negative oxidase-positive bacteria were obtained, each of which produced a fluorescent green pigment, and which were tentatively identified as strains of P . jluorescens. Three of the
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FIG. 3. Growth of a cyanide-utilizing strain of Pseudomonasfluorescensin fed-batch cultures with KCN (0)or NH4Cl ( 0 )as the nitrogen source, or with no source of The source of carbon and energy was 1 0 m glucose. combined nitrogen present (0). From Hams and Knowles (1983a), with kind permission of the Journal of General Microbiology.
bacteria were examined further and shown to be P. fluorescens biotype 11. One of them, which did not clump in liquid medium, was retained for further studies (strain NCIB 11764). Figure 3 shows growth of P.fluorescens in fed-batch culture with KCN or NH4Cl as the limiting nutrient (Harris and Knowles, 1983a). Growth was similar in both cases, and terminated due to depletion of glucose from the medium. The maximal dilution (growth) rate with cyanide as the limiting nutrient was 0.5 h-I. There was no growth in the absence of a source of combined nitrogen. Pulsing 1 mM cyanide into ammonia-grown cultures stopped growth, and there was no conversion of CN- to NH: ions. In contrast, pulsing 1m M cyanide into cyanide-grown cultures resulted in stoicheiometric conversion of it into NH: ; growth resumed when all the cyanide had been degraded. This suggests that cyanide assimilation is by an inducible system. Addition of cyanide to suspensions of cyanide-grown bacteria resulted in stimulation of O2uptake and formation of ammonia. The molar ratio of CNutilized/NH: formed was 1: 1. I (Harris and Knowles, 1983a). Cell-free extracts of cyanide-grown, but not ammonia-grown, bacteria were also able to convert cyanide into ammonia and C02(Harris and Knowles, 1983b).The cyanide-degradingsystem was located in the soluble fraction of the cell, and required NADH and 0,for activity. Both O2uptake and NADH oxidation were stimulated by addition of cyanide to the soluble fraction. NADPH could replace NADH with 40% of the activity. Ammonium sulphate fractionation showed that at least two proteins are involved in the process. The molar proportion of CN- utilization/NH,+ formation/O, uptake/NADH utilization/CO, formation was 1:0.860.96:I .00:0.67. Measurement of CO,
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formation was probably an underestimate, due to technical difficulties. These results suggest that the cyanide utilizing process is: NAD(P)H
-
+ 2H+ + HCN + 0,
C02
+ NH: + NAD(P)+
The enzyme system thus appears to be an inducible, multi-component dioxygenase, requiring a source of reducing equivalents (NAD(P)H), and could be related to some aromatic dioxygenases(Yeh et al., 1977; Crutcher and Geary, 1979). However, the process could be in two stages, with the first stage a mono-oxygenase and with, for example, cyanate as an intermediate; cyanase (Taussig, 1960, 1965) would then degrade the cyanate: NAD(P)H
+ H + + HCN + OzHOCN
+ HzO
-
HOCN CO,
+ NAD(P)+ + H,O
+ NH,
VI. Concluding Remarks
It has been shown that cyanide is a relatively common product of microbial as well as plant metabolism. The biochemistry and physiology of cyanide production by fungi and heterotropic non-photosynthetic bacteria appear to have many similarities. Cyanide production by these micro-organisms has many characteristicstypical of secondarymetabolism. Since it is probably the simplest secondary metabolic system, a continued investigation of cyanide formation should greatly aid a better understanding of microbial secondary metabolism in general which, despite its past and future importance, is lacking. Even under optimal conditions, photosynthetic micro-organisms synthesize much smaller quantities of cyanide than do other cyanogenic microbes. In addition, the biosynthetic pathways involved, and the possible physiological reason for cyanide production, are very different. Because of its great importance, the role of cyanogenesis by these micro-organisms in the regulation of nitrate reductase clearly requires further elucidation. A wide range of micro-organisms, including some cyanogenic species, are able to catabolize cyanide. In addition, some bacteria are known to utilize cyanide as a source of nitrogen for growth. It has yet to be confirmed unambiguously that cyanide can be used as a source of carbon and energy for growth. Clearly it is intriguing to know what significance cyanide metabolism has in the cycling of carbon and nitrogen in the environment, especially as plants, as well as micro-organisms, both produce and catabolize cyanide. In addition to the biological production of cyanide, large quantities of cyanide wastes are generated annually by industry. Inevitably microbial cyanide degradation is being assessed for its applicability to treatment of general or specific types of cyanide-containing wastes. In summary, investigation of this seemingly esoteric area of microbiology
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C.J. Knowles, J. Westley and F. Wissing, eds), pp. 275288. Academic Press, London and New York. Yanese, H., Sakai, T. and Tonomura, K. (1982a). Agriculturaland Biological Chemistry 46,355. Yanese, H., Sakai, T. and Tonomura, K. (1982b). Agricultural andBiologica1 Chemistry 46,363. Yanese, H., Sakai, T. and Tonomura, K. (1982~).Agricultural and Biological Chemistry 46, 2925. Yanese, H., Sakai, T. and Tonomura, K. (1983). Agricultural and Biological Chemistry 47,473. Yeh, W.K., Gibson, D.T. and Liu, T.-N. (1977). Biochemical and Biophysical Research Communications 78, 401. Zabbon, W. and Helwick, R. (1980). Plating and Surface Finishing 67, 56.
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Bacterial Oxidation of Methane and Methanol C . ANTHONY Department of Biochemistry, University of Southampton, Southampton,
UK
I. Introduction . . . . 11. Oxidation of methane to methanol . . . A. Introduction . . B. The methane mono-oxygenasesof Methylococcus capsulafus(Bath) . C . The methane mono-oxygenass of Methylosinus trichosporiwn . . D. The electron donor for methane mono-oxygenase . . E. The substrate specificity of methane mono-oxygenase F. The mechanism of methane mono-oxygenase . . . . 111. Oxidation of methanol to formaldehyde . . . A. Methanol dehydrogenase . . . . . . . B. The prosthetic group and mechanism of methanol dehydrogenase . C. Cytochrome c and its involvement in methanol oxidation . D. The methanol cytochrome c oxidoreductase activity of methanol dehydrogenase . . . . . . . IV. Energy transduction during the oxidation of methane and methanol A. Introduction . B. Electron transport and proton translocation in Methylosinus frichosporiwnand other methanotrophs . . . C. Elwtron transport and proton translocation in Pseudomoms AM 1 . D. Electron transport and proton translocation in Paracoccus denitr$cans . E . Electron transport and proton translocation in Methylophilus methylotrophus F. The coupling of methanol oxidation to synthesis of adenosine triphosphate V. Acknowledgements. . . . References . . . . .
.
.
113 116 116 118 122 126 127 128 129 129 147 162 173 179 179 184
186 189 191 199 203 203
1. Introduction
Bacteria able to oxidize methane and methanol, and to use the energy made available by this process for growth, have been known for about 80 years Since Sohngen’s description of Bacillus methunicus (Sohngen, 1906). Such bacteria have become known as methylotrophs. These are defined as microorganisms able to grow at the expense of reduced carbon compounds conADVANCES IN MICROBIAL PHYSIOLOGY, VOL. 27 ISBN 0-12-027727-1
Copyright 0 1986 by Academic Press London AII rights of reproduetion in my form reserved
114
C.
ANTHONY
taining one or more carbon atoms but containing no carbon-carbon bonds. Obligate methylotrophs grow only on such compounds, whereas facultative methylotrophs are also able to grow on a variety of other organic multicarbon compounds (Colby and Zatman, 1972; Anthony, 1982). In terms of their biology, the methylotrophs can be divided into two well-defined groups; those able to use methane and those unable to do so. Methylotrophic bacteria that are able to grow on methane are also called methanotrophs; they are usually obligately methylotrophic, growing well on methane but often rather poorly on methanol. A few facultative methanotrophs have been described but some of the original isolates have changed markedly since their first description, whereas other isolates have been shown to be mixed cultures (see Anthony, 1982; Lidstrom-OConnor et al., 1983). Most methylotrophs that are unable to grow on methane use methanol or methylated amines (or both) as their C, substrates; other C, substrates, used by relatively few methylotrophs, include formate, formamide, CO, dimethylsulphide or trimethylsulphonium compounds. This review is concerned with the enzymes involved in the bacterial oxidation of methane and methanol to formaldehyde, and the energy transduction systems involved in coupling these oxidations to ATP synthesis by way of electron transport chains, proton translocation and the proton motive force. Bacteria oxidizing methane and methanol do so by the following route:
The first reaction in methane oxidation is a hydroxlyation catalysed by a very unusual mono-oxygenase; this requires a reductant that is probably always NADH. Methanol oxidation to formaldehyde is always catalysed by a methanol dehydrogenase (MDH) having pyrrolo-quinoline quinone (PQQ) as its prosthetic group. When bacteria are oxidizing methane, then the oxidation of formaldehyde must always be coupled to NADH formation. In contrast, during growth on methanol this is not essential, and formaldehyde may be oxidized by alternative “dye-linked’’ dehydrogenases which may be flavoproteins. The oxidation of formate is always coupled to the reduction of NAD+ by formate dehydrogenase. Some bacteria have low concentrations of formate dehydrogenase and they oxidize formaldehyde to C02 by a cyclic variant of their carbon assimilation pathway, which yields two molecules of NAD(P)H (see Anthony, 1982 for a review of the enzymes involved in formaldehydeand formate oxidation). The thermodynamic constants for the reactions involved in methane oxidation are summarized in Table 1.
TABLE 1. Thermodynamic constants for reactions involved in the oxidation of C, compounds. From Ribbons et al. (1970)
-
AG; (PH 7.0) (kJ mol-')
Reaction CH4 CHSOH HCHO HCOOH CH4 CH3OH HCHO NADH H+
+
+ 0.5
0 2
+ 0.5 + 0.5 O2 + 0.5 + 2.0 + 1.5 02+ 1.0 + 0.5 O2 0 2
0 2
0 2
0 2
CH30H HCHO H2O HCOOH COZ H2O C02 2H20 C02 2 H20 C02 H2O NAD+ H 2 0
-
+
+
+ + +
+
Redox couple
E,,' (PH 7.0) (V)
HCHO/CH30H HCOOH+/HCHO COJHCOOH+
-0.182 - 0.450 - 0.460
NAD+/NADH + H+
- 0.320
- 109.7 - 188.2 - 240.0 - 244.7 - 782.6
+
+
- 672.9
-484.7 - 236.8
116
C. ANTHONY
11. Oxidation of Methane to methanol A. INTRODUCTION
Work from many laboratories using a variety of methanotrophs (see Table 2) has led to the general conclusion that the first step in methane oxidation is catalysed by a mixed-function mono-oxygenase system which hydroxylates methane to methanol using molecular 0,and a reductant (AH,), which is probably always NADH: CH,
+ 0, + AH,
-
CH,OH
+ A + H,O
The conclusion that a mixed function mono-oxygenase is involved in methane oxidation was first implied by the work of Leadbetter and Foster (1959) and later confirmed by Higgins and Quayle (1970) who showed that whole cells of Methylomonus methunicu and Methylomonas methano-oxidans incorporate "0, into methanol from "OZbut not from water containing "0. Further confirmation, and understanding of the nature of the methane mono-oxygenase (MMO), has depended on the isolation of cell-free systems able to catalyse methane oxidation. Early work depended on measuring NADH oxidation and 0,consumption occurring on addition of methane to particulate preparations derived from Methylococcus cupsulutus (Texas strain) (Ribbons and Michelover, 1970). Because these preparations were also able to oxidize NADH, methanol, formaldehyde and formate (in the absence of methane) (Ribbons, 1975), interpretation and development of this system presented considerable difficulties. These were eventually avoided by developing alternative methods of assay using using analogues of methane (CO, bromomethane, ethylene) as substrate, or by measuring the product, methanol. Dalton's group at Warwick has provided the most definitive description of MMO from their work on the soluble NADH-requiring system from Methylococcus cupsulutus (Bath) (a Type I methanotroph having the ribulose monophosphate pathway of carbon assimilation). This MMO appeared for some time to be markedly different from the MMO in Methylosinus trichosporium (a Type I1 methanotroph having the serine pathway for carbon assimilation). Higgins and his group (originally working at Canterbury, Kent) had described a completely different particulate system from this organism which, after solubilization, was unable to use NADH as electron donor, but instead had used cytochrome c (reduced by ascorbate or by methanol plus MDH). This confusing situation has now been largely resolved. Both types of methanotroph are now known to produce two types of MMO, both able to use NADH (but not ascorbate) as electron donor in uitro. Particulate types
TABLE 2. Studies of methane oxidation in bacteria
(a) Reviews of methane oxidation Anthony (1980, 1982), Best and Higgins (1983), Colby et al. (1979), Dalton (1980a, b, 1981), Dalton and Stirling (1982), Dalton et al.(1984), Dalton and Leak (1985), Higgins (1979, 1980), Higgins et al. (1981a, b, c, l982,1984a, b), Hou (1984a), Hou et al. (1980a) (b) Methane oxidation in different mefhylotrophs Orgpnism Methylococcus capsulatus (Bath strain)
Methylococcus capsulatus (other strains) Methylomonas methanica (Pseudomom methanica) Methylomonas (other species) Methylomonas methano-oxidans Methylosinus trichosporiwn
Methylobacterium sp. CRL-26
Other methanotrophs Ammonia-oxidizing bacteria
References Colby and Dalton (1976, 1978, 1979), Colby et al. (1977), Dalton (1977), Stirling and Dalton (1977, 1979a, b, 1980, 1981), Stirling et al. (1979), Dalton (1981), Dalton et al. (1981), Leak and Dalton (1983), Stanley et al. (1983), Woodland and Dalton (1984a, b), Prior and Dalton (1985a, b), Lund and Dalton (1985), Lund et al. (1985) Ribbons and Michalover (1970), Ribbons (1975). Patel et al. (1976), Ribbons and Wadzinski (1976), Stirling and Dalton (1977), Leadbetter and Foster (1959), Ferenci (1974, 1976a, b), Ferenci et al. (1975), Colby et at. (1975), Stirling et ul. (1979) Hubley et al. (1974, 1975), Stanley et al. (1983) Higgins and Quayle (1970) Hubley et al. (1974, 1975), Ferenci (1974), Patel et al. (1976), Tonge et al. (1975, 1977a), Thomson et al. (1976), Higgins et al. (1976a, 1979, 1981a), Hammond et al. (1979), Higgins (1979), Stirling and Dalton (1979a), Stirling et al. (1979), Higgins et al. (1981a, b), Best and Higgins (1981), Scott et al. (1981a, b), Jezequel and Higgins (1983), Joergensen and Degn (1983), Stanley et al. (1983), Cornish et at. (1984), Burrows et al. (1984) Patel et al. (1982) Hou et al. (1979a, b, 1980a, b, 1981, 1982a, b), Patel et al. (1979b, 1980), Leak and Dalton (1983), Stanley et al. (1983), Hou (1984b) Jones (1983), Hyman and Wood (1983, 1984)
118
C. ANTHONY
of MMO are formed in conditions of copper sufficiency, and soluble types of MMO are formed when bacteria are grown under conditions of copper insufficiency. Copper sufficiency and insufficiency are governed by both the copper concentration in the growth medium and by the cell density, and so they can appear to be determined by alterations in the growth rate or in the carbon, 0, or nitrogen supply (Dalton et al., 1984; Cornish et al., 1984; Burrows et al., 1984; Prior and Dalton, 1985a). For completeness, the evidence will be presented that relates to both types of MMO in the two methanotrophs studied in most detail. Similar types of MMO have been described in the other methanotrophs listed in Table 2. B. THE METHANE MONO-OXYGENASES OF METHYLOCOCCUS CAPSULATUS (BATH)
I . The Soluble Methane Mono-oxygenase from Methylococcus capsulatus (Bath)
This NADH-dependent, soluble mono-oxygenase is induced during growth under conditions of copper insufficiency (Stanley et al., 1983; Dalton et al., 1984). After breakage of the bacteria in a French Pressure cell, the MMO complex exists free in solution and has three essential components (A, B and C) which have been resolved and characterized (Colby and Dalton, 1978; 1979; Dalton, 1980a, b; Woodland and Dalton, 1984a, b; Dalton and Leak, 1985; Lund and Dalton, 1985; Lund et al., 1985). a. Component A. This component, characterized by Dalton and coworkers (Woodland and Dalton, 1984a, b; Woodland and Cammack, 1985),comprises up to 30% of the soluble cell protein. Its approximate molecular weight is 210,000 and it is made up of two copies each of three subunits of relative molecular weights (M,) of 54,000 42,000 and 17,000. It is an acidic protein having an isoelectric point between pH 5.1 and 5.2. It contains two atoms of non-haem iron per molecule and a small amount of zinc, but no acid-labile sulphur. It is colourless, but sometimes has a shoulder in the absorption spectrum at 406410nm. Electron spin resonance (ESR) spectra of component A do not appear to be like those of iron-sulphur proteins, and they suggest the possibility of a novel active centre. After reduction with dithionite, the ESR signal is altered by addition of ethene or cyanomethane thus suggesting that the reduced form of component A is responsible for substrate binding. Component A does not resemble other oxygenases, and it is suggested that the unusual ESR spectrum, the presence of iron and zinc, and the absence of haem, together with the apparent lack of an extrudable iron-sulphur cluster, indicate a novel iron-containing prosthetic group which clearly requires further elucidation (Woodland and Dalton, 1984a; Woodland and Cammack, 1985).
BACTERIAL OXIDATION OF METHANE AND METHANOL
119
A protein with properties similar to those of component A has also been purified and characterized from Methylobacterium sp. CRL-26 (Patel, 1984).
b. Component B. This component, characterized by Dalton and coworkers (Dalton, 1980b, 1981a; Lund et al., 1985), is a colourless protein comprising a single polypeptide chain with a molecular weight of between 15,000 and 20,000. It lacks prosthetic groups as judged by ESR and ultraviolet/visible spectroscopy. Although not essential for electron flow from component C to component A (the hydroxylase), component B modifies the electron flow to O2catalysed by component A. In the absence of component B, components A plus C catalyse an “NADH oxidase” activity. By contrast, in the presence of component B, the reaction of component A (the hydroxylase) with methane is facilitated and all electron flow is diverted to the oxygenase reaction. Besides being essential for oxygenase function, an important second role of component B is that of preventing “wasteful” oxidation of NADH by “oxidase” activity in the absence of methane (Green and Dalton, 1985). A recent study of the mid-point redox potentials of the redox centres of component C has confirmed that they are consistent with its proposed role in accepting electrons from NADH and donating them to component A (Lund and Dalton, 1985). The sequence of electron transfer in component C has been further elucidated by stopped-flow experiments and by an investigation of the effects of removal and reconstitution of its flavin-adenine dinucleotide (FAD) and Fe2S2 redox centres (Lund et ul., 1985). This investigation has shown that the NADH :acceptor reductase activity requires FAD but not Fe2S2.The results of these experiments were consistent with the order of electron flow: NADH
- - FAD
Fe,S,
component A
This order suggests that component C functions as a 2e-/le- transformase, splitting electron pairs from NADH for transfer to component A by way of the one-electron-carrying FezS2 centre. A protein with properties similar to those of component C has also been purified and characterized from Methylobacterium sp. CRL-26 (Patel, 1984). A possible scheme for the overall hydroxylation process is given in Fig. 1. The site of binding and activation of O2is not yet known, but it is probably first converted into an electron-deficient, metal-bound O2species which then reacts with the substrate. The non-haem iron species of component A is clearly a prime candidate for the metal involved (see Section 1I.F for further discussion of the mechanisms). c. Component C. This component, characterized by Dalton and coworkers (Colby and Dalton, 1978, 1979; Dalton 1980b, 1981a; Lund and Dalton,
120
C.
ANTHONY Component B involved here
Reduced electron acceptors H+
+NADH>:FADx
+
c
[ F e 2 S 2 ] x F e 3 + ~ C H 3 0 H
reduced
H2O 02
NAD+
FADH2
I I
Fez+
[Fe2S2] oxidized
CH4
I
Electron
acceptors Component C
Component A
FIG. 1. Pathway of electron transfer between the components of the soluble methane mono-oxygenase complex during the oxidation of methane to methanol. This Figure is based on the work of Professor H.Dalton and his colleagues.
1985; Lund et al., 1985), is an iron-sulphur flavoprotein with a single polypeptide chain of molecular weight 44,000. Each molecule contains one molecule of FAD, two atoms of iron and two atoms of acid-labile sulphide. Core extrusion and ESR studies have shown that the iron and sulphide are present as a single iron-sulphide centre of the [2Fe-2S*(S-Cys),] type, as found in spinach ferredoxin and putidaredoxin: cys-s,
p,,
,S--CYS
Fe
Fe
cys-s
A/\ S
s-cys
These properties suggest that the single protein has a function analogous to, for example, the combination of putidaredoxin plus NADH-putidaredoxin reductase (a flavoprotein) in the hydroxlyation of camphor. Besides its apparent function in electron transport in methane hydroxylation, component C also has NADH-acceptor reductase activity. Thus its FAD component is reducible with dithionite or NAD(P)H, and it can be oxidized by ferricyanide, 2,6-dichlorophenolindophenol,horse heart cytochrome c or by stoicheiometric amounts of component A. The optimum pH value for this activity (assayed with indophenol) is 8.5-9.0 compared with 6.5-7.0for MMO activity. The & values for NADH and NADPH are 50 pi and 15.5 m, respectively,and the V,, values are 76units mg- ' for NADH and 7 units mg-' for NADPH, thus suggesting that NADH rather than NADPH
BACTERIAL OXIDATION OF METHANE AND METHANOL
121
is the natural electron donor. Unlike MMO activity, the reductase activity is not inhibited by 8-hydroxyquinoline (1 mM) or by acetylene (0.5 mM), and some preparations of component C lose their oxygenase activity but retain their reductase activity. The soluble MMO in cell-free extracts of Methylococcus capsulatus (Bath) is remarkably insensitive to a wide range of metal chelators and other inhibitors, the only potent inhibitors being the metal-chelating agent '8-hydroxyquinoline and the acetylenic compounds ethyne and propyne (Stirling and Dalton, 1977). In addition to these inhibitors, five other compounds inhibit oxidation in whole cells; these are the metal chelator diethylodithiocarbamate, o-aminophenol, ferron, cyanide and CO. Most of these probably act by inhibiting the supply of NADH to the mono-oxygenase within the cells, but the mode of action of CO is very complex (see Ferenci et al., 1975; Stirling and Dalton, 1977). The insensitivity of the MMO to metal-binding compounds other than 8-hydroxyquinoline suggests that if this is acting by chelating metal ions in the enzyme complex then these must be well-shielded from attack by other metal chelators. 2. The Particulate Methane Mono-oxygenase of Methylococcus capsulatus (Bath) Although all the early work on this organism indicated that the MMO is always a soluble enzyme, it is now known that the location and nature of the MMO depends upon the conditions of growth (Stanley et al., 1983; Dalton et al., 1984). In copper-limiting conditions, the activity is all in the soluble form and sodium dodecyl sulphate-polyacrylamide gel electrophoresis of the fractions shows clearly the protein bands corresponding to the three subunits of component A of the MMO. After being transfered to conditions of excess copper, these bands are markedly diminished and three new proteins can be detected in the particulate fraction, which is the only fraction that catalyses MMO activity in these conditions of excess copper. The three new membrane proteins have molecular weights that are completely distinct from those of the soluble MMO components. This indicates that changing the availability of copper does not merely alter the location of an otherwise identical MMO. This conclusion is supported by the demonstration that the sensitivity to inhibitors of the MMO changes with its location. The soluble MMO is inhibited only by ethyne and 8-hydroxyquinoline, but the particulate MMO is also inhibited by other compounds including cyanide, mercaptoethanol, 2,2-dipyridyl and thiourea; these were also effective against the particulate enzyme from Methylosinus trichosporiwn. A further indication that the particulate MMO of Methylococcus capsulatus is different from the soluble form is its substrate specificity. The
122
C.
ANTHONY
soluble enzyme is able to oxidize higher alkanes, cyclohexane and aromatic compounds (Table 3), but these are not oxidized by the particulate enzymes (Dalton et al., 1984). The particulate MMO was the only MMO produced during growth on methanol; both the concentration of MMO and the production of intracellular membranes were induced by raising the copper concentration in the growth medium (Prior and Dalton, 1985).
c. THE METHANE MONO-OXYGENASE OF Methylosinus trichosporium I . The Soluble Methane Mono-oxygenase from Methylosinus trichosporium Methylosinus trichosporiwn contains a soluble MMO which requires NADH as electron donor and which is similar in other respects to the soluble MMO from Methylococcus capsulatus (Stirling and Dalton, 1979a; Stirling et al., 1979). This has been confirmed by Scott et al. (1981a), who showed that the nature of the MMO in Methylosinus trichosporium depends on the growth conditions, which also determine the nature and extent of the internal membrane system in these bacteria. Particulate MMO is associated with the presence of extensive intracytoplasmic membranes, and was observed only under some OJimiting conditions. By contrast, soluble MMO is the only form of the enzyme found when bacteria are grown under most other conditions in which the internal membranes are fewer and less well organized, and vesicles predominate. A recent development of this work has shown that the key factor in Methylosinus trichosporium that governs both membrane synthesis and the site location of the MMO is the availability of copper (Cornish et al., 1984; Burrows et al., 1984); this is the same as is found with Methylococcus capsulatus (see above). The reason that this was not appreciated initially was that, under some conditions of continuous culture, the cell density was low and so the copper concentration was sufficient; whereas under other conditions the cell density was higher and so copper availability was diminished. The inhibitor sensitivity of the soluble MMO of Methylosinus trichosporium (Stirling and Dalton, 1979a; Scott et al., 1981a) is essentially similar to that of the soluble MMO of Methylococcus capsulatus (Colby and Dalton, 1976; Stirling and Dalton, 1977)and of Methylobacterium sp. CRL-26 (Pate1 et al., 1982). Thus, in all cases studied, the soluble forms of MMO are far less sensitive to inhibition by chelating agents, thiol agents and electron-transport inhibitors than are the particulate forms of MMO. The similarity of the soluble forms of MMO from two of these methanotrophs was demonstrated by reconstituting complete MMO activity from components B and C from Methylococcus capsulatus plus a fraction from Methylosinus trichosporiwn (Stirling and Dalton, 1979a).
