Advances in Microbial Physiology Volume 29

Advances in

MICROBIAL PHYSIOLOGY

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Advances in

MICROBIAL PHYSIOLOGY Edited by

A. H. ROSE School of Biological Sciences Bath University, UK

and

D. W. TEMPEST Department of Microbiology Uniuersity of Shefield, UK

Volume 29 1988

ACADEMIC PRESS Harcourt Brace Jovanouich. Publishers London San Diego New York Boston Sydney Tokyo Toronto

ACADEMIC PRESS LIMITED 24-28 Oval Road London NWI 7DX U . S . Edirion published by ACADEMIC PRESS INC San Diego 92101

Copyright $3 1988 by ACADEMIC PRESS LIMITED

AN Righis Reseroed

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

ISSN 0065-291 1

Printed in Great Britain at the Alden Press, Oxford

Contributors M. O’Brian Department of Biology, The Johns Hopkins University, Baltimore. M D 21218, USA G . A. Codd Department of Biological Sciences, University of Dundee. Dundee DDI 4HN, U K M. J. Danson Department of Biochemistry, University of Bath, Claverton Down. Bath BA2 7AY, U K W. Fischer Institut fur Biochemie, Universitat Erlangen-Nurnberg, Fahrstrasse 17, D-8520 Erlangen, FRG L. S. Frost Department of Biochemistry, University of Alberta, Edmonton, Alberta, Canada T6G 2H7 R. J. Maier Department of Biology, The Johns Hopkins University, Baltimore, M D 21218, USA W. Paranchych Department of Biochemistry, University of Alberta. Edmonton, Alberta, Canada T6G 2H7

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Contents Contributors

V

Hydrogen Metabolism in Rhizobium: Energetics, Regulation, Enzymology and Genetics M A R K R. O’BRIAN and ROBERT J. MAIER I. 11. 111. IV . V. VI.

Introduction Regulation Enzymology Energetics Genetics Acknowledgements References

2 6 13 24 38 47 47

The Physiology and Biochemistry of Pili WILLIAM PARANCHYCH and LAURA S. FROST I. 11. 111. IV. V. VI . VII .

Introduction Nomenclature Classification High-resolution studies on pilus structure Organization and expression of pilin genes Structure-function relationships of pili proteins Acknowledgements References

53 54

55 64 68 82 102 102

Carboxysomes and Ribulose Sisphosphate Carboxylase/Oxygenase G E O F F R E Y A. C O D D I. Introduction XI. Distribution and structure of carboxysomes

115 117

...

CONTENTS

Vlll

I ll . Carboxysome composition IV. Ribulose I ,5-bisphosphate carboxylase/oxygenase (RuBisCO) V. Carboxysome function VI. Further aspects of carboxysomes References

124 i32 149 155

157

Archaebacteria: The Comparative Enzymology of Their Central Metabolic Pathways MICHAEL J. DANSON

I.

Introduction

11. Archaebacterial pathways of central metabolism 111. Archaebacterial enzyme diversity

IV. Structure of archaebacterial enzymes V. Concluding remarks VI. Acknowledgements References

166 176 194 217 222 222 223

Physiology of Lipoteichoic Acids in Bacteria W . FISCHER

I.

Introduction

11. Occurrence and structure 111. Metabolism

IV. V. VI. VII.

Cellular location Biological activities Concluding remarks Acknowledgements References

Author index Subject index

233 235 247 275 277 295 296 296

303 327

Hydrogen Metabolism in Rhizobium: Energetics. Regulation. Enzymology and Genetics MARK R . O’BRIAN and ROBERT J . MAIER Department of Biology. The Johns Hopkins Uniuersity. Baltimore. M D 21218. USA

1. Introduction . . A . General background

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. . . . . B . Hydrogen evolution by nitrogenase . . C . Hydrogen oxidation by legume root nodules Regulation . . . . . . . A . Oxygenandcarbon . . . . B . Hydrogenase and carbon dioxide fixation C . Host control . . . . . Enzymology . . . . . . A . Purification and some properties . . B . The K,,, value for hydrogen . . . C. Electron acceptor reactivity . . . D . Oxygen lability . . . . . E . Nickel . . . . . . . F . Lipid requirement . . . . G . Kinetic mechanism of hydrogenase . . Energetics . . . . . . . . A . Physiological considerations . . . B . Electron transport . . . . . Genetics . . . . . . . . A . Mutants . . . . . . . B . Molecular genetics . . . . . Acknowledgements . . . . . . References . . . . . . . .

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ADVANCES I N MICROBIAL PHYSIOLOGY VOL ?Y ISBN 0-12-0?7729-8

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Copyright (3 1988 by Academic Press Limited All rights OF reproduclion in any Form reserved

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M. R. O'BRIAN AND R. J . MAIER

I. Introduction A. GENERAL BACKGROUND

The term hydrogenase refers to enzymes that catalyse Hz consumption or evolution according to the reaction: (1) H2 e2 H + 2eAll hydrogenases are bidirectional to some extent in vitro, but the enzyme appears to catalyse either H2 oxidation or production in uivo. Hydrogen evolution usually occurs in anaerobic micro-organisms, and serves to get rid of excess reductant when protons are the only available oxidant (Schlegel and Schneider, 1978), whereas H2 utilization can occur in aerobic and anaerobic bacteria, and is linked to ATP-producing electron transport systems. Anaerobic bacteria oxidize H2 using sulphate, sulphur, C 0 2 , nitrate or fumarate as the terminal electron acceptor, and the photosynthetic bacteria use H2 and other compounds, rather than H20, as the reductant for COz fixation (Knaff, 1978). Aerobic Hz-oxidizing bacteria can grow with H2 and COZ as the sole energy and carbon sources, respectively. Among these bacteria, some have a soluble and a membrane-bound hydrogenase; the soluble enzyme catalyses Hz-dependent NAD+ reduction which is used for COZfixation, whereas the membrane-bound hydrogenase is linked to electron transport, and is therefore involved in energy production (Adams et al., 1981). The aerobic N2-fixing bacteria evolve and consume HI, and among this group are the rhizobia, the azotobacter and the cyanobacteria. Hydrogen evolution by these micro-organisms is catalysed by nitrogenase, and an uptake hydrogenase is responsible for H2 oxidation. Whereas the cyanobacteria have hydrogenase activity even in cells not fixing NZ(Tel-Or et al., 1977; Eisbrenner et al., 1978), hydrogenase is generally derepressed under Nz-fixing conditions in Azotobacter and in hydrogen uptake positive (Hup+)strains of Rhizobium. Autotrophic growth of R. japonicurn on H2 has been demonstrated in the laboratory (Hanus et al., 1979), and thus H2 oxidation by this bacterium, without concomitant Nz fixation, may conceivably occur in nature under some conditions. +

B. HYDROGEN EVOLUTION BY NITROGENASE

Nitrogenase catalyses the reduction of dinitrogen to ammonia according to the reaction: Nz+ 8H+ +8e-

+ 16ATP-+2NH3+16ADP+ 16Pi+ Hz

(2) As eqn. (2) shows, the physiological substrates include H + as well as N2, and

HYDROGEN METABOLISM IN RHIZOBIUM

3

H2 production is apparently obligatory to the N2 fixation reaction. What this equation does not indicate is that the ratio of Nz reduced to H2 evolved is not constant, and that this ratio can be altered in iiirro, and is variable under physiological conditions as well. Rivera-Ortiz and Burris ( 1975) demonstrated that H2 evolution by partially purified nitrogenase decreases as the N2 concentration increases. Hageman and Burris (1980) showed that the allocation of electrons to NZor H + by purified nitrogenase is dependent on the rate of electron flux through the enzyme, which in turn depends on the ATP concentration, the ratio of component I to component 11, and the concentration of reductant. In those experiments, a high electron flux favoured N2 reduction, whereas H2 production was favoured by a low flux of electrons through nitrogenase. Schubert and Evans (1976) found that only 40-60% of the electron flow to nitrogenase participates in NZreduction in various legume root nodules, and the remainder is lost through HZ evolution. These data indicate that the ratio of N2 fixed to HZproduced by nitrogenase is variable in nature, and is not merely an experimental phenomenon. Although H2 evolution by nitrogenase can be decreased by several experimental manipulations, it cannot be eliminated, and H2 production is apparently an obligatory product of biological NZ fixation. When H2 evolution by nitrogenase is plotted as a function of N2 concentration and extrapolated to infinite pNz, H2 evolution is found to occur at 13-23% of its maximal rate, implying that H2 production cannot be eliminated (RiveraOrtiz and Burris, 1975). This conclusion was confirmed by Simpson and Burris (1984), who showed that HZevolution by nitrogenase occurs at a pNz of 50 atmospheres (5.07 MPa). In those experiments, 27% of the total electron flux through nitrogenase was allocated to HZproduction, and the ratio of NZ fixed to H2 produced was about 1: 1. Thus the stoicheiometry represented in eqn. (2) is apparently an upper limit with respect to the amount of N2 that can be fixed per H2 evolved. It is not known for certain why H2 is produced during N2 fixation or why the amount produced is dependent on the rate of electron flux, but Chatt (1980) proposed a mechanism for Nz reduction by nitrogenase that could explain some of the observations discussed above. In his scheme, molybdenum is assumed to be the active site of the enzyme, and present in the trihydridic state. It is known that NZforms complexes only with transition metals in a strongly reduced state, and thus if the rate of electron flow is slow, the electrons are discharged by protons to form HZbefore the molybdenum moiety can become sufficiently reduced to bind Nz. When the electron flow to molybdenum is faster than the discharge rate to protons, the metal is capable of binding NZ and reducing it. Molybdenum in the highly reduced state can react with N2 and H + , but most of the protons will be discharged by interaction with the N2 bound to molybdenum, which they reach before they can interact with the

4

M . R . O’BRIAN AND R . J . MAIEK

metal. Under maximal N2-fixingconditions, one N2 molecule would have to displace two of the hydrides in order to bind to molybdenum, resulting in the formation of one H2 molecule. When Nz is completely reduced, it is removed to give two NH3 molecules, and two protons then bind to the metal to restore the trihydritic state. The formation of the hydride ions occurs either because there is no N? at the active site, or because it is a necessary condition for the release of NH3. This scheme can explain some features of nitrogenase enzymology, but it should be kept in mind that some of the assumptions on which the hypothesis is built have yet to be demonstrated. It is not known for certain that N? reacts with molybdenum moiety, nor is it known that the metal binds with three hydride ions in its most reduced state. It is also assumed that the molybdenum active site is in a pocket or cleft, and has limited access to protons and other substrates.

C. HYDROGEN OXIDATION BY LEGUME ROOT NODULES

Phelps and Wilson (1941) discovered that pea nodule bacteroids have hydrogen uptake activity, but cultures of R. leguminosarum do not. These findings could not be confirmed (Shug et al.. 1956), however, and the rediscovery of the uptake hydrogenase in pea nodules did not occur until 1967 (Dixon, 1967). Dixon (1972) concluded that the presence of an uptake hydrogenase increases the overall efficiency of the N2 fixation process in three ways: (i) it prevents the inhibition of nitrogenase by hydrogen gas; (ii) it consumes 0 2 and thereby protects nitrogenase from 0 2 inactivation; and (iii) it produces ATP by Hz-dependent oxidative phosphorylation. The latter two functions of hydrogenase have been demonstrated in R . japonicum (Emerich et al., 1979), some Hup+ strains of R . leguminosarum (Nelson and Salminen, 1982), and also in Azotohucter and the cyanobacteria (Adams rt a/., 1981). Hydrogen is known to inhibit nitrogenase, but it is not clear whether the intracellular H2 concentration is high enough to be inhibitory in uiuo (Robson and Postgate, 1980; Dixon et al., 1981). Schubert and Evans (1976) surveyed the magnitude of H2 evolution in leguminous and non-leguminous nodules, and found that, among the legumes, only nodules from Rhizohium spp. strain 32H1 in association with Vigna sinensis evolved very little HI. Schubert et a / . (1977) described a H:! uptake hydrogenase in a strain of R . japonicum and of R. “cowpea”, and Carter et al. (1978) reported that six out of thirty-two R . japonicum strains examined evolved little or no HZ as nodules, which was attributed to an uptake h ydrogenase. “Relative efficiency” is a parameter often used to assess the efficiency of Nz

HYDROGEN METABOLISM IN HHIZOBIC’M

5

fixation by measuring HI evolution by nodules (Schubert and Evans, 1976). I t is defined as:

-[

Rate of Hz evolution in air Rate of CzH2reductionp]

(3)

It can be seen that a low rate of H? evolution compared to C2H2 reduction will yield a high relative efficiency. In strains that lack an uptake hydrogenase (Hup-), the relative efficiency is an indication of the fraction of electrons allocated to Nz reduction, and thus the term “relative efficiency” has literal meaning. However, in strains containing hydrogenase, relative efficiency measurements are only an indication of how much of the Hz evolved by nitrogenase is consumed by hydrogenase; this parameter, in itself, does not say anything about the efficiency of N ? fixation. A relative efficiency of 1 does not mean the total electron flux through nitrogenase is directed toward NI reduction, but rather it means that all of the H? produced by nitrogenase is oxidized by hydrogenase. In this case the relative efficiency, as defined in eqn. (3), is the same regardless of whether hydrogenase actually affects the efficiency of Nz fixation. Albrecht et al. (1979) compared the effects of Hup+ and Hup- R..japonicurn strains on soybean plants grown in a greenhouse. They found that plants inoculated with Hup+ strains contained 16% more dry weight, 10% more N per total dry weight and 26% more total N. In similar experiments comparing a Hup- mutant and a Hup+ revertant, Evans et a/. (1983) found that plants inoculated with the Hup+ strain showed increases in weights of nodules, shoots, total plant material and in total N per culture. These results indicate that the Hup+ trait is beneficial to the Nz fixation process in the R .japonicirmsoybean symbiosis. However, the benefits of H7 oxidation in the R. leguminosarum-pea symbiosis are tenuous at this time, and these data are discussed later in this article. The remainder of this review will deal with selected aspects of H2 metabolism in Rhizohium. particularly R. ,japonicum and R . leguminasartmi. Although Hz metabolism and N2fixation in organisms other than the rhizobia will be discussed, it will be for comparative purposes, and the data presented pertaining to those organisms is not intended to be comprehensive. The enzymology of hydrogenase will concentrate on that of R.,japonicum since the R. leguminosarum enzyme has yet to be purified and characterized. Until very recently, different approaches have been taken in studying genetic problems concerning Hz metabolism in R. ,japonicum and R. lcguminosarum, and thus they are discussed separately.

6

M . R . O'BRIAN AND R. J. MAIER

11. Regulation

A. OXYGEN AND CARBON

Hydrogen uptake (Hup) by nodules or bacteroids of Hup+ R . japonicurn strains can be easily demonstrated, but heterotrophically-grown cells do not normally oxidize H2. The bacteroid environment is very different from that of a cultured cell, and thus there are many external factors which can potentially regulate the expression of hydrogenase activity. These factors include 0 2 , H2, carbon sources accessible to the cell, and trace elements, as well as the more elusive contributions from the plant host. Maier et al. (1978a) demonstrated that H2 uptake activity can occur in freeliving cells under reduced 0 2 and organic carbon concentrations, and in the presence of Hz. The decreased O2 tension and increased H2 concentration necessary for the derepression of hydrogenase activity may imitate the bacteroid environment, at least qualitatively, but the carbon requirement is harder to evaluate. The low carbon concentration required for good Hup activity may also be characteristic of the nodule milieu since photosynthate is believed to be limiting in the nodule (Hardy et al., 1978), but the carbon sources used by nodule bacteroids are not known with certainty, and thus it is difficult to be confident in such speculation. It is also possible that the phenotype of hydrogenase-derepressed cells is similar to that of free-living aerobic hydrogen bacteria, and that the derepression conditions do not actually simulate a nodule environment at all. The ability of R. japonicum to grow autotrophically (Hanus et al., 1979) and express ribulose 1,5-bisphosphate (RuBP) carboxylase (Simpson et al., 1979)lends credence to this notion. It was later found that hydrogenase activity can be derepressed in the complete absence of organic carbon, and is enhanced by the addition of C02 (Maier et al., 1979). Numerous carbon substrates repress hydrogenase expression, and this repression is apparently at the level of hydrogenase synthesis (Maier et al., 1979). The addition of arabinose or gluconate to derepressing cells causes the H2 uptake rates of the cells to level off, whereas the activity of these cell continues to increase in the absence of carbon. The same phenomenon is observed when O2is added to, or H2 is removed from, the derepressing cells. Since the activity of hydrogenase-derepressed cells is not inhibited by the addition of O2 or organic carbon, these substances act by repressing hydrogenase synthesis, and do not inhibit the hydrogenase enzyme. Several types of mutants of R .japonicum have been obtained that are either hypersensitive (Maier and Merberg, 1982) or insensitive (Merberg and Maier, 1983; Merberg et al., 1983) to repression by 0 2 . The 02-hypersensitive mutants were initially characterized by their ability to express Hup activity as

HYDROGEN METABOLISM IN RHlZOBlUM

7

bacteroids, but not under normal H2-derepressing conditions in culture. However, five out of seven such mutants do express Hup activity when the 0 2 tension is lowered from 2% to 0.4%. The kinetics of hydrogenase repression by 2% O2 in these mutants is similar to the repression of activity in the wild type by 20% 0 2 . The addition of 2% 0 2 to the mutants does not cause an inhibition of H2-derepressed cells, but rather it results in the cessation of derepression. This observation means that the mutants do not make a more 02-labile H2 uptake system, nor is the 0 2 sensitivity due to a general toxicity effect. Since the mutants grow heterotrophically, the mutation seems to be specific to H2 metabolism. The second class of R.japonicum mutants are insensitive to 0 2 repression of hydrogenase activity. The mutants were isolated by selecting for cells that can grow chemoautotrophically under 10% 0 2 (Merberg and Maier, 1983). The wild type cannot grow autotrophically under such high 0 2 tensions, nor can it be derepressed for Hup activity at 10% 0 2 . These 02-tolerant mutants have significantly greater 0 2 - and methylene blue-dependent H2 uptake activity as bacteroids compared with the wild type, suggesting that common factors regulate H2 oxidation in free-living cells and bacteroids. Further analyses show that these mutants are extremely interesting in several respects. Whereas the hydrogenase activity of the wild type is repressed by organic carbon, the 02-insensitive strains are considerably less sensitive to carbon repression (Merberg et al., 1983). The addition of arabinose or succinate to derepressing cells at concentrations that repress the wild type 90% or more only inhibit hydrogenase expression 30-50% in the 02-insensitive strains. These data strongly suggest that there is a common regulatory element involved in the control of hydrogenase expression by 0 2 and carbon. This assertion is supported by the observation that mutants hypersensitive to hydrogenase repression by 0 2 are also hypersensitive with respect to carbon repression (Merberg et al., 1983). The 02-tolerant strains also derepress hydrogenase in the absence of added H2 (Merberg et al., 1983), which is required for hydrogenase expression in the wild type (Maier et al., 1979). Since these mutants express hydrogenase activity in heterotrophically-grown cells, and need not be induced to express hydrogenase, they are referred to as hydrogenase-constitutive (Hup") mutants. The nature of the mutations causing the Hupc phenotype is not known, but it is apparently not due to an increase in intracellular cyclic AMP (adenosine 3',5'phosphate) concentration (Merberg et al., 1983), which has been shown to coincide with hydrogenase expression in R. japonicum (Lim and Shanmugam, 1979). It is interesting to note that all R. japonicum HupC mutants isolated thus far produce significantly more cytochrome o than does the wild type (O'Brian and Maier, 1985a). Like hydrogenase, cytochrome o is synthesized under low 0 2 conditions in many bacteria (Poole, 1983),

8

M . K. O'RRIAN A N D R . J. MAIER

suggesting that the regulatory gene altered in the Hup'mutants also affects cell systems not directly related to hydrogenase. Mutants similar to the R . juponicum Hup'strains have also been obtained in a strain of Alculigenes eutrophus (Cangelosi and Wheelis. 1984). Alculigenes eutrophus strain 17707 does not grow chemoautotrophically under 20%)0 2 . but it will grow at 4% O2and synthesize both hydrogenases. Mutants relieved of 0 2 repression (oxygen sensitivity negative, or Osec) can grow chemoautotrophically, and have soluble and membrane-bound hydrogenase activities. The Ose- mutation mobilizes with a self-transmissible plasmid that carries genes necessary for hydrogenase expression. Ose- strains can also be obtained from mutants with plasmid-borne lesions which result in the loss of soluble hydrogenase activity, showing that the Ose- phenotype is independent of soluble hydrogenase activity. Chromosomal lesions resulting in diminished soluble and membrane-bound hydrogenase activities cannot be made Ose- . These data suggest that the Ose trait may act only on the membrane-bound hydrogenase, but the inability to obtain mutants deficient only in the particulate hydrogenase makes this speculation difficult to prove. Like the Hup' R . ,juponirum mutants (Merberg et ul., 1983). the Ose- A . eutrophus mutants are relieved of hydrogenase repression by organic carbon substrates (Cangelosi and Wheelis, 1984). Since H? is known to inhibit heterotrophic growth of A . eurruphus if hydrogenase is synthesized (Schink and Schlegel, 1978; Schlesier and Freidrich, 1982), Cangelosi and Wheelis (1984) proposed that the Ose system serves to minimize this phenomenon (called the hydrogen effect). They also suggest that Hz oxidation by A . eutrophus probably occurs in a mixotrophic, rather than an autotrophic, context in nature, and the down regulation of hydrogenase activity maximizes mixotrophic growth. This idea is supported by the observation that heterotrophic growth on glycerol is inhibited by HI in the Ose- mutants, but not in the wild-type strain (Cangelosi and Wheelis, 1984). This hypothesis is reasonable from the standpoint that Hz oxidation and mixotropic growth may not be advantageous in the presence of adequate organic carbon sources, and thus hydrogenase synthesis would be repressed by organic carbon. The repression of hydrogenase by 02,however, is less obvious in terms of mixotrophic growth since there seems to be no reason to assume that a low 0 2 concentration accompanies mixotrophy in nature. However, an energy deficiency could occur under low 0 2 conditions, and thus a substrate that yields the most ATP per 02consumedwould be beneficial to the cell. Bongers (1967) found that the P/O ratio with Hz as substrate is substantially higher than with succinate or fihydroxybutyrate as substrates in A . eufrophus.This observation indicates that Hz could indeed be a good energy source when 0 2 is limiting, and would explain why hydrogenase expression is derepressed under low 01tensions. It is not known whether 0 2 and carbon regulation of hydrogenase

HYUROGI
9

expression in R .juponicum has the same advantages as it does in A . cutrophus since the hydrogen effect has not been demonstrated in R.,juponicutii. It seems that 0 2 regulation of hydrogenase in R .juponicum ensures that hydrogenase is expressed under N?-fixing conditions, when the 0, tension is low and molecular hydrogen is available to the bacterium. Since photosynthate seems to be limiting in nodules, hydrogenase expression is advantageous symbiotically, and would explain the regulation by organic carbon. Rhizohium ,juponicumcan grow autotrophically (Hanus et d., 1979), however, and thus it cannot be ruled out that this growth mode occurs in the soil under free-living conditions. If R . juponicum does indeed grow autotrophically in nature, then 02 and carbon regulation of hydrogenase may serve the same purpose as it does in A . eutrophus. The proposed reasons for hydrogenase regulation by 0 2 and organic carbon in symbiotic and free-living cells are not mutually exclusive. From an evolutionary perspective, a Hz-oxidizing soil bacterium possesses traits that are advantageous in a Nz-fixing, symbiotic environment, including the regulation of metabolism by 0 2 and organic carbon. B. HYDROGENASE AND CARBON DIOXIDE FIXATION

Maier et a/. (1979) found that Hz uptake by hydrogenase-derepressed cells of R. juponicum is enhanced significantly by the addition of 4% CO2 to the derepression atmosphere, implying that the metabolism of H2 and CO? are related in some way. I t was subsequently found that COzfixation occurs in Hzderepressed cells, as discerned from RuBP carboxylase activity in these cells (Simpson et a/., 1979). Furthermore, hydrogenase and RuBP carboxylase activities are both derepressed by H, and repressed by added succinate, and the Hup- mutants SR2 and SR3 cannot be derepressed for RuBP carboxylase activity (Simpson eta/., 1979). Autotrophic growth of R . juponicum has been demonstrated using H? and COZ as the sole energy and carbon sources, respectively (Hanus et a/., 1979), and RuBP carboxylase has been purified from the autotrophically-grown cells (Purohit et a/., 1982). In contrast to R . juponicum, the expression of hydrogenase and RuBP carboxylase do not appear to be correlated in A . eutrop/ius.Whereas hydrogenase is maximally expressed in A . eutrophus when the availability of electron donors to the cell is limiting, RuBP carboxylase expression is maximal under conditions of carbon starvation and excess reducing equivalents (Friedrich, 1982). Understanding the correlation of hydrogenase and RuBP carboxylase expression in R. juponicum is complicated by the fact that hydrogenase, but not RuBP carboxylase, is expressed symbiotically. This suggests that these enzymes are subjected to multiple forms of regulation. Maier (1981) isolated R.,juponicum mutants that cannot grow autotrophically, and all of the Hupmutants also show diminished or no ability to fix COz (Cfx-). The converse is

10

M. R. O'BRIAN AND R. J . MAIER

not true, however, and mutants were obtained that are Cfx-, but Hup+. Of the four Hup+ Cfx- mutants characterized, two are specifically missing RuBP carboxylase activity. The acquisition of Hup+ Cfx- and Hup- Cfx- mutants, but not Hup- Cfx+ mutants suggest that RuBP carboxylase-mediated C 0 2 fixation ability depends on the expression of hydrogenase, but not vice versa. Physiologically speaking, this mode of regulation makes sense since there are no known growth conditions where C02 fixation occurs in the absence of H2 oxidation. In nodules, however, hydroxygenase is synthesized and RuBP carboxylase is not. Merberg and Maier (1984) found that mutants of R . japonicum which express hydrogenase constitutively (Hup') also express RuBP carboxylase activity in heterotrophically-grown cells, without subjecting the cells to the derepression conditions required by the wild type. Many of these mutants are the result of spontaneous lesions, which strengthens the argument that hydrogenase and RuBP carboxylase are coregulated by a single regulatory factor. Like the wild type, bacteroids of the Hup" mutants express hydrogenase activity, but not RuBP carboxylase activity. This latter finding is extremely interesting in light of the fact that the Hup' mutation is known to exert its effect on bacteroid cells, as discerned by the greater hydrogenase activity expressed by nodule bacteroids of the Hupc mutants compared with the wild type (Merberg and Maier, 1983). The gene mutated in the Hup' strains must therefore be involved in the regulation of hydrogenase expression both symbiotically and in free-living cells, whereas RuBP carboxylase must be regulated by a gene, in addition to the one altered in the Hup' mutants, which represses it symbiotically. C . HOST CONTROL

An aspect of hydrogenase regulation that has not been greeted with the enthusiasm it deserves is the plant host dependence of hydrogenase expression in Rhizobium. Dixon (1972) observed that the Hup phenotype of nodules infected with R . leguminosarum strain ONA 31 1 depends on the legume host from which the nodules are obtained. Peas (Pisum sativum) infected with strain ONA 31 1 produce nodules with good Hup activity, but nodules from Vicia bengalensis and V . faba have diminished and no Hup activity, respectively. Nodules from all three plants hosts fix Nz and evolve Hz. Keyser et a f . (1982) found that hydrogenase expression in some strains of R. japonicum are also dependent on the host. Rhizobium japonicum strains USDA122, USDA6I and USDA74 all form effective nodules on cultivars of soybean (Gfycine max) and cowpea ( V . unguiculata) but only USDA122 bacteroids have hydrogenase activity in both soybean and cowpea nodules. Strains USDA61 and USDA74 only express hydrogenase activity in bacter-

HYDROGEN METABOLISM IN RHIZOBIUM

11

oids from cowpea nodules, and not from soybean nodules. In addition cowpea nodules from USDA61 or USDA74 infection evolve less HZthan do soybean nodules formed from the same bacteria. Since the H2 uptake activities of USDA61 and USDA74 bacteroids from cowpea nodules are too low to account for the differences in H2 evolution between soybean and cowpea nodules, it is possible that the allocation of electrons to N2 and H + by nitrogenase is under host control (Keyser et al., 1982). Phillips and coworkers (Bedmar et af., 1983; Bedmar and Phillips, 1984a,b) found that different pea cultivars significantly affected H2 metabolism in R. leguminosarum strain 128C53, showing that host control of hydrogenase expression also occurs within a plant host species. Nodules formed by the symbiosis of strain 128C53with pea cultivars JI1205 of “Feltham First” show that the Feltham First nodules evolve 20 times more Hz per plant than JI1205 nodules. The dramatic difference in H2 evolution rates is largely due to an uptake hydrogenase expressed by the JI1205 nodules which is almost completely absent from nodules from Feltham First (Bedmar et af., 1983). Rhizobium leguminosarum strain 128C53 isolated from Hup- Feltham First nodules can be cultured and used to inoculate JI1205 peas; the resulting nodules are Hup+, showing that the plant host does not exert a permanent effect on the bacterium, such as promoting the loss of plasmid-borne hup genes (Bedmar and Phillips, 1984b). Grafting experiments show that shoots of JI1205 or “Alaska” can cause Feltham First nodules formed by strain 128C53 to express H2 uptake activity (Table 1; Bedmar and Phillips, 1984b). The reverse experiment of grafting Feltham First shoots to roots of JI 1205 or Alaska shows that the JI 1205 and Alaska nodules retain their Hup+ phenotypes. These fascinating data suggest that a transmissible shoot factor(s) can increase the H2 uptake activity of the bacterial symbiont, and that the root genotype is also important in Hup expression. Similar experiments using the Hup- R. leguminosarum 300 strain show that JI1205 or Alaska shoots grafted to Feltham First roots result in a decrease in H2 evolution by the Feltham First nodules, and a concomitant increase in the relative efficiency (Table 2; Bedmar and Phillips, 1984b). These results indicate that the allocation of electrons to Nz or H + reduction by nitrogenase is altered by the plant host. It is not known whether this phenomenon occurs in strain 128C53 as well since this strain has H2 uptake activity, but it is very possible that the lower rate of H2 evolution by strain 128C53 in JI1205 or Alaska nodules is partially due to a shift in the electron flux through nitrogenase favouring N2 reduction over H2 evolution. Hageman and Burris (1980) reported that a high electron flux through purified nitrogenase results in N2 being the preferred substrate, whereas low electron flux favours H + reduction. In those experiments, electron flux was controlled by varying the ATP concentration, the amount of reductant, or the ratio of

TABLE 1. Effect of shoot and root genotype of peas on physiological functions of root nodules by Rhizobiurn lcguniinosururn 128C53" Acetylene Hydrogen Hydrogen Relative Shoot Root reduction" evolution" uptake' efficiency ~

__ __

~

--

A A FF FF A FF FF A LSD (0.05)

12.3 14.6 8.6 10.9 3.9

1.6 5.6 2.5 0.8 I .3

5.3 I I .22 5.78 3.81 1.39

0.87 0.62 0.71 0.93 0.08

JI JI FF FF JI FF FF JI LSD (0.05)

25.6 14.0 17.9 6.0 4.4

0.6 5.5 2.3 0.1 I .o

7.77 0.89 6.13 8.97 1.09

0.98 0.61 0.87 0.98 0.03

PeacultivarsareAlaska(A), Feltham First(FF)and 511205(J1). Adapted from Bedmar and Phillips (1984b)with permission. *pnol h - ' plant-'. ' nmol H2 h-I (mg nodule fresh weight)-'. (I

TABLE 2. Effect of shoot and root genotype of peas on physiological function of root nodules formed by Rhizobium lcgurninosarurn 300" Acetylene Hydrogen Hydrogen Relative Shoot Root reduction" evolution" uptake' efficiency A A FF FF A FF A FF LSD (0.05)

10.2 9.4 5.7 6.4 2.4

6. I 5.9 2.3 4.7 1.8

0.04 0.0 I 0.04 0.04 0.01

0.40 0.37 0.60 0.27 0.07

JI JI FF FF JI FF FF JI LSD (0.05)

11.0 9.4 9.7 10.3 2.7

7.6 5.9 4.9 7.1 1.8

0.06 0.0 I 0.05 0.02 0.02

0.3 I 0.37 0.49 0.3 1 0.08

' Pea

cultivars are Alaska (A), Feltham First (FF). and J11205

(51). Adapted from Bedmar and Phillips (1984b) with permission.

" p n o l h - ' plant-'. ' nmol H2 h - ' (mg nodule fresh weight). I .

HYDROGEN METABOLISM IN RHIZOBID’M

13

component I to component I1 of nitrogenase. I t is conceivable, therefore, that JI1205 and Alaska regulate H2 evolution by providing more photosynthate to the nodules than does Feltham first, and thereby act to increase the ATP and/ or reductant available to the bacteroids. This simplistic hypothesis is supported by the grafting experiments described above using the Hup- strain 300 (Table 2) where the grafts taken on the shoot trait with respect to H? evolution and the relative efficiency. I t would be interesting to measure rates of photosynthesis in the leaves of the various pea cultivars which exhibit host control of HZ metabolism of nodule bacteroids. It is generally assumed that increased photosynthesis by the plant improves N2 fixation by providing energy for the ATP-dependent nitrogenase reaction, but it should be considered, in light of the experiments described above, that increased photosynthesis also improves the efficiency of NH1 production by nitrogenase. 111. Enzymology A. PURIFICATION AND SOME PROPERTIES

The hydrogenase from R . ,juponicum bacteroids was first reported to be a monomer with a molecular weight of about 65,000 (Arp and Burris, 1979). The enzyme was purified anaerobically from membranes by detergent solubilization, poly(ethy1ene glycol) fractionation, DEAE-cellulose and Sephadex G- 100 column chromatography. This hydrogenase preparation is similar to the hydrogenase from A . uinelundii (Kow and Burris, 1984) with respect to molecular weight composition, PI, temperature optimum, electron acceptor reactivity, K,,, value for Hz and activation energy. Subsequent purifications of the R . juponicum hydrogenase from bacteroids (Arp, 1989, chemolithotrophically-grown cells (Harker et a/., 1984) and heterotrophically-grown cells (Stults et a/., 1986) all revealed that the enzyme contains two subunits with molecular weights of about 65,000 and 35,000. Hydrogenase from bacteroids and chemolithotrophically-grown cells was purified under strict anaerobic conditions using ion-exchange and sizing columns, whereas the enzyme from a R. juponicum strain that synthesizes hydrogenase heterotrophically was purified aerobically, using a “Reactive-Red” affinity column. The molecular weight of the native enzyme is 104,000, as determined by sucrose density gradient centrifugation (Arp, 1989, indicating that one of each subunit is present in the native state. The subunit stoicheiometry of 1 : 1 was confirmed by densitometric scans of sodium dodecyl sulphate (SDS)polyacrylamide gels of the purified enzyme (Harker ef ul., 1984; Arp, 1985). Antibodies raised against each of the two subunits do not cross-react with the other subunit, showing that the 35 kDa subunit is not a degradation product

14

M. R. O'BRIAN A N D R. J. MAIER

of the larger subunit (Harker et al., 1984; Stults et al., 1986). The low molecular-weight subunit appears to be quite sensitive to proteolysis (Harker et al., 1984; Arp, 1985; Stults et al., 1986) which would explain why it eluded detection in previous studies. The role of each subunit in H2 oxidation is not known, nor is it known whether one or both subunits has catalytic activity. The purified monomer has substantial hydrogenase activity (19.6 pmol H2 oxidized min-' (mg protein)-'), but is several-fold more active when purified as a dimer (40-77 pmol H2 oxidized min-' (mg protein)-'). This increase in activity may be due to the presence of the 35 kDa subunit, or it may merely reflect the added precautions taken during the purification necessary to discern the more labile subunit. The amino acid composition of the R. japonicum hydrogenase is similar to that of A . latus, and to a lesser extent to those of A . eutrophus, A . vinelandii and Desulfovibrio vulgaris (Arp et al., 1985). Furthermore, antibodies raised against R. japonicum hydrogenase cross-react with the hydrogenase of A . latus, A . eutrophus and A . vinelandii to varying degrees. Anti-R. japonicum hydrogenase antibodies inhibit H2 uptake activity of the R. japonicum, A . eutrophus and A . latus hydrogenase, but it does not inhibit the activity of the hydrogenase from A . vinelandii (Arp et al., 1985). It is clear from these and other criteria that hydrogenase from many organisms are similar in many respects. It is difficult, however, to categorize the enzyme with respect to a particular structural or functional criterion without contradicting a separate classification based on a different parameter. Given the functional and structural similarities of the hydrogenases from R . japonicum and A . vinelandii, it is surprising that the R. japonicum hydrogenase is antigenically more similar to the A . eutrophus hydrogenase than to that of A . vinelandii. By the criterion of amino acid composition, R . japonicum hydrogenase is more similar to that of the obligate anaerobe D . vulgaris than it is to the A . vinelandii hydrogenase, yet the enzymes from R . japonicum and D . vulgaris are very different with respect to metal content, subunit composition, HZevolution rates, and their physiological roles in situ. Based on the diversity of hydrogenases found in micro-organisms, it seems that studying model systems may not be a lucrative approach to understanding the enzyme of interest. Arp (1 985) found that hydrogenase from bacteroids contains about six iron atoms per molecule, and the absorption spectrum (Fig. 1) is typical of an ironsulphur protein. The features of this spectrum are almost identical with the membrane-bound hydrogenase of A . eutrophus (Schink and Schlegel, 1979). The spectrum of the HI-reduced enzyme shows that there is no heme iron associated with the enzyme, as evident by the lack of absorption peaks in the 500-600 nm region of the spectrum. A b-type cytochrome is associated with the hydrogenase of Xanthobacter autotrophicus (Schink, 1982), but this is not

HYDROGEN METABOLISM IN RHIZOBIUM

8 0.3 c

e

5:

s 0.2 0.1

0

300

400

500

600

Wavelength (nm)

FIG. 1. Absorption spectrum of purified hydrogenase from Rhizobium japonicum bacteroids. Spectrum taken in the presence of 101 kPa Hz. Reproduced with permission from Arp (1985).

typical of hydrogenases and is not the case for the R. japonicum hydrogenase as was once speculated (Eisbrenner and Evans, 1982b). Unfortunately, the iron-sulphur clusters of R. japonicum hydrogenase have not been characterized by core extrusion or electron paramagnetic resonance (EPR) studies, and thus the nature of these centres is not known. Based on current knowledge of iron-sulphur proteins, it seems that there are three possible arrangements for six Fe atoms in Fe-S clusters: (i) three [2Fe-2S] clusters; (ii) a [4Fee4S] cluster and one [2Fe-2S] cluster, or (iii) two clusters of the [3Fe-xS] type; all three cluster types have been found in hydrogenases of other bacteria (see Moura et al., 1984; and references therein). It will be interesting to see how nickel interacts, both structurally and functionally, with the Fe-S clusters, and how the redox reactions of these groups allow catalytic activity. B. THE K, VALUE FOR HYDROGEN

The K, value of purified hydrogenase for H2 is 1-2 PM (Arp and Burris, 1979, 1981); this is fairly typical for hydrogenases from N2-fixing organisms (Kow and Burris, 1984; Benson et al., 1980; Houchins and Burris, 1981), but is very low compared with the hydrogenases from bacteria that do not fix N2. The hydrogenases from Paracoccus denitrijicans (Sim and Vignais, 1979), A.