BACTERIAL OXIDATION OF METHANE AND METHANOL
123
2. The Particulate Methane Mono-oxygenase from Methylosinus trichosporium OB3b Although Methylosinus trichosporiwn is able to synthesize a soluble MMO (above), the first description by Tonge et al. (1975,1977a) of the MMO from Methylosinus trichosporiwn indicated that it was particulate and that the solubilized protein components were very different from those of the soluble system . These workers purified a three-component system from Methylosinus trichosporium able to catalyse the oxidation of methane to methanol, the first step in its isolation being its removal from cell membranes by phosopholipase treatment. The purified components were protein 1 (molecular weight 47,000), containing one atom of copper per molecule, protein 2 (molecular weight 9 W ) , and soluble cytochrome c (molecular weight 13,000), which contained variable amounts of copper and which was able to react with CO (this cytochrome is discussed in Sections 1II.C and 1V.B). Equal amounts of each component were required for maximum activity (6 pmol min-' (mg of protein)-') measured at the optimum pH value for the system (pH 7.0). All components were inactivated by freezing. The K,,, value for methane was 6 6 p and ~ CO, ethane, propane and n-butane also were oxidized. Methane oxidation was highly sensitive to cyanide, chelating agents (especially those chelating copper), 2-mercaptoethanol and dithiothreitol. Similar inhibition by some of these compounds was also reported for the particulate types of MMO of Methylosinus CRL-15 (Pate1 et al., 1979b), Methylomonas methanica (Ferenci et al., 1975; Colby et al., 1975) and Methylococcus capsulatus (Texas) (Ribbons, 1975). In crude preparations, NADH was able to act as electron donor, but after purification it was necessary to use ascorbate which could be replaced by a mixture of methanol and partially purified MDH. These results led to the suggestion that, in intact bacteria, the cytochrome c is not only an oxidase (see Section IV.B) but also the electron donor to the oxygenase. It was also suggested that the cytochrome was able to bind methane during the hydroxylation reaction (Hammond et al., 1979). There is little direct evidence to support any of these proposed roles for cytochrome c in methane oxidation. Furthermore, recent attempts to purify this three-component MMO from Methylosinus trichosporiwn have been unsuccessful (Higgins et al., 1981c), and attempts to use ascorbate or methanol plus MDH as electron donors to cytochrome c, and hence to the MMO, have been uniformly unsuccessful (Pilayashenko-Novokhatnyi et al., 1979; Scott et al., 1981a; Stirling and Dalton, 1979a; Dalton et al., 1984). In summary, although Methylosinus trichosporium is able to form soluble or particulate MMO, the only electron donor for both these enzymes in vitro is NAD(P)H. Similarly, the only electron donor to the soluble MMO from
TABLE 3. The substrate specificity of the soluble methane mono-oxygenase from Methylococm capdatw (Bath). From Colby et al. (1977). Per cent of rate Substituted methane Per cent of rate CI-Cs derivatives with methane n-alkanes with methane Chloromethane Bromomethane Iodomethane Dichloromethane Trichloromethane Tetrachloromethane Cyanomethane Nitromethane Methanethiol Methanol Trimethylamine Carbon monoxide
99 78 0 97 41 0 39 53 75
289 0
72
Products
Ethane Propane Butane Pentane Hexane Heptane
81 82 92 87 48 87
Ethanol Propan-1-01 and propan-2-01 Butane-1-01 and butan-2-01 Pentan-1-01 and pentan-2-01, not pentan-3-01 Hexan-1-01 and hexan-2-01, not hexan-3-01 Heptan-1-01 and heptan-2-01; not heptan-3-01 or heptan4ol
Octane
11
Octan-1-01 and octan-2-01; not octan-3-01 or octan-4-01
C&, n-alkenes
Per cent of rate with methane
Ethene Propene But-1-ene cis-But-2-ene
176 99 58 68
trans-But-2-ene
168
Ethers Dimethyl ether Diethyl ether
295 54
Products
Alicyclic, aromatic Per cent of rate and heterocyclic compounds with methane
Epoxyethane 1,2-Epoxypropane 1,2-Epoxybutane
Cyclohexane Benzene Toluene
14 14
cis-2,3-Epoxybutane and cis-buten-1-01 trans-2,3-Epoxybutane and trans-2-buten-1-01
Styrene Pyridine L-Phenylalanine
56 35 0
63
Products Cyclohexanol Phenol Benzyl alcohol and cresol Styrene epoxide Pyridine N-oxide None
Not known Ethanol and ethanal
The enzyme system was crude soluble extract; NADH was electron donor; O2 was an absolute requirement, and oxidation of all substrates was inhibited by the 'specific' inhibitor ethyne (acetylene). The values are expressed as a percentage of the value with methane (85nmol of product formed mh-' (mg protein)-'). It should be noted that the relative rates are for the highest rates values were not determined. The products of oxidation measured with various amounts of substrate; they are not V,, values, and K,,, of substituted methane derivatives were not identified; rates of hydroxylation of these substrates were determined from the rate of disappearance of the substrates.
126
C.
ANTHONY
the facultative methanotroph Methylobacterium sp. CRL-26 was N AD(P)H, with neither ascorbate nor methanol plus MDH acting as electron donor. D. THE ELECTRON DONOR FOR METHANE MONO-OXYGENASE
There now seems little doubt that all forms of MMO, when measured in vitro, require NAD(P)H as electron donor, and there is no reason to consider that , this does not remain true in whole bacteria which contain only the soluble form of the enzyme. The situation may be different, however, in bacteria growing under conditions of high copper availability in which the MMO is particulate. It is, of course, very difficult to be certain of the nature of the electron donor in whole bacteria. The products of the activity of MMO must be further oxidized to provide electron donor for hydroxylation, or a second substrate must be made available for this purpose. As mentioned in previous sections, inhibitor studies of particulate forms of MMO have indicated that either electron transport is involved or that an extra inhibitor-sensitive component mediates between NADH and the oxygenase system. One obvious possibility is that electrons from MDH are used to reduce NAD’ to NADH by “reversed electron transport” from cytochrome c. There is some evidence that this might occur from studies using ethanol, which is oxidized by MDH, as electron donor in whole bacteria (Ferenci et al., 1975; Leak and Dalton, 1983; Dalton et al., 1984). Leak and Dalton (1983) concluded that their results are also consistent with the possibility that the donor to the particulate MMO might be an intermediate between cytochrome c and NADH, such as an iron-sulphur protein. That “reversed electron transfer” from MDH must occur in some photosynthetic methylotrophs has already been demonstrated (Anthony, 1982). It should be emphasized that growth yields of methanotrophs, in which NADH is an essential reductant for the MMO, are likely to be NADH limited. The growth yields would therefore be markedly greater if NADH could be produced by reversed electron transport from methanol by way of MDH and cytochrome c, or if NADH could be replaced by an alternative electron donor reduced via MDH (see van Dijken and Harder, 1975; and Anthony, 1978, 1982, for extensive discussions of the relationship between growth yields and the nature of the system for methane hydroxlyation). Final questions to raise, but not to answer here, in this context are: why do the bacteria respond to copper insufficiency by producing soluble MMO? Is this secondary to their failure to produce extensive internal membranes? Which MMO system, if any, is more “natural”? And is either MMO system better for biotechnological purposes?
BACTERIAL OXIDATION OF METHANE AND METHANOL
127
E. THE SUBSTRATE SPECIFICITY OF METHANE MONO-OXYGENASES
The substrate specificity of soluble MMO is different from that of particulate MMO in the same organism (see Dalton et ul., 1984; Burrows et ul., 1984), and this conclusion is consistent with the difference in specificity previously shown by those workers using Methylococcus cupsulutus (mainly soluble MMO) and those using Methylosinus trichosporium (mainly particulate
MMO).
The soluble MMO of Methylococcus cupsulutus (Bath) is very non-specific, and many of its substrates show little or no structural resemblance to methane (see Table 3 taken from Colby et al., 1977, in which there is extensive discussion of the significance of these results). The mono-oxygenase catalyses the hydroxylation of primary and secondary alkyl C-H bonds, the formation of epoxides from internal and terminal alkenes, the hydroxlyation of aromatic compounds, the N-oxidation of pyridine, the oxidation of CO to CO, and the oxidation of methanol to formaldehyde. The list of products in Table 3 demonstrates that some substrates can be attacked at more than one position (e.g. the but-Zenes and toluene). In addition to these substrates, the MMO is also able to oxidize methyl formate to formaldehyde and formate, and ammonia to hydroxylamine (Dalton, 1977; Stirling and Dalton, 1980). This last observation is analogous to the observation that methane and ethylene are oxidized by the ammonia mono-oxygenase of Nitrosomonas europueu (Hyman and Wood, 1983, 1984). The substrate specificity of the mono-oxygenase in crude extracts of Methylosinus trichosporium (soluble enzyme, measured with NADH as reductant) was very similar to that of Methylococcus cupsulutus (Bath), whereas that of the system in Methylomonus methunica was more limited and aromatic, alicyclic and heterocyclic compounds were not oxidized (Stirling et al., 1979; Burrows et ul., 1984). The only substrates oxidized by the mono-oxygenase in whole cells of Methylococcus capsulutus (out of those listed in Table 3) were those whose further oxidation yields NADH; these are methane, methanol, ammonia, chloromethane, bromomethane, dimethyl ether, ethene and propene (Stirling and Dalton, 1979b). The last three substrates were oxidized more rapidly in the presence of formaldehyde, whose oxidation yields NADH. This suggests that, in the absence of an exogenous supply of reducing power, the rgte of initial hydroxylation of these substrates was limited by the poor generation of NADH arising from their further oxidation. In addition to the eight substrates listed above, seven further substrates were oxidized by whole cells, but only in the presence of formaldehyde as a source of reductant (NADH); these were CO, diethyl ether, ethane, butane, but-l-ene, cis-but-2-ene and trans-but-Zene. The oxidation of substrates that are unable to support growth has been
128
C. ANTHONY
termed co-oxidation, this being a special case of the phenomenon of co-metabolism. Stirling and Dalton (1979b) have discussed the ambiguities associated with these terms and have proposed that co-metabolism be redefined as “the transformation of a compound, which is unable to support cell replication, in the requisite presence of another transformable compound (co-substrate)”. Thus, in the case of Methylococcus capsulatus (Bath), those seven compounds that are only oxidized in the presence of formaldehyde are co-metabolic substrates, the co-substrate being formaldehyde. It is suggested that oxidation of substrates (in the absence of co-substrate) that are unable to support growth (e.g. chloromethane, bromomethane, dimethyl ether, ethane and propene) is merely a reflection of the non-specific nature of the MMO, and that this oxidation should be termed “fortuitous oxidation”. This phenomenon is quite common and is analogous, for example, to the oxidation by whole cells of Pseudomonas M27 of about 20 primary alcohols which are unable to support growth, but which are good substrates for the MDH (see Table 6). With respect to the MMO, Higgins et al. (1980b) have suggested that such “extraordinary lack of enzyme specificity would be extremely unusual if it were entirely fortuitous” and they have argued that “this phenomenon has developed and been retained because of its survival value to the species”. It remains a matter of debate whether or not these oxidations and co-oxidations are entirely fortuitous, or whether some of them can be considered to be “supplementary metabolism” enabling these obligate methanotrophs to co-utilize other carbon and energy sources (see Stirling and Dalton, 1981; Higgins et al., 1981a, b, 1984b). Because of the wide substrate specificity of the MMO and because its substrates are relatively intractable to limited chemical oxidations, it offers a potentially valuable industrial catalyst for effecting these oxidations. This, and other aspects of substrate specificity of MMO forms found in a variety of methanotrophs, are discussed extensively in the following references: Colby et al., 1977; Stirling and Dalton, 1977; Higgins et al., 1979, 1980a, b, 1981b, 1982; Stirling et al., 1979; Hou et al., 1979a, b, 1980a, b, 1981, 1982a, b; Dalton, 1980a, b; Hazeu and de Bruyn, 1980; Patel et al., 1980, 1982; Anthony, 1982; Best and Higgins, 1983; Cornish et al., 1984; Hou, 1984a, b; Patel, 1984; Burrows et al., 1984; Dalton and Leak, 1985. F. THE MECHAMSM OF METHANE MONO-OXYGEWASE
Most of the discussion below concerns work on soluble MMO. The relationship of this work to particulate types of MMO is discussed by Dalton and Leak (1985). As demonstrated in the previous section, a remarkable feature
BACTERIAL OXIDATION OF METHANE AND METHANOL
129
of these mono-oxygenasesis the wide variety of substrates whose oxygenation they catalyse. The main categories to consider in this context are (a) alkanes yielding primary and/or secondary alcohols, (b) monosubstituted aromatic compounds yielding para-hydroxy derivatives and (c) alkenes yielding epoxides, which are sometimeschemically stable. This makes the mechanism particularly interesting, and it provides some useful approaches to solving the mechanism although, as for all hydroxlation reactions, a definitive mechanism is difficult to prove. Some substrates are hydroxylated in more than one position, probably by more than one mechanism. For example, the saturated side chain of ethylbenzene is oxidized to the primary alcohol, whereas the aromatic ring is oxidized to give a hydroxy group in the para position. The simpler type of hydroxylation occurs when a double bond is involved (aliphatic or aromatic; sp2 hydroxylation). In this case, the most likely mechanism involves direct attack by an enzymically activated electrophilic oxygen species on the electron-rich system, giving rise to an epoxide intermediate. Evidence for such a mechanism has been obtained by demonstrating a hydride (NIH) shift during the hydroxyation of aromatic substrates by the MMO of Methylococcus cupsulutus (Dalton, 1981; Dalton et ul., 1981, 1984) and Methylosinus trichosporium (Jezequel and Higgins, 1983; Higgins et ul., 1984a; Dalton and Leak, 1985). This conclusion is supported by the demonstration that stable epoxide products are formed during the oxidation of some unsaturated substrates by MMO (Colby et ul., 1977; Higgins et al., 1979). A different mechanism must occur during hydroxylation of a saturated carbon atom (sp3hydroxylation). Two such mechanisms might occur; either a concerted single-step insertion process, or a two-step mechanism consisting of a preliminary hydrogen abstraction followed by hydroxylation (Dalton and Leak, 1985). Jezequel and Higgins (1983) have concluded that the latter (two-step)mechanism probably occurs during the hydroxylation of saturated carbon atoms by MMO; and these authors suggest that the same enzyme component involved in one of these two steps might also be involved in the hydroxlylation of unsaturated (sp2)substrates. By contrast, Dalton and Leak (1985) have argued that a concerted mechanism (rather than a two-step mechanism) is more likely.
HI. Oxidation of Methanol to Formaldehyde A. METHANOL DEHYDROGENASE
I. Introduction Methanol oxidation in bacteria is usually catalysed by the NAD+-independent alcohol dehydrogenase described originally in Pseudomonus M27 (Anthony
130
C. ANTHONY
and Zatman, 1964a, b). Although not specific for methanol, its usual functions is to catalyse methanol oxidation and so it is usually referred to as methanol dehydrogenase (MDH; EC 1.1.99.8). Possession of this enzyme is one feature that appears to be common to almost all methane and methanoloxidizing bacteria; Table 4 gives references to the enzyme from a wide range of different methylotrophs and Table 5 summarizes their remarkably similar properties. Typically, these enzymes oxidize a wide range of primary alcohols using phenazine methosulphate as artificial electron acceptor and ammonia or methylamine as activator. The pH optima are pH 9 or higher, and they are often stable at pH 4.0. They are usually dimers of identical subunit molecular mass of 60,000 Da. The specific activity of MDH in crude extracts from different bacteria varies over a wide range (between 4 and 1300nmolmin-'mg-', but usually between 60 and 600nmolmin-'mg-'); and the specific activities of the purified enzymes also vary considerably (0.3-18 pmol min-' (mg protein)-'; see Goldberg, 1976; Bamforth and Quayle, 1978a, b). These results reflect to some extent the variety of growth conditions and methods of cell breakage and enzyme assay, but they also suggest a genuine range of activities in methylotrophs. MDH is often induced during methylotrophic growth, when it usually constitutes between 5 and 15% of the soluble protein. This indicates its importance to the growth of methylotrophs, and this is confirmed by the isolation of mutants that lack the dehydrogenase and have lost the ability to grow on methane or methanol (Heptinstall and Quayle, 1970; Dunstan et ul., 1972; OConnor and Hanson, 1977). A MDH that differs from those described in Table 5 has recently been isolated from Nocurdia sp. 239, a Gram-positive organism that appears to lack a typical MDH (Hazeu et ul., 1983). The only methanol-oxidizing activity detected was due to a PQQ-containing MDH that was present in a multi-enzyme complex, together with NAD' -dependent aldehyde dehydrogenase and NADH dehydrogenase (Duine et ul., 1984a). Methanoldependent dye reduction catalysed by this complex required NAD'; but this was either not reduced to NADH by methanol, or it was not released from the enzyme after reduction. The preliminary results published at present do not necessarily demonstrate that the MDH in this organism catalyses the methanol-dependent reduction of NADH, but if the enzyme does do so then this might have important consequences with respect to the bioenergetics of any bacteria having such an enzyme. 2. The Primary Electron Acceptor The physiological electron acceptor for MDH is probably always
BACTERIAL OXIDATION OF METHANE AND METHANOL
131
cytochrome c (Section III.C), but for assay of the extracted enzyme it is usually necessary to use the artificial electron acceptor phenazine methosulphate. In the enzyme assay, the re-oxidation of reduced phenazine methosulphate is coupled to reduction of 2,6-dichlorophenolindophenol(measured spectrophotometrically) or 0,(measured with an O2electrode). The only conventional electron acceptor to replace the phenazine methosulphate is phenazine ethosulphate. It has been suggested by Ghosh and Quayle (1979) that phenazine ethosulphate should always be used as electron acceptor in assaying dye-linked dehydrogenases (rather than phenazine methosulphate) because it does not reduce the indophenol non-enzymically. They showed that phenazine derivatives form free radicals in alkaline solution, and it has been suggested that the free radical may be the true electron acceptor in the MDH assay system (Duine et al., 1978; Ghosh and Quayle, 1979). This suggestion is supported by the demonstration that alternatives to phenazine methosulphate and phenazine ethosulphate, as electron acceptors, are the free radicals produced by the one-electron oxidation of NNN' N'tetramethyl-p-phenylenediamine(Wurster's blue, TMPD) and of 2,2'-azinodi-(3-ethylbenzthiazoline-6-sulphonicacid) (Duine et al., 1978).
3. Substrate Specificity of Methanol Dehydrogenase The dehydrogenase has a wide but well-defined specificity; only primary alcohols are oxidized and their steric configuration is more important in determining whether or not they are oxidized than the presence or absence of atoms or groups producing electron-displacement effects (Anthony and Zatman, 1965; Sperl et al., 1974; Duine and Frank, 1980a).Using the enzyme from Pseudomunas M27, it was shown that a second substituent on the C-2 atom appears to prevent binding, the general formula for an oxidizable substrate being R.CH,OH where R may be H, OH (as in hydrated R RC=CH(Anthony and Zatman, 1965; aldehydes), R CH,-or Table 6). The rate of oxidation of these substrates is usually at least 30% of that with methanol, which is the best substrate. The K, value for methanol , the affinity for the enzyme often decreases with is usually low (10-20 p ~ )and increasingsize of the alcohol. Whole bacteria usually oxidize the same range of alcohols as are oxidized by the pure enzyme, and their oxidation is inhibited by ethylenediaminetetra-acetate (EDTA), phenylhydrazines and high concentrations of phosphate which are inhibitors of methanol oxidation in whole cells. Transport of the larger substituted alcohols into the organism is unlikely to be a problem if, as is probably always the case, the MDH is on the outer surface of the cytoplasmic membrane, or in the periplasmic space. Although most of these dehydrogenaseshave a similar substrate specificity to that of Pseudomonas M27, there are some minor differences. For example, it was shown that Hyphomicrobium sp., Pseudomonas TP-1 and Pseudomonas
TABLE 4. Methanol dehydrogenase Source of methanol dehydrogenase Methad-ntilizers (facdtative) Pseudomonas M27 Pseudomonas AM1 Pseudomonas extorquens Protaminobacter ruber Pseudomonas PP Pseudomonas TP-1 Pseudomonas RJ1 Pseudomonas 526 Pseudomonas 294 1 Pseudomonas S25 Strain S50 (Acinetobactersp.) Organism PAR Parococw denitrijicans
References Anthony and Zatman (1964a, b, 1965, 1967a, b), Pate1 et al. (1972, 1973) Johnson and Quayle (1964), Heptinstall and Quayle (1969), OKeeffe and Anthony (1980a, b), Bolbot and Anthony (1980), Beardmore-Gray et al. (1983); BeardmoreGray and Anthony (1984), Froud and Anthony (1984a, b), Ford et al. (1985) Johnson and Quayle (1964) Johnson and Quayle (1 964) Ladner and Zatman (1969) Sperl et al. (1974) Mehta (1973) Michalik and Raczynska-Bojanowska (1976) Yamanaka and Matsumoto (1977a, b; 1979), Yamanaka (1981) Yamanaka and Matsumoto (1977a, 1979), Yamanaka (1981) Yamanaka and Matsumoto (1977a, 1979), Yamanaka (1981) Bellion and Wu (1978) Bamforth and Quayle-(1978a),Alefounder and Ferguson (1981); Beardmore-Gray et al. (1983) Sped et al. (1974), Harder and Attwood (1975), Duine et al. (1978, 1980, 1981), Duine and Frank (1980a, 1981a); de Beer et al. (1983) Sahm et al. (1976), Bamforth and Quayle (1978b, 1979), de Beer et al. (1979) .
Hyphomicrobium Rhodopseudomonas acidophila Methanol-otilizers (obligate) Pseudomonas W 1 Pseudomonas C Methylomonas P11 Methylophilur methylotrophus
I
Sperl et al. (1974) Goldberg (1976) Michalik and Raczynska-Bojanowska (1976), Drabikowska (1977) Ghosh and Quayle (1981), Cross and Anthony (1980b), Ghosh (1980), BeardmoreGray (1982), Beardmore-Gray et d. (1983), Froud and Anthony (1984a, b)
Strain 4025 Methylomonus J Metbanotrophs Methylococcus capsulutus Methylomonas methanica Methylosinus sporium Methylobacterium organophilum Methylobacterium R6
Vrdoljak and Froud (1982) Ohta et al. (1981), Ohta and Tobari (1981)
Patel and Hoare (1971), Patel et al. (1972, 1973), Wadzinski and Ribbons (1975) Johnson and Quayle (1964), Patel et al. (1978a), Mincey et al. (1981), Parkes and Abeles (1984) Patel and Felix (1976) Wolf and Hanson (1978) Patel et al. (197813)
The MDH enzymes from these bacteria are similar in most respects; they all oxidize a wide range of primary alcohols for which they usually have a very high affinity; they use phenazine methosulphate as primary hydrogen acceptor; they use ammonia or methylamine as activator; and they have a high pH optimum. The properties of these enzymes are summarized in Table 5.
TABLE 5. Summary of properties of the methanol dehydrogenases listed in Table 4 Source of methanol dehydrogenase Group A Pseudomonas AM 1 Pseudomonas 294 1 Pseudomonas S25 Pseudomonas M27, Pseudomonas W1, Hyphomicrobium and strain 4025 Pseudomonas RJ1 and Pseudomonas TP-I Methylophilus methylotrophus Methylococcus capsulatus Methylomonas J Group B Methylobacterium organophilum Pseudomonas C Diplococcus PAR Group C Paracoccus denitrijicans Strain S50 Group D Methylomonas methanica and Methylosinus sporium Group E Rtiodopseudomonas acidophila
Molecular weight
Subunit molecular weight
120,000 128,000 128,000 120,000
60,000 62,000 62,000
8.8 7.38 9.4 High
120,000 I 15,000 120,000 135,000
62,000 62,000 60.000
High High 9.3
135,000 128,000 112,000
62,000 60,000 56,000
151,000 158,000
76,000 76,000
3.7 3.82
60,000
60,000
High
1 16,000
63,000
Oxidation of secondary alcohols
+ + +
+
Isoelectric point
High
9.35
The division into groups is rather arbitrary and sometimes based on incomplete or preliminary descriptions. Molecular weights of whole enzymes are based on gel filtration. A “high” isoelectric point is above 7.0 and is sometimes based only on observations during ion-exchange chromatography.
cd
+
9
X I 0-v-X I
X
0
dc
X
0,
8I I
X S u-u
8
X
0,
B I
X S u-u I 3 u
1,2-Propanediol
Glycerol
CHZ-CH--CH,OH AH
Pentafluoropropanol
CF3CF2CH20H
AH
This table demonstrates the wide range of alcohols oxidized by this one enzyme. They are usually oxidized at similar rates to those measured with methanol, but their affinities for the MDH vary (see also Sped et al., 1974). Representative alcohols are given; not all of those tested are included. Almost all those alcohols that are oxidized by the pure enzyme are also oxidized by whole bacteria. Those marked with an asterisk have a low afl6nity for this enzyme; but this affinity is increased by a modifier protein (see the text). No secondary or tertiary alcohols are oxidized.