16

M . K . O’BKIAN A N D K . J . MAIEK

eutrophus (soluble and membrane-bound; Schneider and Schlegel, 1976; Schink and Schlegel, 1979), and Clostrirlium pastrurianum (hydrogenase 11; Adams and Mortenson, 1984) have K,, values for H, that are approximately an order of magnitude higher than those found in the N, fixers. An exception to this is found in the hydrogenase of AIca1igene.s latus (Pinkwart et al., 1983), an organism that fixes nitrogen when grown autotrophically or heterotrophically with Nz as the sole nitrogen source. The K,,, values of the A . 1atu.s hydrogenase for H, is 20-25 PM in organisms cultured under either growth condition, which is more typical of the non-N:! fixers in this respect. The high affinity of R . juponicum hydrogenase for H:! is consistent with the role of hydrogenase as an HI scavenger in these cells. Since the only source of H2 in R . ,japonicurn bacteroids is from Hz evolution by nitrogenase, the H? concentration is probably low in the cell, and a high affinity for H,, by hydrogenase, would be essential in order to use the available H2 as an energy source. Emerich et al. ( 1 980) measured the apparent K,,, value of hydrogenase for Hz in whole bacteroids and found it to be about 50 nM; it is not surprising, therefore, that no H2 is evolved by nodules of hydrogen uptake positive strains (Carter et a[., 1978), and that H, oxidation by these strains is very efficient. C. ELECTRON ACCEPTOR REACTIVITY

Rhizobium japonicum hydrogenase is capable of catalysing the Hz-dependent reduction of methylene blue, dichlorophenol-indophenol (DCPIP), ferricyanide, phenazine methosulphate (PMS) and cytochrome c, and also benzyl viologen at a slow rate (Table 3; Arp and Burris, 1979). NAD(P)+, FAD, FMN, methyl viologen and 0 2 cannot accept electrons from hydrogenase. Similar, but not identical, electron acceptor reactivities have been observed in the hydrogenases from Azotohacter chroococcum (van der Werf and Yates, 1978), A . ilinelmdii (Kow and Burris, 1984), A . eutrophus (membrane-bound; Schink and Schlegel, 1979) and A . /atus (Pinkwart et a/., 1983). The electron acceptors that can support hydrogenase activity in purified R . japonicum also work well in membrane preparations of free-living cells derepressed for hydrogenase activity (Table 3; Mutaftschiev et al., 1983). The membranes also contain an electron transport system that, unlike the purified preparation, allows 02-dependent Hz oxidation (O’Brian and Maier, 1982). Assuming the electron acceptor studies have some physiological significance, it seems that the electron acceptor of hydrogenase in situ has a midpoint potential close to 0 mV since artificial electron acceptors with more negative potentials cannot be reduced by HI via hydrogenase. Since the Hr/H+ redox couple has a standard midpoint potential of -420 rnV, it may appear that H2 oxidation via hydrogenase is an extremely inefficient mechanism since the reducing power of Hr via the enzyme is only about 0 mV. It should be

17

HYDROGEN METABOLISM I N RNIZOSIi'Ai

TABLE 3. Activity of hydrogenase in the purified state and in membranes with various electron acceptors Activity (%)

Electron acceptor

Purified" Membranesb ~~~

Methylene blue

Phenazine methosulphonate (PMW Ferricyanide

11

100

I00

80 360

N.D. 134

182 141

217 816 250

78 0 80

53 40 18

Dichlorophenol-indophenol

(DCPIP) Oxygen Cytochrome c Flavin mononucleotide (FMN) Benzyl viologen Methyl viologen

- 122 - 360

0 0.3

440

<0.1

-


" Hydrogenase purified from bacteroids. Adapted from Arp and Burris (1979) with permission. Membranes from hydrogenase-derepressed cells. Adapted from Mutaftschiev et ( I / . (1983) with permission.

considered, however, that the reducing power of Hz depends on the ratio of H2 to H + in the bacteroid, thus the actual reduction potential may be quite different from the standard reduction potential of - 420 mV. The apparent K,, value for HZin the bacteroid is estimated to be 50 nM (Emerich e f al., 1980), suggesting that the intracellular HZconcentration is very low. If the actual H2 concentration in the bacteroid is close to its K,,, value (50 n M ) and the pH is 7 (Hf = M), then the reduction potential of the H2/H+ couple is about - 200 mV as calculated from the Nernst Equation; this figure is considerably higher than the standard reduction potential. If these considerations are true, then the efficiency of H2 oxidation via hydrogenase is more efficient than routine enzyme assays would suggest. Hydrogen-oxidizing bacteria with high intracellular HZ concentrations possess hydrogenases that are capable of reducing low-potential electron acceptors. For example, C . pasreurianunz hydrogenase I1 can rapidly catalyse the reduction of methyl viologen (Eo' = -440 mV) by H2 (Adams and Mortenson, 1984), and the physiological electron acceptor of several Desulfovibrio species is cytochrome cj, which contains four hemes with midpoint potentials ranging from - 339 to - 235 mV (Odom and Peck, 1984). The low-potential electron acceptors probably reflect the low potential of hydrogenase in these organisms, which can

18

M . R . O'BRIAN AND R . I. MAIER

efficiently exploit the greater potential energy of H2 compared with the N2fixing aerobes. D. OXYGEN LABILITY

Hydrogenases are generally treated as 02-labile enzymes during purification and analyses. The 0 2 sensitivity of hydrogenases and other iron-sulphur proteins is presumably due to the oxidation of the acid-labile sulphide moieties and the cysteine residues of the iron-sulphur clusters (Adams et ai., 1981). Hydrogenase I of C . pasteurianum has a half-life of only two to three minutes in air (Erbes and Burris, 1978), and the purification of this enzyme under strict anaerobic conditions (Chen and Mortenson, 1974) has set a precedent in the purification of hydrogenase from other organisms. There are, however, quite a few reports showing that some hydrogenase preparations are rather 02-tolerant during purification and storage (Adams et ai., 1981). The soluble hydrogenase of A . eutrophus is actually stabilized by 0 2 ; the activity decreases by only about 25% in three weeks when stored in air, but loses all enzymatic activity in five days when stored under H2 or other reducing agents (Schneider and Schlegel, 1976). The hydrogenase from R .japonicum purified anaerobically is quite sensitive to 0 2 with a half-life of about one hour in air (Arp and Burris, 1979, 1981). Oxygen can also act as a reversible inhibitor of this hydrogenase preparation if the 0 2 concentration is low and the exposure time short (Arp and Burris, 1981). This 0 2 inhibition of activity is non-competitive versus methylene blue, and uncompetitive versus H2 with methylene blue as the electron acceptor. Mutaftschiev et al. (1983) found that 0 2 is a reversible inhibitor of methylene blue-dependent H2 oxidation in R . japonicum membrane preparations, and that the activity continues to increase for about five minutes after the O2 is depleted by respiration. Recently, Stults et al. (1 986) purified hydrogenase under completely aerobic conditions using a Reactive-Red 120-agarose affinity column. This preparation was homogeneous by the criterion of SDS-polyacrylamide gel electrophoresis, and had very good H2 uptake activity. It was found that, unlike the anaerobic preparations of hydrogenase, the aerobically purified hydrogenase was very stable in air. However, when this preparation was treated with dithionite and subsequently exposed to air, the hydrogenase activity became quite 0 2 sensitive (Fig. 2), and the half-life of activity was about one hour (Stults et al., 1986), as was found with the anaerobic preparations (Arp and Burris, 1979). A similar phenomenon has been reported for the aerobically-stable hydrogenase from D . uulguris (van der Westen, et ul., 1980); this enzyme was purified aerobically, and was stable in air, or when reduced with HZor dithionite, but the reduced form of the enzyme

19

HYDROGEN METABOLISM IN RHIZOBIUM

100

80

.-cx. 0

60

-

0 .c ..-E c

c

40

L

cit: 20

0

0

1

2

3

4

Time (hours)

FIG. 2. Effect of air and dithionite on the stability of hydrogenase activity. Hydrogenase was purified aerobically from heterotrophically-grown Rhizobium juponicum, and incubated in air (a), or air plus dithionite (0).Reproduced from Stults et al. (1986).

was rapidly inactivated when exposed to air. The addition of reducing agents to hydrogenase in the presence of air is likely to cause the partial reduction of 02 to form very reactive oxygen species, such as H202, 0 2 -and , OH., which will inactivate the enzyme. Likewise, the purification of hydrogenase under strict anaerobic conditions and in the presence of reducing agents would result in an 02-sensitive preparation due to the formation of oxygen radicals. Schneider et al. (1979) added ferricyanide to crude extracts of A. eutrophus during purification to stabilize the soluble hydrogenase. This stability is probably due to the oxidation of endogenous reducing agents by ferricyanide, thus preventing the formation of reactive oxygen species. E. NICKEL

Nickel is now known to play a role in HZmetabolism in a wide variety of

20

M. R . O'ERIAN AND R . J . MAIER

organisms. With hindsight, this is not surprising since nickel readily reacts with Hz to form reactive nickel hydride (Thauer ef al., 1980), and nickel catalysts can be used in hydrogenation reactions that involve nickel hydride complexes (Sacconi et al., 1974). Nickel has been shown to be required for chemolithotrophic growth in several H2-oxidizing bacteria (Tabillon et al., 1980), and is required for hydrogenase synthesis or activity in Anabaena cylindrica (Daday and Smith, 1983), A . variabilis (Almon and Boger, 1984). Azotobacter chroococcum (Partridge and Yates, 1982) and A . eutrophus (Friedrich et af., 198la). More specifically, nickel has been shown to be a part of the hydrogenase enzyme of Escherichiu coli (Ballantine and Boxer, 1985), Chromatium vinosum (Albracht et ul., 1983), Methanobacterium thermoautotrophicum (Graf and Thauer, 1981 ). M . bryantii (Lancaster, 1982). Desuljiovibrio desuljiuricans (Kruger et al., 1982), D . gigas (LeGall et al., 1982), A . eutrophus (Friedrich et ul., 1982; Schneider et a[., 1983) and Rhodopseudomonus capsulata (Colbeau and Vignais, 1983). There is evidence that nickel acts as a redox centre in the hydrogenase of D . gigas (Cammack et a[., 1982; LeGall et al., 1982), and that Hz may bind to the nickel centre (LeGall et al., 1982). The role of nickel in the catalytic site of hydrogenase is discussed by Moura et al. (1984). Klucas et ul. (1983) studied the effects of nickel on urease and hydrogenase activities in soybean leaves and root nodule bacteroids, respectively; urease is known to be a nickel-containing enzyme. Urease activity in the leaves of 64day-old soybean plants increased over 10-fold when 0.1 p~ NiClz was added to the nutrient media. The effect of nickel on the hydrogenase activity of nodule bacteroids depended on the age of the plant in those experiments. Bacteroids from 29-day-old plants had good hydrogenase activity regardless of whether nickel was added to the nutrient media. However, bacteroids from 48-day-old plants treated with nickel had 33% more activity than those from plants deprived of nickel. Added nickel had the greatest affect on the hydrogenase activity of bacteroids from nodules that had begun to senesce; although the activity was poor, it was about 70% higher in bacteroids from nickel-treated plants than from plants not treated with nickel. The same investigators reported that free-living cells can only be derepressed for hydrogenase activity if nickel is added to a derepression medium that was chemically treated to remove trace metals. Although this restored activity was low, it was higher than if nickel (5 p ~ was ) replaced by chromium, tin, vanadium, or lead ( 1 PM) in the depression medium. Stults et al. (1984) found that the metal chelator EDTA inhibits the derepression of hydrogenase activity in free-living R .juponicum cells, and that Ni can overcome the EDTA inhibition. The metal ions Co, Cu, Fe, Mg, Mn or Zn cannot replace Ni in overcoming this inhibition. These data show that nickel is important in the expression of hydrogenase in a manner unique to

HYDROGEN METABOLISM IN RHIZOBI(IM

21

that metal. They also showed that 63Nico-migrates with hydrogenase activity in native polyacrylamide gels of solubilized membranes from a hydrogenaseconstitutive strain of R. japonicum. The latter result shows that nickel is a structural component of the hydrogenase enzyme, which has been subsequently confirmed in the R. japonicum hydrogenase from autotrophicallygrown (Harker et al., 1984) and symbiotically-grown cells (Arp, 1985). Arp (1985) determined the nickel and iron contents of the purified hydrogenase from bacteroids by atomic absorption spectroscopy, and found 0.59 mol Ni and 6.5 mol Fe per mol hydrogenase. It is not known how nickel is arranged in the enzyme, or if the two metals interact with each other, or even if Ni undergoes any redox changes during catalysis, as has been found in hydrogenases of other bacteria (Cammack et al., 1982; LeGall e f al., 1982). EPR studies show that the nickel is distant from the EPR-detectable Fe-S cluster in D . gigas, although both groups may be involved in catalysis (Cammack et al., 1982). Lindahl e f al. (1984) found that sulphur is the dominant scatterer in the nickel EXAFS (Extended X-ray Absorption Fine Structure) spectrum of the oxidized Fd2~-reducinghydrogenase of M . thermoautotrophicum; this scattering is due to three sulphur atoms at about 2.25 A. The Fe EXAFS spectra show that Fe-S and Fe-Fe distances are nearly identical with those found in well characterized iron-sulphur proteins. The EPR and EXAFS studies give the impression that Ni and the Fe-S clusters d o not form a single species within the enzyme. Using EPR to study the catalytic properties of D.gigas hydrogenase, it was found that the Ni and Fe-S centers are reduced to the same extent by a chemical reductant, but in the presence of H2 the Ni centre can be fully reduced without full reduction of the Fe-S centres are reduced to the same extent by a chemical reductant, but in the presence of H2 the Ni centre can be fully reduced without full reduction of the of nickel in hydrogen metabolism is complicated by the fact that Ni is not obligatory in H2 consumption or evolution by hydrogenase since the hydrogenases of C. pasteurianum (Adams and Mortenson, 1984) and that of D.vulgaris (Huynh e f a/., 1984) are very active, yet they d o not contain nickel. F. LIPID REQUIREMENT

Several membrane-bound enzymes have a general or specific lipid requirement for optimal activity (Isaacson et al., 1979). It has been observed that phosphatidyl choline, a rare lipid in most bacteria, is a major phospholipid component in a number of Hroxidizing bacteria (Thiele and Oulevey. 1981). including R. japonicum (Bunn and Elkan, 1971). Moshiri and Maier (1983) studied the effect of lipids on hydrogenase activity in R. ,japonicum. It was found that acetone extraction removes over 8 5 % of the phospholipid of membranes from hydrogenase-derepressed cells, but only diminishes hydro-

22

M. R . O'BRIAN AND R. J . MAIER

TABLE 4. Effect of lipid extraction procedures on hydrogenase activity in membranes from hydrogenase-derepressed cells. From Moshiri and Maier (1983). Extraction treatment None Acetone (90%) Acetone plus ammonia Acetone, twice Acetone, then acetone plus ammonia Ammonia in buffer

Lipid phosphorus Hydrogenase Total protein recovery" activity" recovered" 100 15

100

86

100 53

5

N.D.h

32 62

49 42

N.D.

13

40

100

98

100

Percent of control (no treatment). N.D.,not determined.

gen uptake activity 10-15% (Table 4). However, extraction of membranes with acetone plus 0.05% ammonia decreases hydrogenase activity 65-70%, and removes 95% of the total phospholipid. Thin-layer chromatography of the residual lipid from the extracted membranes shows that acetone-treated membranes are significantly enriched for cardiolipin compared with acetone plus ammonia treated samples. Unfortunately, attempts to reconstitute delipidated membranes have been unsuccessful, and thus the specific lipid requirements for hydrogenase has not been ascertained. The failure to reconstitute hydrogenase activity in delipidated membranes may be a result of irreversible denaturation of hydrogenase on removal of tightly bound lipids. The presence of phosphatidyl choline in H2-oxidizing bacteria may be due to an evolutionary relatedness of these organisms rather than being a lipid required for hydrogenase per se. This speculation is based on the fact that phosphatidyl choline is a major lipid in H2 oxidizing bacteria, but hydrogenase makes up only a small portion of the total protein in the membrane. Cardiolipin, however, is only a minor component in the R. japonicum membrane (Moshiri and Maier, 1983), and extraction procedures that preferentially remove this lipid (Fleischer and Fleischer, 1967) result in a significant decrease in hydrogenase activity. G. KINETIC MECHANISM OF HYDROGENASE

Arp and Burris (1981, 1982) studied the kinetic mechanism of hydrogen

HYDROGEN METABOLISM IN RHIZOBIUM

23

oxidation by hydrogenase from R. japonicum bacteroids. Isotope discrimination experiments showed that hydrogenase oxidizes H2 about 25% faster than it does DZ(2H2).This small isotope effect on the oxidation rate shows that the breaking of the dihydrogen bond is not the rate-limiting step in catalysis. They proposed a two-site, ping-pong mechanism for the enzyme in which H2binds and is activated at one site, and electron carriers bind at a second site (Fig. 3). In this scheme, H2 binds to the oxidized enzyme (E), resulting in the release of protons and the reduction of the enzyme (E'). The reduced enzyme can then react with the oxidized electron carrier (B) at a site different from the HZbinding site, and electrons are transferred from the enzyme to the electron carrier. The data for the mechanism shown in Fig. 3 are essentially as follows: (i) H2is a strong inhibitor of H2 evolution, and although H2 and the reduced electron carrier (methyl viologen) both react with the oxidized enzyme, the inhibition of H2evolution by H2 is non-competitive, showing that H2 and the reduced electron carrier bind to different sites on the enzyme. (ii) The inhibition of H2 oxidation by oxidized methylene blue is competitive versus H2.Although this could be interpreted to mean that H2 and methylene blue bind at the same site, the authors point out that this is unlikely given the very different chemical structures of the two substrates, and suggest that the binding of oxidized methylene blue to the enzyme prevents the binding of H2 at another site. This conclusion implies that H2 must bind to the enzyme before the binding of the electron acceptor. (iii) Carbon monoxide is a competitive inhibitor of H2 and non-competitive versus methylene blue, which means that CO and H2 bind at the same site on the enzyme, which is different from the methylene blue binding site. (iv) Oxygen is an uncompetitive inhibitor versus H2, thus 0 2 is shown to EB

ECO

FIG. 3. Model for the kinetic mechanism of the Rhizobium japonicum hydrogenase. E, oxidized enzyme; E , reduced enzyme; B, oxidized electron carrier; B , reduced electron carrier. k l , kz etc. are rate constants. Reprinted with permission from Arp and Burris (1981). Biochemistry 20, 2234. Copyright 1981, American Chemical Society.