138
C. ANTHONY
W1 contain enzymes that oxidize alcohols substituted with a methyl group on the second carbon atom; these enzymes have a relatively low affinity for the substituted alcohols but still oxidize them at high rates (Sperl et al., 1974). Furthermore, preliminary observations suggest that some secondary alcohols may be substrates for the dehydrogenasesfrom the facultative methylotroph Methylobacteriumorganophilum (Wolf and Hanson, 1978),from Pseudomonas C (Goldberg, 1976)and from an uncharacterizedmethanol utilizer (organism PAR; Bellion and Wu, 1978). The most unusual MDH with respect to substrate specificity, and indeed with respect to other properties, is that from the photosynthetic methylotroph Rhodopseudomonas acidophila (Sahm et al., 1976; Bamforth and Quayle, 1978a, 1979);it oxidizesprimary and secondary alcohols at similar rates, although its affinity for secondary alcohols is lower. This enzyme has the greatest affinity for ethanol (&6 q) and the lowest affinity for methanol (K, 57m~),which is rather remarkable for an enzyme initiating the oxidative attack on the growth substrate. As expected from the general formula for an oxidizable substrate (see above), 1,2-propanediol, having two substituents on its C-2 atom, is not oxidized by pure MDH. In spite of this, studies on the metabolism of 1,Zpropanediol by Pseudomonas AM 1 unexpectedly demonstrated that MDH is involved in the oxidation of this substrate; mutants lacking MDH or cytochrome c neither grew on propanediol nor oxidized it. Preliminary results indicated that a second protein modifies the pure dehydrogenase, enabling it to bind primary alcohols with a substituent hydroxyl residue on the C-2 atom and to oxidize them in two consecutive steps to the carboxylic acid (Bolbot and Anthony, 1980). We have now shown that this unexpected oxidation of 1,Zpropanediol to lactate requires two proteins in addition to MDH (Ford et al., 1985). The first is a “dye-linked‘’ aldehyde dehydrogenase, able to oxidize lactaldehyde to lactate. The second is a modifier protein (M protein) that binds to MDH, increasing its affinity for a number of alcohols, including 1,2-propanediol. In the absence of M protein, the affinity of MDH for propanediol is so low that this substrate fails to protect the dehydrogenase against inactivation by phenazine methosulphate (PMS); in the presence of M protein, the affinity of the MDH for propanediol increases, the alcohol binds to the enzyme which oxidizes it to lactaldehyde: pM(
CHI CHOH
I
CH,OH
P M C JH;
JSH-l:kH
CHI CHOH
I
CHO Methanol dehydrogenase plus ‘Mprotein’
I
COOH Aldehyde dehydrogenase
BACTERIAL OXIDATION OF METHANE AND METHANOL
139
Other alcohols whose affinity for MDH is increased by the M protein are l,Zbutanediol, 1,3-propanediol, 1,3-butanediol, 3-methylbutan-1-01 and Chydroxbutyrate. This last substrate is a growth substrate for Pseudomonas AMI, and its metabolism has been shown to be very similar to that of 1,2propanediol; that is, MDH appeared to be essential for its oxidation but pure enzyme is unable to oxidize it (Cox and Quayle, 1976). It thus appears that both 1,Zpropanediol and 4-hydroxybutyrate are oxidized by MDH in the presence of M protein, which thus allows growth on these substrates. Despite this apparent physiological role of the M protein, further evidence suggests that its main function is not to facilitate growth on these multicarbon compounds, but to regulate MDH with respect to the oxidation of formaldehyde (see below). That the role of this protein in the growth of Pseudomonas AM1 on propanediol is fortuitous, and secondary to its main role, is supported by the observation that the M protein occurs in a range of methylotrophs, including obligate methylotrophs (Methylophilus methylotrophus and Methylosinus trichosporium) which are unable to grow on multicarbon compounds such as 1,2-propanediol and 4-hydroxybutyrate. A common characteristicof MDHs is their ability to catalyse the oxidation of formaldehyde to formate, first shown by Ladner and Zatman (1969) and Heptinstall and Quayle (1969). The rate of formaldehyde oxidation is usually similar to that of methanol oxidation, and the affinity of the ezyme for the two substrates is often similar. Although formaldehyde is often the only aldehyde oxidized, acetaldehyde, trifluoroacetaldehyde and trichloacetaldehyde are also sometimes substrates. The MDH from Rh. acidophila is again unusual, catalysing the oxidation of formaldehyde, acetaldehyde and propionaldehyde. Although the rates are similar to those measured for ethanol, the best substrate, the affinity of the enzyme for aldehydes is relatively low (1% of that for ethanol; Sahm et al., 1976; Bamforth and Quayle, 1978b). It has been suggested that the actual substrate during formaldehyde oxidation is the gem-diol hydrated aldehyde and that the extent to which other aldehydes are oxidized may be related to their degree of hydration (Sperl et al., 1974). Duine and Frank (1981a) have argued, however, that the rather sharp break observed in the aldehyde substrate specificity spectrum of those methanol and alcohol dehydrogenases able to oxidize aldehydes cannot be accounted for by the extent of hydration because this follows a gradual change through the range of aldehydes. Instead, they suggest that the results can best be explained by assuming a dual-substrate specificity, one for the alcohol and the other for the aldehyde substrate (Duine and Frank, 1981b). It is worth mentioning that the oxidation of formaldehyde is a feature not only of the enzyme when measured in the dye-linked assay, but also in the physiological system using cytochrome c as the electron acceptor
140
C. ANTHONY
(Beardmore-Gray et al., 1983; Section 1II.D). The oxidation of formaldehyde by whole bacteria, however, is not inhibited by inhibitors of methanol oxidation, and mutants lacking MDH still oxidize formaldehyde. This demonstration that MDH is not responsible for oxidizing formaldehyde in methylotrophs indicates that the dehydrogenase must be regulated to prevent any formaldehyde oxidation which might otherwise occur (see below, Section A.8). A final point to make about the substrate specificity of MDH is that, even after purification and extensive dialysis, the enzyme catalyses a rapid and unexpectedly large reduction of added artificial electron acceptor in the absence of any added substrate (Anthony and Zatman, 1964b; Goldberg, 1976; Bamforth and Quayle, 1978b; Duine et al., 1978). That this is not all bound methanol has been shown using the enzyme from M . methylotrophus (Ghosh and Quayle, 1981); each dimeric molecule was shown to have two bound molecules of methanol (or formaldehyde) and, in addition, a further 90 molecules of an unidentified endogenous reductant. Incubation of the enzyme with phenazine methosulphate and ammonia, in the absence of substrate or alternative protecting molecules such as cyanide, led to oxidation of the endogenous reductant but always yielded an inactive enzyme (Duine and Frank, 1980a). The unusual endogenous reduction catalysed by MDH is not only seen when it is assayed with artificial electron acceptors; when pure MDH is used to catalyse the reduction of cytochrome c by methanol, it is always necessary to first oxidize the endogenous reductant before any methanol-dependent reduction of the cytochrome c can be measured (see Section 1I.D). 4. Activators of MethanoI dehydrogenase
When prepared aerobically, MDH, assayed with phenazine methosulphate, has an absolute requirement for ammonium salts, which can be replaced by methylamine but not by di- or tri-amines nor by long-chain alkylamines. The relatively higher concentrationsof ammonium salt required at lower pH values suggests that the free base is the active species (Anthony and Zatman, 1964b). The MDH from Methylomonas PI I is exceptional in being active only with ammonia and not with methylamine, and that from Methylobacterium orgunophilum has considerable ammonia-independent activity (2040% of that in its presence; Wolf and Hanson, 1978). The MDH from Rh.acidophila is unusual in being activated by ammonia and a wide range of primary alkylamines. Although the highest rate is obtained with ammonia, the atfinity for the amine activator increases with increasing chain length, the II;, value for nonylamine being 26 p~ and that for ammonia being 42 m ~ . The MDH from Hyphomicrobium X is activated by ammonia and by esters
BACTERlAL OXIDATION OF METHANE A N D METHANOL
141
of glycine or 8-alanine, but not by lysine esters nor by aliphatic amines or amino acids; the K, value for glycine (0.8 mM) was 30 times lower than that for ammonia (Duine and Frank, 1980~).A similar result was obtained using the MDH from M . methylotrophus; in this case the K , value for ammonia was 8m~ and the K, value for ethylglycine was 1.4mM (M. Beardmore-Gray and C. Anthony, unpublished results). Although ammonia activation is such a well-defined and universal characteristic of MDH, this may not be a feature of the activity of the MDH within the cell. The first evidence for this was obtained using extracts of Hyphomicrobium X which were prepared anaerobically and partially purified under anaerobic conditions (Duine et al., 1979a). The MDH in these extracts no longer required ammonia for activity until they were exposed to air for some time, after which ammonia became essential. A similar result was obtained using some anaerobic preparations of Pseudomonas AM 1 but, more frequently, results with extracts of this organism resembled those obtained with extracts of M . methylotrophus; anaerobic preparations of this organism always required ammonia in the dye-linked assay (Beardmore-Gray and Anthony, 1984). It has now been shown, using completely pure proteins, that the natural physiological electron acceptor for MDH is a cytochrome c (BeardmoreGray et al., 1983; Beardmore-Gray and Anthony, 1984). When assayed at pH 7.0, using this electron acceptor, no activator was required. It is possible therefore that the amine groups required to measure activity of MDH at high pH values, using phenazine methosulphate as electron acceptor, may be “replacing” the lysine residues that usually form an important part of the binding face of cytochrome c. Whatever the mechanism of ammonia activation, it appears that high pH values and ammonia are only essential for re-oxidation of the reduced enzyme, and not for the reduction of enzyme by its substrate (Duine and Frank, 1980a, 1981a). 5. Inhibitors of Methanol Dehydrogenase
Very few inhibitors of MDH are known. The oxidation of methanol by whole bacteria is inhibited by EDTA, p-nitrophenylhydrazine and high phosphate concentrations, but these compounds do not inhibit the isolated enzyme when assayed with the artificial electron-acceptor phenazine methosulphate (Anthony and Zatman, 1964b; Anthony, 1975). It is thought that the EDTA acts either by removing a divalent metal ion essential for binding MDH and cytochromes c to the bacterial membrane (Carver and Jones, 1984; Carver et al., 1984), or that it affects the binding of dehydrogenase to cytochrome c (Beardmore-Gray and Anthony, 1984; this is discussed further in Section 1II.D. 1).
142
C. ANTHONY
Oxidized phenazine methosulphate is a potent irreversible inhibitor of most MDH enzymes, complete inactivation occurring within a matter of seconds (Anthony, 1963; Cross, 1980; Beardmore-Gray, 1982). Methanol completely protects the enzyme from inactivation, and it is possible that the endogenous reductant usually present may also afford some protection before being removed by its oxidation. A second inhibitor of the dehydrogenaseis cyclopropanol, which irreversibly inactivates the enzyme by reacting with the oxidized form of the prosthetic group (Dijkstra et al., 1984; Groen et al., 1984; Groenveld et al., 1984; Section III.B.5). Cyanide reversibly inhibits some MDH enzymes, but the mechanism is not always the same and it is sometimes only observable when the enzyme is assayed in the presence of low concentrations of substrate or activator. The enzyme from Hyphomicrobium X is inhibited by KCN ($ l m ~ ) ,the inhibition being competitive with respect to substrate (Duine and Frank, 1980a). This is also true for the M . methylotrophus enzyme, the & value for KCN being 1 2 m ~ this ; concentration also completely abolished the oxidation of endogenous reductant, and it was sufficient to stabilize the enzyme against inactivation during purification in the absence of methanol (Beardmore-Gray, 1982; Beardmore-Gray et al., 1983). The enzyme from Paracoccus &nitr@cans showed a similar sensitivity to KCN, 50% inhibition being measured at 5.6 mM KCN (Bamforth and Quayle, 1978a). As is the case with respect to many other characteristics, the MDH from Rh. acidophila differs from other dehydrogenases with respect to inhibition by KCN; although the inhibition is competitive in nature, the KCN competes not with substrate but with the amine activator for its binding site (Bamforth and Quayle, 1978b). This site on the dehydrogenaseis also responsible for binding the metal-chelating agent 2,T-bipyridine (Ki 1.5 m~), which is a less potent inhibitor of other MDH enzymes. The MDH from Rh. acidophila also differs from the other dehydrogenases in that O2 is a competitive inhibitor with respect to methanol. Bamforth and Quayle (1978b) concluded that there is an electrophilic site on the enzyme which binds both an amino group from the activator and certain metal-chelating agents, and that there is also an electrophilic site which binds the alcohol and which can also competitively . speculated from this that “both sites are at the active-centre bind 0 2They of the enzyme which contains a metal group, this metal group being involved in binding of the above four ligands”. Although consistent with their observations, however, there has been no further evidence published that supports the view that a metal group is included at the active site of this or any other MDH.
BACTERIAL OXIDATION OF METHANE AND METHANOL
143
6. Molecular Weight, Isoelectric Point and Amino Acid Composition of Methanol Dehydrogenases, and Serological Relationships Between them
Methanol dehydrogenases have molecular weights between 112,000 and 158,000 and can be dissociated by low pH values or by sodium dodecyl sulphate to two identical subunits of molecular mass 56,000-76,000 Da (see Table 5). The dehydrogenases from the methanotrophs Methylomonas methanica and Methylosinus sporium are exceptions in being monomers of molecular weight about 60,000 (Patel and Felix, 1976; Patel et al., 1978a). It has been pointed out that these monomeric enzymes are only likely to have one prosthetic group per molecule and that they might not therefore be able to oxidize methanol to formate, but only to formaldehyde (Duine and Frank, 1981a). Most of these dehydrogenaseshave high isoelectric points (7-10.5), and the exceptions happen to be those having exceptionally high molecular weights (Table 5). They are acid-stable at pH 4.0, except for those from Methylosinus sporium (Patel and Felix, 1976), Pseudomonas Wl (Sperl et al., 1974) and Methylomonas J (Ohta et al., 1981). Amino-acid compositions of the dehydrogenases from the following bacteria have been published; Pseudomonas M27 (Anthony and Zatman, 1967a), Methylosinus sporium (Patel and Felix, 1976), Pseudomonas C (Goldberg, 1976), Methylomonas methanica (Patel et al., 1978a), Rh. acidophila (Bamforth and Quayle, 1979), Methylomonas J (Ohta et al., 1981) and M . methylotrophus (Beardmore-Gray, 1982). Amino-acid compositions are similar in most respects but, as might be expected, they shed little light on the mechanism of MDH. All are deficient in free thiol groups, a property reflected in their insensitivity to inhibitors such as iodoacetic acid and p-chloromercuribenzoate (Anthony and Zatman, 1965; Goldberg, 1976; Bamforth and Quayle, 1978b). A consideration of the relationship between isoelectric points and amino-acid compositions shows, again as expected, that dehydrogenases having lower lysine contents tend to be those with lower isoelectric points (Yamanaka and Matsumoto, 1977a, b; Yamanaka, 1981). Antisera prepared against pure MDH from Methylococcus capsulatus (a methanotroph having the ribulose monophosphate (RuMP) pathway) and from Pseudomonas M27 (a typical pink methanol-utilizer with the serine pathway) have been used to allocate a number of methane- and methanolutilizing bacteria into various groups that are satisfyingly similar to those proposed by conventional methods (Patel et al., 1973; 1978a;Patel and Felix, 1976). The type I methanotrophs, having the RuMP pathway, were similar to one another, but different from the type I1 methanotrophs having the serine pathway; these showed some similarity to the facultative methanol-utilizers which also have the serine pathway. The MDH from the obligate methanolutilizer organism W 1,having the RuMP pathway, was serologically distinct
144
C. ANTHONY
from all other such dehydrogenases. A similar study, using antisera produced against pure MDH from the facultative methane-utilizer Methylobacteriwn organophilum (serine pathway), showed, as expected, that this enzyme is more similar to the enzymes from the facultative methanol-utilizers, and from serine pathway methanotrophs, than to those from Methylococcus capsulatus (an RUMP pathway methanotroph) or from the facultative autotroph Rh. acidophila (Wolf and Hanson, 1978). A further serological study of the MDH from Rh. acidophila by Bamforth and Quayle (1979) emphasized the uniqueness of this enzyme; there was no cross reaction between the antiserum raised against it and the purified enzyme from P . denitriJicans and M . methylotrophus or crude extracts of Pseudomonas AM 1 and Hyphomicrobium X. Similarly, the antisera raised against the purified MDH from P. denitrijicans and M . methylotrophus showed no cross reaction with the pure enzyme from Rh. acidophiia. 7. Location of Methanol Dehydrogenase
Although MDH from most bacteria is found in the soluble fraction after cell breakage, it is probable from its function that it binds to the membrane in which the components of the electron transport chain are situated. As the MDH often constitutes about 10% of the protein in the cell extracts, it is perhaps unlikely that it all arises from the membrane during cell breakage. Breakage of cells in a French Press, or by sonication, usually provides sufficientenzyme for purification, and the membrane fraction is often discarded and not assayed for the presence of MDH. When it is measured, however, some activity is found in the membrane fraction, the amount remaining being affected by the treatment of the membrane fraction. MDH has been found on membranes, or perhaps enclosed in membrane vesicles, in a wide variety of bacteria including Methylococcus capsulatus (Wadzinski and Ribbons, 1975), Hyphomicrobium X (Duine et al., 1978), Pseudomonas AM1 (Netrusov and Anthony, 1979) and M . methylotrophus (Cross and Anthony, 1980b; Froud and Anthony, 1984a). Probably all “methanol oxidases” reported to be present in crude membrane preparations of methylotrophs have been due to membrane-bound MDH. A particularly important study of membrane-bound dehydrogenases is that of Wadzinski and Ribbons (1975; Ribbons and Wadzinski, 1976) who have shown that MDH released by detergent treatment of membranes (about 60% of the total MDH) of Methylococcus capsulatus has identical properties after purification to those of the soluble form. A similar result has been obtained using MDH purified from Triton extracts of membranes from the obligate methylotroph M . methylotrophus (Froud and Anthony, 1984a). A comparison of the binding of MDH to membranes of Type I and Type
BACTERIAL OXIDATION OF METHANE AND METHANOL
145
II methanotrophs showed that about 60% remained on the membranes prepared by French Press extraction of Type I methanotrophs (Methylococcus, Methylobacter and Methylomonas spp.), whereas all is found in the soluble fraction of the Type I1 methanotrophs (Methylosinus and Methylocystis spp.) (Pate1 and Felix, 1976). Once again, this result probably reflects the methods used for growth, or for extraction, because other workers have found that some of the MDH of Methylosinus trichosporium remains bound to membranes (Tonge et al.; 1975, 1977a). When bacteria are treated with lysozyme and EDTA, in the presence of osmotic stabilizers such as mannitol, much of the outer wall can be removed leaving intact sphaeroplasts and releasing the periplasmic fraction-that part of the bacteria between the inner cytoplasmic membrane and the outer wall. When the methanol utilizers P . denitrificans and M . methylotrophus were treated in this way, most of the MDH was released in the periplasmic fraction together with much of the soluble cytochrome c (Alefounder and Ferguson, 1981; Jones et al., 1982; Burton et al., 1983). This result shows that the MDH is either freely diffusible in the periplasm or, more likely, loosely bound to the outer side of the cytoplasmic membrane. An alternative approach to the localization of dehydrogenases, by radiochemically labelling ['4C]isothionylacetamide,also led to the conclusion that MDH is periplasmic in Hyphomicrobium X and bacterium W3A1 (Kaspnak and Steenkamp, 1983). This approach has also been used by Quilter and Jones (1984) to confirm the periplasmic location of MDH and soluble cytochrome c in M. methylotrophus. This conclusion is consistent with all work on electron transport and energy transduction in these methylotrophs (for discussion of the location of MDH, in the context of energy transduction, see Section 1V.F). It should be noted that, if the MDH of methanotrophs is also in the periplasmic space of the bacteria, and if the MMO is in the cytoplasm or on the cytoplasmic side of the cytoplasmic membrane, then the methanol produced by the oxidation of methane will have to pass out of the cytoplasm into the periplasmic space. After oxidation by the MDH, the formaldehyde produced will then have to reenter the bacteria. There appears to be no information on the possibility of a periplasmic location of MDH in methanotrophs. A similar situation must also occur during oxidation of trimethylamine to formaldehyde by way of methylamine dehydrogenase in M . methylotrophus (Burton et al., 1983). 8. Regulation of Methanol Dehydrogenase Activity Because MDH is coupled to the electron transport chain at the level of cytochrome c, the rate of oxidation of methanol to formaldehydewill depend on the concentrations of methanol and of oxidized cytochrome c. Because the
146
C. ANTHONY
affinityof the enzyme for methanol and cytochrome is so high, and because the rate of flux through electron transport chains is usually determined by the activity of the primary dehydrogenases, it is probable that MDH needs to be regulated in some more specific manner. The mode of regulation likely to be involved is not immediately obvious because of the variety of possible “end products” of its activity. In some organisms the immediate product of MDH activity, formaldehyde, is oxidized by way of formate to C02, whereas in others the route for its oxidation is a cyclic modification of the assimilation route. Assimilation routes also vary; in some cases the methanol growth substrate is assimilated at the level of C 0 2(ribulose bisphosphate pathway); in others it is assimilated at the level of formaldehyde by way of an enzyme that uses formaldehydeitself as substrate (hexulose phosphate synthase in the RUMPpathway); and, in the serine pathway, part of the carbon is assimilated as CO, and part as formaldehyde bound to tetrahydrofolate (methylenetetrahydrofolate). Clearly, if MDH is regulated by the concentrations of its end products, then MDHs, although similar in most other respects, are likely to vary considerably with respect to their regulation. This being the case it is noteworthy that nothing of substance has ever been published about its regulation. In the context of regulation of MDH, it is worth emphasizing the peculiar importance of formaldehyde, a potentially lethal metabolite reacting nonenzymicallywith many metabolites and polymers in living organisms. During growth on methane, or methanol, it is a product of MDH but it is also a substrate for this enzyme (Section II.E), which in some conditions can oxidize methanol to formate without the formation of detectable formaldehyde. Whether or not this means that the formaldehyde remains bound to the active site between the two sequential oxidations is not known; if it is bound then it may have to become hydrated before being oxidized to formate. If, on the other hand, it is released, then it will compete with methanol for the active site of the enzyme; clearly some regulation of this is likely to be necessary. The nature of this regulation appears to be very unusual, requiring the presence of a second “modifier” protein. As mentioned above (Section III.A.3), we have recently isolated and purified a modifier protein (M protein) from Pseudomonas AM1 and M . methylotrophus. This M protein acts by increasing the affinity of MDH for various substrates such as 1,2-propanediol (Ford et al., 1985). We have now found that the M protein occurs in a wide range of methylotrophs (Pseudomonas AM 1,M.methylotrophus, strain 4025, Methylosinus trichosporium), although only one of these, Pseudomonas AM 1, is able to grow on propanediol. Because the M protein affects the affinity of the MDH for a number of substrates, we investigated its effect on the oxidation of formaldehyde. The conclusion drawn form this work is that the function of the M protein might be to bind to MDH and thus prevent the
BACTERIAL OXIDATION OF METHANE A M ) METHANOL
147
oxidation of formaldehyde by this enzyme to formate (M. D. Page and C.Anthony, unpublished observations). In its presence, methanol is oxidized only to formaldehyde; and formaldehyde is no longer oxidized when added as the sole substrate. It is clear that for this regulation to occur, both the M protein and the MDH must be present in the same part of the bacterium. In those organisms in which we have been able to separate clearly cytoplasm from periplasmic material, it was shown that more than 90% of MDH was in the periplasm together with more than 70% of the M protein. Whether or not the modifier protein has a more complex role in the regulation of MDH than of merely preventing formaldehyde oxidation must await further investigation. B. THE PROSTHETIC GROUP AND MECHANISM OF METHANOL DEHYDROGENASE
1. Chemical Characterization of the Prosthetic Group (Pyrrolo-Quinoline Quinone, PQQ)
Pure MDH has a characteristic absorption spectrum (Anthony and Zatman, 1967b; Fig. 2). The absorption due to the protein has a peak at 280 nm and a shoulder at 290 nm; that due to the prosthetic group has a peak at about 345 nm and a shoulder at about 400 nm. It has been shown that this yellow/ green enzyme is a partially reduced form of the dehydrogenase. The slight
‘.Or
wOValsngm(Ill?l)
FIG. 2. Absorption spectra of different forms of methanol dehydrogenase. From Duine et al. (1981). The enzyme concentration is 5mg of protein ml-’. The reduced form (-) corresponds to MDHred in Fig. 7; it contains PQQHzand has a similar, but not identical, appearance to spectra of methanol dehydrogenases as they are usually isolated. The oxidized form (---) corresponds to MDHox, in Fig. 7. It is produced by addition of electron acceptor to MDHred in the presence of activator (NH,Cl) and cyanide. Further spectra of the various forms of MDH are given in Duine et al. (1981) and in de Beer et al. (1983).
148
C.