24

M . R . O’BRIAN A N D R . J . MAIEK

bind to the reduced enzyme in Fig. 3 . The inhibition by 0 2 in non-competitive versus methylene blue, indicating that 0 2 and methylene blue bind at different sites on the reduced enzyme. The latter result is consistent with the observation that 0 2 cannot act as an electron acceptor of the purified enzyme. (v) The enzyme can catalyse a D?-HzO exchange reaction in which H D and H2 are formed. This shows that H? can bind to, and protons be released from, the enzyme without the presence of an electron carrier. The release of product (protons) without the binding of the electron carrier is consistent with a pingpong mechanism. If the mechanism were sequential, both substrates would need to bind to the enzyme before any product was released.

IV. Energetics A. PHYSIOLOGICAL CONSIDERATIONS

The obligatory evolution of Hz by nitrogenase is very expensive in terms of the ATP and reducing equivalents spent on the reduction of protons to Hz. The reduction of Component I of nitrogenase by Component I 1 requires 2 ATP molecules per electron (Tso and Burris, 1973; Ljones and Burris, 1978), thus the two electrons involved in one Hz molecule costs at least four ATP molecules. Considerations of the energy required to produce the low-potential electron equivalents that reduce Component I1 bring the HS production budget up to seven ATP molecules according to some estimates. Calculations of the energy efficiency of N2 fixation assume 25% of the electron flow through nitrogenase is committed to H, production, when actually the allocation of electrons to proton reduction may be as high as 60% (Schubert and Evans, 1976). The H2 molecule contains a tremendous amount of potential energy, and thus the conservation of some of this energy via Hz oxidation, as postulated by Dixon ( I 972). would be advantageous. The recovery of Hz potential energy could conceivably occur via electron transport to 0 2 and concomitant ATP synthesis, or by providing reducing equivalents directly to nitrogenase. There is evidence that Hr oxidation in Anahaena cylindrica and Azorohacter chroococcum does donate reducing equivalents to nitrogenase (Bothe et al., 1977; Walker and Yates, 1978). Hydrogen oxidation-dependent ATP synthesis has been demonstrated in P . denitr$can.v (Knobloch et a/., I97 I ; Porte and Vignais, 1980), Pseudomonas sacchurophiliu (Ishaque et al., 1971) and A . eutroplius (Bongers, 1967; Ishaque and Aleem, 1970) by measuring P/O ratios or H+/O ratios in vesicles or crude extracts. In Rhizohium, energy conservation of HI oxidation has been studied by examining steady-state levels of ATP in cells in the absence and presence of Hr (Emerich et a/., 1979; Nelson and Salminen, 1982). The oxidation of Hz by

HYDROGEN METABOLISM IN RHlZOBIUM

25

isolated R.japonicum bacteroids increases the steady-state pool of ATP by 2040% (Emerich et al., 1979). In the presence of iodoacetate, a respiratory inhibitor that blocks endogenous respiration to a greater extent than it does H2 oxidation, the steady-state cellular ATP concentration is 500% greater in the presence of H2than in its absence. These data indicate that HZoxidation is coupled to ATP synthesis. However, Nelson and Salminen (1982) found that only five out of fourteen Hup+ R. leguminosarum strains studied were coupled to ATP synthesis as judged by ATP levels of bacteroids in the presence and absence of Hz. The increase in the intracellular ATP concentration in these five strains ranged from 15 to 70% in the presence of Hz. Interestingly, the five strains that show H2-dependent oxidative phosphorylation also have the highest H2 uptake rates of the 14 strains examined. It was also shown that H2 oxidation by R. leguminosarum results in an increase in the 0 2 concentration optimum for C2H2reduction by the bacteroids regardless of whether the strain couples Hz uptake to ATP synthesis. From these data, it was concluded that the major role of H2 oxidation in R. leguminosarum is to protect nitrogenase from damage by 0 2 . Hydrogen oxidation in R.japonicum also appears to raise the 0 2 concentration at which CzH2 reduction is maximal (Emerich et al., 1979). There has been speculation recently that hydrogenase may actually be detrimental to Nz fixation in R. japonicum based on the assertion that H2 oxidation consumes 0 2 that would otherwise be used more efficiently by another energy source (Drevon et al., 1985). This speculation assumes that H2 oxidation produces less ATP per molO2 consumed than does the oxidation of other available substrates, and that 0 2 , rather than photosynthate, is the limiting factor in the nodule. These two considerations are discussed below. The conclusion that H2 yields less energy than other substrates available in root nodules comes from experiments done with succinate-limited chemostat cultures of Rhizohium ORS 571. Stam et al. (1984) observed a slight decrease in the molar growth yield on 0 2 (Yoz)when H2 was added to Nz-fixing chemostat cultures. Considering that the decreases in Yo2 on the addition of H2 were only 4% and 15% in two experiments, it is difficult to apply these data confidently to R.japonicum grown in soybean root nodules. Unlike symbiotic R. japonicum, Rhizohium ORS 571 assimilates the N2 it fixes, and in the chemostat cultures described above, NZis the sole nitrogen source. Given this fact, factors that affect the efficiency of NZ fixation would be expected to influence the Yo2.In the experiments of Stam.et al. (1984), HZwas added as a gas containing 4-1 5% H2, which could inhibit the nitrogenase enzyme. The inhibition of nitrogenase by H2 results in a decrease in the efficiency of N2 fixation (Dixon et al., 1981), and therefore may affect the YOZin Rhizobium ORS 571. It should also be seriously considered that the addition of H2 to the chemostat cultures may induce the synthesis of RuBP carboxylase and other

26

M. R. O'BRIAN A N D R . J. MAIER

C02 fixation enzymes, as occurs with R. japonicum (Simpson et al., 1979) and many hydrogen bacteria. The induction of RuBP carboxylase could account, in part, for the observed increase in the molar growth yield on succinate by the addition of H2. Also, the oxygenase activity of RuBP carboxylase would decrease the Y02, and would also mitigate the importance of a Yo2 measurement. It seems that growth yield studies in nitrogen-fixing organisms should be approached with caution because a tremendous amount of energy is expended in reducing N2 to ammonia, which does not contribute to an increase in the biomass in most symbiotic N2 fixers. Rhizobium ORS 571 does assimilate the fixed nitrogen, but the energy requirement is so great (42 ATP/ N2 fixed; Stam et al., 1984) that growth yield measurements are likely to be insensitive to energy fluctuations in the cell, The second assumption made in speculating that H2 oxidation is detrimental to N2 fixation is that 02,rather than photosynthate, is limiting in the nodule. This controversy has not been resolved, but the evidence thus far suggests that photosynthate is the limiting factor in N2 fixation. Hardy and coworkers (Hardy et al., 1978,and referencestherein) have demonstrated that N2 fixation can be increased significantly in nodules from soybeans, peas and peanuts by enriching the C02 content in the atmosphere of the plant. The higher C02/02 ratio shifts the ratio of carboxylase to oxygenase activities of RuBP carboxylase in favour of carboxylase activity, resulting in an improved efficiency of photosynthesis. The greater availability of photosynthate to the nodule results in increased nodule mass, increased specific N2-fixingactivity, and delayed senescence. Furthermore, Minchin et al. (1985) reported that long-term exposure of pea and soybean plants to various 0 2 concentrations does not result in significant differences in N2 fixation ability or rates of plant growth, suggesting that 0 2 is not limiting. The question of 0 2 limitation can also be addressed at the molecular level. The 0 2 concentration inside a soybean root nodule is estimated to be 1 1 nM as calculated by the concentration of leghaemoglobin in the peribacteroid space, the equilibrium dissociation constant of leghaemoglobin for 02,and the partial oxygenation of leghaemoglobin in the nodule (Appleby, 1984). The low O2concentration in the R. japonicum bacteroid is compensated for by a terminal oxidase with an extremely high affinity for 0 2 (Bergersen and Turner, 1980); the apparent dissociation constant of the oxidase for 0 2 is about 5 nM. Furthermore, the high 0 2 affinity system is coupled to ATP synthesis, and the bacteroid ATP concentration is greatest when the 0 2 tension is between 20100 nM in the presence of leghaemoglobin (Bergersen and Turner, 1975). Appleby (1984) estimated that the ratio of bound (as oxyleghaemoglobin) to free O2is about 55,000 in the nodule cytoplasm at a free O2concentration of 1 1 nM, thus leghaemoglobin has an enormous O2buffering capacity. In addition, the 0 2 dissociation rate constant is relatively fast (5.5 s-', Appleby et al.,

HYDROGEN METABOLISM IN RHIZOBIVM

27

1983), therefore the 0 2 concentration in the bacteroid should be maintained during vigorous respiration. Despite the buffering capacity of leghaemoglobin and the high O2 affinity terminal oxidase in R .japonicum, there are conceivably situations in which 0 2 may be limiting in the nodule. Sheehy et al. (1985) proposed a mathematical model to explain 0 2 diffusion, and resistance to diffusion, in a legume nodule. They asserted that 0 2 may be limiting under situations where the diffusion resistance of the nodule is invariable. Criswell et al. (1976) found that exposing intact soybeans to low O2tensions results in an initial decrease in N2 fixation followed by a recovery of activity after 4-24 hours. These data are interpreted by Minchin et al. (1985) as a reduction in the diffusion resistance by the nodule, presumably caused by a change in nodule structure. Furthermore, 2deprivation does not affect NZfixation in soybean or pea nodules long-term 0 (Minchin et al., 1985). It seems that the nodule is able to “adjust” its diffusion resistance to maintain an internal 0 2 concentration optimal for N2 fixation. These experiments indicate that 0 2 is not limiting in intact nodules since decreasing the atmospheric 0 2 concentration to levels much lower than are likely to occur in nature does not result in any long-term inhibition of N2 fixation. B. ELECTRON TRANSPORT

1 . General Background

The electron transport components that oxidize H2 have been studied in P . denitriJicans(Porte and Vignais, 1980; Knobloch et af., 1971), A . eutrophus (Bongers, 1967; Ishaque and Aleem, 1970; Bowien and Schlegel, 1981) and A . vinelandii (Laane et al., 1979; Wong and Maier, 1984); spectral studies and P/O measurements show that electron flow from H2 may follow a different route from other substrates. Paracoccus denitr$cans grown autotrophically with H2 contains cytochromes aa3 and o as the terminal oxidases but H2 is oxidized only by cytochrome o (Porte and Vignais, 1980). Hydrogen is oxidized by the cytochrome d branch of the electron transfer system in N2fixing A . uinelandii,and cytochrome o does not seem to be involved (Wong and Maier, 1984). The H2-oxidizingelectron-transport system of R.japonicum has been studied in free-living cells under various growth conditions (O’Brian and Maier, 1982; 1985a,b; Eisbrenner and Evans, 1982b; Eisbrenner et al., 1982), and in bacteroids (Eisbrenner and Evans, 1982a; OBrian and Maier, 1983). Despite the 0 2 lability of the H2 oxidation system in R .japonicum, membranes from bacteroids and free-living cells with good 02-dependent hydrogenase activity can be obtained provided that certain precautions are taken to exclude 0 2 during the membrane isolation procedure (O’Brian and Maier, 1982,1983;

28

M. R. O’BRIAN AND R . J. MAIER

Mutaftschiev et al., 1983). Such membrane particles are ideal for studying Hz oxidation in R. juponicum since whole cells contain large amounts of endogenous substrates which cannot be readily exhausted, making H2dependent reactions difficult to discern. Furthermore, membrane preparations are generally more sensitive to electron-transport inhibitors than are whole cells. 2. Free-living Rhizobium japonicum

Oxygen-dependent H2 uptake by membranes from free-living cells are derepressed for hydrogenase activity is sensitive to the classical oxidase inhibitors cyanide, azide and hydroxylamine, and also to antimycin A (O’Brian and Maier, 1982). The latter inhibitor is not effective in blocking H2 oxidation in P . denitrificans at concentrations sufficient to inhibit NADH and succinate oxidase activities (Porte and Vignais, 1980). Rotenone is a good inhibitor of NADH oxidase activity in R.japonicum membranes, but does not inhibit H2 uptake (O’Brian and Maier, 1982), indicating that H Zoxidation is not NADH-linked, as it is in the H2 oxidation pathway associated with the soluble hydrogenase of A . euttophus (Bowien and Schlegel, 1981). Cytochrome spectra of membranes from H2-derepressed cells show that HZ reduces b- and c-type cytochromes, and cytochrome aa3 (Fig. 4; O’Brian and Maier, 1982). Unlike NADH or succinate, H2 does not reduce any flavin moiety, showing that R. juponicum is not a flavoprotein, nor does it reduce NADH dehydrogenase, as occurs with the soluble hydrogenase of A . eutrophus (Bowien and Schlegel, 1981). The spectra are consistent with the observations that Hz oxidation is rotenone-insensitive (O’Brian and Maier, 1982), and hydrogenase does not donate electrons to NAD+ in membranes (Mutaftschiev et al., 1983) or in the purified state (Arp and Burris, 1979). Cytochrome aa3 is present in cells derepressed for hydrogenase activity (OBrian and Maier, 1982), as well as in heterotrophically-grown cells, and has been shown to be an oxidase by the criterion of photochemical action spectra (Appleby, 1969b). Since derepression of hydrogenase requires that cells be incubated under 1YOO2for 48-72 hours, the presence of cytochrome aa3 in these cells is somewhat surprising. Previous studies show that this oxidase disappears, or is greatly diminished in cells grown under 2% 0 2 (Daniel and Appleby, 1972), and in cells that are incubated under restricted aeration (Avissar and Nadler, 1978). Also, cytochrome ua3 expression is repressed in 0,-limited cultures of P . denitrificans (Sapshead and Wimpenny, 1972) and Rhodopseudomonas sphaeroides (Sasaki et al., 1970),suggesting that this oxidase is not suited to function at low 0 2 tensions. Also, P . denitrijicans cells grown autotrophically with H2 as the sole energy source contain cytochrome uu3, but at a lower concentration than in cells grown heterotro-

29

HYDROGEN METABOLISM IN RHIZOBIUM

428

551.5

463

420

460

4$2 , , 1 500 460 500 540 Wavelength (nm)

1

1

580

I

I

I

620

FIG. 4. Substrate-reducedminus 02-oxidized absorption spectra of membranes from hydrogenase-derepressed cells of Rhizobium juponicum. A, dithionite; B, Hz; C, succinate; D, NADH. Reproduced from O’Brian and Maier (1982).

phically (Porte and Vignais, 1980). Cytochromes uu3 and o both act as terminal oxidases in autotrophically-grown cells of P. denitri$cuns, but Hz uses only the electron-transfer pathway to 0 2 terminated by cytochrome o (Porte and Vignais, 1980). Hydrogen oxidation in derepressed R. juponicum involves both cytochromes uu3and o (O’Brian and Maier, 1982), although the extent to which each oxidase participates has not been determined. The persistence of cytochrome uu3 in H2-derepressed cells of R. juponicum may be due to the fact that the cells do not grow during the derepression period, and thus turnover rates of some systems may be slow. Alternatively, cytochrome uu3 may be required for cell viability during the energy-stressed condition of hydrogenase derepression. Although the 6-type cytochrome can only be discerned as a shoulder at 560 nm in H2-reduced difference spectra of membranes from hydrogenasederepressed cells (Fig. 4), CO and cyanide spectra reveal two 6-type cytochromes involved in H2 oxidation (OBrian and Maier, 1982). The CO-

30

M. R. O'BRIAN AND R. J. MAIER

and CN--binding b-type cytochrome is cytochrome 0;this component has been shown unequivocally to be a terminal oxidase, as discerned by photochemical action spectra (Appleby, 1969b). Cytochrome o is also reducible by ascorbate, whereas the other b-type cytochrome is not (OBrian and Maier, 1982). The reduction of cytochrome o by ascorbate is presumably due to the relatively high redox potential of this oxidase compared to the other b-type cytochrome. Mutants of R. japonicum have been isolated that express hydrogenase activity constitutively (HupC) in free-living culture, and need not be derepressed for hydrogenase activity as does the wild type (Merberg and Maier, 1983; Merberg et al., 1983). Cytochrome spectra of membranes from Hup' mutants grown heterotrophically show that these strains express significantly more 6-type cytochrome than does the wild type, and further analyses show that the additional component is cytochrome o (O'Brian and Maier, 1985a). Carbon monoxide spectra of membranes show that the Hup' mutants contain over three times as much cytochrome o as does the wild type, and the dissociation constant of CO for cytochrome o in the membrane is about 6 p~ for the wild type and two Hup' strains examined. The cytochrome o expressed in the Hupc strains is readily reducible by NADH and succinate, as well as H2, and is not unique to the H2 oxidation system. However, hydrogenase and cytochrome o expression may be regulated by common factors such that a lesion resulting in hydrogenase synthesis also results in the expression of cytochrome 0 . In several bacteria that possess cytochromes aa3 and o as the terminal oxidases, the expression of cytochrome o is greatly enhanced when the cells are grown at low 0 2 tensions (Sapshead and Wimpenny, 1972; Sasaki et al., 1970; Sone and Yanagita, 1982), suggesting that cytochrome o is better suited to function at a low 0 2 concentration than is cytochrome uu3. In R. juponicum, cytochrome o reduction is not observed until the membrane suspension is nearly anaerobic (OBrian, 1984), indicating that cytochrome o can bind 0 2 with a high affinity. Also, H2 and NADH oxidations involve cytochrome o, and not cytochrome ua3, at an 0 2 concentration of 5 p~ (O'Brian and Maier, 1985a), showing that cytochrome o is the functional oxidase in R . japonicum when the 0 2 concentration is low. Since hydrogenase is normally expressed only under restricted aeration, and cytochrome o expression is enhanced at low 0 2 concentrations, it seems that the Hup' mutants are altered in regulation by 0 2 . resulting in the expression of both cytochrome o and hydrogenase at O2concentrations which 2 is would normally repress them. It is likely, however, that regulation by 0 only a partial explanation of the cytochrome o expression in the HupC mutants. The cytochrome pattern of the Hup' strains (OBrian and Maier, 1985a) and cells grown autotrophically with H2 under 2% 0 2 (Eisbrenner et al., 1982)are similar to each other in that the ratio of b- to c-type cytochromes