ANTHONY
wovSlsn(lth(rm)
FIG. 3. Absorption spectra of the quinol (-) and quinone (---) forms of the prosthetic group of methanol dehydrogenase. The quinol form (PQQH,) was o b tained by reducing the quinone form (PQQ) with hydrogen in the presence of platinum oxide. The spectra were measured in anaerobic conditions in 50 m potassium phosphate, pH 7.0. These spectra were kindly provided by Dr. Duine and Dr. Frank. Spectra of various adducts of PQQ are published in Dekker et al. (1982).
variations in spectra from one enzyme to another, particularly with respect to the extent of absorption at 400 nm, are probably due to variations in the proportions of partially and fully reduced enzyme present in the preparation (see Section III.B.5). After storage of the MDH from M . methylotrophus at low temperatures, we have found that a red inactive enzyme sometimes is produced. The spectrum of this form of the monomeric dehydrogenase from Methylomonas methanica has been described and discussed by Mincey et al. (1981). In this case, however, the red form of the enzyme was active; after extraction, the properties of the prosthetic group were the same whether it was extracted from the yellow or the red form of the dehydrogenase. Although MDH has only a typical protein fluorescence, on boiling or treatment with acid or alkali the green fluorescent prosthetic group is released with concomitant loss of enzyme activity. This prosthetic group is reddishbrown in colour, highly polar, acidic and it has a low molecular weight. Maximum fluorescence occurs at low pH values, excitation maxima are at 255 nm and 365 nm and the fluorescence maximum is at 470 nm (Anthony and Zatman, 1967b; see Fig. 3 for its absorption spectrum). Because these fluorescence characteristics are typical of pteridines, it was originally concluded that the novel prosthetic group of MDH might be an unusual pteridine (Anthony and Zatman, 1967b). The structure of this molecule then resisted all attempts at elucidation for more than a decade, during which time the similarity of the fluorescence characteristics of the prosthetic group to those of pteridines was confirmed (Urushibara et al., 1971; Sperl et al., 1973). The first demonstration that the prosthetic group is not a pteridine derivative was published by Duine and Frank and their colleagues. Using a wide range of chemical and physical techniques, they showed that it is a multicyclic
BACTERIAL OXIDATION OF METHANE AND METHANOL
PQQ
(quinone)
I e-
n*
* PQQH'
(free radical)
1 e-
n+
149
* PQQH, (quinol)
FIG. 4. The prosthetic group of methanol dehydrogenase. The full name of PQQ is 2,7,9-tricarboxy-1H-pyrrolo[2,3-flquinoline-4,5-dione. The trivial name methoxatin was initially proposed (Salisbury et uf., 1979) but the abbreviation PQQ (pyrroloquinoline quinone)emphasizes the functionalimportanceof the orthoquinonepart of the structure (Duine and Frank, 1981a). PQQ, PQQH' and PQQH2are all involved in the reaction cycle as indicated in Fig. 7. Adducts with water, methanol, acetaldehyde, acetone and ammonia are formed by addition at C-5. The mid-point redox potential of the PQQ/PQQH, couple is + 90 mV at pH 7.0 and + 419 mV at pH 2.0 indicating that PQQ is a 2e-/2H+ redox carrier (Duine et af., 1981).
ring compound with two uncoupled aromatic protons, an inner ring orthoquinone, two nitrogen atoms and one or more carboxyl groups (Duine and Frank, 1981a;Table 7 and Fig. 4). These proposals are all consistent with the structure published by Salisbury et al. (1979) based on X-ray diffraction analysis of a crystalline acetonyl derivative of the presumed prosthetic group extracted from whole cells of Pseudomonas TP-1. This showed that the prosthetic group is a novel and complex orthoquinone derivative of fused quinoline and pyrrole rings. Although a trivial name (methoxatin) has been suggested for the prosthetic group, a more informative name is pyrroloquinoline quinone (PQQ). The prosthetic group is sometimes referred to as a co-enzyme; this is a less than ideal terminology, however, because it might be inferred from this that it is similar to NAD+ in dissociating from the dehydrogenase during the catalytic cycle. Although PQQ is not usually covalently bound, it remains firmly attached to the enzyme during its catalytic cycle and so is more properly called a prosthetic group. On the basis of electron neutron double resonance (ENDOR) measurements, it has been suggested that the noncovalently bound prosthetic group is situated in a hydrophobic site on the dehydrogenase (Duine et al., 1984b). Although it has not been possible to reconstitute active MDH from the isolated PQQ prosthetic group plus apoenzyme, such reconstitution has been achieved using the apoenzyme of the glucose dehydrogenase of Acinetobacter calcoaceticus (Duine et al., 1979b). This assay system has been recently used to confirm the identity of the PQQ prosthetic group synthesized chemically from 2,3-dimethoxytoluene (Gainor and Weinreb, 1981; Kilty et al., 1982).
TABLE 7. The pyrrolo-quinoline quinone (PQQ) prosthetic group" of methanol dehydrogenase -~
X-ray crystallography and structure proposal Ultraviolet and visual absorption spectra of MDH Fluorescence and absorption spectra of PQQ High-pressure liquid chromatography characterization and quantitative analysis Reconstitution of a PQQ enzyme Electron-spin resonance, electron neutron double-resonance and circular dichroism spectrometry Nuclear magnetic resonance and mass spectrometry Formation of adducts with water, methanol, aldehydes, acetone, ammonia and amines The mechanism of quinoprotein dehydrogenases Chemical synthesis
~
Salisbury et al. (1979) Anthony and Zatman (1967b), Duine et al. (1979a, 1980, 1981), Mincey et al. (1981) Anthony and Zatman (1967b), Duine et al. (1978, 1980), Duine and Frank (1980b), Mincey et al. (1981), Ohta et al. (1981), Dekker et al. (1982), Ameyama et al. (1984a) Duine et al. (1980, 1981), Duine and Frank (1980b, 1981b), Duine et al. (1983) Duine et al. (1979b, 1980), Ameyama et al. (1981a), Kilty et al. (1982), Duine et al. (1983) Duine et al. (1978, 1981), Westerling et al. (1979), de Beer et al. (1979, 1980), Bamforth and Quayle (1979), Mincey et al. (1981); Ohta et al. (1981), Duine et al. (1984b) Duine et al. (1980, 1981), Gainor and Weinreb (1981), Dekker et al. (1982) Salisbury et al. (1979), Duine and Frank (1980b), Duine et al. (1981); Ekkert et d.(1982), Dekker et al. (1982), Dijkstra et al. (1984) Duine et af. (1980, 1981); Forrest et al. (1980), Duine and Frank (1981a), Oshiro et al. (1983), Dijkstra et al. (1984), Parkes and Abeles (1984) Gainor and Weinreb (1981, 1982), Corey and Tramontano (1981), Oshiro et 01. (1983), Hendrickson and de Vries (1982)
"This novel prosthetic group was first described by Anthony and Zatman (1967b); its structure (Fig. 4) was determined by Salisbury et al. (1979), and Duine and Frank and their colleagues (see references in the Table).
BACTERIAL OXIDATION OF METHANE AND METHANOL
151
dOH
Pyrroloquinoline quinone (PQQ)
Dihydroquinol derivative of PQQ
(0.2pM, 100%)
(IWpM. < 0.2%)
COOCH,
0
\
H s:@ :& -c
0
N
CH,OOC
0
0
(4.9 mM, 17%)
Phenanthroline dione analogues of pyrroloquinoline quinone
FIG. 5. Biological activities of the prosthetic group and related compounds. From Duine et al. (1980). These compounds were tested for activity in reconstitutingglucose dehydrogenase when mixed with the apoenzyme from Acinetobacter calcoaceticus.The activities measured in this test are expressed as percentages of that measured with the prosthetic group PQQ.
It has also facilitated the testing of analogues of PQQ in order to determine which structural elements of the prosthetic group are essential for its functioning or its binding to the enzyme (Duine et al., 1980).The structures of the analogues used, together with the relative rates measured in the glucose dehydrogenase assay system, are shown in Fig. 5. It was concluded that the orthoquinone in PQQ is essential for activity because the dehydrogenase was inactive when reconstituted with the 4,5-dihydroquinol derivative (PQQH,). By contrast,'it was shown that the pyrrolo ring and the 9-carboxylic acid are not essential for activity because they could be replaced with a pyridinol ring and a 9-hydroxy group, respectively, in the phenanthroline-dione analogue of
152
C.
ANTHONY
PQQ. It should be noted that it has proved impossible to reconstitute active glucose dehydrogenase using these analogues with the presently available strain of Acinetobacter (the original Hauge strain has been lost; Dr. J. Frank, personal communication).
2. Chemical Reactions of the Pyrrolo-Quinoline Quinone Prosthetic Group It might be expected that a knowledge of the structure of the prosthetic group would rapidly lead to an understanding of its function with respect to binding to apoprotein, electron transfer, activation and inhibition. This, however, has not been the case although a great deal of information is becoming available on the reactions between PQQ and various compounds known to have some effect on enzyme activity, such as alcohol, aldehydes, ammonia, amines and cyanide (Dekker et al., 1982). Although the relevance of such studies to an understanding of the enzyme mechanism is at present uncertain, they are shedding light on some of the problems encountered in earlier investigations of the prosthetic group and its absorption and fluorescence spectra. For example, it has been shown by Dekker et al. (1982) that PQQ in water consists of two compounds (at least); these are PQQ itself, and PQQ hydrated at the C-5 position (PQQ-H,O), this being the fluorescent species. These two species are in equilibrium, low temperature favouring the formation of the fluorescent PQQ-H,O. The presence of an equilibrium mixture of the two species in aqueous solvents explains the observation that the absoprtion spectrum of the prosthetic group is different from the excitation spectrum at room temperature, whereas at low temperature the spectrum becomes similar (Anthony and Zatman, 1967b; Dekker et al., 1982). Besides PQQ and PQQ-H,O, it appears that a small amount of a third species, the dihydrate (PQQ-2H20), also exists in solution. This was deduced from the observation that absorption spectra in borate buffers were quite different from spectra in other buffers at comparable pH value, indicating that additional hydration of PQQ-H,O takes place at the C:4 position, resulting in the dihydrate having vicinal diol groups able to form a complex with borate (Fig. 6). Other compounds forming adducts with PQQ include acetone, acetaldehyde, methanol, cyanide, ammonia and amines (at pH 9); the spectra of all such addition compounds are similar, indicating that addition is at the C-5 position (see Fig. 6). It is thus clear that PQQ is able to react with all the substances that play a role in the enzymic reactions of MDH, although, as mentioned above, whether or not the reactions demonstrated are part of the enzyme mechanism is not yet known. Even when pure PQQ, or its analogues, are shown to catalyse apparently relevant reactions, these are not necessarily models of dehydrogenase activity. For example, the demonstration that PQQ, in micelles with hexadecyltrimethylammonium bromide, catalyses the
G HO
O
HO $30; OH
OH
QooH 0-1 B-OH
HO
I
PO0
OH
+O
NR \
CH,
FIG. 6. Adducts of pyrrolo-quinoline quinone (PQQ). These adducts and their significance are discussed extensively by Dekker et al. (1982). The production of these adducts is reversible in aqueous solutions.
154
C. ANTHONY
oxidation of amines and alcohols to aldehydes or ketones is obviously interesting (Oshiro et al., 1983); but it is not necessarily relevant to the mechanism of the dehydrogenase,although speculative mechanisms in which amines or alcohols become covalently bound to the dehydrogenase have been proposed (see Fig. 8). It should be noted that some chemical studies can be misleading because of false assumptions about the nature of the MDH and its environment in the cell. For example, the thorough study of PQQ and its analogues with respect to redox potentials (Eckert et al., 1982) must be interpreted with caution because the authors assume that the dehydrogenase resides in an aprotic lipophilic cell-membrane environment, and that the carboxyl substituents on the PQQ remain undissociated. It is more likely that MDH is only loosely bound to the outer periplasmic side of the bacterial membrane, or that it is free in the periplasm (Section 1V.F). This does not make interpretation straightforward, however, because the prosthetic group has been shown to be, at least partially, in a hydrophobic site on the dehydrogenase (Duine et al., 1984b). 3. The Detection and Determination of Pyrrolo-Quinoline Quinone
PQQ can be determined quantitatively after release from purified enzymes by its absorption at 249nm ( E 18,40O~-'cm-'), and use of this method led originally to the suggestion that there is one PQQ molecule in each dimeric molecule of MDH (Duine et al., 1980). Subsequent knowledge of the properties of PQQHz, however, necessitated a re-evaluation of this result and it has now been established that two prosthetic groups can be extracted from each molecule of the dimeric MDH, one being the oxidized form (PQQ) and the other the quinol form (PQQH,) (Duine et al., 1981). The most likely interpretation of this observation is that the MDH, as isolated, contains PQQH', two molecules of which disproportionate to one PQQ plus one PQQH2 during the extraction procedure. The methods used in these studies were based on measurement of ultraviolet spectra and ultraviolet detection of high-pressure liquid chromatography (HPLC) eluates. These methods are suitable for use with clean samples (such as extracts of purified quinoprotein) but not for more complex samples such as culture fluids; in such cases, fluorescence detection is more appropriate. Any method depending on the fluorescence properties of the prosthetic group needs to take into account the fact that preparation procedures may lead to a variety of adducts of PQQ differing in their fluorescence characteristics. PQQ itself is not fluorescent but, as discussed above, it occurs in equilibrium in water with the hydrated form (PQQ-H,O) which is fluorescent at appropriate temperatures and pH values.
BACTWAL OXIDATION OF METHANE AND METHANOL
155
The chemical methods described by Duine et al. (1983), for extracts of purified enzymes, depends on ion-pair chromatography on HPLC reversephase columns with ultraviolet detection. For analysis of culture supernatants they have described a fluorescence detection system. Proof of the presence of PQQ is obtained by treating samples with butyraldehyde, which converts the prosthetic group into a stable adduct having a suitable retention time in the HPLC system. The sensitivity and selectivity of the analysis was further enhanced by reducing samples with NaBH,, thus producing PQQH, which is then oxidized with NaIO, to a strongly fluorescing compound of unknown structure. Ion-suppression chromatography, instead of ion-pair chromatography, worked satisfactorily for analysis of the butyraldehyde adduct and of the fluorescent oxidation product. The lowest detection level for PQQ analysed by way of this product was 80 nM. For detection of lower concentrations of PQQ, a biological test system is more appropriate. This is based on an easily prepared apoenzyme from the unpurified quinoprotein glucose dehydrogenase from Pseudomonas aeruginosa (Duine et al., 1983). The PQQ sample is preincubated with apoenzyme, and then assayed by addition of glucose (substrate) and Wurster’s blue (TMPD, an artificial electron acceptor). This method is both straightforward and sensitive ( 2 n ~ PQQ being the lowest detection level); but adducts of PQQ have a low activity in this test. 4. Pyrrolo-Quinoline Quinone as the Prosthetic Group of Other Quinoproteins
Although thought for many years to be both novel and unique, it has now been shown that PQQ is the prosthetic group of a range of different dehydrogenases which Duine and Frank have aptly called quinoproteins (Table 8). This has been achieved by chemical studies of the extracted and purified prosthetic groups, and by reconstituting pure prosthetic groups with apoprotein to yield active enzyme (Duine et al., 1979b; Ameyama et al., 1981a). This approach is impossible with MDH because release of the prosthetic group always denatures the enzyme (Anthony and Zatman, 1967b). By contrast, it is readily achieved using the glucose dehydrogenase from Acinetobacter calcoaceticus (Duine et al., 1979b, 1980; Kilty et al., 1982) and from Pseudomonas aeruginosa (Ameyama et al., 1981a; Duine et al., 1983). The apoprotein from this dehydrogenase is able to form active enzyme when mixed with its own purified prosthetic group, or with the prosthetic group from other quinoproteins. A similar glucose dehydrogenase has now been demonstrated in the enteric organism Klebsiella aerogenes after growth in glucose-sufficient (K+-limited) conditions (Neijssel et al., 1983). The apoenzyme is also produced by some strains of Escherichia coli ( H o m e s
156
C. ANTHONY
et al., 1984). This organism appears to be unable to synthesize PQQ which must therefore be preincubated with extracts prior to demonstration of glucose dehydrogenase activity. Mutants of E. coli that lacked one of the proteins of the usual glucose utilization system (enzyme I of the phosphoenolpyruvate phosphotransferase system) were only able to grow on glucose when cultures were supplied with PQQ as a growth factor. A similar requirement for PQQ as a growth factor has recently been demonstrated in a range of different bacteria and yeasts. In some cases, the PQQ stimulated growth rates and yields, whereas in other organisms its function appears to be as a growth initiator, diminishing the lag period prior to onset of growth (Shimao et al., 1984; Ameyama et al., 1984b, c). Table 8 lists those dehydrogenases now known to be quinoproteins. It should be noted that although PQQ is probably the prosthetic group of primary amine dehydrogenase, in this case it is covalently bonded to the polypeptide chain, part of which remains attached to the prosthetic group during its isolation (de Beer et al., 1980; Ishii et al., 1983). Because of this it has not yet been shown to be identical with PQQ from other quinoproteins (see Kenney and McIntire, 1983). A second enzyme having PQQ as a covalently bound prosthetic group is bovine serum amine oxidase, which also contains copper ions essential for its activity (Lobenstein-Verbeek et al., 1984). This enzyme is especially noteworthy as it is the first example of a quinoprotein oxidoreductase discovered in a eukaryotic organism. A second example may be the amine oxidase of Aspergillus niger (Ameyama et al., 1984a). Another quinoprotein that is especially relevant in this review of MDH is the membrane-bound quinoprotein alcohol dehydrogenase from an organism originally thought to be A. calcoaceticus (Duine and Frank, 1981b). Although methanol is a very poor substrate for this enzyme, it resembles MDH in other respects; that is, it has a pH optimum of 9.5, it has a requirement for an amine activator, it oxidizes primary alcohols, it shows the characteristic absorption spectrum, it has a PQQ prosthetic group and it has a molecular weight of about 120,000. The activator and, to some extent, the substrate specificity can best be compared with those of the atypical MDH from Rh. acidophila, but the physicochemical properties of the two enzymes are quite different. MDH is an exceptional dehydrogenase in reacting with the electron transport chain at the level of cytochrome c, thus bypassing cytochrome b (see Section I1I.C). It might be expected, therefore, that all quinoprotein alcohol dehydrogenases would interact with the cytochrome chain in a similar manner. Acinetobacter species, however, typically contain no cytochrome c. This suggested that either the alcohol dehydrogenase studied by Duine and Frank (1981b) was not from A. calcoaceticus or that there might be an unexpected diversity in the manner of coupling quino-
BACTERIAL OXIDATION OF METHANE AND METHANOL
157
protein alcohol dehydrogenases to the electron transport chain. Further investigations (Beardmore-Gray and Anthony, 1983) confirmed that the strain used by Duine and Frank contained the unusual alcohol dehydrogenase, but they also showed that this unidentified organism (now lost) contained cytochrome c. Genuine strains of Acinetobacter contained neither cytochrome c nor the membrane-bound alcohol dehydrogenase, but instead contained NAD+-linked alcohol dehydrogenase that could couple to the respiratory chain at a lower redox potential by way of NADH dehydrogenase. This conclusion is consistent with our expectation that all quinoprotein alcohol dehydrogenases are likely to be similar to MDH in being coupled to the electron transport chain at the level of a high-potential cytochrome c. Support for this conclusion is provided by the properties of a membranebound quinoprotein alcohol dehydrogenase (from Gluconobacter suboxyduns) which has now been solubilized, purified and crystallized. The enzyme crystallized as a dehydrogenase+ytochrome c complex, which also contained a third small protein of unknown function (Adachi et al., 1982). 5. The Mechanism of Methanol Dehydrogenase
After the first description of MDH and its prosthetic group, further elucidation of its mechanism was hampered by the fact that the pure enzyme is always isolated in a partially reduced state, and so addition of substrate has no effect on the spectrum of the enzyme. Furthermore, prior oxidation of the reduced enzyme with electron acceptor (phenazine methosulphate) inactivates the enzyme, and this also prevents observation of substrate-induced spectral changes. This problem was overcome by Duine and Frank (1980a, b) who oxidized the pure dehydrogenase by addition of phenazine methosulphate and activator (ammonia) in the presence of KCN which, being an inhibitor competitive with respect to methanol, binds to the active site and protects it against inactivation. On oxidation, the characteristic 345 nm peak of the partially reduced form was shifted to about 400nm, and additions of stoicheiometric amounts of substrate changed the spectrum back to that of the reduced form (see Fig. 2). Similar changes occurred on reduction by mercaptoethanol or catalytic hydrogenation of the fluorescent prosthetic group; the fluorescence was lost and the higher absorption peak decreased in wavelength (see Fig. 3). The reduced prosthetic group could be re-oxidized by molecular 0, back to the oxidized fluorescent form. The absorption maxima in the whole enzyme were 20 to 50nm higher than in the isolated prosthetic group. The three forms of the prosthetic group that are involved in catalysis are shown in Fig. 4. That the free radical (PQQH') is an intermediate is indicated by the observation that the quinone free radical, measured by ESR on the
TABLE 8. Dehydrogenases having a pyrrolo-quinoline quinone (PQQ) prosthetic group; quinoproteins Enzyme
Organism
Methanol dehydrogenase
Methylotrophs
Primary arnine dehydrogenase (the PQQ is covalently bound)
Pseudomonas AM I
Bacterium W3A1 Thiobacillus versustus (A2)
Alcohol dehydrogenase
Unknown bacterium Pseudomonas aeruginosa Acetobacter pasteurianum Gluconobacter suboxydans
Polyol dehydrogenases (?) Alcohol (long-chain) dehydrogenase Glucose dehydrogenase
Glucose dehydrogenase (apoenzyme)
Gluconobacter oxydans Alkane-grown Pseudomonas aeruginosa Acinetobacter calcoaceticus Gluconobacter oxydans Gluconobacter suboxydans Pseudomonas aeruginosa Pseudomonas Jltlorescens Klebsiella aerogenes Escherichia coli
References Duine and Frank (1980b, 1981a), (references in Table 4) de Beer et a/. (1980), Anthony (1982), Ishii et al. (1983) Kenney and McIntire (1983), Ameyama et al. (1984a) Duine and Frank (1981c), Haywood et al. (1982) Duine and Frank (1981b), Beardmore-Gray and Anthony (I 983) Groen et al. (1984) Duine et al. (1979b) Ameyama et a/. (1981a); Adachi et al. (1 982) Duine et al. (1979b) Duine et al. (1979b) Duine et al. (1979b; 1980) Duine et al. (1979b) Ameyama et al. (1981a, b) Ameyama et al. (1981a), Duine et al. (1983) Ameyama et al. (198 1 a) Neijssel et al. (1983) Hommes et al. (1984)
Aldehyde dehydrogenase Lactate dehydrogenase (?) Amine oxidase
Gluconobacter suboxydans Propionibacterium pentosaceum Mammalian (bovine serum)
Tryptophan side chain oxidase (?)
Aspergillus niger Pseudomonas ATCC 29514
Ameyama et a f . (1981a) Duine and Frank (1981~) Lobenstein-Verbeek et al. (1984); Ameyama et al. (1 9 8 k ) Ameyama et al. (1984~) van der Graaff et al. (1 984)
160
C. ANTHONY
isolated enzyme, is the same as that obtained with the half-reduced form of the isolated prosthetic group (Duine et al., 1978; Bamforth and Quayle, 1979; Westerling et al., 1979; de Beer et al., 1979). The reactivity of the MDH with one-electron acceptors, during assay and oxidative titration, is consistent with the proposal that electron transfer from the reduced dehydrogenase proceeds by way of the free radical intermediate, and it is also consistent with the results of kinetic studies. In analysing the catalytic cycle, it would be ideal if the form of the prosthetic group, and related absorption spectra, during each phase of catalysis were known. At present, however, this ideal cannot be completely achieved. Although PQQ, PQQH' and PQQH, can all be extracted from MDH, their proportions in the enzyme cannot be known with absolute certainty because comproportionation and disporportionation reactions may take place after extraction. Furthermore, spectra are difficult to interpret because the dimeric enzyme contains two prosthetic groups, each able to be in one of the three redox forms, some of which are able to form adducts having spectra similar to those of the unmodified prosthetic group. Figure 7 is a summary of the reaction cycle in vitro first proposed by Duine and Frank (1981a), and now supported by further evidence presented by de Beer et al. (1983) and Dijkstra et al. (1984). The enzyme as isolated is in an oxidation state intermediate between the fully oxidized and the fully reduced state; it is represented as MDHox, (PQQH') in Fig. 7. MDHox*(PQQ), the fully oxidized form, is produced by oxidation of the isolated enzyme (MDHox, ) in the presence of activator (ammonia). The fully oxidized form is the only form able to react with substrate, which converts it into MDHred (PQQH,) with concomitant formation of product. The MDHox, (PQQH') is then regenerated by oxidation with electron acceptor. The reaction cycle is more readily demonstrated in vitro if KCN is used as stabilizer during the initial oxidation of MDHox, (the enzyme as isolated). The KCN converts the unstable MDHox* (PQQ) into the stable form MDHox, (PQQ). This may contain the cyanide adduct of PQQ; if it does then the formation of this adduct must be a rapidly reversible process. An important point about the mechanism proposed in Fig. 7 is that the product of reaction must be released before any re-oxidation of the prosthetic group takes place (Duine and Frank, 1980a). There is no evidence for the existence of intermediates between the fully oxidized form and the substrate reduced form. The free radical form, MDHox, (PQQH.), is only likely to be an intermediate during the re-oxidation of the fully reduced form and the fully oxidized form that acts as electron acceptor for substrate. A completely different mechanism from that described above has been proposed for the monomeric MDH from Methylomonas methanica by Mincey et al. (1981). These authors concluded that only those enzyme
I
I
I
MDHof(PQQ)
7 Ammonia
1
KCN(ammonia)
MDHox, (PQQH’)
- -------;------ - + MDHox~(PQQ)
\, e-
l
I
‘1
I
Electron acceptor
II
Formaldehyde
Formaldehyde I
Methanol
t
_ _ _ _ _ _ _ _ _ ‘>---d%,L-
MDHred(pQQH2)+
,I’
I
Methanol
FIG. 7. Reaction cycle for the activity of methanol dehydrogenase in vitro. This is based on a scheme presented in the review of Duine and Frank (1981a) and slightly modified by de Beer et al. (1983). The solid line indicates the “normal mechanism”, and the dashed line indicates the route most readily demonstrated in vitro (see the text). The probable redox forms of the prosthetic groups in each form of the enzyme are given in parentheses. The isolated enzyme is mainly MDHox, (sometimestogether with some MDHred). MDHox, may be produced by oxidation of the isolated enzyme by addition of electron acceptor in the presence of KCN and activator (ammonia). MDHred is produced from MDHox, by titration with substrate and MDHox, regenerated by titration of MDHred with electron acceptor. MDHox* is an unstable form produced from MDHred by titration with electron acceptor in the presence of ammonia; subsequent addition of KCN then yields the stable oxidized form MDHox,. Spectra of the some of these forms of the enzyme are given in Fig. 2.
molecules that contain semiquinone are catalytically active, producing a three-electron paramagnetic reduced form of the enzyme on reduction with substrate. It was proposed that the aldehyde product is only released after the enzyme is re-oxidized by electron acceptor. Many aspects of this mechanism are not in agreement with observations published by other workers, and it has been suggested by de Beer et al. (1983) that some of the complicated ESR spectra forming the basis of the mechanism are due to an artefact that arises on ageing of the enzyme. These authors concluded that the properties of the monomeric dehydrogenase, as described by Mincey et al. (1981) do not indicate a reaction mechanism different from that based on their own work with the enzyme from Hyphomicrobium sp., as summarized in Fig. 7. Further experiments on the mode of action of cyclopropanol and its derivatives (Dijkstra et al., 1984) have confirmed that the mechanism outlined in Figs 4 and 7 is consistent with considerably more of the evidence than is the alternative mechanism proposed by Mincey et al. (1981). Dijkstra et al. (1984) have shown that MDH is inactivated by cyclopropanol and cyclopropanone, and, if the enzyme is also able to oxidize secondary alcohols, by cyclopropanone ethyl hemiketal. Only enzyme molecules containing the oxidized prosthetic group (PQQ), and not those containing the free radical (PQQH’), were inactivated. The inactivation required stoicheiometric amounts of enzyme and proceeded without reduction of electron acceptor
162
C. ANTHONY
and without proton production. The inactivated enzyme contained no free radical, but a modified prosthetic group could be extracted from it. All the evidence is consistent with cyclopropanol reacting with the enzyme by way of a ring-opening mechanism, thus producing a free radical of propionaldehyde which then forms an adduct at the C-5 position (see Fig. 6). The observation that the monomeric enzyme required only one molecule of cyclopropanolfor complete inactivation, and that the dimeric enzyme needed two molecules, demonstrates that the two catalytic sites on typical dimeric MDH act completely independently. Although the catalytic cycle as proposed in Fig. 7 is of great value in interpreting experiments on the mechanism of the isolated dehydrogenase, some important questions remain. These included the nature of the binding of methanol to the enzyme and the role of the activator molecules (ammonia or amines). For example, it is clear that activator is required for the oxidation of MDHox, (PQQH' ), when using phenazine methosulphate as electron acceptor, and yet no activator is required for the methanol-dependent reduction of the physiological acceptor (cytochrome c) by MDH (Duine et af., 1979a; Beardmore-Gray et al., 1983). A second question concerns the ease of formation of adducts at the C-5 position of PQQ, and whether or not this has any significance in the catalytic activity of the dehydrogenase. In this context it should be noted that Forrest et al. (1980) have proposed a mechanism in which activation by ammonia or amines involves covalent bonding at C-4. The alcohol is bonded to the same C-4 atoms and then released as the aldehyde, the 2H being passed to the prosthetic group which is then oxidized by the electron acceptor (Fig. 8). This scheme does not appear to be consistent, however, with the observation that the ammonia or amine activator is readily removed by dialysis, nor with the conclusion that a mechanism involving a PQQ-alcohol adduct, which is subsequently oxidized by an electron acceptor, is not consistent with much previously published experimental evidence (Duine and Frank, 1980a; Duine et al., 1981). Intermediates similar to those in Fig. 7 are perhaps more likely to be involved in the mechanism of catalysis by the quinoprotein methylamine dehydrogenase (Anthony, 1982). C. CYTOCHROME C AND ITS INVOLVEMENT IN METHANOL OXIDATION
1. Evidence for the Involvement of Cytochrome c in Methanol Oxidation by
Whole Bacteria and Membrane Preparations
Dehydrogenases catalysing the oxidation of all organic substrates except methanol, and sometimes methylamine, interact with the lower redox potential part dfthe electron transport chain, prior to cytochrome b. It is thus
BACTERIAL OXIDATION OF METHANE AND METHANOL
COOH
163
COOH
NHR
0
PQQ COOH
HOOC
NHR
R.CH,OH
"2H"
HOOC R .CHO
FIG. 8. A mechanism proposed for the involvement of pyrrolo-quinoline quinone in catalysis by methanol dehydrogenase. From Forrest et ul. (1980).
evident that the conclusion that electrons from methanol are donated by MDH to the electron transport chain at the more positive level of cytochrome c is one of the most unexpected conclusions to be drawn from work on electron transport in methylotrophs. The extensive, but indirect, preliminary evidence for this conclusion is summarized below. (a) Some methylotrophs, able to grow on methylamine but unable to grow on methane and methanol, are devoid of cytochrome c. By contrast, all species of methyltrophic bacteria, from a wide range of different genera, that are able to grow on methane and methanol, and which therefore need to oxidize methanol, contain at least one soluble cytochrome c having a high mid-point redox potential (250-350 mv).Mutants lacking cytochrome c no longer oxidize or grow on methanol, but oxidize and grow on other substrates. Such mutants have been isolated from Pseudomonus AM 1 (Anthony, 1975; Widdowson and Anthony, 1975), Methylobacterium organophilum (OConnor and Hanson, 1978) and P. &nitr@cuns (Willison and John, 1979). All oxidizable substrates, but not methanol, are able to reduce the cytochromes b and aa, in the cytochrome c-deficient mutant of Pseudomonus AM 1, whereas all substrates, including methanol, are able to reduce both cytochromes c and uu, in the wild-type bacteria (cytochrome b
164
C. ANTHONY
is obscured by the cytochrome c in wild-type bacteria and so cannot be measured). (b) Membrane vesicles of Pseudomonas AM1 oxidize NADH, succinate and methanol, and all these substrates are able to reduce cytochrome c. Respiration, and reduction of cytochrome c by NADH and succinate, is inhibited by antimycin A, whereas the oxidation of methanol, and the reduction of cytochrome c by methanol, is not affected by this inhibitor (Netrusov and Anthony, 1979). Antimycin A is known to inhibit the oxidation of cytochrome b, and therefore of substrates donating electrons to sites prior to cytochrome b in the electron transport chain. Our results strongly suggest, therefore, that methanol donates electrons to the chain after cytochrome b (Netrusov and Anthony, 1979). Similar studies, using whole cells or extracts, have c o n k e d that the oxidation of NADH and succinate, but not of methanol, is inhibited by antimycin in Pseudomonas extorquens (Higgins et al., 1976a, b; Tonge et al., 1977b), P. denitri$cans (Bamforth and Quayle, 1978a; van Verseveld and Stouthamer, 1978a), Methylosinus trichosporium (Higgins et al., 1976a) and Pseudomonas sp.2 (Netrusov et al., 1977). (c) During the respiration-coupled ATP synthesis catalysed by membrane vesicles by Pseudomonas AM1, the P/O ratio with methanol is the same as measured with ascorbate/TMPD, but it is markedly more when succinate or NADH is the substrate (Netrusov and Anthony, 1979; Netrusov, 1981). 2. The “Functional Coupling” of Methanol Dehydrogenase to Cytochrome c in Crude Bacterial Extracts Although it was clear for some time that MDH is coupled to the electron transport chain at the level of cytochrome c, it was impossible to demonstrate this using pure MDH and pure cytochrome c. When these proteins were incubated together, the cytochrome was always found to be in the reduced state, even in the absence of methanol; and after oxidation of the cytochrome c with femcyanide, methanol-dependent cytochrome reduction could not be demonstrated (Anthony, 1975; O’Keeffe and Anthony, 1980b). This result suggested that either some extra component is required for activity, or that one or other of the proteins become damaged during extraction; and this appears to have been confirmed using enzyme prepared from Hyphomicrobium X (Duine et al., 1979a). Extracts prepared anaerobically, and partially purified under anaerobic conditions, contained both MDH and cytochrome c which, after oxidation with femcyanide, could be reduced with methanol at pH 6.5 in the absence of ammonia as activator. Similarly, methanoldependent reduction of the artificial electron acceptor (Wurster’s blue, TMPD) did not require ammonia activator. After exposure of the “anaerobic
BACTERIAL OXIDATION OF METHANE AND METHANOL
165
Time (minutes)
FIG. 9. The reduction of cytochrome c by methanol dehydrogenase in anaerobically prepared extracts of MethyloPhilus methylotrophus before (a) and after (b) aeration. Anaerobic reduction of cytochrome c was measured spectrophotometriclly (at 550 nm) using crude extracts containing MDH and cytochrome c after oxidation with 0.5 jmol femcyanide. What was actually measured in this type of experiment was the rate of ferricyanide reduction by the MDH/cytochrome “complex”; it is only after the ferricyanide has been completely reduced that the reduced cytochrome c starts to accumulate. The rate of reduction of cytochrome c in the presence of methanol ( 0 ) was twice the rate measured in its absence (0). After exposure to air for 20 minutes, the rate in the absence of methanol increased to the rate measured in its presence. The rate of reduction was directly proportional to the concentration of MDH present. When larger amounts of ferricyanide were used, the rates of the “endogenous” reaction were markedly diminished (to zero when sufficient was used) and the rates with added methanol also decreased to some extent. It appeared that the ferricyanide was destroying the MDH and/or cytochrome in the absence of partially protecting methanol. From results discussed in Beardmore-Gray and Anthony (1984).
preparation” to air for an hour, however, ammonia became essential for dye reduction, the cytochrome c became oxidized, and its reduction by methanol could no longer be demonstrated by using difference spectrophotometry. Similar results to these have been obtained using some preparations of Pseudomonus AM 1, but most preparations differed in some important respects and gave results mare like those obtained with M . merhylotrophus (Beardmore-Gray and Anthony, 1984). Anaerobic preparations of this organism always required ammonia for activity in the dye-linked assay, and aeration did not lead to cytochrome c oxidation. In the presence of methanol,
166
C. ANTHONY
the rate of cytochrome reduction was greater than in its absence when using crude extracts containing anaerobically prepared MDH and cytochrome c (Fig. 9). After aeration, however, methanol-dependent reduction of cytochrome c could no longer be demonstrated. This was because the rate of endogenous reduction increased to the level occurring in the presence of methanol. Ammonia was not required as activator for the reduction of cytochrome c by MDH, either before or after aeration. In summary, (1) by using anaerobic preparations, methanol-dependent cytochrome c reduction could be demonstrated at pH 6.5-7.0 in all the systems studied; this activity never required ammonia and it was always inhibited by EDTA. (2) Anaerobic preparations catalysed methanoldependent dye reduction but did not always require ammonia. (3) Aeration of the extracts led to changed MDH although the nature of the change was not the same in all systems. Thus, in extracts of Hyphomicrobium X, ammonia-dependent dye-linked activity was destroyed, and the cytochrome c became oxidized and was no longer susceptible to reduction by either methanol or endogenous reductant. In aerated extracts of M . methylotrophus, by contrast, the cytochrome c remained reduced and, after oxidation with ferricyanide, MDH was still able to catalyse its reduction, but with the endogenous rate now being increased to that occurring in the presence of added substrate methanol. Perhaps the most important conclusion from these studies is that there was not always a clear relationship between the loss of “functional coupling” of MDH to cytochrome c, and the production of “classical” ammonia-requiring MDH. One possible explanation for the variety of observations described here is that the enigmatic endogenous reduction usually found on MDH may protect the enzyme from 0, inactivation, the extent of protection perhaps depending on the amount of reductant present on a particular dehydrogenase. It is probable that more complete conclusions will have to await more information on the various states of the PQQ prosthetic group occurring during enzyme catalysis, and the effects of 02,phenazine methosulphate, ferricyanide and ammonia on them. A second important conclusion is that the free radical species of cytochrome c proposed in Fig. 11 may be susceptible to damage by ferricyanide (see OKeeffe and Anthony, 1980b). 3. The Cytochromes c of Methylotrophs (a) Introduction. Having shown that MDH interacts with the electron transport chain at the level of cytochrome c, it is necessary to consider whether the same cytochrome c is involved in all electron transport in methy!otrophs, or whether there is a particular cytochrome c with special properties required for
BACTERIAL, OXIDATION OF METHANE AND METHANOL
167
reaction with MDH. In this context, the first point of note is that all methylotrophs studied in detail contain at least two soluble cytochromes c. This has been shown in Pseudomonas AM1 (OKeeffe and Anthony, 1980a, b), M. methylotrophus (Cross and Anthony, 1980a, b), Methylomonas J (Ohta and Tobari, 1981), P. denitr8cans (van Verseveld and Stouthamer, 1978a), strain 4025 (Vrdoljak and Froud, 1982), Hyphomicrobium X (Beardmore-Gray, 1982) and Acetobacter MB58 (E. J. Elliott and C. Anthony, unpublished observations). Earlier reports of a single cytochrome c in some methylotrophs probably reflects a similar oversight that led to an erroneous preliminary description of a single cytochrome c in Pseudomonas AM1 (Anthony, 1975). The cytochromesc that have been described in most detail from methylotrophs are those from Pseudomonas AM1 and M . methylotrophus, and these have been shown to be similar in many important respects: they have a single polypeptide chain which bears a single haem, having histidine and methionine as the two axial ligands. They both have a high mid-point redox potential (more than 250 mv), characteristic reduced a bands at about 550nm, and they are low spin in both the oxidized and reduced states (see Table 9). The two cytochromes c are labelled according to their isoelectric points, cytochrome cH having the higher isoelectric point and cytochrome cL the lower. Although applicable to the cytochromes c from Pseudomonas AM 1, M . methylotrophus and Hyphomicrobium X , because their isoelectric points differ by about 4pH units, the distinction is not so marked for the cytochromes from Methylomonas J and P. denitrijicans. Their cytochromes c with the lower isoelectric point appear to correspond to the cytochromes cL of other methylotrophs in their function, but the differences in isoelectric points are not so great and both are below pH 7.0. Amino-acid analysis of the two types of cytochrome c from Pseudomonas AM 1 and from M . methylotrophus clearly showed that the difference in isoelectric points is not due to differences in lysine content, but is probably due to differences in the extent of amidation of aspartate and glutamate (Beardmore-Gray et al., 1982). The division of the two types of cytochrome c according to their isoelectric points correlates with differences in many other characteristics (Table 9). Cytochrome cL is larger, has a lower isoelectric point, a lower mid-point redox potential, reacts more completely with CO, has an extra cysteine residue, and is the most sensitive to digestion by proteinases. By contrast, cytochrome cH is smaller, has a higher isoelectric point, higher mid-point redox potential, only two cysteine residues (binding the haem), reacts less well with CO, and is relatively resistant to proteolytic digestion. Analysis of the amino-acid composition, and the products of partial proteolytic digestion, shows that cytochrome cH from Pseudomonas AM1 is very similar to that from M. methylotrophus, and that the two cytochromes cL
TABLE 9. Properties of cytochromes c, and cLof methylotrophs. From OKeeffeand Anthony (1980a), Cross and Anthony (1980a) and Beardmore-Gray et al. (1982, 1983)
Pseudomonas AM 1 Cytochrome cH Cytochrome cL Relative proportions in crude extracts (YO) Isoelectric point Molecular weight Redox potential (Em,; mV) CO binding (YO) No. of cysteine residues Sensitivity to proteolytic digestion No. of lysine residues mol-' Stimulation of autoreduction by MDH Electron acceptor for MDH
72
28
8.8
50
294 36 2 Insensitive
4.2 20,900 256 72 3 Sensitive
11
13
11,OOO
+
Methylophilus methylotrophus Cytochrome c, Cytochrome cL
8.9 8500 373 7 2 Insensitive I1
+
+
42
4.0-4.4 17,000-2 1,000 310
60 3 Sensitive 12
+ +
After purification of cytochrome cL from this organism it is present in two forms, one of which may arise from the other by loss of a 4000 Da fragment (Cross and Anthony, 1980a).
BACTERIAL OXIDATION OF METHANE AND METHANOL
169
are also similar to each other (Beardmore-Gray et al., 1982). There is, however, little similarity between the cytochromes cH and the cytochromes c,. These studies also demonstrated that the two types of cytochrome c found in methylotrophs are completely distinct proteins; one type is not a dimer or degradation product of the other. This being the case, it is perhaps surprising that the cytochrome c-deficient mutant of Pseudomonas AM 1 (Anthony, 1975) lacks both the cytochrome cH and cytochrome cL, and also the membrane-bound cytochrome c. That the mutation is not in a gene responsible for haem biosynthesis is indicated by the normal concentrations of cytochromes a and b in the mutant (Anthony, 1975; Widdowson and Anthony, 1975). It is thus possible that the mutation is in a gene affecting the incorporation of haem into the apoproteins of the various cytochromes c. It is interesting that all cytochromes c, both soluble and membrane-bound forms, were also lost by a single mutation in P. denitr9cans (Willison and John, 1979), and in Rh. capsulata (Michels and Haddock, 1980). (b) Reaction of Cytochromes c with Carbon Monoxide. A remarkable characteristic of most cytochromes c of methylotrophs is their reaction with CO, which has led to them being referred to as cytochromes cco. I have avoided this terminology in order to avoid implying too great a physiological significance to the slow, incomplete reaction with CO. The CO binding of the cytochrome c of Methylosinus trichosporium has led to the speculation that it might have an oxidase or oxygenase function during the oxidation of methane, and an oxidase function has also been proposed for the cytochrome c of the facultative methanol-utilizer Pseudomonas extorquens (Tonge et al., 1975, 1977a, b; Higgins, 1979, 1980). There is, however, no evidence that it functions as an oxidase and, in other methylotrophs containing CO-binding cytochromes c, all the evidence is against such a function (Widdowson and Anthony, 1975; O’Keeffe and Anthony, 1980a, b; Cross and Anthony, 1980a, b; Dawson and Jones, 1981~). It is well known that damaged cytochromes can sometimes become reactive with respect to CO. This is unlikely to be the case with the cytochromes c from methylotrophs because they react with CO in whole cells, and because neither the rate nor extent of CO binding alters during purification of the cytochromes. The estimated CO binding of less than 100% is not a reflection of a mixed population of cytochrome, some binding and some not; it is probably because the cytochrome reacts slowly with CO to form a complex (absorption maximum at 412nm) having a high dissociation constant (Widdowson and Anthony, 1975; OKeeffe and Anthony, 1980a). The reaction with CO of these c-type cytochromes probably reflects the structure around the haem pocket that allows a more readily dissociable iron-methionine bond. That the haem environment is slightly unusual is
170
C. ANTHONY
indicated by the unusual response of the mid-point potential of the cytochromes c of Pseudomonus AM1 to changing pH values (OKeeffe and Anthony, 1980a). Both cytochromes c have two ionizing groups affecting the mid-point redox potentials, the pK values being 3.5 and 5.5 in the oxidized forms, and 4.5 and 6.5 in the reduced forms. If these dissociations arise from the haem, then the higher of the pK values is likely to be due to the rear (inner) haem propionate in the hydrophobic environment of the haem cleft, and the lower pK due to the front (outer) propionate in its more hydrophilic environment. These pK values are sufficiently different from the pH value within the bacteria to preclude a proton-translocating function for the cytochrome c. (Similar results with the cytochrome cZ5,of Pseudomonas ueruginosu have been reported by Moore et al. (1980).) (c)Autoreduction of Cytochrome c in Methylotrophs. A second, more unusual, characteristic of the cytochromes c of methylotrophs is their capacity for rapid autoreduction (OKeeffe and Anthony, 1980b; Beardmore-Gray et al., 1982, 1983). Autoreduction is the reduction of the haem iron of femcytochrome c occurring in the absence of an added reducing agent. This characteristic must be discussed if only because it is responsible for some of the problems encountered in studying MDH-cytochrome c interactions (see below); but it may also constitute a clue to the mechanism of these interactions. The phenomenon of autoreduction occurs also in horse heart cytochrome c (Brady and Flatmark, 1971), but it occurs at about 100 times the rate in the cytochromes c of methylotrophs (Fig. 10). The autoreduction of these cytochromes was not inhibited by p-chloromercuribenzoate or iodoacetamide, indicating that free thiol groups are not involved, a conclusion differing from that for the autoreducible cytochrome f from horseradish (Tanaka et ul., 1978). The autoreduction process is a first-order intramolecular reaction that occurs at high pH values, the pK for this process being greater than pH 10. A mechanism that is consistent with all the available evidence, and which involves electron transfer between a dissociable group (XH) and the haem iron of ferricytochrome c, is presented in Fig. 11. The weakly acidic group (XH)dissociates at a high pH value to give a negatively charged species able to donate an electron to the haem, the free radical produced by this process being stabilized by sharing an electron with the haem iron. This proposal has the advantage that ekckron transfer to the iron, and skabilizakion of the resulting radical, do not have to be explained separately. The electrondonating group must be within the usual atomic distance to the iron, and it is conceivablethat, at high values, it might indeed replace the usual methionine as the sixth ligand to the iron. That the sixth ligand is methionine (at pH 7.0), as is usually the case in cytochrome c, is indicated by the 695 nm absorption
pH Value
FIG. 10. Autoreduction of the cytochromes c of Methylop..:lus methylotrophus. From Beardmore-Gray et ul. (1983). 0,Cytochrome cH (with and without MDH); B, cytochrome c,; 0, cytochrome cL plus MDH; A, horse heart cytochrome c (with and without MDH).
[ri
+I:[
(1)
~
pKI\
(3)
(2)
~
(autoduction)
n+ (The pK of this dissociation is lowered in the prrsma of methanol dehydrogenase) Methanol dehydrogenase (oxidized)
Methanol dchydrogenase (red=d)
FIG. 1. A speculative mechanism for the reduction of cytochrome c by methand dehydrogenase. From OKeeffe and Anthony (1980b) and Beardmore-Gray et ul. (1983). In this scheme the electron donor for autoreduction is a weakly acidic group O(H) on the cytochrome that dissociates to give a negatively charged species able to donate an electron to the haem. Species (1) is the undissociated ferricytochrome c which becomes dissociated in the presence of MDH (or at high pH values) to species (2). Species (3) is the radical complex of ferrous iron, isoelectronic with the ferric Species 2. Species (4) is the ferric form of the radical and can only be produced in the presence of added electron acceptor; this would be cytochrome oxidase (or perhaps cytochrome cH)in physiological conditions, of ferricyanide (as in Fig. 9), or mammalian cytochrome c (as in Fig. 12). Species (4) can be reduced to species (2) to complete the cycle by electron transfer from methanol dehydrogenase, the electrons arising from methanol or from the endogenous reductant.
172
C. ANTHONY
band shown to be present in the methylotroph ferricytochromes c. This absorption band is lost, as expected, on reduction with dithionite at pH 7.0, or on autoreduction of ferricytochrome cL at high pH values. Results of experiments using magnetic circular dichroism (MCD) spectrometry confirm that the autoreduction at high pH values of ferricytochromecLdoes not lead to a marked change in the nature of the haem ligation. The iron in this cytochrome c appears to be low spin in both oxidized and reduced states at pH 7.0, and when it is autoreduced at pH 10.0. It is clear from these results that if the sixth ligand does change during autoreduction, then the methionine must be replaced by an alternative strong-field ligand. On raising the pH value of ferricytochrome cL, the disappearance of the 695nm band occurs more rapidly than the appearance of the typical Q band at 550nm; this suggests that displacement of the methionine ligand does occur, that it is more rapid than the intramolecular autoreduction of the cytochrome, and that the two processes may occur independently (Beardmore-Gray et al., 1982).