HYDROGEN METABOLISM IN RHIZOBIUM

31

is greater in these cells compared to cells grown heterotrophically. However, cells grown heterotrophically under 2% 0 2 synthesize significantly more band c-type cytochromes than cells grown in air, but the ratio of b- to c-type cytochromes is lower in the cells grown under restricted aeration (Daniel and Appleby, 1972). In other words, the expression of cytochrome o in the Hup‘ mutants is probably a low 0 2 response but the cytochrome pattern of these cells does not look like cells grown heterotrophically under restricted aeration. Although not rigorously examined, it seems that the carbon and energy sources must also regulate cytochrome expression in R.japonicum. Cox et af. (1978) found that P. denitrzjicans cells grown aerobically in minimal media with succinate contain cytochromes o and aa3, but cells cultured in a rich nutrient medium lack cytochrome aa3 even though the latter growth condition was also aerobic. When this bacterium is grown autotrophically, a-type cytochrome expression is enhanced when methanol is the energy source (van Verseveld and Stouthamer, 1978), whereas synthesis of this cytochrome is diminished when H2 is the substrate (Porte and Vignais, 1980). It should be considered, therefore, that changes in the cytochrome pattern of the Hup‘ strains and chemoautotrophically-grown cells of R. japonicum are also controlled by carbon and energy sources. Although the Hup‘ mutants are grown heterotrophically, the regulation of these cells by H2 and organic carbon has been altered (Merberg et al., 1983), which may affect the cytochrome pattern in a way similar to that postulated for 0 2 . Ubiquinone is present in R.japonicum cells grown symbiotically, aerobically and anaerobically, and it participates in 02-dependent NADH oxidation (Daniel, 1979). Ubiquinone involvement in H2 oxidation in hydrogenasederepressed cells was implicated by the observation that H2 uptake is greatly diminished in crude extracts irradiated with ultraviolet light, and that activity can be restored by the addition of ubiquinone (O’Brian and Maier, 1982). Direct evidence for the participation of ubiquinone in H2 oxidation was obtained by extracting the quinone from Hz-treated membranes such that the redox state of ubiquinone is preserved during the extraction procedure (O’Brian and Maier, 1985b). It was found that H2 reduces 75-85% of the extractable ubiquinone in the membrane (O’Brian and Maier, 1985b), similar to the NADH-dependent reduction level in aerobically-grown cells (Daniel, 1979). In the presence of HQNO, H2 is able to reduce ubiquinone, whereas H2dependent cytochrome reduction and 02-dependent H2 uptake are blocked by this inhibitor (O’Brian and Maier, 1985b). These data show that HQNO inhibits Hrdependent electron transport between ubiquinone and the cytochromes, and therefore ubiquinone is located on the substrate side of the cytochromes in the electron-transport system, with no cytochrome component mediating electron flow from hydrogenase to ubiquinone. There appears to be only one HQNO-binding site, with a Ki value of about 7 /AM(OBrian and

32

M. R . O’BRIAN A N D R . J. MAIER

- -

HZ

H2ase

0 -

I

I

CN-

Ascorbate

HQNO

b(560nm)

: I

-

co I

1 c-a-

a3-

I

O2

I I

Lbt (0,558nm) I

Ascorbate

I I

*0 2

I

CN -

co

FIG. 5. Proposed electron-transport pathway from H2to 0 2 in membranes from hydrogenase-derepressedcells of Rhizobiumjaponicum. Reproduced from OBrian and Maier (1982).

Maier, 1985b), indicating that the branching point of the electron-transport system occurs after ubiquinone. Cytochrome spectra of membranes taken immediately after the addition of substrate shows that cytochromes c and aa3 are reduced whereas the b-type cytochromes remain oxidized (O’Brian and Maier, unpublished observations), implying that cytochromes c and aa3 are part of a different branch from the 6-type cytochromes. Also, mutants of R. japonicum lacking cytochromes c and aa3 are able to grow well in culture (El Mokadem and Keister, 1982), suggesting that oxidases are present that do not oxidize cytochrome c, and that cytochromes c and aa3 are functionally distinct from the other cytochromes. The data obtained thus far indicate that the branching point of the electron-transport system in membranes of free-living R.japonicum is at the level of cytochrome b, with one branch terminating with cytochrome aa3 and the other with cytochrome o (Fig. 5). 3. Bacteroids

The electron-transport system of bacteroids also contains b- and c-type cytochromes, as well as numerous CO-reactive haemoproteins that may be terminal oxidases (Appleby, 1969a). Despite the excellent work on the electron-transport system in R. japonicum bacteroids (Appleby, 1969a, 1978; Appleby et af.,1975; Bergersen and Turner, 1975, 1980), not a single terminal oxidase has been identified by photochemical action spectra. The failure to obtain these spectra is probably caused by the high affinity of the terminal oxidases for 0 2 , making the binding of CO to the oxidase impossible in the presence of 0 2 . The acquisition of very low free 0 2 concentrations would require the presence of leghaemoglobin or myoglobin, which have their own CO-binding properties. Studies using whole bacteroids show that b- and c-type cytochromes are reduced faster in the presence of Hz plus endogenous substrates than by

33

HYDROGEN METABOLISM IN RHIZOBIUM

endogenoussubstrates alone (Eisbrenner and Evans, 1982a). Also, bacteroids preincubated anaerobically for one hour in the presence of the plastoquinone antagonist dibromothymoquinone show diminished rates of H2 uptake (Eisbrenner and Evans, 1982a); this was interpreted to mean that ubiquinone is involved in H2 oxidation. Cytochrome spectra of membranes from bacteroids show that H2 reduces band c-type cytochromes, but no cytochrome uu3 is observed in H2- or dithionite-reduced samples (Fig. 6; O’Brian and Maier, 1983). In addition, a distinct trough is observed at 456 nm of the H2-reduced difference spectrum which is indicative of a flavoprotein. The flavoprotein inhibitor atebrin inhibits Hz uptake and exogenous cytochrome c oxidation with similar Ki values, indicating that atebrin inhibits the oxidation of both substrates at the same site (O’Brian and Maier, 1983). Since cytochrome c has a high midpoint potential (+260 mV), the flavoprotein must also have a high redox potential.

C

I 300

1

I

I

I

420

460

500

540

I 500

I

620

I 660

Wavelength (nml

FIG. 6.Substrate-reducedminus 02-oxidizedabsorption spectra of membranes from Rhizobium japonicum bacteroids. A, dithionite; B, H,; C, argon. Reproduced from OBrian and Maier (1983).

34

M. R. O’BRIAN AND R . J . MAIER

Appleby (1978) made essentially the same observation for the cytochrome c oxidase activity in a “soluble” fraction of R . japonicum cells ruptured in low ionic strength buffer. Although atebrin inhibits H2 uptake in bacteroid membranes, it does not inhibit H2-dependent cytochrome reduction, indicating that atebrin is located on the oxygen side of the cytochromes (O’Brian and Maier, 1983); this is consistent with the assertion that the flavoprotein has a high redox potential. The cytochrome c oxidase activity of R . japonicum membranes is also inhibited by cyanide (Appleby, 1978; O’Brian, 1984), and the H2-dependent flavoprotein reduction is inhibited by this ligand (O’Brian and Maier, 1983), suggesting that the flavoprotein is an 02-binding component; this observation, as well as the implications that the flavoprotein has a high redox potential, indicates that the flavoprotein is an oxidase. Appleby (1975) speculated that the flavin and metal prosthetic groups may act together to transfer the four electrons required for the complete reduction of molecular oxygen to water, as occurs with the heme and copper moieties of the mitochondria1 cytochrome c oxidase. The cytochrome c oxidase activity of the flavoprotein is not inhibited by CO (Appleby, 1978) and therefore may be the terminal oxidase of the uncoupled, CO-insensitive electron-transport pathway described by Bergersen and Turner (1975). If this is true, then the involvement of the flavoprotein in H2 uptake (O’Brian and Maier, 1983) indicates that hydrogen oxidation can indeed be an 02-protective mechanism by consuming O2via the ATP-uncoupled system. Hydrogen oxidation in R . japonicum bacteroid membranes involves three CN--reactive components, as discerned by three distinct phases of CNinhibition of H2 uptake by membranes (O’Brian and Maier, 1983). The three phases correspond to CN- binding sites with Ki values of about 1, 10 and 90 PM. About 20% of the H2 uptake activity of membranes persists in the presence of 1 mM cyanide, similar to that observed for endogenous respiration in whole bacteroids (Appleby, 1969a). The component with the highest affinity is most likely the flavoprotein based on the CN- sensitivity of cytochrome c oxidase activity in membranes (OBrian, 1984). The other two CN- binding sites have not been assigned to their respective components, but H2-reduced difference spectra of membranes in the presence of cyanide show that some b- and c-type cytochrome cannot be reduced in the presence of this ligand. Furthermore, CO spectra with H2 or dithionite as the reductant show that CO-reactive b- and c-type cytochromes are present in the membrane, and are reducible by H2 (OBrian and Maier, 1983). The CN-- and CO-reactive cytochrome c was identified as cytochrome c-552 by Appleby (1 969a), which is possibly involved in the electron-transport pathway coupled to ATP synthesis (Appleby, 1984). The CO- and CN--reactive b-type cytochrome was tentatively identified as cytochrome 0,which has been reported to be present in bacteroids in one study (Keister et al., 1983), and absent from another (Appleby, 1969a).

HYDROGEN METABOLISM IN RHIZOEIUM

35

The presence of multiple oxidases in R . japonicum seems to allow the regulation of specific pathways of electron transport by 0 2 (Appleby et al., 1975; Bergersen and Turner, 1975, 1980). Under low 0 2 concentrations, electron transport is coupled to ATP synthesis with concomitant consumption of 0 2 ; this branch of the electron-transport system must be terminated by an oxidase with an extremely high affinity for 0 2 . Under high 0 2 conditions, respiration is not coupled to ATP synthesis, and electron transport may act as an 0 2 protection mechanism for nitrogenase. The involvement of multiple oxidases in H2 metabolism is consistent with the data showing that H2 oxidation provides ATP to the bacteroid, and can protect the 02-labile nitrogenase from inactivation. Unfortunately, it is not known whether cytochrome P-450 plays a role in H2 oxidation. This cytochrome, although not proven to be an oxidase, is almost certainly involved in the electron-transport system coupled to ATP synthesis at low O2 concentrations (Appleby et a/., 1975; Bergersen and Turner, 1980). It would, therefore, be extremely beneficial to know if H2 is oxidized by cytochrome P-450 in order to assess more accurately the value of H2 oxidation to Nz fixation.

4 . Is a Unique b-type Cytochrome Involved in Hydrogen Oxidation?

Evans and coworkers (Eisbrenner and Evans, 1982b; Eisbrenner et al., 1982) observed that R . juponicum cells grown chemoautotrophically contain significantly more 6-type cytochrome than do cells grown heterotrophically, and concluded that the additional b-type cytochrome is a low-potential component unique to the H2 oxidation system. They also claimed that this cytochrome, called “component 559-H2”by these authors, is present in bacteroids, although at a much lower concentration than in cells grown autotrophically. O’Brian and Maier (1985a) found that strains of R . japonicum that express hydrogenase constitutively also express more b-type cytochrome, but the additional component in these cells is cytochrome 0, which is not unique to H2 oxidation, and does not have a low redox potential. Several arguments can be marshalled against the conclusion that a lowpotential cytochrome b, solely reducible by H2, is involved in H2 oxidation. Furthermore, some of the kinetic data presented to argue the case for component 559-H2 (Eisbrenner and Evans, 1982b; Eisbrenner et al., 1982) are actually more consistent with the 6-type cytochrome in question being an 02binding protein with a high redox potential, as is cytochrome 0.Some of the arguments are as follows: (i) O’Brian and Maier (1985a) found that cytochrome o is reduced more slowly than the other 6-type cytochrome in the membrane, and that cytochrome o can be distinguished from the other 6-type cytochrome in the

36

M. R. O'BRIAN A N D R . J. MAIER

membrane based on these rate differences. The slower rate of cytochrome o reduction is characteristic of an 02-binding cytochrome since it remains oxidized in the presence of 0 2 , whilst the other cytochromes are reduced. It was also observed that cytochrome o will be reduced faster by a good reductant than by a poorer one, as is expected for any cytochrome. Eisbrenner et al. (1982) also observed that some of the b-type cytochrome in chemoautotrophically-grown cells is reduced more slowly than the other cytochromes using either endogenous substrate or endogenous substrate plus Hz as the reductant(s). They found that H2 plus endogenous substrate facilitated the rate of the slowly reduced cytochrome b compared to endogenous substrate alone, and thus concluded that there is a b-type cytochrome unique to the H2 oxidizing electron-transport system. This conclusion is not correct, however, and the data only show that the addition of a good substrate will increase the rate of cytochrome reduction. The rate of 02-dependent H2 oxidation in chemolithotrophically-grown cells is extremely fast ( > 17,000 nmol h- (mg protein)-'), and the addition of HZ to these cells increases the overall respiration rate more than eightfold (Eisbrenner et af., 1982). It is important to note that the addition of H2 increases the reduction rate of the total b- and ctype cytochrome in chemolithotrophically-grown cells (Eisbrenner et af., 1982), and thus selecting a particular cytochrome as being unique to the H2 oxidation system is arbitrary and unwarranted. (ii) The cytochrome spectra of chemolithotrophic cells presented by the Evans group shows that the H2 reduces more c-type, as well as b-type, cytochrome than does succinate (Fig. 2 of Eisbrenner and Evans, 1982b). On comparing these spectra, there appears to be no basis for concluding that only a b-type cytochrome is unique to H2 oxidation. Since succinate is a very good reductant (Appleby, 1969a,b; O'Brian and Maier, 1982), a reasonable and simple explanation for the lack of succinate-reduced cytochromes is that the spectrum was taken only 20 seconds after the addition of succinate. (iii) Eisbrenner and Evans (1982b) quantified component 559-H2by taking Hz-reduced minus oxidized difference spectra of cells in the presence of cyanide. Although the rationale for adding cyanide was not stated, it is clear that what was actually being measured, at least in some of the spectra shown (Fig. 2 of Eisbrenner and Evans, 1982b, for example), was total b-type cytochrome, and not just component 559-H2. By this quantitation method, Eisbrenner and Evans (1982b) correlated H2 uptake activity with component 559-H2 from cells grown symbiotically, heterotrophically or chemolithotrophically, and in Hup- strains. They reported a correlation coefficient of 0.890.98 and concluded that component 559-H2 is closely related to hydrogenase function or structure, and is perhaps associated with the enzyme per se. Since cytochrome expression in Rhizobium, and in other bacteria, has been shown repeatedly to be dependent on growth conditions, comparison of cells grown

'

HYDROGEN METABOLISM IN RHIZOBIUM

31

under drastically different conditions is not meaningful. The correlation data also extends to Hup- strains; since component 559-H2 is, by definition, reducible only by H2, it cannot be valid to examine the extent of Hz-dependent cytochrome reduction in a strain that does not oxidize H2. Also, there is evidence that hydrogenase and cytochrome o are regulated by common factors (OBrian and Maier, 1985a), and thus a positive correlation would not be surprising. Such a correlation, however, would not warrant the conclusion that cytochrome o is structurally or functionally related to the hydrogenase enzyme in any direct way. (iv) The claim that component 559-H2 has a low redox potential, and receives electrons from hydrogenase, is based on the conclusion that this cytochrome is unique to the H2 oxidation system (Eisbrenner and Evans, 1982b). However, there are no precedents which mandate that component 559-H2 has a low potential in order to participate solely in H2 oxidation. Also, the kinetic data show that component 559-H2 is reduced more slowly than the other cytochromes (Eisbrenner et al., 1982), whereas a low potential cytochrome is expected to be more reduced when in a steady state than the components with higher potentials. Therefore, the kinetic behaviour of component 559-H2 is similar to that of cytochrome o, which is not a lowpotential cytochrome. It has been found that H2 does not reduce more 6-type cytochrome than does succinate or NADH in H2-derepressed wild type cells or in heterotrophially-grown cells that express hydrogenase constitutively (0’Brian and Maier, 1982, 1985a), showing that component 559-H~is not present in these cells. It has also been shown that there is no 6-type cytochrome which mediates electron flow from hydrogenase to ubiquinone in hydrogenase-constitutive cells (O’Brian and Maier, 1985b), ruling out a low-potential cytochromeb that accepts electrons directly from hydrogenase. Furthermore, hydrogenaseconstitutive strains express significantly more 6-type cytochrome than does the wild type, and this component is cytochrome 0,not component 559-H2. Although hydrogenase and cytochrome o may be regulated by common factors, cytochrome o is reducible by NADH and succinate, as well as by H2, and participates in substrate oxidation (O’Brian and Maier, 1985a).Cells of P . denitrificans grown autotrophically with H2 also express more cytochrome o than do heterotrophically-grown cells (Porte and Vignais, 1980), supporting the assertion that hydrogenase and cytochrome o are regulated by common factors. Despite the data showing component 559-H2 to be absent from H2oxidizing R .japonicum, the Evans group insists that this cytochrome is present in bacteroids and chemoautotrophically-grown cells, and in membranes of chemolithotrophic cells (Lambert et al., 1985b). Figure 7 shows difference spectra of wild-type bacteroid membranes using H2 or succinate as the

38

M. R. O'BRIAN AND R. J. MAIER

551.5

FIG. 7. Comparison of H2- and succinate-reduced cytochrornes in bacteroid membranes. The absorption spectra are substrate-reduced minus 02-oxidized mernbrane samples.

reductant. It is clear from these spectra that H2 reduces no more cytochrome b than does succinate, showing that component 559-Hz is not present in bacteroid membranes.