(d) The Effect of Methanol Dehydrogenase on the Autoreduction of Cytochrome c. When pure MDH is mixed with pure oxidized cytochrome c the cytochrome becomes rapidly reduced, the presence of added substrate being unnecessary for this reaction. The “reduction” is a first-order reaction with respect to ferricytochrome c, indicating that ’an intramolecular oxidiation-reduction reaction is occurring. The first-order rate constants for this autoreduction of cytochrome c induced by MDH are shown in Fig. 10. These data suggest that addition of MDH to femcytochrome c lowers the pK value of the group whose initial dissociation provides the electron for the autoreduction process; the binding of MDH to the cytochrome c thus allows the intramolecular autoreduction of the cytochrome c to occur at a lower pH value (pH 7.0) than it otherwise would (Fig. 11). Consistent with this proposal is the demonstration that the 695 nm absorption band of the ferricytochrome c disappears on reaction with MDH at pH 7.0, and that the MCD spectrum of the ferricytochromec is the same whether it is autoreduced at pH 10.0 or reacted with MDH at pH 7.0. It should be noted that MDH had no effect whatsoever on the autoreduction of mammalian cytochrome c. The MDH from M . methylotrophus only induced the autoreduction of cytochrome cLand not of cytochrome c H ,but the MDH from Pseudomonas AM 1 was not so specific and induced autoreduction of both cytochromes c from this organism. Whether or not the autoreduction process is involved in the physiological function of MDH and cytochrome c is not yet proven, but a speculative mechanism for such an involvement is presented in Fig. 11. This mechanistic scheme is discussed at greater length in Section III.D.4 in which some
BACTERIAL OXIDATION OF METHANE AND METHANOL
173
evidence is presented to support it, and predictions arising from the scheme are also described. D. THE METHANOL :CYTOCHROME C OXIWREDUCTASE ACTIVITY OF METHANOL DEHYDROGENASE
I . The Demonstration of Methanol: Cytochrome c Oxidoreductme Activity This section describes the experiments leading to the conclusion that pure MDH catalyses the methanol-dependent reduction of pure cytochrome c. There were three independent problems to be overcome in demonstrating this. The first was the instability of the MDH during purification; the second was the presence of “endogenous reductant” on the MDH (Section III.A.3); and the third was the MDH-induced autoreduction of cytochrome c. The first of these problems was avoided by using MDH from M . methylotrophus, protected during aerobic preparation by using KCN, which acts by binding to the substrate binding site. The other problems were overcome by using bacterial cytochrome c as the primary electron acceptor, coupled to a large excess of mammalian cytochrome c as terminal electron acceptor (Beardmore-Gray et al., 1983). A typical experiment using the pure proteins from M . methylotrophus is depicted in Fig. 12. When MDH and mammalian cytochrome c were mixed the cytochrome remained oxidized but addition of a small amount of cytochrome cL led to reduction of the mammalian cytochrome c (present in 50-fold excess over the cytochrome cL). This was presumably due to oxidation of endogenous reductant on the MDH. Added mammalian cytochrome oxidase was used to rapidly oxidize all the ferrocytochrome c, after which
Time (minutes)
FIG. 12. Demonstration of methanol-dependent reduction of cytochrome c by methanol dehydrogenase. From an experiment described by Beardmore-Gray et al. (1983). The reduction of cytochrome c was measured spectrophotometrically at 550 run.
174
C . ANTHONY
KCN was added to inhibit the oxidase. Addition of methanol then led to complete reduction of all the cytochrome c present, the final rate of this methanol-dependent reduction being directly proportional to the concentration of MDH. The rate of reduction of cytochrome c was independent of the pH value, between pH 7.0 and 9.0, and ammonia had no effect on it. Cytochrome cL was essential for the reaction and could not be replaced by cytochrome cH . The initial rates of reduction obeyed hyperbolic kinetics with respect to the concentration of cytochrome cL. The K, value was 1.2p~ and the V,,, value was 1.5 nmol of cytochrome reduced min-l (nmol of MDH)-'. These values were the same during the endogenous reduction observed in the first part of the experiment. That is, the V,, value and K,,, value for cytochrome cL were independent of the substrate responsible for the initial reduction of MDH. No endogenous reduction occurred when cytochrome c, was used instead of cytochrome c,. After growth on methanol, M. methylotrophus contains a small amount of a blue copper protein, probably similar to that found in Methylomonus J (Tobari and Harada, 1981; Tobari, 1984). This protein, partially purified, could not replace cytochrome cL in the system described in Fig. 12, and it had no effect on the methanol :cytochrome cL oxidoreductase activity (Beardmore-Gray, 1982). Table 10 summarizes the results of an investigation of the specificity of the cytochromes c as electron acceptors for various MDH enzymes. No cytochrome c from a non-methylotroph was active and, of the two soluble cytochromes c found in each methylotroph, only one was able to accept electrons from MDH with concomitant production of formaldehyde. This was the cytochrome cL from M. methylotrophus and Pseudomonas AM 1, and one of the cytochromes c from P . denitrif?cans, which was presumably the extra cytochrome c, induced during growth on methanol (van Verseveld and Stouthamer, 1978a). It is of particular importance to note that the methanol :cytochrome c oxidoreductaseactivity demonstrated here with completely pure proteins was inhibited by EDTA, as is the case in whole cells. The obvious target to consider for EDTA action is a divalent cation, perhaps involved in binding the MDH and cytochrome together, but we have no evidence to support such a mechanism; the proteins were purified in the absence of divalent cations and these had no effect when added to the oxidoreductase assay system. Our preliminary experiments have shown that the reversible competitive inhibition occurs at very low concentrations of EDTA (K,is less than 20 p ~ ) , which acts by decreasing the affinity of MDH for cytochrome c,. These results suggest that EDTA reversibly binds to a site on either MDH or on cytochrome cL crucial for formation of an active complex. If the carboxyl groups of the PQQ prosthetic group are involved in binding MDH to
TABLE 10. The specificity of cytochromes c from methylotrophs as electron acceptors for various methanol dehydrogenases. From Beardmore-Gray et al. (1983) Reduction of cytochromes by methanol dehydrogenase from the following bacteria: Methylophilus Paracoccus Pseudomonas AM 1 methylotrophus denitrifcans
Methylophilus methylotrophus Cytochrome c, @I 8.9) Cytochrome c, @I 4.4) Paracoccus denitrifcans Cytochrome c @I acidic) Cytochrome c @I more acidic) Pseudomonas AM 1 Cytochrome cH @I 8.8) Cytochrome cL (PI 4.2)
-
+ -
-
-
+
-
-
+
+
-
+
Reactions with MDH from M.methylotrophus and P. denitrifcans were measured at pH 7.0 in the absence of NH4Cl, whereas reactions with the MDH from Pseudomonas AM1 were measured at its pH optimum @H 9.0) in the presence of NH4Cl which stimulated the rate fourfold. The isoelectric points @I values) of the dehydrogenases from P. denitrifcans, Pseudomonas AM1 and M . methylotrophus were 3.7,8.8 and greater than 8.0, respectively. This information is provided to show that there is no clear pattern of reaction between the cytochromes and dehydrogenasesmerely in terms of their PI values. No MDH was able to reduce horse heart cytochrome c (PI 10.4), or the cytochrome cggl@I 4.7) from the non-methylotroph Pseudomonas aeruginosa. Indicates methanoldependent reduction of cytochrome c with concomitant production of formaldehyde; - indicates no methanol-dependent reduction of cytochrome c or production of formaldehyde.
+
176
C. ANTHONY
cytochrome cL then it is possible that EDTA, whose four carboxyl groups can take up many configurations, may bind to the same site on the cytochrome. These conclusions, using the pure MDH and cytochrome cL , appear to be at variance with those of Carver et al. (1984) and Carver and Jones (1984). These authors have described an extensive study of the mode of action of EDTA on methanol oxidation by whole cells and crude sonic extracts of M . methylotrophus. Their first conclusion was that EDTA inhibits methanol oxidation at a site between methanol and cytochrome c; and this is in agreement with our conclusion, and with results of our similar experiments with this organism and with Pseudomonus AM1. They concluded that EDTA exerts its effect by chelating divalent metal ions (probably M g + ) , which they suggest are involved in the functional association of MDH with the respiratory membrane. This is clearly not the mode of action of EDTA on the soluble methanol :cytochrome c oxidoreductase activity. Some of the results of Carver et al. (1984) could be interpreted in terms of the non-chelating model that we have proposed (above), but certainly not all of them. Furthermore, nothing in our results argues against a role for MgZ+ in binding MDH to the outer surface of the bacterial respiratory membrane. A second effect on methanol oxidation appears to be due to release of soluble cytochromes cH and cL from the periplasm into the medium surrounding the bacteria (Carver and Jones, 1984). The possibility thus remains that EDTA acts on methanol oxidation in more than one way. 2. The Products of Methanol; Cytochrome c Oxidoreductuse Activity Because of the problems arising from the “endogenous reductant” on MDH, and the MDH-induced autoreduction of cytochrome c, it was of interest to determine the stoicheiometry of methanol oxidation during cytochrome c reduction. To achieve this the MDH from M . methylotrophus was purified using KCN instead of methanol as a protective agent during purification, thus ensuring that bound methanol was absent or kept to a minimum. When this MDH was incubated with cytochrome cL plus an excess of mammalian cytochrome c, in the absence of methanol, no formaldehyde was produced, but there was considerable reduction of the cytochrome c (Fig. 13). As most of the cytochrome present in these experiments was mammalian, this endogenous reduction cannot have been due to MDH-induced autoreduction of the cytochrome cL; it must have been due to oxidation of endogenous reductant. In the presence of methanol there was no lag period before production of formaldehyde, indicating that added methanol is used in preference to the “endogenous reductant”. The ratio of molecules of cytochrome c reduced per molecule of formaldehyde produced, increased from an initial value of two throughout
BACTERIAL OXIDATION OF METHANE AND METHANOL
177
Tlme (minuter)
FIG. 13. Production of formaldehyde ( 0 )during methanol oxidation by methanol: cytochrome c oxidoreductase measured as cytochrome c reduction (0) in the presence (-) or absence (---) of methanol. From Beardmore-Gray et al. (1983)
the experiment because some methanol was oxidized completely to formate during the reaction (Fig. 13). This was confirmed by demonstrating that formaldehyde was also oxidized by MDH, the rate of its oxidation being similar to that with methanol as substrate (Fig. 14). The stoicheiometry was two molecules of cytochrome c reduced per molecule of formaldehyde oxidized, and this value was constant throughout the reaction. After complete disappearance of substrate formaldehyde, the cytochrome c became further reduced, the extent of reduction being the same as that occurring in the absence of added substrate. This confirms that added substrate is oxidized in preference to the mysterious “endogenous substrate”.
FIG. 14. The formaldehyde: cytochrome c oxidoreductase activity of methanol dehydrogenase measured by formaldehyde consumed (0) and cytochromec reduction (0).From Beardmore-Gray et al. (1983).
178
C . ANTHONY
It has been known for some time that MDH can catalyse the oxidation of formaldehyde to formate in the dye-linked assay. That it can do so in the cytochrome-coupled system is clearly of some physiological importance. As formaldehyde is required for assimilation, there must be some regulation of the methanol :cytochrome c oxidoreductase activity with respect to formaldehyde oxidation and this is discussed in Section III.A.8. 3. The Relationship Between the Methanol :Cytochrome c Oxidoreductase Activity of Methanol Dehydrogenase and Methanol Oxidation in vivo
Having demonstrated that pure MDH can catalyse methanol-dependent cytochrome cL reduction with concomitant product formation, it is necessary to consider how this finding relates to the oxidation of methanol in vivo. The most important point that must be stressed in this context is that the rates of electron transfer in vitro were far too low (less than 1%) to account for the rates of electron transfer in whole bacteria. The specificity of the system with respect to both cytochrome c and MDH, and its sensitivity to inhibition by EDTA confirm, however, that the oxidoreductase activity in vitro is not completely spurious. We consider that the marked discrepancy between the rates of respiration in vivo and the activity of the purified and reconstituted system may be related to the high concentrations of MDH and cytochrome c found in methylotrophs (Beardmore-Gray et al., 1983). In M . methylotrophus for example, the concentration of both these proteins is about 0 . 5 m ~ , assuming that the periplasmic volume is about 20% of the cell volume and that all the cytochrome cLand MDH is periplasmic (Beardmore-Gray, 1982; Jones et al., 1982). Furthermore, all the cytochrome cL and MDH will be in the associated form most of the time because the K, value of MDH for cytochrome cL is very low. It is possible that the two proteins completely cover most of the outer surface of the bacterial membrane, and that the marked change in environment occurring on release of the two proteins on cell disruption may contribute to their low activity when measured in the reconstituted system (see also Carver et al., 1984). 4. The Relationship Between Methanol :Cytochrome c Oxidoreductase Activity of Methanol Dehydrogenase and its Induced Autoreduction of Cytochrome c.
In this context, the scheme in Fig. 11 provides a mechanistic model that incorporates three key observations. (1) Cytochrome c, is autoreducible, the intramolecular redox reaction obeying first-order kinetics. (2) In the absence of added electron acceptor, the reduction of cytochrome cL by MDH obeys first-order kinetics with respect to the oxidized cytochrome cL and does not
BACTERIAL OXIDATION OF METHANE AND METHANOL
179
nequire added methanol. (3) In the presence of added electron acceptor (mammalian cytochrome c, ferricyanide, or oxidase) the rate-limiting step in the reaction is the reduction of cytochrome cL by MDH. The simplest interpretation of these observations is that, in the absence of added electron acceptor, no electron transfer occurs between the MDH and the cytochrome cL.Under these conditions, the whole effect of the MDH is to stimulate autoreduction by lowering the pK value of the dissociating group on the cytochrome. In the presence of added electron acceptor, however, the autoreduced cytochrome species is able to donate an electron to the acceptor, thus generating an oxidized form of the cytochrome. This can be subsequently reduced to the original species with electrons derived from the oxidation of either methanol or the “endogenous reductant”. This model is consistent with our failure to demonstrate methanoldependent reduction of cytochrome cL when this cytochrome is the sole electron acceptor, even under conditions where there is more than enough to oxidize all of the endogenous reductant on the MDH. Furthermore, this model allows us to make the following predictions, which must be tested to raise the model from conjecture to mechanism. (1) A free radical, due to cytochrome c, should be detectable during the reaction with MDH. (2) The rate of reduction of the external final electron acceptor is unlikely to be greater than the rate of MDH-induced autoreduction. (3) In the presence of methanol, but in the absence of a terminal electron acceptor, no formaldehyde should be produced during the reduction of cytochrome cL by MDH. IV. Energy Transduction During the Oxidation of Methane and Methanol A. INTRODUCTION
The energy available from the oxidation of methane a d methanol is harnessed as ATP by way of proton-translocating electron transport chains consisting of dehydrogenases, iron-sulphur proteins, quinones (Co-Q), cytochromes and cytochrome oxidases. Very little is known about the ironsulphur proteins of methylotrophs, although Co-Q has been investigated in some Gram-negative species. The type of Co-Q operating in electron transport depends on the type of methylotroph. All those studied are Gramnegative and contain ubiquinone; the obligate methylotrophs have Co-Q, (8 isoprenoid units), the Hyphomicrobia have Co-Qgand the pink facultative methylotrophs, Microcyclus species and P . denitrficans have Co-Q,, (Drabikowska, 1977, 1981; Natori et ul., 1978; Urakami and Komagata, 1979). In aerobic bacteria using O2as terminal electron acceptor, the cytochrome
180
C.
ANTHONY
chains do not vary greatly in their composition. Such variations that do occur concern the presence or absence of cytochrome c, the nature of the terminal oxidase (which is usually cytochrome aa, or the most common alternative cytochrome oxidase, cytochrome 0 ) and the branch point to alternative oxidases which occurs at the level of cytochrome b or cytochrome c (Jones, 1977; Haddock and Jones, 1977). That methylotrophs may differ from this normal pattern, and that some variation might be expected within this diverse group of bacteria, is indicated by the following special features of the metabolism of methane and methanol. (1) Every molecule of growth substrate (methane and methanol) is oxidized by way of MDH, including those that are eventually assimilated into cell material. The MDH is a novel type of quinoprotein that reacts directly with cytochrome c (see Section 1II.D). This raises two important questions relating to electron transport from methanol to 0,. The first concerns the possibility that there is an electron transport chain involved in methanol oxidation that is separate from that involved in NADH oxidation; and the second question concerns the coupling of methanol oxidation to proton translocation and ATP synthesis. (2) Between 50 and 90% of the 0,used as terminal electron acceptor for electron transport chains is for the oxidation of MDH (Table 11). (3) Methylotrophs growing on methane have a particularly high requirement for NADH; these bacteria will tend to be NADH-limited and very little electron transport will occur from NADH as electron donor compared with that in typical heterotrophic bacteria (Table 11). (4) The hydroxlyation of methane under some conditions, or in some bacteria, might require a reversed electron transport system to provide NADH, or it might involve “recycling” of electrons from methanol to the oxygenase by way of cytochrome c. Reversed electron transport is also likely to be essential in phototrophs using methanol as a source of reducing power (Anthony, 1982). Table 12 lists those methylotrophs that have now been studied with respect to their electron transport systems, and some generalizations made as a result are discussed below. All methylotrophs grown on methane and methanol have cytochromes b and c, and the cytochrome c is often present in exceptionally high concentrations which mask the cytochrome b in spectra of whole bacteria when recorded at room temperature. The terminal oxidase is either cytochrome aa, or an o-type oxidase; both may be present in a single organism. Although the cytochrome complements of different methylotrophs grown on methane or methanol are fairly similar, the proportion of cytochrome types measured in a single species depends on the growth substrate and
TABLE 11. Electron flow from each dehydrogenase expressed as a proportion of total electron transport. From Anthony (1982)
Growth substrate
Methanol
Electron flow from each dehydrogenase (YO) Formaldehyde Flavoprotein
NADH
~~
Methane Serine pathway bacteria (e.g. Methylosinus trichosporium) Ribulose monophosphate pathway bacteria (e.g. Methylococcus capsulatus) Methanol Serine pathway bacteria (a) Formaldehyde yields one NADH molecule (e.g. Pseudomonas AMl) (b) Formaldehyde oxidation yields two NADH molecules Ribulose monophosphate pathway bacteria (e.g. Methylophilus methylotrophus) Ribulose bisphosphate pathway bacteria (e.g. Paracoccus denitr$cms) Conventiod slrbstrates (iiioding formate)
78-88
WAD+-linked)
e12
0-1 3
84-92
(NAD+-linked)
0
8-1 6
53-56
35
10
0-3
53-64
(NAD+-linked)
9-1 5
21-40
57-70
(NAD' -linked)
0
3w3
52-62
(NAD+linked)
0
3W8
0
0
&26
74-100
The range of values predicted is for a range of assumed P/O ratios; the lowest is assumed to be one for each oxidation step and the highest P/O ratios are assumed to be 1,2 or 3 for MDH, flavoprotein and NADH, respectively. Higher P/O ratios lead to a lower proportion of electron transport from NADH dehydrogenase and a higher proportion from other dehydrogenases. During growth on methane only 25 to 40% of the total O2consumed is by way of electron transport and oxidases, the remainder being used in the initial hydroxylation reaction (assumed here to be NADH-linked). In serine pathway bacteria the oxidation of acetyl-CoA to glyoxylate is assumed to involve a flavoprotein.
182
C.
ANTHONY
TABLE 12. Cytochromes, electron transport systems and proton translocation in methylotrophs able to grow on methane or methanol Organism
References
(a) Cvtocbromes and electron transport systems
Metcanotrophs Methylomonas albus Methylomonas agile Methylomonas methanica Methylococcus capsulatus Methylosinus trichosporiwn
Methylosinus sp. GB2 Methy lobacterium organophilum Obligate methanol-utilizers Methylomonas P11 Methy lophilus methylotrophus
Pseudomonas W6 (MB53) Methylomonas J Organism 4025 Facultative methanol- or methylamine-utilizers Pseudomonas extorquens Pseudomonas AM 1
Protaminobacter spp. Pseudomonas methylica sp 2 Acetobacter sp. MB58 Hyphomicrobiwn
Davey and Milton (1973). Monosov and Netrusov (1975). Tonge et al. (1974), Ferenci et al. (1975), Ferenci (1976a), Babel and Steudel (1977), Pate1 et al. (1979a) Ribbons et al. (1970), Grozdev et al. (1983) Davey and Mitton (1973), Weaver and Dugan (1979, Monosov and Netrusov (1975), Tonge et al. (1974, 1975, 1977a), Higgins et al. (1976a), Higgins (1979, 1980), Hammond et al. (1979) Babel and Steudel(l977) OConnor and Hanson (1978), Wolf and Hanson (1978) Drabikowska (1977, 1981) Cross and Anthony (1980a, b), Dawson and Jones (1981c), Beardmore-Gray et al. (1982, 1983), Burton et al. (1983), Jones et al. (1982), Carver and Jones (1983, 1984), Carver et al. (1984), Froud and Anthony (1984a, b), Quilter and Jones (1984) Babel and Steudel(l977) Ohta and Tobari (1981), Tobari (1984) Vrdoljak and Froud (1982), Vrdoljak et al. (1984), Lawton and Anthony (1985a, b)
Tonge et al. (1974, 1977b), Higgins et al. (1976a, b) Tonge et al. (1974), Anthony (1975), Widdowson and Anthony (1979, OKeeffe and Anthony (1978, 1980a, b), Netrusov and Anthony (1979), Keevil and Anthony (1979a, b), Ivanovsky et al. (1980), Beardmore-Gray et al. (1982, 1983), Tobari (1984), Froud and Anthony (1984b), Fukumori et al. (1985) Vrdoljak et al. (1984) Netrusov et al. (1977) Steudel and Babel (1982) Hirsch et al. (1963), Tonge et al. (1974), Widdowson and Anthony (1975), Babel
BACTERIAL OXIDATION OF METHANE AND METHANOL
183
TABLE 12. (continued) Organism
References
and Steudel (1977), Large et al. (1979), Duine et al. (1979a) Paracoccus denitrijicans Bamforth and Quayle (1978a), van Verseveld and Stouthamer (1978a), Willison and John (1979), Boogerd et al. (1980), Vignais et al. (1981), Willison and Haddock (1981), Willison et al. (1981a, b), van Verseveld et al. (1983), Ludwig et al. (1983), Davies et al. (1984), Froud and Anthony (1984b) (a) Proton translocation and ATP synthesis in metblotrophp (Papers indirectly related to this topic are included in parentheses) Methylosinus trichosporium Tonge et al. (1977b) Pseudomonas extorquens Higgins (1980), Hammond et al. (1981) Pseudomonas AM 1 OKeeffe and Anthony (1978), Keevil and Anthony (1979b), Netrusov and Anthony (1979); Netrusov (1981), Hammond et al. (1981), Fukumori et al. (1985) Methylophilus Cross and Anthony (1978), Dawson and methylotrophus Jones (1981a, b, c, 1982), Patchett et al. (1985) Pseudomonas EN (NClB Drozd and Wren (1980) 11040) Paracoccus denitrijicans van Verseveld and Stouthamer (1978a, b), van Verseveld et al. (1978, 1981, 1983), Alefounder and Ferguson (1981), (Kell et al., 1978; McCarthy et al., 1981; Willison and Haddock, 1981; Solioz et al., 1982; Ferguson, 1984)
growth conditions. For example, the concentration of soluble cytochrome c is higher in methylotrophically grown Pseudomonas extorquens and Pseudomonas AM 1 than when they are grown heterotrophically; furthermore, the concentrations of all cytochromes (a, b and c) on membranes is markedly lower during methylotrophic growth (Tonge et al., 1974; Widdowson and Anthony, 1975; Higgins et al., 1976a; Keevil and Anthony, 1979b). This is probably related to the greater capacity for NADH oxidation required during heterotrophic growth (Table 11) and to the particular importance of cytochrome c during methanol oxidation. The relative proportions of the two main soluble cytochromes (cytochromes cH and cL) do not usually vary greatly with varying growth conditions, and they are similar in a variety of methylotrophs that differ with respect to their carbon-assimilation pathways and terminal oxidases (Froud and Anthony, 1984b).
184
C.