V. Genetics A. MUTANTS

The generation of mutants to study a complex and highly regulated system such as H2 oxidation is obviously desirable, but the acquisition of such strains is diffcult when the trait of interest in only expressed symbiotically, and screening mutants is cumbersome, and often impossible. Fortunately, conditions have been found in which R. japonicum expresses hydrogenase activity in free-living culture (Maier et al., 1978a), and thus mutants with lesions affecting Hz metabolism have been obtained. Hup- mutants were first selected by enriching a bacteroid suspension for cells unable to reduce PMS (phenazine methosulphonate) with HZ, and then screening for colonies that failed to reduce triphenyltetrazolium chloride in the presence of H2 (Maier et al., 1978b). Hup- mutants obtained from this screening cannot oxidize HZ

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39

symbiotically or in free-living culture, and nodules formed from these mutants evolve H2, whereas the wild type does not. The triphenyltetrazolium chloride screening method has been used to isolate Hup- mutants of mungbean and urd bean Rhizobium (Pahwa and Dogra, 1981, 1983). Maier (1981) isolated over 100 ethyl methane sulphonate (EMS)-induced mutants of a USDA 122 derivative (SR) based on the inability of these mutants to grow autotrophically with H2 and C02 as the sole energy and carbon sources, respectively. The phenotypes of these mutants fall into various classes. About 70% of the mutants cannot oxidize H2, but two strains, SR106 and SR166, do have hydrogenase activity in the presence of artificial electron acceptors such as methylene blue or PMS. These mutants presumably synthesize an active hydrogenase enzyme, but may be missing some other component in the electron-transport system to 0 2 . Preliminary experiments indicate that, as bacteroids, these strains synthesize b- and c-type cytochromes similar to the wild type (M.R. O’Brian and R.J. Maier, unpublished observations), but some other component not discernible by absorption spectroscopy of fractionated cells could be missing. Four strains were isolated that have hydrogenase activity, but have no C02 uptake ability. Two of these mutants lack ribulose 1$bisphosphate (RuBP) carboxylase activity; this enzyme is the primary carboxylase functioning during autotrophic growth of R.juponicum (Lepo et al., 1980). The two carbon-fixation mutants that do not have RuBP carboxylase activity may be deficient in another enzyme in the Calvin cycle. Hydrogenase activity can be reconstituted when two EMS-induced Hupmutants are mixed together (Maier and Mutaftschiev, 1982). Soluble fractions of the Hup- mutants SRI 18 and SR146 were incubated together under 100% H2, and methylene blue-dependent HZ uptake by this mixture could be observed within two hours, with maximal activity seen after 10 hours. The kinetics of H2 uptake during the incubation period corresponded with an increase in turbidity over the same period; electron micrographs of the reconstituted system showed that membrane vesicles formed during the incubation period. Furthermore, the reconstituted vesicles could be sedimented, and the resulting pellet had Ordependent hydrogenase activity which was sensitive to cyanide. It was later found that hydrogenase activity of S R l l 8 could be partially restored by adding a chloroform-methanol extract of the wild type to crude extracts of the mutant (T.-Y. Wong and R.J. Maier, unpublished observations). It is not known yet whether a lipid or some other component of the extract is responsible for complementing the mutant. Moshiri et al. (1983) found two Hup- mutants that also lack N2 fixation ability. One of the Hup- Nif- mutants, SR143, produces green nodules, and has much less leghaemoglobin than nodules produced by the wild type. The hydrogenase activity of this mutant can be partially restored by the addition of

40

M. R. O'BRIAN AND R. J . MAIER

heme plus ATP. Apparently, SR143 makes apocytochromes, but is deficient in some aspect of heme biosynthesis. The addition of heme to crude extracts or membranes of E. coli (Haddock and Schairer, 1973) and Staphylococcus aureus (Lascelles, 1979) heme-negative mutants also restores respiratory activity in those bacteria. The failure of SR143 nodules to synthesize leghaemoglobin supports the current notion that heme for leghaemoglobin is synthesized by the bacteria rather than by the plant host (Cutting and Schulman, 1969; Avissar and Nadler, 1978). The Nif- Hup- mutant SR139 produces pink nodules, and is therefore a different mutant from SR143. Mutant SRl39 has no methylene blue-dependent Hz uptake activity, suggesting that it does not synthesize an active hydrogenase enzyme. Furthermore, nitrogenase activity in SR139 cannot be restored by the addition of components I or I1 of nitrogenase, showing that this mutant lacks both of the nitrogenase components. Since SR139 reverts to the wild type at a high frequency, the Nif- Hup- phenotype must be due to a single genetic lesion that affects the expression of three enzymes. The mutated gene may be regulatory, or it may be a structural gene that codes for a factor common to the enzymes, such as iron-sulphur clusters. It is unlikely, however, that SRl39 has a problem with general iron or sulphur metabolism since these cells grow normally under heterotrophic conditions. A 23 kb DNA fragment has been isolated that can complement SR139 (Hom et al., 1985, see Section V.B), and at least one other gene involved in the expression of hydrogenase is on this DNA segment. The gene mutated in SR139 may be unique to Nif and Hup expression since it is located near other hup genes. Several mutants have been isolated that affect the expression of hydrogenase by 02; these mutants are discussed in detail in Section 11. One class of mutants expresses hydrogenase activity in nodules, but can only be derepressed for activity in liquid culture if the 0 2 tension is reduced to 0.4%.The wild-type strain can be derepressed for activity under 2% 02, and thus these mutants seem to be hypersensitive to 0 2 repression. The very low 0 2 tension inside the nodule would explain the Hup+ phenotype of these mutants as bacteroids. The second class of mutants are insensitive to 0 2 repression, and express hydrogenase activity under atmospheric conditions. These mutants have greater methylene blue- and 02-dependent hydrogenase activities as bacteroids and free-living cells when compared with the wild type. Twodimensional polyacrylamide gels show that the 02-insensitive (Hupc) mutants synthesize at least six peptides not found in the wild-type parent strain. B. MOLECULAR GENETICS

1. Genetic Techniques Used to Study Rhizobium Rhizobium has been refractory to genetic analyses and manipulations using

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standard techniques developed in E. coli. There have been no transducing phages isolated that can be used to package foreign DNA and infect Rhizobium, and although sphaeroplasts of R . japonicum have been described (Child and Sietsma, 1975; Berry and Atherly, 1984), it is difficult to transform Rhizobium efficiently. Furthermore, the standard cloning vectors used in E. coli do not replicate in Rhizohium, and finally, R . japonicum, R . lupini and Rhizohium spp. grow slowly, making genetic manipulations extremely tedious in these organisms. Despite these drawbacks, several ingenious methods have been devised to transfer DNA into Rhizobium and to create mutants using transposon and site-directed mutageneses. Plasmid RK2 is a broad host range plasmid that can be mobilized into, and replicated in, Rhizobium and other Gram-negative bacteria, but its large size (56 kb) makes it an impractical cloning vector. Ditta et al. (1980) developed a cloning system that separates the transfer and replication functions of RK2 onto separate plasmids. The actual cloning vehicle, pRK290, is 20 kb in size and contains the replication functions. The plasmid pRK20 13 contains the RK2 transfer genes cloned onto a ColEl replicon (thus it replicates in E. coli), and serves to complement pRK290 in trans for mobilization. This system can be used to mobilize relatively large DNA fragments, cloned into pRK290, into Rhizobium via conjugation. Friedman et al. (1982) refined this technique by cloning the lambda phage cos site into pRK290, making it possible to ligate 15-30 kb fragments into this cosmid vector (pLAFRl), package it into lambda phage heads and infect E. coli. The greater efficiency of DNA transfer by transduction, compared with transformation, allows the construction of much larger gene banks, and the inserts of these banks are approximately uniform in size. Ruvkun and Ausubel (1981) used pRK290 to develop a method for sitedirected mutagenesis. A Tn5-mutated nif region of R . meliloti borne on pRK290 was introduced into a R . meliloti wild-type strain. Recombination of the plasrnid-borne n i j region::Tn5 DNA with the chromosome results in a Nif- mutant if the Tn5 is inserted into a DNA region required for the expression of nitrogenase. This technique has been used to study hup genes in R.japonicum (Haugland et al., 1984). Simon et al. (1983a) modified an E. coli strain and several E. coli plasmids such as pBR325, pACYC177 and pACYC184 so that the vectors can be mobilized into Gram-negative bacteria as a means of delivering Tn5, or some other piece of foreign DNA, into the cell. This was achieved by integrating the transfer genes of plasmid RP4 into the chromosome of E. coli strain S49-20, and cloning the RP4 “mob” site into the E. coli vector. The mob site is believed to include the origin of transfer (ori T) and the recognition site for the transfer gene products. Since the vectors cannot replicate in Rhizobium, the introduced DNA can only be expressed if it is incorporated into the genome by recombination, or by transposition in the

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M. R. O'BRIAN AND R. J. MAIER

case of Tn5. It is possible, therefore, to create a population of Tn5 mutants in Rhizobium, or do site-directed mutagenesis experiments by recombining a mutated gene into the genome of a wild-type recipient. If the inserted gene is mutated by a transposon, selection for the recombinants by antibiotic resistance is straightforward.

2. Hup Genes on Indigenous Plasmids Since nif and nod genes of several Rhizobium species are found on plasmids (Nuti et al., 1979; Krol et al., 1981; Hombrecher et al., 1981; Hooykaas et al., 1981; Prakash et al., 1981), it is reasonable guess that hydrogenase genes may be on a plasmid as well. Spontaneous hydrogen oxidation mutants (Hox-) of Alcaligenes eutrophus strain TF931 lack plasmids, but they can recover the ability to express hydrogenase activity and grow autotrophically when mated with the wild-type strain HI6 (Friedrich et al., 1981b). Transfer of hydrogenase activity to the mutants occurs in the absence of a mobilizing plasmid such as RP4. Strain HI6 and the transconjugants of the strain TF93 mutants contain a large plasmid with a molecular weight of 270- lo6. These data show that A . eutrophus harbours a self-transmissible plasmid that contains genes necessary for the expression of hydrogenase activity. Further studies by the same group (Hogrefe et al., 1984; Friedrich et al., 1984) showed that genetic determinants of the membrane-bound and soluble hydrogenases are plasmidborne, but hydrogenase determinants are located on the chromosome as well. Brewin et al. (1980) investigated the possibility that hydrogen uptake genes (Hup)are on a plasmid in R . leguminosarum.Since nodgenes are known to be found on a plasmid, they transferred the nod genes of a Nod+ Hup+ strain into the Nod- Hup- strain 16015 to see if the Hup and Nod determinants were cotransferred. The Nod-containing plasmid R16JI was not self-transmissible, but could be mobilized after recombination with a transmissible R. leguminosarum plasmid. Over half of the R. leguminosarum 16015 cells that received the recombinant plasmid could nodulate peas, and all the Nod+ transconjugants had hydrogenase activity comparable to that of the wild type. They concluded that determinants of nodulation and hydrogen oxidation are genetically linked, and are cotransferred due to the Nod and Hup genes being located on the same plasmid. Attempts to locate hydrogenase genes on an indigenous plasmid in R. japonicum have not been successful, nor has any plasmid been found in a strain with substantial hydrogenase activity. Cantrell et al. (1982) looked for plasmids in Hup+ and Hup- strains, and in Hup- mutants of R . japonicum. They found that six out of eight naturally occurring Hup- strains contained plasmids, with molecular weights ranging from 44-180. lo6, and one out of seven Hup+ strains harboured a plasmid; this latter strain had very poor

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hydrogenase activity. It was also found that some Hup- mutants of the Hup+ strain SR contained multiple plasmids. The non-revertible mutants SR 1, SR2 and SR3 contain two plasmids each, whereas the revertible mutants PJ 17, PJ18 and PJ20 do not contain any plasmid. The reason for these phenomena is not known, but several possibilities exist. The hup genes may be located on a plasmid too large to be detected by the procedures used, and a mutation causes the formation of two smaller plasmids that can be discerned. Alternatively, hup DNA may be excised from the chromosome to create plasmids and a Hup- phenotype. The latter event is possible if insertion sequencesare located near the hup locus, as is found in the nifgene region of R. japonicum (Kaluza et a f . , 1985), and thus allowing chromosomal mobilization. Either possibility would explain why SRl, SR2 and SR3 are nonrevertible. Assuming these three mutants are not siblings, the similar plasmid patterns found in them imply that the event causing the mutation is not random, although a larger population of Hup- mutants needs to be examined to verify this. This study also raises the question of whether naturally occurring Hup- strains are actually mutants of Hup+ strains, and if hup genes are contained on the plasmids of Hup- strains. Hybridization experiments using isolated hup genes would directly address this question.

3. Hup Genes in Rhizobium japonicum

A gene bank of R . japonicum was constructed into the broad host range cosmid pLAFR1, and mated en mane with Hup- recipients in order to isolate DNA fragments that complement the mutants (Cantrell et al., 1983). Hup+ transconjugant colonies were selected based on their ability to reduce methylene blue in the presence of Hz and respiratory inhibitors (Haugland et al., 1983). Hup+ colonies arose at a frequency of 6 - lop3per transconjugant when the Hup- mutant PJ17nal was the recipient, but the mutant PJ 18nal could not be complemented in trans by the gene bank. Cosmids isolated from eleven of the Hup+ transconjugants of PJ 17nal contained three common EcoRl fragments of 13, 2.9 and 2.3 kb in size. Five out of six of the Hup+ transconjugants expressed hydrogenase activity as nodules, but only at 525% of the wild-type level. One of the cosmids, pHU 1, was mutagenized with Tn5 in various regions of the insert DNA, and the mutated DNA was introduced into the wild-type genome by homologous recombination to determine the hup-specific regions of the cosmid (Haugland et al., 1984). It was found that hup-specific sequences spanned 15.5 kb as determined by the Hup phenotype of the Tn5 mutants created by the homologous recombination. The mutant PJ18nal can be complemented by fragments of pHU 1 cloned into pBR325 if the entire recombinant plasmid is integrated into the chromosome, presumably by a single cross-over event. This observation is interpreted to

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M. R . O’BRIAN AND R . J . MAIER

mean that the PJ18nal mutation site occurs within one of the subcloned fragments (several EcoRl fragments of pHUl were cloned onto one plasmid in a random order), and that the mutation is dominant. Lambert et al. (1985a) isolated additional cosmids that complement a Hup- mutant other than PJ17nal. One such cosmid, pHU52, was introduced into several naturally occurring Hup- strains of R .japonicum and into strains of R . meliloti and R . leguminosarum. These transconjugants can be derepressed for hydrogenase activity in free-living culture, and thus the authors concluded that all the determinants required for the expression of Hup activity in the free-living and symbiotic states of R. japonicum are on pHU52. Although these results are exciting, we do not agree that the data show that all the Hup determinants are on pHU52, nor is there reason to believe that pHU52 does not regulate structural genes of the host, and vice versa. Cantrell et al. (1982) provided evidence that the difference between a Hup+ and HupR.Japonicum strain is due to the rearrangement of the DNA to form multiple plasmids from the genome or from larger plasmids. If this is true, then hup genes may be present in Hup- strains which can be expressed in the presence of pHU52. Hybridization experiments are needed to see if the Hup- strains contain DNA homologous to pHU52, and further characterization of the Hup- strains is necessary to determine the nature of these strains. Rhizobium meliloti strain 102F51 has Hup activity in the free-living and symbiotic states (Ruiz-Argueso et al., 1979), and thus transconjugants of this strain must possess hup genes not borne on pHU52. Finally, R. leguminosarum strain 128C53is a Hup+ strain symbiotically, and certainly contains indigenous hup genes (Brewin et al., 1980). In fact, Tn5-induced Hup- mutants of strain 128C53 can be complemented by pHUl (Kagan and Brewin, 1985); this plasmid insert has about 25 kb in common with the pHU52 insert. It is entirely possible that pHU52 contains genes that allow the expression of hup genes in free-living culture, but it is premature to assume that the Hup+ phenotype of transconjugants harbouring pHU52 is entirely due to the expression of genes on that plasmid. Hom et al. (1985) complemented the Nif- Hup- mutant SR139 with cosmids from a R.japonicum gene bank constructed in pLAFR 1. It was found that the Hup+ transconjugants of SR139 also have nitrogenase activity both symbiotically and under conditions where nitrogenase is induced in free-living cells; SRI 39 has no nitrogenase activity under either growth condition. The nitrogenase activity of the SR139 transconjugants are much higher in freeliving cells than in nodules, relative to those of the wild type, presumably due to segregational loss of the plasmid in nodules. One of the cosmids that complements SR139 (pSH22) was tested to see if it could complement other autotrophic growth mutants. Six Hup- Nif+ mutants could be complemented by pSH22, but a carbon-fixation mutant, and a Hup- Nif- mutant different

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from SR139 were not complemented by the plasmid. The ability of pSH22 to complement both a Hup- Nif- and a Hup- Nif+ mutant suggests that this cosmid harbours more than one gene essential for the expression of hydrogenase activity. This conclusion is strengthened by the fact that SRl39 is a revertible mutant (Moshiri et al., 1983), and one of the Hup- Nif+ mutants (SU306-47) is a Tn5-induced mutant, therefore each of these mutants almost certainly has a single lesion in a different region on the genome. These data also suggest that the niflhup gene mutated in SRl39 and the hup gene mutated in SU306-47 are located on different operons, or the hup gene is located downstream from the nif/hup gene on the same operon since the polar mutation in SU306-47 does not result in a Nif- phenotype. It should also be noted that pSH22 does not complement seven Hup- Nif+ mutants tested, thus there must be genes involved in Hup expression other than the ones discussed above. It is not known yet where on pSH22 the specific hup genes are located, but it is known that a 13.2 kb EcoRl fragment of pSH22 can complement SRl39 (P. Novak and R.J. Maier, unpublished observations). 4. Hup Genes of Rhizobium leguminosarum

As stated above, hup genes of R. leguminosarum are carried on an indigenous plasmid along with nod and nif genes. This plasmid, pRL6J1, is nontransmissible, but it can mobilize into other cells by recombining with mobilizable R.leguminosarum plasmids such as pVWJ3 1 or pVWJ51 (Brewin et al., 1980, 1982). When the recombinant plasmid pIJ1008 (recombinant of pVW5JI and pRL6JI) is introduced into Nif+ Nod+ Hup- strains, the resulting Hup+ transconjugants have superior symbiotic properties than the parent strain, as judged by plant dry weight, nitrogen content, leaf area and nitrogen concentration (DeJong et a/., 1982). One of the transconjugants, strain 3960, has better symbiotic properties than the Hup+ strain 128C53, from which the hup determinants in pRL6JI were derived. It was concluded that the increased effectiveness of strains harbouring pIJ1008 may be due to the ability of these strains to recycle the H2 evolved by nitrogenase via hydrogenase. They do not rule out, however, the possibility that some other determinant on pIJ 1008 confers superior symbiotic qualities on the transconjugants. Indeed, other experiments described below suggest that determinants other than Hup may be responsible for the symbiotic effectiveness of the Hup+ transconjugants of Hup- strains. Kagan and Brewin (1985) mutagenized a Hup+ R.leguminosarum strain to create Hup- mutants using a technique that minimizes the number of mutants that have to be screened on plants. The Hup+ strain 128C53was mutagenized with Tn5-mob, a Tn5 derivative that contains the mobilization site of plasmid RP4 (Simon et al., 1983b). The Tn5-mob insertion allows a plasmid to be

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M. R. O’BRIAN AND R. J. MAIER

mobilized, thus plasmid-linked mutations can be enriched for by mating the Tn5-mob mutants with a plasmidless recipient and selecting for kanamycinresistant (due to Tn5) transconjugants. These recipients are screened further for the presence of the symbiotic plasmid pRL6JI (strain l28C53 contains multiple plasmids) by scoring for colonies that cross-react with an antibody made against a protein encoded by pRL6JI that is expressed in free-living culture. Using this selection procedure, eight mutants were isolated that have no hydrogenase activity symbiotically. All eight mutants can be complemented with cosmid-borne R. japonicum genes that can complement a HupR.japonicum mutant (see Cantrell et al., 1983). Surprisingly, this R .japonicum DNA fragment has only weak homology with strain 128C53, whereas it is strongly homologous with several other Hup+ R. leguminosarum strains (Nelson et al., 1985, see below). Cunningham et al. (1985) used the Hup- R. leguminosarum mutants described above to study the effect of the Hup phenotype on nitrogen fixation in pea nodules. Three pea cultivars were each inoculated with one of six Hupmutants, the Hup+ parent strain, or the Hup+ parent strain with a Tn5-mob insertion on pRL6JI. It was found that, for each cultivar, at least one of the six Hup- mutants fixed as much N2 as the wild-type strain. Since previous experiments (DeJong et al., 1982) suggested that pRL6JI confers superior symbiotic properties on nodules from strains harbouring this plasmid, it was concluded that some trait encoded by pRL6J1, other than the Hup+ phenotype, is responsible for the improved symbiotic performance. At the time of writing this review, these results are only present in abstract form, and thus the data are not available for analysis. It would be interesting to examine the Hup- mutants that did not fix as much N2 as the wild type to see if these differences are significant. Since at least five of the six Hup- mutants have the Tn5-mob insertion in different sites on the plasmid (Kagan and Brewin, 1985) and have different N2-fixing capabilities, it seems that the mutation of some hup genes may affect N2 fixation despite the observation that the Hupphenotype is not always detrimental. The genetic analyses of R. leguminosarum and R.japonicum are consistent with the physiological data which show that the hydrogen oxidation systems of the two bacteria are different in several respects. There is a considerable amount of data indicating that Hz oxidation does not increase Nz fixation ability in R.leguminosarum (Cunningham et al., 1985; Nelson, 1983; Truelsen and Wyndaele, 1984),whereas most of the data for R.japonicum indicates the Hup+ phenotype confers a symbiotic advantage on the strain (Albrecht et al., 1979; Evans et al., 1983). The most obvious explanation is that the R. japonicum hydrogenase is more efficient at recycling the H2 evolved by nitrogenase than is the R. leguminosarum hydrogenase (Carter et al., 1978; Truelsen and Wyndaele, 1984). The variability in the H2 oxidation systems is

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also reflected in the level of homology of the hup-specific DNA in R. leguminosarum and R.japonicum. Nelson et al. (1985) found that hup-specific DNA from R. japonicum and R. leguminosarum hybridize weakly or not at all with total and plasmid DNA from four out of twelve R. leguminosarum Hup’ strains examined. These observations not only reflect intergenic variability of hup-specific DNA, but also differences among R. leguminosarum strains. Rhizobium leguminosarum strain 128C53, the strain on which most of the genetic and mutant studies have been performed, shows only weak hybridization with hup genes from R.japonicum and from R. leguminosarum strain BIO. Interestingly, three of the four strains that have little or no homology to the hup DNA probes are known to have H2 oxidation systems coupled to ATP synthesis; most Hup+ R. leguminosarum strains seem to have Hz uptake activities not coupled to ATP synthesis (Nelson and Salminen, 1982). This is ironic since the R.japonicum strain from which the hup probes were derived has a H2 oxidation system that is coupled to ATP synthesis (Emerich et al., 1979). It is clear from these studies and from those discussed above that the regulation of hup genes in Rhizobiurn is extremely complex, and is controlled by many factors, including the host, of which almost nothing is known.

VI. Acknowledgements Work from the laboratory of Robert Maier has been supported by grants from the United States Department of Agriculture and from Allied Corporation. The authors thank Dr Robert Burris and Dr Daniel Arp for permission to reproduce their data. Thanks are also extended to Patricia Novak and Farhad Moshiri for their help with this manuscript. REFERENCES

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The Physiology and Biochemistry of Pili WILLIAM PARANCHYCH and LAURA S. FROST Department of Biochemistry. University of Alberta. Edmonton. Alberta. Canada T6G 2H7

. . . I. Introduction . . . . . . . . . I1. Nomenclature . . . . . . 111. Classification . . . . . . . . . A . Criteria for classification . . . . . . . . . B. Morphology . . . . . . . . C . Function and biochemical properties . . . . IV . High-resolution studies on pilus structure . . . . V . Organization and expression of pilin genes A . Conjugative pili . . . . . . . . B. Type 1 pili . . . . . . . . C . Pili designated Pap . . . . . . . D . Pili designated CFA/I and CFA/II (CSl, CS2 and CS3) E. Pili designated K88 and K99 . . . . . F. Pili designated NMePhe . . . . . . . . VI . Structure-function relationships of pili proteins . A . Conjugative pili . . . . . . . . . . . . B . Adhesive pili of Escherichia coli . C . Pili designated NMePhe . . . . . . VII . Acknowledgements . . . . . . . . References . . . . . . . . .

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I. Introduction Bacterial “fimbriae” or “pili” are thin (2-1 2 nm diameter) non-flagellar protein filaments found on the surfaces of many types of bacteria . They are variable in length (0.5-10 pm) and the number per cell ranges from one or two to several hundred . Most functions attributed to pili may be ascribed to their adhesive properties . These allow them to bind to other bacteria, bacteriophages. mammalian cells and inert surfaces. Piliated bacteria that adhere to ADVANCES IN MICROBIAL PHYSIOLOGY. VOL . 29 ISBN 0-12-0277294

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mammalian cells are often more pathogenic than their non-piliated counterpart because the pili enable them to become anchored to the host tissue and resist elimination by body fluids. Numerous reviews have already dealt with various aspects of the structure and function of pili (Brinton, 1965, 1967, 1971, 1977; Ottow, 1975; Tomoeda et al., 1975; Pearce and Buchanan, 1980; Gaastra and de Graaf, 1982; Jones and Isaacson, 1983; Klemm, 1985; Mooi and de Graaf, 1985). This article will attempt to update these reviews and focus on the structure and function of pili in light of recent advances employing biochemical, immunological and genetic approaches.

11. Nomenclature

Since their discovery by Anderson ( I 949) and Houwink (1949), bacterial nonflagellar filamentous appendages have been referred to as threads, filaments, bristles, cilia, fibrillae, fuzz, colonization factor antigen, adhesins, fimbriae and pili. The designation “fimbriae” (Latin for thread or fibre) was introduced by Duguid and his coworkers in about 1955 (Duguid el a/., 1955), while Brinton (1959) introduced the name “pili” (Latin for hair-like structure) four years later. Until 1964, pili were distinguished primarily on the basis of morphology, but Crawford and Gesteland (1964) found by electron microscopy that the RNA “male-specific” phage R17 adsorbed selectively to a new kind of pilus, distinct from the more numerous “type 1” (or “common”) pili. Brinton et al. (1964) confirmed this observation and named the filaments F pili after demonstrating that these filaments were required for genetic transfer as well as F-specific phage susceptibility, and that the genes for their formation were encoded by the F plasmid. Ottow (1975) subsequently suggested that the term “pili” be used for conjugative filaments encoded by self-transmissible plasmids, and the term “fimbriae” be reserved for non-conjugative filaments, many of which promote adherence to mammalian tissues. Some of the latter (K88 and K99) were initially thought to be capsular antigens (Orskov et al., 1961, 1979, while others (CFA/I, CFA/II) were described as “colonization factor antigens” (Evans et al., 1975; Evans and Evans, 1978). Structures designated CFA/II were subsequently found to consist of three different types of pili that were named “coli surface antigens” 1,2 and 3 (CSI, CS2 and CS3; Smyth, 1982). Brinton (1965) originally distinguished six types (type I-V and F) of pili, while Duguid et al. (1966) identified seven types (types 1-6 and F). However, these classifications were inconsistent with various types of pili described by other workers (Bradley, 1966; Fuerst and Hayward, 1969; Mayer, 1971;

PHYSIOLOGY AND BIOCHEMISTRY OF PILI

55

Schmidt, 1971; Weiss, 1971), and now only the type 1 classification is still in common use. One type of pilus was recently named Pap (pyelonephritis associated pili; Normark et al., 1983) because it was found to be prevalent among uropathogenic strains of Escherichia coli. These same pili were named P fimbriae by Kallenius et al. (1981) when they discovered that the adhesins bind to the P blood-group substance which contains the active receptor disaccharide a-D-Gal-( 1+4)-b-~-Gal. O’Hanley et al. (1985) later named these adhesins Gal-Gal pili because the synthetic analogue of this disaccharide inhibits haemagglutination by these pili. A number of other pili types were named according to the strain from which they were isolated, e.g. 978P (Isaacson et al., 1977), F41 (J. A. Morris et al., 1980), PAK (from Pseudomonas aeruginosa; Frost and Paranchych, 1977), GC (for “gonococcal”; Buchanan et al., 1973) among others. An attempt by Orskov and Orskov (1983) to introduce a new nomenclature based on serology of fimbrial antigens in which only F (fimbrial) designations would be used (Fl-Fn) has not been widely accepted. This nomenclature would lead to confusion between conjugative and non-conjugative pili, since the most widely studied conjugative pili are of the F type whose incompatibility groups (Inc) have been designated FI-FV (see p. 58). For purposes of simplicity, and because there is no general agreement regarding the nomenclature of bacterial non-flagellar surface filaments, the term pili is used throughout this review article to describe all non-flagellar surface appendages, including conjugative and non-conjugative types.

111. Classification A. CRITERIA FOR CLASSIFICATION

To date, there has been no general agreement on any specific classification scheme for pili. Three criteria that lend themselves to the classification of pili are: (1) morphology (e.g. thin flexible, thick flexible, rigid), (2) function (e.g. distinct from conjugative as adhesive pili), and (3) biochemical properties (e.g. type-related pili with free amino termini as distinct from those with NMePhe at the amino terminus). The three major groups discussed in this article are termed conjugative, adhesive and NMePhe pili, and electron microscope photographs of representative bacteria are shown in Fig. 1. B. MORPHOLOGY

A large number of electron microscope studies have described the morpho-

FIG. 1. Electron micrographs of representatives of the three pilus groups: conjugative, adhesive and NMePhe. The conjugative pili are represented by the F plasmid in Escherichiu coli HBl I and the pili have been labelled with R17 bacteriophage. The adhesive pili are represented by CFA/I pili found on Escherichiu coli H 10407 (Evans et ul., 1975). The NMePhe pili are represented by the multipiliated mutant Pseudomonar ueruginosu K/ZPfs (Bradley, 1966).

PHYSIOLOGY AND BIOCHEMISTRY OF PILl

57

logical characteristics of a bewildering array of pili types. Diameters in the range of 2-12 nm have been reported, primarily for pili from Gram-negative bacteria, although some types of filaments on Gram-positive organisms have also been described (Ottow, 1975). Bradley (1980a, b, 1983a, 1984)has published a useful system for classifying pili on the basis of electron-microscope appearance. He examined conjugative pili encoded by plasmids from 37 incompatibility groups, and found them to fall into three morphological classes: thin flexible (6-7 nm diam.), thick flexible (8-10 nm diam.), and rigid (8-1 1 nm diam.) (Table 1). In addition, they can be differentiated by the presence of tapered tips at the distal end of the pilus or knobs that represent a pilus-associated structure at the base of free pili derived from the cell surface. Conjugative pili also display a varying propensity for aggregation which aids in their identification. Many non-conjugative pili such as Type 1, CFA/I, 987P, CS 1, CS2 and Pap can also be classified within this system, since they all have the appearance of thin, rigid rods with diameters of about 7 nm (Gaastra and de Graaf, 1982; Levine et al., 1984; Klemm, 1985). Non-conjugative pili from Ps. aeruginosa (Frost and Paranchych, 1977), Moraxella sp. (Bovre and Froholm, 1972; Pedersen et al., 1972), Neisseria sp. (Salit, 1981; Stephens and McGee, 1981; Swanson et al., 1971) and Bacteroides nodosus (Every, 1979) all produce thin flexible pili (about 6 nm diam.); hence, this group can also be classified within the Bradley system. In contrast, the K88, F41 and CS3 pili are visualized by the electron microscope as very thin, flexible threads with diameters of around 2 nm (Levine et al., 1984; Klemm, 1985), while K99 pili are also flexible and quite thin (about 4.5 nm diam.; de Graaf et al., 1980). Thus, a fourth morphological class, called “very thin”, should be created to accommodate this group of pili, as well as those of 3-4 nm diameter that are produced by Bordetellapertussis (Blom et al., 1983) and B. bronchiseptica (Lee et al., 1986).

C . FUNCTION AND BIOCHEMICAL PROPERTIES

The simplest classification of pili on the basis of function is the division into two broad groups: “conjugative” and “adhesive” pili. The biochemical properties help to identify subpopulations of these pili.

I . Conjugative Pili Conjugative pili are generally encoded by self-transmissible plasmids, which are capable of passing a copy of their genetic material to a recipient bacterium through a process known as conjugation. In Gram-positive bacteria, conjugative pili have not been identified; the donor cell releases sex

TABLE 1. Morphological properties, bacteriophage sensitivity and mating type of conjugative pili

References

IncompatiRepresentative bility plasmid($ group

Escherichia coli 36,37,41,42,30 14,42,30 18,31,35,39 6.26 6.26 6.26 20.26 12.1 3,23,24,40,30,43

FI FII N HI I HI2 HI3 HI1 I1

F R1-19,R100-1 Folac.pED208 R27 R478 MIP233 pHH I457 R64

6,13,30

II + B

RIM

6.13

B

R16,R621a

6.13

K

R387

13.46

I5

pIE360

13,47

Z

pIE545

22.24.28

I2

R72 I

6,2 I 3 I ,27 6,21

I 17,30,33,48 6 6,16,30 4.8.10 32.38 5.6.25 19 2.3.5.29.32.61

X X

RAI R71 I b,R778b R391 R71"' pIN25 R753 R6K R48Y

N M U

N3 R446b RA3,pAR-32

W

sa

C D J cod T

V

Pilus morphology' Thick flexible Thick flexible Thick flexible Thick flexible Thick flexible Rigid Thick flexible Thin flexible Rigid Thin flexible Rigid Thin flexible Rigid Thin flexible Rigid Thin flexible Rigid Thin flexible Rigid Thin flexible Rigid Thick flexible Thick flexible Thick flexible Thick flexible Thick flexible Thick flexible Thick flexible Thick flexible +thin flexible Rigid Rigid Rigid Rigid

Approx. diam. Basal (nm)b knobs' 9 9 9 II II 11 9

6 II 6 II 6 6

+

+ + + + + + + -

+ + + -

6 10

9 9.5 10

9 9 9 9 9 5

II I1

++ + ? +

+ + + + + + +-

Bacteriophage sensitivity*

Distal Aggre- Serological pointsd gation' relatedness

-

+ + + + + + + + ++ + ?

+

+ + + + + + +

+

++ + + + + + + + + --? -

--

F1,FII

fl .RI 7,QB fl *,R I7*,QB* fl ,UA6,Folac pilHa pilHa pilHa pilHa Ifl,Ia,PR&IFS PR64FS

Universal Universal Universal Universal Universal Universal Surface preferred Universal' Surface obligatory Universal'

PR64FS

Universal'

PR64FS

Universal'

15,B.K,Zh PR64FS I I +B,B,K,15,Zh PR64FS

Universal'

HII,HI2.HII

IIJI +B,B~ WI +B,B,K~

121 12h CJ

Ifl ,PR64FS PR4JKe.12-2,X C-l ,C-2,J' fl,MS2*,J J

N t,tf-l

X

+

--

Mating typeg

Universal' Universal' Surface preferred Surface preferred Universal Universal Surface preferred' Universal Surface preferred Surface obligatory

IKe.PR4.X' M,X X

Surface obligatory Surface obligatory Surface obligatory

PR4.XK7

Surfaceobligatory

TABLE 1 (continued) Pseudomonos aeruginose 1.5.1 1.32.45 P- I II P-2 II P-3 I1 P-5 !I P-7 II P-8 11-34 P-9 11 P-I0 11 P-I 1 11 P-12 11 P-13

RPI R931,CAM RIP64 Rmsl63 Rms148 FP2 R2,TOL R91.5 RPI-I R716 pMG26

Rigid Rigid Thick flexible Thick flexible Rigid Thick flexible Rigid Rigid Rigid ? Thick flexible +thin flexible

8

9

+ ? ?

? ?

+ + +

+ + +

+ +

+

-

-

?

?

+

+

9 6

-

+

+ +

-

8.5-10

+ +

-+

?

IncCk

PRRl ,Pf3,1Ke,X,PR4 Surface obligatory Surface obligatory Surface obligatory Universal Surface preferred Universal PR4 Surface preferred X* Surface preferred Surface preferred Surface preferred Surface preferred

Pilus morphology describes pili according to 7.9.1 I . 'The pilus diameters are approximate values obtained from negatively stained EM samples. Knobs refers to any structure at the base of the pilus and includes plates and discs as well as round knobs. These knobs appear to contain pilus material. dOnly pili with visibly pointed tips are marked +. 'The aggregation of pili into long fibres or bundles is pilus-specific. (?) indicates that too few pili were present in a culture to determine this property. /Bacteriophages that either attach without infecting the cell or propagate but do not produce plaques are marked with an asterisk. Mating type has been defined ( I 5) as the ratio of mating ability on an agar surface as compared to that in a liquid culture. Universal has a ratio near I .O, surface preferred 45450, surface obligatory > 2000. In Pscudomonads, the mating types can be defined universal or surface preferred ( I I). There are three serotypes for the thin and thick pili of the 1 complex. The 1 complex plasmids permit universal mating when both thin and and thick pili are produced. 'The thin flexible pili of R485 have a wavy appearance with a pitch of 4.6 nm. IncP3 plasmids are the equivalent of In& plasmids in E. c o / i The plasmids Folar (IncFV), R71 (comg), TP224 and pPLS have been grouped into lncS (48) on the basis of phage sensitivity. This table is principally a compilation of the information in 6,7,9, I I , 13. The references are: Bradley, 1974b(1), 1975(2), 1976(3), 1978a(4), 1978b(5), 1980a(6), 1980b(7), 198Oc(8), 1981(9), 1982(10), 1983a(l I), 1983b(12), 1984(13), 1985(14); Bradleyeral., 1980(15), 1981a(16), 1981b(17), 1981c(18), 1982419), 1982b(20), 1982~(21).1983(22); Coetz.ee ef a/, 1980(23), 1982(24), 1983(25), 1985a(26), 1985b(27); Bradley and Coetzee, 1982(28); Bradley and Cohen, 197q29); Bradley and Fleming, 1983(30); Bradley and Meynell, 1978(3I); Bradley and Rutherford, 1975(32); Bradley and Whelan, 1985(33); Bradley and Williams, 1982(34); Armstrong et a/., 1980(35); Caro and Schnoss, 1966(36); Crawford and Gesteland, 1964(37); Dennison and Baumberg, 1975(38); Falkow and Baron, 1962(39); Feruichi et 01.. 1984(40); Lawn et a/., 1967(41);Lawn and Meynell. 197q42); Meynell and Lawn, 1968(43); Moms ef a/., 198q44); Olsen and Shipley, 1973(45); Tschape and Tietze, 198I(46). 1983(47); D.E.Bradley, personal communication (48).