ANTHONY
In methylotrophs it is usually found that all three types of cytochrome (a, b and c) are able to react with CO to some extent, as shown by measurements of spectra of whole bacteria. Because the reaction of haem iron with CO is analogous to its reaction with O,, the reaction of CO with cytochromes is used as a preliminary indication of the presence of cytochrome uu3 and cytochrome o (an oxidase containing haem b). This is, in itself, insufficient evidence for the operation of a particular type of oxidase, and some demonstrations of “cytochrome 0’’ in the literature are probably erroneous. It is especially difficult to be certain that a CO-binding b-type cytochrome is cytochrome o if any other oxidase is also present. The cytochromes c of all methylotrophs examined are able to react with CO. This is an unusual feature for a cytochrome c; it does not necessarily indicate an oxidase function, and it is discussed in detail in Section III.C.3. All the methanotrophs tested have cytochrome uu3 as oxidase, as have many of the methanol-utilizers. Some methylotrophs also contain the alternative oxidase, cytochrome 0, the amount of which may depend on growth conditions (e.g. M . methylotrophus, see Section 1V.E). A few methanol-utilizers contain cytochrome o as the sole oxidase under all growth conditions; these include the obligate methylotrophs Methylomonus P11 (Drabikowska, 1977, 1981) and organism 4025 (Vrdoljak and Froud, 1982). In summary, all methylotrophs grown on methane or methanol have band c-type cytochromes; they usually have a cytochrome ua3,and sometimes an o-type oxidase which may be the sole oxidase in some species. A second generalization is that MDH reacts with cytochrome c, thus bypassing cytochrome b in the electron transport chain (see Section 1II.D). This leaves the following questions to consider: are the cytochromes arranged in the conventional mitochondria1 sequence as found in many heterotrophic bacteria? Are the electron transport chains arranged to translocate protons as conventionally described? Is there a typical ATP synthetase coupled to a proton motive force? Is electron transport by way of the unusual MDH/ cytochrome system arranged to create a proton motive force and, if so, how? What I have done below is to select some examples of electron transport pathways in methylotrophs in order to highlight any points that might make them especially interesting, and I have discussed proton-translocation studies in relation to the work on electron transport in the same organism, where these two approaches have complemented each other. B. ELECTRON TRANSPORT AND PROTON T R A N S ~ A T I O NIN Methylosinus trichosporium AND OTHER METHANOTROPHS
Methylosinus trichosporium is an obligate methane-utilizer, assimilating its carbon substrate by the serine pathway and having a Type I1 internal
BACTERIAL OXIDATION OF METHANE AND METHANOL Amytal NADH+Cyt
M e I h a n ?H20 ~ Methanol
,'
/
'\
vwylow KCN
-. -
Anttmycm A
b+ *
CyI C
185
low KCN
Cyt
U U ~
02
\
Methanol
dehydrogmmw
Cy; cco -)to,
I
High KCN
FIG. 15. Tentative scheme for electron transport in Methylosinus trichosporium and Pseudomonus extorquens. After Higgins (1980). A notable feature of this scheme is that the cytochrome c is proposed as electron donor to the MMO of Methylosinus trichosporium and as the predominant oxidase during growth on methanol by both bacteria. As emphasized in the text, there is little evidence for an oxidase function for cytochrome cc0 in methylotrophs. An alternative electron donor to the MMO is likely to be NADH (see Section 1I.C).
membrane system. As with all methane-utilizers, less than 40% of the O2consumed during growth is by way of oxidases, the rest being used in the initial hydroxylation of methane. A second general point is that in all methaneutilizers that use NADH as the reductant for the MMO, between 80 and 90% of electron transport is coupled to oxidases by way of MDH, NADH dehydrogenase being relatively unimportant (see Table 11). All studies of electron transport during methane oxidation are very difficult to interpret because there are at least two 0,-consuming sites, the oxygenase and the oxidase, and these may be sensitive to the same or different inhibitors. A further complication is that electron donors to the oxygenase are required, and their production may depend on the further metabolism of methanol arising from the methane hydroxylation. Thus mid-chain inhibitors such as amytal or antimycin A may inhibit a conventional electron transport chain between NADH and O,, or they may inhibit the production of reductant for the oxygenase. The relatively few studies of electron transport in methanotrophs have emphasized some points of similarity with the methanol oxidizers. In particular, they contain cytochromes b and c and have cytochrome au3 as a potential oxidase; the MDH is typical and some, at least, is bound to membranes in the intact bacteria; and the MDH almost certainly donates electrons to the cytochrome chain at the level of cytochrome c (see Table 12 for references). The electron transport scheme proposed, tentatively, for Methylosinus trichosporium is shown in Fig. 15. This is based on inhibitor studies, and on the properties of the soluble cytochrome c, whose CO-binding characteristics have led to the suggestion that it is the main oxidase during methanol
186
C. ANTHONY
oxidation (Higgins et ul., 1976a; Tonge et al., 1977a). In considering an oxidase function for the cytochrome c it should be recalled that, provided it is reducible by substrate (methanol, NADH or ascorbate) and is also slightly autoxidizable,an apparent (non-physiological) activity is bound to be observed when the activity of the normal oxidase such as cytochrome a, is inhibited by low concentrations of cyanide. In the absence of any further evidence, it is probably not justified to ascribe an oxidase function to the cytochrome c of Methylosinus trichosporium. Proton translocation measurements with Methylosinus trichosporium have shown that the highest H + / O ratio obtainable is two with methane, methanol, formaldehyde or formate as respiratory substrates (Tonge et al., 1977b). Whether or not this reflects a maximum P/O ratio of only one for each oxidation step is not known, but the failure to measure any respirationcoupled proton translocation at all in Methylococcus cupsulatus, which has the alternative internal membrane arrangement, might suggest the existence of special technical and interpretive difficulties in measuring proton translocation of bacteria having complex internal membrane systems (D. T. O’Keeffe, H. Dalton and C. Anthony, unpublished results). C. ELECTRON TRANSPORT AND PROTON TRANSLOCATION IN
Pseudomonus AM 1
;?’seudomonus AM1 is a pink facultative methylotroph able to grow on methanol but not on methane. It assimilates methanol by the serine pathway, and formate dehydrogenaseis the only NAD+-linked enzyme involved in the oxidation of methanol to C02. Growth thus tends to be NADH-limited and less than 5% of electron transport to O2during growth on methanol is from NADH; 50% is from MDH and the remainder from formaldehyde dehydrogenase and flavoproteins (Table 11). Its electron transport system, and that of its cytochrome c-deficient mutant, have been investigated using a range of approaches including cytochrome characterization (Anthony, 1975;Widdowson and Anthony, 1975;Keevil and Anthony, 1979b;O’Keeffe and Anthony, 1980a, b; Beardmore-Gray et al., 1982, 1983; Tobari, 1984; Froud and Anthony, 1984b), studies of proton translocation (O’Keeffe and Anthony, 1978; Keevil and Anthony, 1979a; Hammond et al., 1981), measurement of ATP synthesis in vesicle preparations (Netrusov and Anthony, 1979; Netrusov, 1981), and determination of growth yields (Goldberg et al., 1976; Keevil and Anthony, 1979a; Tsuchiya et al., 1982). Some of the results with respect to the site of interaction of MDH and the cytochrome chain have already been discussed in Section 1I.A. Pseudomonas AM1 contains at least two soluble cytochromes c (cytochromes cH and cL), some membrane-bound cytochrome c, two cytochromes b and cytochrome au,, which is probably the only oxidase.
BACTERIAL OXIDATION OF METHANE AND METHANOL ( 0 ) Electron transport
+
in wiid-type bocteno (carbon-limited cmditrons)
tci
NADH Rotenone
-
* cyt c
/i
Antimycin A
Methanol
-
Methanol d e h y d - 7
cyt 003
f L
O2
KCN
cyt cL---cyt
2H
187
cn
+
(b) Electron transport in wild-type bacierio (corbon-excess conditions) and in a cytochrome c-deficient mutant
NADH Rotenone
Cyt b
I
-..--.
Cyt oo3
-
Antimycin A Methanol
Methanol cyt cL--* dehydrogeq\-
Cyt CH
2H+
FIG. 16. Electron transport and proton translocation in Pseudomonas AM1 (based on work of author and colleagues). Cytochrome c may be able to mediate between cytochrome b and the oxidase in all conditions but it does not appear to be involved in proton translocation in conditions of carbon excess (4or ammonia limitation). It is not known if the cytochrome c mediating between cytochromes b and aa, is the same as that involved in methanol oxidation (cytochrome cL)or if it is the alternative soluble cytochrome c (cytochrome cH). There is some evidence that cytochrome c, is the immediate electron donor to the oxidase (Tobari, 1984; Fukumori et al., 1985).
Cytochrome cL is the electron acceptor from MDH (Beardmore-Gray et al., 1983; and Section 1II.D.). The proportions of cytochromes cH and cL do not vary greatly with growth conditions (Froud and Anthony, 1984b). Because all oxidizable substrates are able to reduce all of the cytochrome c and cytochrome uu3, it was initially concluded that the “conventional” cytochrome sequence occurs in Pseudomonas AMl, as indicated in Fig. 16. However, it appears that cytochrome c is not necessarily involved in the oxidation of NADH, or of cytochrome b, because in the mutant lacking cytochrome c all substrates, except for methanol and methylamine, are oxidized, and are capable of reducing cytochromes b and uu3. If cytochrome c is not involved in the oxidation of NADH, then a maximum H+/O ratio of only four, equivalent to two proton-translocating segments, would be expected during meausrements of respiration-coupled proton translocation (Jones, 1977;Haddock and Jones, 1977). This was indeed the result found for substrates oxidized by way of NADH, and the value was the same in the mutant lacking cytochrome c; this appeared therefore to confirm the pathway shown in Fig. 16 (OKeeffe and Anthony, 1978). Further confirmation was
188
C. ANTHONY
obtained by measuring ATP synthesis in membrane vesicles of batch-grown Pseudomonus AM 1 and its cytochrome c-deficient mutant (Netrusov and Anthony, 1979; Netrusov, 1981). It was shown that the oxidation of methanol to formaldehyde is coupled to the synthesis of ATP, the P/O ratio being similar to that observed during the oxidation of ascrobate/TMPD, and lower than that found with NADH and succinate. The P/O ratios for succinate and NADH oxidation were identical in vesicles prepared from wild-type bacteria and from the mutant lacking cytochrome c (Netrusov, 1981), thus further supporting the scheme in Fig. 16 in which cytochrome c is not involved in proton translocation during the oxidation of cytochrome b. This conclusion cannot hold for all growth conditions, however. All the results described above were obtained with batch-grown bacteria (nutrient-sufficient conditions), but higher H+/O ratios (equivalent to three proton-translocating segments) were measurable in wild-type bacteria grown in continuous culture under carbon-limited conditions. By contrast, lower H+ / O ratios (two proton-translocating segments) were found in the wildtype bacteria grown in continuous culture under conditions of carbon excess, and in the cytochrome c-deficient mutant under all growth conditions. These results suggest that the cytochrome c may be involved in electron transport and proton translocation from NADH under carbon-limited conditions, but not during growth with an excess of carbon substrate (Keevil and Anthony, 1979a). In our work on proton translocation in Pseudomonus AM 1, thiocyanate ions were used to abolish the membrane potential, and this ion also inhibited methanol oxidation, so preventing determination of H+/O ratios with this important substrate. The inhibitory effect of thiocyanate was confirmed by Hammond et ul. (1981), who overcame the problem by using a mixture of thiocyanate plus valinomycin. Their results with methanol as substrate were all consistent with the proton-translocating electron transport chain shown in Fig. 16. Hammond et ul. (1981) also demonstrated that the state of the bacteria used for the determination of H+/ O ratios can markedly affect the values obtained. They confirmed our conclusion that the stoicheiometries vary with growth conditions, but they also showed that other factors were important and that interpretation of the variations described above must be treated with caution. Our demonstration that cytochrome c may be involved in ATP synthesis during the oxidation of NADH was confirmed by demonstrating that, during carbon-limited growth on succinate, the yields measured with the wild-type bacteria containing cytochrome c were higher than those meausred during growth of the cytochrome c mutant on this substrate. It should be noted, however, that although incorporation of cytochrome c into the electron transport chain between cytochromes b and uu3gave these higher yields on
BACTERIAL OXIDATION OF METHANE AND METHANOL
189
succinate, an increase in the P/O ratio for NADH oxidation would give little benefit to the organism during growth on methanol, because relatively little NADH is oxidized during growth on this substrate (see Table 11). In summary, all of the experiments described here embody considerable difficulties of execution and interpretation. The simplest conclusion that can be drawn is that in Pseudomonas AM1 cytochrome c is essential for the oxidation of methanol, and for coupled ATP synthesis (P/O ratio of one); and that it may also be involved in electron transport, proton translocation and ATP synthesis during oxidation of NADH, but perhaps only during growth under carbon-limited conditions. A final point to mention with respect to the electron transport in Pseudomonas AM1 is the function of the blue copper proteins found in this organism. Tobari and Harada (1981) have shown that a blue copper protein, which they have called amicyanin, is the immediate electron acceptor from methylamine dehydrogenase. It has been proposed that this protein then donates electrons to cytochrome c,, and thence to cytochrome cH and the oxidase (Tobari, 1984). It has also been proposed that a second, azurin-type protein might mediate between amicyanin and the oxidase, or between cytochrome c and the oxidase, during the oxidation of methanol. Although we have shown blue copper proteins to be present, none of our results at present support the proposal that a blue copper protein is essential during growth on methanol; although amicyanin is certainly involved during growth on methylamine (Lawton and Anthony, 1985and unpublished observations). D. ELECTRON TRANSPORT AND PROTON TRANSLOCATION IN
Paracoccus denitriJcans
The facultative autotroph, P. denitriJicansgrows on methanol as sole source of carbon and energy, oxidizing the methanol to C02 and assimilating this CO, by the ribulose bisphosphate pathway. Methanol is oxidized by MDH, and formaldehyde and formate by NAD+-linked dehydrogeanses. It can be estimated that about 40% of electron transport to 0, will be from NADH and 60% from MDH during growth on methanol (see Table 11). This organism is also capable of typical heterotrophic growth on multicarbon compounds and of typical autotrophic growth on H, plus CO,. Growth on all these carbon sources may be either aerobic, or anaerobic with nitrate as terminal electron acceptor. It is perhaps this wide variety of growth conditions that is responsible for the complexity of electron transport systems described in this organism (see Table 12 for references). Paracoccus denitriJcans always contains cytochrome c, cytochrome c, , at least two cytochromes b, cytochrome aa3 and usually cytochrome 0.There is some doubt about the nature and function of this potential oxidase (see, for
190
C.
ANTHONY
(a) Aerobic electron h o n m
Fcnroldehyde
>= NADHRotenone -*
[ ] Cyt. 6562
cyt. b56S Antimycin A
dH
115pM
KCN
-
Methonol -Methanol dehydrogenase
-77
Cyt.cC0
(b) Anaerobic elecfron transpoot
COP
Formoldehyde
f Methanol dehydrogenase
I
N O S (-NO*-
1
FIG. 17. The respiratory chains of Paracoccus denitrificans. (a) from van Verseveld and Stouthamer(1978a, b); (b) from Bamforth and Quayle (1978a). The hatched bars indicate the site of action of inhibitors (with concentrations for 50% inhibition). It should be noted that no evidence was presented for the involvement of more than one cytochrome c in methanol oxidation. It is possible that the cytochrome c,, is oxidized directly by cytochrome a,. This inducible cytochrome c,, is presumably equivalent to cytochrome c, in other methylotrophs (see the text and Table 10).
example, Vignais et ul., 1981), and there is now some evidence for a second alternative oxidase, cytochrome a, (Van Verseveld et ul., 1983). Cytochrome uu3appears to be the only oxidase involved in the oxidation of methanol and NADH during growth of P . denitrificuns on methanol (van Verseveld and Stouthamer, 1978a; Bamforth and Quayle, 1978a). The scheme presented in Fig. 17 (based on van Verseveld and Stouthamer, 1978a) shows the electron transport chain branching from cytochrome b to the alternate oxidases. This differs from the pathway in M. methylotrophus, in which the branch point to the alternative oxidases is at cytochrome c rather than cytochrome b. It is presumably for this reason that P . denitrificuns cannot oxidize methanol by way of the 0-type oxidase. P. denitrificuns contains at least two soluble cytochromes c during growth on methanol, only one of which reacts with MDH (Beardmore-Gray et ul., 1983). This cytochrome is the one referred to as cytochrome cco in Fig. 17 and it corresponds to cytochrome cL in other methylotrophs (see Section 1II.D). This cytochrome differs from those in other methylotrophs in being induced during growth on methanol. The reason for this is perhaps that P. denitrificuns has a separate membranebound cytochrome c, which may be replaced in other methylotrophs by being
BACTERIAL OXIDATION OF METHANE AND METHANOL
191
a tightly bound form of cytochrome c, and which must, therefore, be synthesized under all growth conditions (Froud and Anthony, 1984b; and Section IV.E). As found with Pseudomonas AMl, mutants lacking cytochrome c no longer grew on methanol (Willison and Haddock, 1981). Measurements of cell yields and proton translocation in P. denitrijicans suggest that this organism differs from other methylotrophs in translocating between three and four protons during methanol oxidation (van Verseveld and Stouthamer, 1978a). This high number is probably because the cytochrome oxidase also pumps protons (van Verseveld et al., 1981; Solioz et al., 1982). The results of these studies confirmed that the oxidation of methanol to formaldehyde is coupled to synthesis of one molecule of ATP per molecule of methanol oxidized. During anaerobic growth of P . denitrijicuns, methanol is oxidized by the same MDH as operates during aerobic growth, and it has been proposed that this dehydrogenase reduces the electron transport chain at the level of cytochrome c, which is oxidized by nitrite as the terminal electron acceptor (Fig. 17). Because nitrate usually accepts electrons from cytochrome b, it is proposed that during growth on nitrate the reductant for nitrate reductase is NADH, produced during subsequent oxidation of formaldehyde and formate. The nitrite produced from this process is then used as the terminal electron acceptor for MDH (Bamforth and Quayle, 1978a). E. ELECTRON TRANSPORT AND PROTON TRANSLOCATION IN
Methylophilus methylotrophus Methylophilus methylotrophus is an obligate methanol-utilizer which, because it can express high growth rates and yields, has been selected by ICI as the organism of choice for production of singlecell protein. It assimilates methanol by the RUMP pathway and produces two molecules of NAD(P)H during oxidation of methanol to C02. Less than 35% of electron transport to O2is likely to be from NADH, and more than 65% from MDH (Table 11; Anthony, 1982). 1. The Electron Transport Chains of Methylophilus methylotrophus
The electron transport chains that operate in this organism are summarized in Fig. 18; they are based on the work described below which, unless stated otherwise, was first published by Cross and Anthony (1980a, b) and Froud and Anthony (1984a, b). Methylophilus methylotrophus contains at least three c-type cytochromes, all remarkable for their high mid-point redox potentials (3W360 mV). The properties of the two main soluble cytochromes(cytochromes c, and cL) have
192
C.
ANTHONY
(01 Methonol- limited conditions HQNO Antimycin A CYt bso
NADH +[Cyt Rotenone
[, * I
+ *-
cyt c
b,,,]
/
2 p M KCN 5 O p ~ozide
uos++oe
-cyt CYt
I
0 -02
HQNO
Methonol -+Methanol dehydmgenase
2f’MKCN
4 p azide ~
(b) Methanol-excess conditions (Opor niiroqen limited)
NADH
+[
Rolenone
CYf b60
1
HQNO Antimycin A
[Cyl
++ .
CYt
410
/
c
-*I
cyt
I
0
HQNO
w
+
0
2
2FMKcN 4 p azide ~
Methanol+Methonol dehydrogermse
FIG. 18. Electron transport in Methylophilus methylotrophus. From Cross and Anthony (1980a, b). The hatched bars indicate the site of inhibitors (concentration for 50% inhibition). The thickness of the lines indicates the relative importance of the alternative oxidases. * denotes that a single cytochrome c is shown for convenience. The only soluble cytochrome c reacting with MDH is cytochrome cL; whereas cytochrome cH is the probable donor to the o-type oxidase (which itself has a cytochromec component). The soluble cytochromes c can react with each other. It is not known which cytochrome c accepts electrons from cytochrome b nor which donates electrons to cytochrome aa,. Fig. 19 shows a more complete view of the respiratory chain operating under methanol-excess growth conditions.
been described in Section III.C.3. The proportions of these two cytochromes do not vary markedly with variations in the growth conditions, and they are similar to those measured in other methylotrophs. The soluble forms of these cytochromes are located exclusively in the periplasm, and can be readily removed from the bacteria by treatment with EDTA (Jones etal., 1982; Carver et al., 1984; Carver and Jones, 1984). Cytochrome cH, the smaller of the two soluble cytochromesis the most readily released from whole bacteria, the permeability of the outer membrane to this cytochrome leading to the release of up to 40% of the total cytochrome c into the culture medium during growth (Cross and Anthony, 1980b). Of the cytochrome c measured in these bacteria, 3&50% is membrane bound, and redox potential measurements originally indicated that most of it was of the same type as was found in solution after breakage of the bacteria (Cross and Anthony, 1980b). It is now known that this is not the case (Froud and Anthony, 1984b). Less than 6% of the cytochrome cH is membranebound, together with about 30% of the cytochrome cL which appears to be identical with the soluble cytochrome cL . The second major membrane-
BACTERIAL OXIDATION OF METHANE AND METHANOL
193
bound cytochrome c is the cytochrome c component of the o-type oxidase, cytochrome co (see the next section). This cytochrome c (55% of the cytochrome c in membranes) has a redox potential of about 350mV, which happens to be the same as that of the soluble cytochrome cH, from which it differs, however, in all other respects (Froud and Anthony, 1984b). The soluble cytochrome cLis the electron acceptor from MDH (Section III.D), but the role of the membrane-bound cytochrome C, is not known; it is possible that it is involved in the oxidation of NADH, forming part of a cytochrome bc, complex analogous to the cytochrome bc, complex in mitochondria. The soluble cytochrome cH is the preferred electron donor to the o-type oxidase (see the next section), but it is not known if it is also an electron donor to cytochrome uu3.If it is, then it may be at a branch point, mediating between soluble cytochrome c, and the terminal oxidases; and also between the cytochrome bc, complex and the terminal oxidases. In this context it should be noted that the soluble cytochromes c are able to interact with each other in vitro, and that all substrates are able to reduce all of the cytochrome c present in whole cells of M . methyfotrophus (Cross and Anthony, 1980b; Carver and Jones, 1984). Membranes of M. methyfotrophus always have two cytochromes b with mid-point redox potentials of 60 mV and 110mV. Under carbon-excess growth conditions (0,or ammonia limitation) a third cytochrome b (midpoint potential + 260 mv), which binds CO, is induced 10-fold. The synthesis of this new potential oxidase occurs in parallel with a similar 10-fold increase in the rate of oxidation of TMPD by membranes, and in a large increase in sensitivity of respiration to azide. There is no cytochrome uu3 under these conditions, and it is reasonable to assume that the induced cytochrome b is a genuine alternative oxidase and hence it has been called cytochrome o. Both this oxidase and cytochrome uu3can be involved in the oxidation of methanol and NADH; and all the evidence indicates that cytochrome c is the point of interaction of MDH with the electron transport chain. These two facts, together with evidence from inhibitor studies, suggest that the branch point for electron flow to the alternative oxidases is at the level of cytochrome c. This conclusion is supported by the results of work on proton translocation occurring in the presence of a variety of substrates and inhibitors in M. methyfotrophus(Dawson and Jones, 198lc). The nature of the terminal parts of the electron transport chains of M. methyfotrophus is determined by the growth conditions (see Fig. 18). Cytochrome uu3 is the predominant oxidase in bacteria growing in continuous culture under conditions of methanol limitation; whereas under conditions of methanol excess (0,or ammonia limitation) the 0-type oxidase is induced 10-fold and cytochrome uu3 is completely absent. Furthermore, during growth in batch culture (exponential phase) the o-type oxidase is the
+
+
194
C.
ANTHONY
predominant oxidase. These observations would appear to rule out the possibility discussed by Dawson and Jones (198 lc) that M. methylotrophus contains two respiratory chains with functionally distinct pools of cytochrome c such that NADH is oxidized only via cytochrome uu, and methanol is oxidized only by way of cytochrome 0.It would also rule out the possibility discussed by Carver and Jones (1984) that the o-type oxidase is specifically involved in the oxidation of NADH and various flavin-linked substrates. Why M. methylotrophus has alternative oxidases, whose syntheses are regulated by the carbon supply, is not clear. It is more usual for regulation of oxidase synthesis to be determined by O2concentration, but even here the rationale for possession of alternative oxidases is not particularly obvious (Harrison, 1976;Jones, 1977; Haddock and Jones, 1977). The affinity of both values could not be determined oxidases for O2 is high, although the K,,, because they were below the limit of sensitivity of the conventional oxygen electrode. If the terminal part of the respiratory chain, having the o-type oxidase, is uncoupled from ATP synthesis, then under conditions of carbon excess, when cytochrome uu3is absent, the cell yield would be lower; but the advantage of this to the bacteria is unclear unless uncoupling permits a higher growth rate in the presence of plentiful carbon substrate. That electron transport is not uncoupled from ATP synthesis with either oxidase is clear, however, from the demonstration that the overall H+ and K + translocation stoicheiometries measured in carbon-limited bacteria were independent of the nature of the oxidases responsible for respiration (Dawson and Jones, 1981c). In the context of the relationship between growth efficiency and oxidase function in methylotrophs, it is worth mentioning that in two other obligate methylotrophs growing on methanol the sole oxidase under all growth conditions is an o-type oxidase. These bacteria, similar in most respects to M . methylotrophus, are Methylomonus P11 (Drabikowska, 1977, 1981), and organism 4025, an obligate methylotroph selected for its potential as a fast-growing, high-yielding source of single-cell protein (Vrdoljak and Froud, 1982). 2. The o-Type Cytochrome Oxidase of Methylophilus methylotrophus The work described in the previous section and in Section 1II.C has shown that M . methylotrophus has at least two soluble cytochromes c which are able to react with each other, that only one of these, cytochrome cL,is the electron acceptor from MDH, and that the branch point to the oxidases is at the level of cytochrome c. This raises two obvious questions. Are both soluble cytochromes c able to act as electron donors to both the oxidases, and are
BACTERIAL OMDATION OF METHANE AND METHANOL
195
there any proteins required for the oxidation of methanol other than MDH, cytochrome cL and the oxidase? In order to answer these questions it was necessary to purify one of the oxidases. To avoid the complication of starting with material containing two oxidases, we chose to purify and characterize the o-type oxidase from the membranes of M . methylotrophus grown on excess methanol under conditions of O2 limitation (Froud and Anthony, 1984a). The oxidase was extracted from the membranes using Triton X-100 and was purified to homogeneity (15-fold). Assuming that no inactivation had occurred during purification, then it could be concluded that the oxidase constitutes about 7% of the membrane protein. The pure active oxidase consists of equal amounts of b-type and c-type cytochromeswhich have molecular weights of 31,500and 23,800 respectively. The active oxidase complex probably consists of two cytochrome b units plus two cytochrome c units. The cytochrome c subunit does not correspond to either of the two predominant soluble cytochromes c, nor to the membrane-bound cytochrome cL (Froud and Anthony, 1984b). Although of similar size to cytochrome c,, the pattern of peptides produced by proteolytic digestion of the cytochrome c component of the oxidase was sufficiently different to conclude that it is not the same as soluble cytochrome cL (H.M. Ashworth and C. Anthony, unpublished observations). This differs from the preliminary conclusion of Carver and Jones (1983), who first showed this oxidase to have cytochrome c and cytochrome b components. In their description of the partially purified oxidase they suggested that the cytochrome c component might be the same as the soluble cytochrome cL. A problem of terminology of the o-type oxidases has been raised by the discovery that the active oxidase contains not only a CO-binding cytochrome b but also a cytochrome c component that is tightly bound and essential for activity. This was discussed at length by Froud and Anthony (1984a) who concluded that a suitable name is one analogous to cytochrome ua3, the proposed name for the o-type oxidase being cytochrome co. Although this is the most suitable name, to avoid confusion when discussing earlier work this oxidase is sometimes referred to as cytochrome o. When this is done in the present review, reference to cytochrome o of M . methylotrophus is always to the complete active oxidase and not merely to the cytochrome b component. The pure 0-type oxidase (cytochrome co) from M . methylotrophus (Froud and Anthony, 1984a) oxidized TMPD very rapidly, the turnover number (262 s- ' ) being almost identical to that measured using membrane preparations. It was inhibited non-competitively by azide (Ki 1.13 /AM)and KCN (40.2 p ~ ) the , 4 values being similar to those measured during respiration by whole bacteria. The purified oxidase was insensitive to inhibition by hydroxy-quinoline N-oxide (HQNO) (Carver and Jones, 1983; Froud and
196
C. ANTHONY
Anthony, 1984a). This suggests that HQNO inhibits TMPD oxidation by whole membranes (Cross and Anthony, 1980b) at a site not available on the Triton-solubilized oxidase complex. Although the affinityof the oxidase for the two soluble cytochromes c from M. methylotrophus was similar (K, about 1 6 p ~ )the , cytochrome cH was oxidized 50 times faster than the cytochrome cL, thus suggesting that the physiological electron donor to the oxidase complex is the smaller basic cytochrome cH. Preliminary observations indicated that the affinity of the oxidase complex for O2is very high, the K , value being less than 1p ~ . The observation that an essential part of the oxidase complex is a cytochrome c component raised the possibility that this component is the electron acceptor from MDH, but this was shown to be unlikely. When mixed with MDH, the oxidase did not catalyse methanol-dependent O2 consumption. This is a further confirmation that the cytochrome c component is not, as suggested by Carver and Jones (1983), identical with the soluble cytochrome cLfrom this organism. This demonstration that “methanol oxidase” activity could not be reconsituted from pure MDH plus the pure oxidase suggested that a separate cytochrome c must be essential for such reconstitution. This was confirmed by demonstrating that incubation of methanol, MDH and oxidase with pure cytochrome cL led to “methanol oxidase” activity. When low concentrations of oxidase were used, the oxidation of cytochrome c became rate-limiting and addition to the system of the preferred electron donor (cytochrome cH) increased the overall rate. All activity of the reconstituted “methanol oxidase” was abolished by a i d e or EDTA, as found in studies of methanol oxidation by whole bacteria. During growth under methanol-limited conditions, when the predominant oxidase is cytochrome aa3, the o-type oxidase is also present (Cross and Anthony, 1980b), and its properties observed after partial purification suggest that it is the same as that induced during growth under conditions of methanol excess (Carver and Jones, 1983). Whether or not the o-type oxidase continues to function in the same way in the presence of cytochrome aa3 is at present uncertain (Dawson and Jones, 1981c; Carver and Jones, 1983, 1984). In summary, the work presented above, together with that described by Cross and Anthony (1980a, b), Jones et al. (1982) and Beardmore-Gray et al. (1983), suggests that electron transport in M. methylotrophus, when grown under conditions of O2 limitation, is as depicted in Fig. 19. Because cytochrome C, is oxidized at 50 times the rate of soluble cytochrome cL, which is the specific electron acceptor from MDH, the electron transport chain between methanol and O2in whole organisms is likely to include both of the soluble cytochromes c. The demonstration that about half of the cytochrome c bound tightly to membranes is cytochrome cL, the rest being the
BACTERIAL OXIDATION OF METHANE AND METHANOL
cytopkn H++NADH
Fwipksm
197
NAD+
Methanol -Methanol
FIG. 19. The electron transport chain of Methylophilus methylotrophus grown under conditions of O2 limitation. After Froud and Anthony (1984a, b). This electron transport chain also operates under ammonia limited conditions and, to some extent, in batch culture and in continuous culture under conditions of methanol limitation. Solid lines indicate reactions that have been demonstrated in these bacteria, the relative thickness indicating the probable relative importance of the routes for oxidation of the soluble cytochromes c. Dotted lines are alternative routes for oxidation of cytochrome b which have not yet been investigated. About 40% of the cytochrome cLis firmly attached to the membrane and this may form a cytochrome bcLcomplex analogous to the cytochrome bc, complex in mitochondria and in P . denitrificans. It should be noted that the MDH and soluble cytochromes c are shown having a periplasmic location; this does not preclude the possibility that they are loosely bound to the periplasmic surface of the bacterial membrane. TMPD can probably donate electrons directly to all the c-type cytochromes and not only to the cytochrome c component of the oxidase as drawn here for convenience. N.B. The “0-type” oxidase contains 6- and c-type cytochromes; an alternative name for it is cytochrome co (see the text).
cytochrome c component of the oxidase, suggests that it might have a role analogous to that of cytochrome c, in mediating between the b-type cytochromes and the soluble cytochrome c during the oxidation of NADH (Froud and Anthony, 1984b). It will clearly be of interest to determine the roles of cytochrome cH and cytochrome c, (membrane-bound and soluble) as potential electron acceptors from cytochrome b during NADH oxidation, and as potential electron donors to the alternative cytochrome aa, when this oxidase is induced during growth under methanol-limited conditions. 3. Proton Translocation, the Proton Motive Force and Phosphorylation Potential in Methylophilus methylotrophus
By analogy with other proton-translocating electron transport chains, the involvement of cytochrome c in electron transport in M . methylotrophus (Figs. 18 and 19) indicates that up to three proton-translocating segments might operate during the oxidation of NADH, and only one during the oxidation of methanol. This has been confirmed in an extensive investigation
198
C.
ANTHONY
of the bioenergetics of M. methyfotrophusby Dawson and Jones (1981b, c, 1982). In this study the bacteria were grown under conditions of methanol limitation, and so had the electron transport chain shown in Fig. 18, in which cytochrome uu, is the predominant oxidase. The stoicheiometries of proton translocation during the oxidation of methanol and NADH (produced from formate) were determined by using both the O2 pulse and the initial-rate methods. This latter method was also used to measure K+/O ratios, in order to determine the charge/O ratios. It was concluded that 6H+/O are translocated during NADH oxidation, and that 2H+/ O are translocated during the oxidation of methanol to formaldehyde. There was no evidence for underestimation of the H + / O ratios due to H+/anion symport, except by the movement of formic acid during formate oxidation (as also shown in Pseudomonus AM 1 by OKeeffe and Anthony (1978)). By comparing their results with the known growth efficiences of this organism, a H+/ATP ratio of close to 2g-ions of H + (mol of ATP)-' was calculated. It was thus proposed that the respiratory chain of M. methyfotrophusis arranged such that there are three sites of energy conservation for NADH oxidation, each translocating 2H+ and each linked to the synthesis of one molecule of ATP, and that only the third site of energy conservation is involved in methanol oxidation. In order to investigate further this third site of energy conservation, the artificial substrate TMPD, which donates electrons to cytochrome c, was used, and appropriate concentrations of KCN were added to selectively inhibit cytochrome o (Dawson and Jones, 1981~).The kinetics of inhibition confirmed that the respiratory chain of this organism is branched at the level of cytochrome c to two major oxidases, cytochrome uu, and the o-type cytochrome oxidase. These kinetics also confirmed previous conclusions (Cross and Anthony, 1980b) that neither of the CO-binding cytochromes c is a physiologically significant oxidase. During the oxidation of TMPD the terminal oxidase of M. methylotrophus exhibits a net inward translocation of 2e-, but no net proton translocation when a pair of electrons are passed from cytochrome c to 02;and the overall translocation stoicheiometries are achieved by the two oxidases functioning similarly. These results demonstrate that neither of the terminal oxidases in M. methyfotrophus acts as a proton pump. For ATP synthesis to occur the electron transport chain described above must produce a proton motive force, the measurable component of which is the AjiH+ (the bulk-phase transmembrane electrochemical potential difference of protons). This is composed of ApH plus the AY, these being the bulk-phase transmembrane pH difference and the bulk-phase transmembrane potential difference, respectively. A measure of the work that can be done in the cell by ATP is the phosphorylation potential AGp. The relationship between this phosphorylation potential and the two components of the
BACTERIAL OXIDATION OF METHANE AND MFTHANOL
199
proton motive force at various pH values has been measured by Dawson and Jones (1982) during the oxidation of methanol by M. methylotrophus. They concluded that this organism is able to sustain a AFH’ of up to - 165mV during respiration, with methanol as substrate. This was composed of a AY and ApH, the values for which depended on the external pH value, ApH being maximal at low external pH values and decreasing to zero at an external pH of 7.0, which is the same as the internal pH value of the bacteria, a value that is rather lower than is usual for neutrophiles. Either the AY or the ApH values alone was shown to be fully competent to drive ATP synthesis in M. methylotrophus, and a AGp of up to -45.8 kJmol-’ was sustained during oxidation of methanol at pH 7.0. F. THE COUPLING OF METHANOL OXIDATION TO SYNTHESIS OF ADENOSINE TRIPHOSPHATE
All of the work on electron transport and proton translocation, described for the four different types of methylotroph and summarized in the previous section, has led to the conclusion that methanol is oxidized to formaldehyde by way of the MDH, cytochrome c and a terminal oxidase which is either cytochrome aa, or an o-type oxidase. This oxidation is coupled to an acidification of the external suspending medium (2H+ for each mole of methanol oxidized to formaldehyde) and production of a proton motive force, which is coupled to synthesis of one molecule of ATP. These conclusions raise two questions. (1) How are the components of the “methanol oxidase” system arranged with respect to the membrane in order to produce the necessary proton motive force? (2) Why do bacteria have this system; why is the “end product” of methanol oxidation only one molecule of ATP? (1) The two obvious options for arrangement of the “methanol oxidase” system are shown in Fig. 20. The first is a classical proton-translocating loop. The key point in such an arrangement is that methanol in the cytoplasm reacts with the MDH which must be integrated with the membrane and actually move protons across it. The MDH and its prosthetic group are, however, very hydrophilic and it is therefore unlikely to span a lipoprotein membrane. An unlikely variation of this scheme, possible only if MDH and cytochrome c do not react directly, would be to have an intermediary hydrogen carrier between MDH on the inside and cytochrome c on the outer side of the membrane. The second option (Fig. 20) is one which we first proposed for the MDHcytochrome c interaction in Pseudomonm AM1 (O’Keeffe and Anthony, 1978). This is similar to the first option in having cytochrome c on the outer side of the membrane, but in this case protons do not actually move across the membrane. They are released from the PQQH, prosthetic group of the
C. ANTHONY
200 OUTSIDE
MEMBRANE
(a) Proton motive redox"loop"
Fmldehyde 2 Hi -Cyt
c
/
\
2 H++ $0,
or cyt 0 ' p H 2 0
(b) Proton mo$e redox "arm
Methonol MDHiPQQ) Formaldehyde
1
2 H++ i 0 2
' cyt 1 003 or
cyt h
0
H
2
0
FIG. 20. Possible arrangements of methanol dehydrogenase and cytochrome c in the bacterial membrane. From Anthony (1982). In both schemes 2 Hf are effectively moved from the inside to the outside for every molecule of methanol oxidized to formaldehyde. In (a) the MDH is transmembranous, and in (b) it is on the outer surface.
dehydrogenase when it reacts with cytochrome c on the outer side of the membrane. Indirect, but compelling, evidence based on measurements of proton translocation (see above) all supports the second of these options (Fig. 20; O'Keeffe and Anthony, 1978; Dawson and Jones, 1981a, b, c, 1982; van Verseveld et al., 1981). More direct evidence has come from measurements of the periplasmic proteins of P. denitrgcans (Alefounder and Ferguson, 198l), M . methylotrophus (Jones et al., 1982; Burton et al., 1983; Quilter and Jones, 1984) and organism 4025 (Lawton and Anthony, 1985). In all cases, a high proportion of the MDH and cytochrome c was released into the growth medium when sphaeroplasts were made by lysozyme treatment, thus demonstrating that these proteins are located predominantly in the periplasmic space, or loosely bound to the outer surface of the cytoplasmic membrane.
BACTERIAL OXIDATION OF METHANE AND METHANOL
201
Although it has not been possible to do such experiments with Pseudomonas AM 1, which is resistant to lysozyme treatment, it is likely that the MDH and cytochrome cL are similarly located, because of the acidification of the suspending medium occurring during the oxidation of ascorbate plus TMPD (OKeeffe and Anthony, 1978). (2) The second question is, why make only one ATP during the oxidation of methanol to formaldehyde? The mid-point redox potential for the formaldehyde/methanol couple is - 0.182 V, which is sufficiently low to support the synthesis of two ATP molecules for each molecule of methanol oxidized by 02. For some methylotrophs the answer is immediately obvious; in those methylotrophs that are NADH-limited, rather than ATP-limited (this includes most methanotrophs), increasing the P/O ratio for methanol oxidation would have a negligible effect on cell yields (Anthony, 1978, 1980, 1982,1983).In these bacteria, however, if the MDH were to become NADHyielding, then the cell yields would be much higher, even if the P/O ratio were to remain low. In those methanol-utilizers such as M.methylotrophus having the RUMP assimilation pathway, higher yields would be obtained if the P/O ratio for “methanol oxidase” were to be increased, or if the MDH were to be NADHyielding. The extent of the expected increases can be estimated from Fig. 21. These data have been calculated using methods previously described (Anthony, 1978, 1982, 1983). The formula for calculation of yields is: 12CH,OH
+ 1.5NADH + 3NH3 + xATP-
(C,H,O,N),
+ IZPQQH,
To use this formula for predicting yields, a value for the ATP requirement for biosynthesis (x) must be chosen and values for the P/O ratios assumed for oxidation of NADH and PQQH, (reduced MDH). The value for x can be calculated, or the measured yield (or Carbon Conversion Efficiency) can be used for estimating a value for x for a given organism from curve A in Figs 21 and 22. Using this same value for x, the effect of changing the nature of the system for methanol oxidation can be determined by using the alternative curves in these Figures. It can be seen that the greatest potential increase in yield would be achieved by changing the MDH to an NAD+-linked dehydrogenase. It should be borne in mind that the very high yield that is predicted in this case might not be obtainable in practice on thermodynamic grounds. Such a modified organism might only be able to exist if it also “adapted itself’ by lowering the P/O ratio for NADH oxidation from 3 to 2. Even if this did occur, then the predicted yield would still be higher than in the organism having the normal MDH. (For further discussion of this problem see Carver and Jones (198Ic)).
202
C . ANTHONY
‘E
NAD
-
3
;\B 35 40 45 50 55 A a M l e d ATP requiremat (I1 for cell rynlhe*s (306p)
mo
1
z
5
, , , ,
[, 30
FIG. 21. The variation of carbon conversion efficiency with ATP requirement. From Anthony (1983). Curve A is the usual situation.
‘r
MDH NAD
mo
P/oro+ios NAN
mot+ 2
-
NAD w p 1 m 2 m 1
3
3
2 3
2
2
Aswmed ATP require^ ( x ) for cell synthesis (306g)
FIG. 22. The variation of yield on O2(Yo,) with ATP requirement for growth. From Anthony (1983). Curve A is the usual situation.
In summary, bacteria growing on methanol by way of the RUMP pathway are some of the most efficient organisms known (in terms of carbon conversion efficiency), and the “altered” organisms discussed here would be even
BACTERIAL OXIDATION OF METHANE AND METHANOL
203
more efficient. The free energy change during such efficient growth would tend to be relatively small, and it must be concluded that rapid growth of such altered organisms might not be possible on thermodynamic grounds.
V. Acknowledgements I should like to thank the SERC for supporting my work described in this review, and Dr. M. Beardmore-Gray for valuable discussions during its preparation. REFERENCES Adachi, O., Shinagawa, E., Matsushita, K. and Ameyama, M. (1982). Agricultural Biological Chemistry 46, 2859. Alefounder, P.R. and Ferguson, S.J. (1981). Biochemical and Biophysical Research Communications98, 778. Ameyama, M., Matsushita, K., Ohno, Y.,Shinagawa, E. and Adachi, 0. (1981a). FEES Letters 130, 179. Ameyama, M., Shinagawa, E., Matsushita, K. and Adachi, 0. (1981b). Agricultural and Biological Chemistry 45, 851. Ameyama, M., Hayashi, M., Matsushita, K., Shinagawa, E. and Adachi, 0. (1984a). Agricultural and Biological Chemistry 48, 561. Ameyama, M., Shinagawa, E., Matsushita, K. and Adachi, 0.(1984b). Agricultural and Biological Chemistry 48, 2909. Ameyama, M., Shinagawa, E., Matsushita, K. and Adachi, 0. (1984~). Agricultural and Biological Chemistry 48, 3099. Anthony, C. (1963). Ph.D. Thesis, University of Reading. Anthony, C. (1975). Biochemical Journal 146, 289. Anthony, C. (1978). Journal of General Microbiology 104, 91. Anthony, C. (1980). In “Hydrocarbons in Biotechnology” (D.E.F. Harrison, I.J. Higgins and R. Watkinson, eds), pp. 35-37. Heyden and Sons, London. Anthony, C. (1982). “The Biochemistry of Methylotrophs.” Academic Press, London. Anthony, C. (1983). Acta Biotechnologica. 3, 261. Anthony, C. and Zatman, L.J. (1964a). Biochemical Journal 92, 609. Anthony, C. and Zatman, L.J. (1964b). Biochemical Journal 92, 614. Anthony, C. and Zatman, L.J. (1965). Biochemical Journal %, 808. Anthony, C. and Zatman, L.J. (1967a). Biochemical Journal 104, 953. Anthony, C. and Zatman, L.J. (1967b). Biochemical Journal 104,960. Babel, W. and Steudel, A, (1977). Zeitschrifr fur Allgemeine Mikrobiologie (Berlin) 17, 267. Bamforth, C.W. and Quayle, J.R. (1978a). Archives of Microbiology 119, 91. Bamforth, C.W. and Quayle, J.R. (1978b). Biochemical Journal 169,677. Bamforth, C.W. and Quayle, J.R. (1979). Biochemical Journal 181, 517. Beardmore-Gray, M. (1982). Ph.D. Thesis, University of Southampton. Beardmore-Gray, M. and Anthony, C. (1983). Journal of General Microbiology 129,2979. Beardmore-Gray, M. and Anthony, C. (1984). In “Microbial Growth on C, Compounds” (R.L. Crawford and R.S. Hanson, eds), pp. 97-105. American Society for Microbiology, Washington.
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Occurrence, Biochemistry and Physiology of Phenazine Pigment Production JOHN M. TURNER Department of Biochemistry, University of Liverpool, Liverpool L69 3BX, UK
and
ANN. J. MESSENGER Department of Biochemistry, University of Hull, Hull HU6 7RX, UK
I. Introduction .
.
11. Natural Occurrence and some properties of phenazines .
.
A. Pseudomonads . . B. A bacterium of unknown identity. C. Actinomycetes other than Sfreptomyces species . . . . D. Streptomycetes . . E. Sorangium (= Polyangium) species . . . 111. Biosynthesis . A. Phenazineorigins . B. Phenazine metabolism . IV. Secondary metabolism and the physiological significance of phenazine production A. Phenazines as secondary metabolites . . . . B. Possible physiological functions of phenazines . V. Acknowledgements. . References .
21 1 218 218 232 232 235 24 1 242 243 249 260 260 265 268 268
I. Introduction Naturally occurring phenazines are pigments formed exclusively by bacteria. Those known longest are pyocyanine (Fordos, 1859), chlororaphine (Guignard and Sauvageau, 1894) and iodinin (Clem0 and McIlwain, 1938), coloured blue, green and purple, respectively. ADVANCES IN MICROBIAL PHYSIOLOGY, VOL. 27 ISBN g12-027727-1
Copyright 0 I986 by Acodemic Press London AN rights of reproduction in any form reserved
212
JOHN M. TURNER AND ANN J. M. MESSENGER
0
0-
OH
HO 0 Iodinin
Pyocyanine
-
CO NH?
H Chlororaphii
Well over 50 phenazines of bacterial origin are now known. A summary of their structural features, nomenclature and origin is given in Table 1. These phenazines, only a few of which have been given trivial names (Table 2), represent every colour of the visible spectrum. The absorption spectra of phenazines are characteristic, with an intense peak in the range 250 to 290 nm and-aweaker peak at 350 to 400nm (Gerber, 1973). At least one main band occurs in the visible region (400 to 600 nm) to which the phenazines owe their colours (Britton, 1983). Unlike the various carotenoid and other pigments that colour many bacteria, growing as yellow, orange or red colonies on nutrient agar, the phenazine pigments are mostly water soluble and are excreted into the medium. Thus, pyocyanine produced by Pseudomonas aeruginosa, diffuses readily into agar-solidified media which become stained blue. Some phenazines are only sparingly water soluble and precipitate. Chlororaphine, a mixture of phenazine-1-carboxamide (oxychlororaphine) and its dihydro derivative, produced by .Pseudomonas chlororaphis, accumulates as isolated emerald-green crystals at the base of agar slants. Iodinin crystallizes on the surfaces of old colonies of Brevibacterium iodinum, giving them a dark-purple appearance, and phenazine- 1-carboxylic acid is deposited as golden yellow crystals in colonies of Pseudomonas aureofaciens and in the surrounding medium (Kluyver, 1956; Haynes et a/., 1956). Production of phenazines is of obvious taxonomic value, particularly when relatively few genera are concerned (Table 1). It should be noted, however, that the same pigment may be produced by unrelated bacteria and “achromogenic” strains of many phenazine-producers are common. Considerableprogress has been made in elucidating the biosynthetic routes to individual phenazines during the last 15 years, although only about half-a-dozen bacteria have been used in such work. No biosyntheticstudy has been made of the majority of compounds listed in Table 1.
0
X"
z
X 0
X 0
0
x
0
0
x-sx x U.u.0 0
0 0
0
X C ou.8 0
0
2" c!
0
0
x
0
0
xx 00
xx
00
xx
00
u.Xn
0
g
TABLE 1. (continued)
Name
1
2
3
4
Positions of suhstituents 5 6
Bacterial 7
8
9
10
sourceb
B. Plmwhea witb m cubon eomtitnmt chektemqckrirg3J92em
Phenazine-l-carboxylic acid Phenazine-lcarboxylic acid methyl ester Phenazine-l-carboxamide 5-Methyl-Famino-I carboxyphenaziniwn betaine 5-Methyl-7-amino-I carboxy-3sulphophenazinium betaine 2-Hydroxyphenazine-1carboxylic acid Di-(Zhydroxy- 1-phenazinyltmethanec 4Hydroxyphenazioe-l-carboxylic acid CHydroxyphenadne-1carboxylic acid CMethoxyphenazine- I carboxylic acid methyl ester 9-Hydroxyphenazine-l-carboxylic acid 2.3-Dihydroxyphenazine- I carboxylic acid 2,CDihydroxyphenazine-1carboxylic acid 2.9-Dihydroxyphcnazine-lcarboxylic acid 2,3,4Trihydroxyphenazine-1carboxylic acid
-
COOH
1, 2, 3, 5. 16, 19, 26 22 I, 2
CO0.CH3 co.NH, COOH
1 1
COOH COOH
3 3 3 3, 5 22
OH OH
COOH COOH COOCH3
OH 0.CH, OH
COOH COOH
OH
COOH
OH
COOH
OH
COOH
OH
OH OH
3
OH OH
5. 6
3
OH
5, 6
3
c. pbeludees with two arbon sllbstitllmts ~*betemeydiemsystem Phenazine-l ,E-dicarboxylic acid
6-Hydroxymethylphenazine-1carboxylic acid a(I-H ydroxyethy1)-phenazine-I carboxylic acid methyl ester 6-( I-Melhoxynhyl)-pbenazizine-I carboxylic acid methyl ester
COOH COOH
COOH CH20H
COO.CH,
CH(OH)CH,
28
CO0.CH3
CH(OCH3)CH3
28
3, 4, 5, 6 17
(343
I
a(3-Methyl-2-butenyl)-I -phenadne carboxylic acid
COOH
6-~l-(2-Hydroxy-6-methyl)benzoylox~lethylphenazine-lCarboxylic acid
COOH
2,3-Di-( 1-methoxycarbonyl-6phenaziny1)butane 2-Hydroxyphenazine-l,6dicarboxylicacid 4-Hydroxyphenazim-l,6dicarboxylicacid dimethyl ester 6-Hydroxymethyl-9-hydroxyphenazinc-Icarboxylic acid 6Hydroxymethyl-9-melhoxyphenazine-1carboxylic acid
26
CH,CH: c
I
25.21
[COO.CH3
CH-CH31,
28
COOH COO .CH,
COOH COO.CH,
3
OH
OH
4
COOH
CH,OH
OH
23
COOH OH
CH,OH
0.CH3
17
CHzOH
0.CH,
24
6-Hydroxymethyl-9-methoxyphenazine1 Carboxylic acid 2.5dihydroxyqdopent-1me-I-amide dH