60

W. PARANCHYCH AND L. S. FROST

pheromones which cause clumping of cells into mating aggregates where transfer takes place. This process has been reviewed by Clewell (1981). In Gram-negative bacteria, the transfer region of the plasmid invariably encodes a pilus, which recognizes a suitable recipient cell and brings it into contact with the donor cell in a poorly understood process (Achtman and Skurray, 1977; Willetts and Skurray, 1980, 1986; Willetts and Wilkins, 1984; Clark, 1985; Silverman, 1986; Ippen-Ihler and Minkley, 1986). Some broad host-range plasmids can transfer both inter- or intragenerically within the Enterobacteriaceae. A convenient method for cataloguing plasmids takes advantage of the fact that closely related plasmids are incapable of co-residing in the same cell, a property that is termed incompatibility (Inc) (Datta, 1975). Previous attempts to classify plasmids on the basis of pilus type (Lawn et ul., 1967), bacteriophage sensitivity and pilus serology (Meynell et al., 1968; Lawn and Meynell, 1970; Feilberg-Jorgansen et ul., 1982) or entry exclusion (Datta and Hedges, 1972) were unsatisfactory. As evidence accumulates, incompatibility appears to be the best scheme for classifying plasmids, since in general terms, plasmids within an incompatibility group exhibit similar plasmid size, similar organization of their transfer regions, similar pili in terms of morphology, serology and phage sensitivity, similar host ranges and similar genetic markers such as drug resistance, resistance to heavy metals, virulence determinants and degradative functions (Table 1). There are currently over 20 incompatibility groups for plasmids borne by E. coli (Shapiro, 1977; Bradley, 1980b). The IncP plasmids are found in Ps.ueruginosu and have been subdivided as PI,P2, etc. (Bradley, 1983b; Shapiro, 1977; Jacoby, 1977; Jacoby and Matthew, 1979). Pili associated with a particular incompatibility group can be further differentiated on the basis of serology, phage sensitivity and morphology into various pilus types. Conjugative pili show variable preferences for promoting bacterial mating in liquid or on solid media (Table 1). Flexible pili generally confer the “universal mating type” and promote mating in liquid media and on solid surfaces equally well, whereas rigid pili are usually associated with “surface preferred” (mating efficiency is higher on solid media than in liquid cultures) or “surface obligatory” (mating occurs only on solid media) mating types (Bradley, 1980b). This is illustrated by the Inci plasmids, where the presence of thick and thin pili allow these cells to become proficient at mating in broth and on plates, whereas the presence of thick pili alone allow mating only on solid surfaces. Thin I pili are thought to stabilize mating pairs formed by the thick, rigid I pili allowing mating in liquid media. Indeed, thin I pili can convert IncP and IncW mating pairs from a “surface obligatory” into a ‘‘L..iversal” system by stabilizing the fragile mating pairs formed by the rigid pili encoded by these plasmids (Bradley, 1984). IncF pili have been characterized on the basis of phage sensitivity patterns

PHYSIOLOGY AND BIOCHEMISTRY OF PILI

61

(Paranchych, 1975; Willetts and Maule, 1986) and serological differences (Lawn and Meynell, 1970; Finlay et al., 1984; Worobec et al., 1983) as well as, at the molecular level, by sequencing the pilin gene. The sequence of the gene encoding F pilin revealed that the pilin subunit had a molecular weight of 7200 and that the pilin gene encoded an exceptionally long signal peptide of 51 amino-acid residues (Frost et al., 1984). The sequence of four other serotypes of F-like pilins revealed that this long signal peptide was common to all of these pilus types and that differences in serology and phage sensitivity, attributable to the pilin subunit, were localized in the N- and C-terminal regions of the pilin protein (Frost et al., 1985). Furthermore, these studies showed that in all IncF pili studied, the N-terminus was acetylated and this group was important in defining the major antigenic determinant of the pilus protein (Finlay et al., 1985;Worobec et al., 1985; Frost et al., 1985, 1986). As yet, no other pilin genes from plasmids of other incompatibility groups have been sequenced and it is not clear whether this long signal sequence, the acetylated N-terminus and the variable sequences at the N- and C-termini of the pilus proteins are characteristic of conjugative pili. 2. Adhesive Pili of Escherichia coli

Pili from E. coli that promote adherence to mammalian cells are broadly classified into mannose-sensitive (MS) and mannose-resistant (M R) classes, depending on the ability of D-mannose to inhibit binding to erythrocytes and epithelial cells. They are also distinguishable by their preference for binding to intestinal or urinary epithelial cells. Some pili that are specific for intestinal epithelial cells show preferential binding to the intestinal epithelium of humans, cattle, pigs or sheep (Gaastra and de Graaf, 1982; Klemm, 1985). Mannose-sensitive binding is mediated by Type 1 pili, which have been described as non-conjugative, chromosomally encoded thin rigid rods (around 7 nm diam.) that are plentiful (lOCr500per cell), and peritrichously arranged on the surface of the cell (Brinton, 1965; Ottow, 1975). They characteristically cause agglutination of guinea-pig erythrocytes (Salit and Gotschlich, 1977) as well as attachment to yeast cells, buccal epithelial cells, and a mannose-containing urinary glycoprotein, the Tamm-Horsfall protein (Ofek and Beachey, 1980; Orskov et al., 1980a). The function of these pili is unclear, since they are often present on both pathogenic and non-pathogenic isolates of E. coli (Orskov et al., 1980b). The majority of E. coli strains causing urinary-tract infections are able to produce both MS and MR pili (Klemm et al., 1982; Salit et al., 1983; Rhen et al., 1983a). However, virulence of uropathogenic strains of E. coli has been attributed to MR (P, Pap or Gal-Gal) pili, which bind globoseries glycolipids containing the a-D-Gal-(1+4)-P-~-Galunit of the P blood-group substance

62

W. PARANCHYCH AND L. S. FROST

(Kallenius et al., 1980; Leffler and Svanborg-Eden, 1980; O’Hanley et at., 1985). Immunological studies and amino-acid sequencing on Pap and Type 1 pili from a variety of uropathogenic isolates of E. coli have shown significant amino-acid homologies at the N-terminus of these proteins (Orskov et al., 1980b; Klemm et al., 1982; Rhen et al., 1983a; Salit et a[., 1983; Baga et a/., 1984; Klemm, 1984; van Die and Bergmans, 1984; Hanley et al., 1985). However, virtually all isolates of Pap pili were serospecific, suggesting that a wide variety of homologous pili species exist among naturally occurring strains of E. coli,and that the genes encoding these pili are polymorphic with respect to antigenic determinants. A pathogenic strain of E. coli that produces MR pili which bind specifically to human intestinal epithelium was first reported by Evans et al. (1 975). These pili were initially designated “colonization factor antigen” or CFA. However, when a second, apparently similar, antigen was discovered and designated CFA/II, the name of the first antigen was changed to CFA/I (Evans and Evans, 1978). Enterotoxigenic E. coli strains positive for CFA/I and CFA/II pili are responsible for a significant number of cases of E. coli diarrhoea in humans (Gross et al., 1978; Craviato et al., 1979). The primary structure of the 15,058 Da CFA/I pilin subunit, which contains 147 amino-acid residues (Klemm, 1982), bears no significant homology with the known sequences of other pili proteins (Klemm, 1985), although CFA/I pili are morphologically similar to Type I and Pap pili (thin rigid rods of about 7 nm diam.). The CFA/II pili originally described by Evans and Evans (1978) were subsequently shown by Smyth (1982) to consist of three distinct coli surface antigens which he named CSI, CS2 and CS3. These three antigens appear to be identical to compounds 1,2, and 3 reported earlier by Craviato et al. (1979), and have molecular weights of 16,300,15,300 and 14,800, respectively (Smyth, 1982). Pili designated CSI and CS2 are morphologically similar to Type 1 pili (thin and rigid), whereas the thin flexible CS3 pili belong to the “very thin” category (Levine et al., 1984; Mullany et al., 1983). The N-terminal aminoacid sequence of the CS2 pilus protein was recently found to be almost identical to that of the CFA/I protein, although the two antigens are immunologically distinct (Klemm, 1985). A number of animal-specific MR pili adhesins ‘have also been described. Those designated K88 and 987P bind to the gastrointestinal mucosa of piglets (Orskov et al., 1961; Sellwood et al., 1975; Isaacson et al., 1978; Isaacson and Richter, 1981; Gaastra and de Graaf, 1982), whereas K99 and F41 pili are associated with gastroenteritis in calves, lambs and newborn pigs (Smith and Linggood, 1972; Orskov et a/., 1975; de Graaf and Roorda, 1982). The K88 protein exists in several antigenic variants, three of which -K88ab, K88ac and K88ad - have been characterized immunologically (Orskov et ul., 1964) and in terms of their primary structures (Gaastra et al., 1979, 1983;

PHYSIOLOGY AND BIOCHEMISTRY OF PlLl

63

Klemm, 1981). Both K88 and F41 pilin subunits are large (around 28,OOG 29,000 Da) relative to other types of pilins, and the intact pili of both are morphologically very thin (approx. 2 nm diam.) (Gaastra and de Graaf, 1982; de Graaf and Roorda, 1982; Klemm, 1985). It is not yet known whether K88 and F41 pili are related in terms of sequence homology, although de Graaf and Roorda (1982) have shown that the two pilins are serologically distinct. It would be most interesting if the two pili types should turn out to be related, since K88 pili are plasmid encoded (Orskov and Orskov, 1966; Smith and Linggood, 1972; Shipley et al., 1978), whereas F41 pili are encoded by the chromosome (de Graaf and Roorda, 1982). Enterotoxigenic strains of E. coli positive for the K99 antigen cause severe diarrhoea in calves, lambs and piglets because the antigen facilitates bacterial adhesion to the intestinal epithelium (Burrows et al., 1976; Moon et al., 1977). The molecular weight of the K99 pilin subunit is 18,400, with PI 9.5 (de Graaf et al., 1981). Some K99-positive strains of E. coli belonging to the 0-serogroups 9 and 101 also express F41 pili, but these are chromosomally encoded and the pilin subunit is an acidic protein (PI 4.6) of 29,500 Da (de Graaf and Roorda, 1982). The nucleotide sequence of K99 pilin is known (Rosendaal et al., 1984),and has revealed that the subunit protein of these thin rigid pili (about 7 nm diam.) have extensive homology in the N- and C-terminal regions with type 1 (Klemm, 1984), Pap (Baga et al., 1984) and F72 (van Die and Bergmans, 1984) pili. The porcine small intestine can be colonized by yet another enterotoxigenic E. coli strain which expresses MR pili designated 987P (Nagy et al., 1976, 1977). These pili are chromosomally encoded, and the 20,000-Da subunit has PI 3.7 (Isaacson and Richter, 1981;de Graaf and Klaasen, 1986). Morphologically, they belong to the thin rigid group of pili, but there is no homology with other known sequences of pili proteins (Klemm, 1985). 3. N-Methylphenylalanine (NMePhe) Type Pili Adhesins

Bacteria such as Pseudomonas aeruginosa (Weiss, 1971; Bradley, 1972a), Neisseriagonorrhoeae (Jephcott et al., 1971; Swanson et al., 1971;Hermodson et al., 1978), Neisseria meningitidis (Hermodson et al., 1978), Moraxella nonliquefaciens (Froholm and Sletten, 1977), Moraxella bovis (Marrs et al., 1985), Bacteroides nodosus (Every, 1979) and Vibrio cholera (J. Mekalanos, personal communication) produce thin flexible pili, around 6 nm in diameter, which have a mainly polar distribution on the cell. These pili are involved in such processes as adhesion of the bacteria to host mucosal surfaces (Punsalang and Sawyer, 1973; Brinton, 1977; Woods et al., 1980), twitching motility (Henrichsen, 1975; Brinton, 1977; Bradley, 1980c), and bacteriophage adsorption (Bradley and Pitt, 1974). The pili are composed of identical

64

W. PARANCHYCH AND L. S. FROST

pilin subunits, which have molecular weights ranging from about 15,000 for Ps. aeruginosa (approx. 145 amino-acid residues; Watts et al., 1983a; Sastry et al., 1983) to about 18,000 for N . gonorrhoeae (approx. 160 amino-acid residues) (Robertson et al., 1977; Hermodson et al., 1978; Haas and Meyer, 1986). Pilin proteins from these bacteria show extensive amino-acid homology in the N-terminal region, which consists almost entirely of hydrophobic residues, and begin with the modified amino acid N-methylphenylalanine (NMePhe) (Frost et al., 1978; Paranchych et al., 1978; Hermodson et al., 1978; Elleman et al., 1986). For this reason, these pili are referred to as the NMePhe type. Nucleotide-sequence studies have shown that the mature protein is initially produced as a propilin with six or seven additional N-terminal residues (Elleman and Hoyne, 1984; Meyer et al., 1984; Marrs et al., 1985; Pasloske et al., 1985; Sastry et al., 1985b). Sequence similarity is extremely high within the first 30 residues, while only slight homology exists between residues 30-55 and within 50 or so residues at the C-termini (Elleman et al., 1986). There is virtually no homology in the central region of these pilin molecules, which contains the immunodominant type-specificepitope in pilins of Ps. aeruginosa (Watts et al., 1983b; Sastry et al., 1985a) and Bacteroides nodosus (McKern et al., 1985). In pilin from N . gonorrhoeae, the major serotype-specific antigenic determinant is located within a large C-terminal cysteine loop near the Cterminus (Rothbard and Schoolnik, 1985), whereas the domain that appears to be involved in adhesion of pili to mammalian cells is found in two separate regions of the central portion (residues 41-50 and 69-84; Rothbard et al., 1985). It is not yet clear how antigenic variation is achieved in Ps. aeruginosa, Moraxella sp. and Bacteroides nodosus. However, a series of studies from several laboratories have shown that antigenic variation in pilin from N . gonorrhoeae is achieved through gene rearrangement between silent genes and expression sites in the chromosome (Meyer et al., 1982; Hagblom et al., 1985; Haas and Meyer, 1986; Bergstrom et al., 1986).

IV. High-Resolution Studies on Pilus Structure The first serious attempt to examine the fine structure of pili by a combination of electron microscope and X-ray fibre diffraction techniques was that of Brinton (1965). An examination of Type 1 pili pseudocrystalline arrays revealed angle layer crystals whereby one layer of pili intersect a second layer at a constant angle of 41.5'. Amino-acid compositional analyses on purified preparations of Type 1 pili had already shown that the minimum molecular weight of Type 1 pilin was approximately 17,000 (Brinton, 1965). Having

PHYSIOLOGY AND BIOCHEMISTRY OF PILI

65

determined by electron microscopy that the diameter of Type 1 pili is 7 nm. and knowing the layer angle to be 41.5", Brinton predicted that the cylindrical pilus is composed of identical subunits arranged in a helical array with a pitch distance of 2.4 nm. This prediction was tested with X-ray fibre diffraction studies on pilus fibres that had been prepared in thin-walled capillary tubes. Reasonably good diffraction patterns were obtained, and Brinton interpreted these patterns with the 17,000 Da pilin subunits arranged in a helical manner with 3.125 subunits per turn of2.3 nm pitch. It was also deduced from electron microscope observations that the cylindrical structure contained an axial hole of approximately 2 nm. Mitsui et al. (1973) carried out a similar X-ray fibre diffraction study on pili purified from a different strain of E. coli. These workers did not identify the type of pili they were studying, nor the molecular weight of the pilus subunit. However, it is likely they were also working with Type 1 pili, since their results were similar to those of Brinton (1965). The arrangement of subunits in the pili rods was found to be strictly simple-helical with 3.14 units per turn of helix and a pitch of 2.54 nm. A structural study on the pili of the non-starforming (sta-) mutant strain 3/7 of Ps.echinoides was carried out by Mayer and Schmitt (1 971) by means of direct and optical diffraction analyses of high-resolution photographs of negatively stained specimens. These pili appeared to be hollow rods of 6.5 nm outer and 2.5 nm inner diameter. The arrangement of the 8,000 Da pilin subunits was described in an array of three helices with a common axis having a pitch of about 20" and six subunits per turn. Folkhard et al. (1979) proposed a model for F pili based on their studies using fibre diffraction. They suggested that F pilli are hollow fibres with an 8.0 nm outer and a 2.0 nm inner diameter. They calculated a mass per length value of 3000 Da per angstrom, and a density of 0.77 Da per angstrom. Their original model was based on a molecular weight of 11,200. Recently, Marvin and Folkhard (1986) have re-examined their model using the correct molecular weight of 7200 for F pilin. They proposed that the subunits in F pili are related by a fivefold rotation axis around the helix axis. The helix symmetry is 25 units in two turns of a helix with a pitch of 16 nm and a crystallographic repeat of 32 nm. This parallels the structure of fd bacteriophage (Gray et al., 1981) which also has fivefold rotational symmetry and has five attachment proteins at one end of the phage (Webster and Lopez, 1985). This seems to be a recurring motif in the structure of filamentous organelles that are capable of disassociating into the bacterial membrane and has been proposed for Pf phage (Makowksi and Caspar, 1981), Pf3 phage (Day and Wiseman, 1978) and perhaps is true for the polar pili of Pseudomonus species (Folkhard et al., 1981; Watts et al., 1983a) (see p. 81). A representation of this model with respect to F pili is shown in Fig. 2(a).

320

256

(b) 2oo

150

2

ci, 100

ILI

i

50

I

0

12.8

i

0

52 A 80 A FIG. 2. Models of F and PAK pili. (a) The lattice diagram of F pili is based on information in Folkhard et al. (1979) and reinterpreted by Marvin and Folkhard (1986). The model shows a pilus diameter of 8 nm with five subunits per turn arranged with fivefold symmetry. There is a rise of 1.28 nm per subunit and the helix symmetryis 25 units in two turns of the helix with a rise of 32 nm. The sketch of the two layers of subunits illustrates the arrangement of subunits with respect to each other and the approximate surface area of a subunit exposed on the pilus surface. At present there is no information on the shape of the subunit or the packing of the subunits in the pilus. (b) The lattice diagram of PAK pili is based on information in Folkhard et a!. (1981) and Watts et al. (1983a). The pilus diameter is 5.2 nm and there are five subunits per turn but there is no fivefold rotational symmetry. There is a rise per subunit of 4.1 nm. The shape of the subunit has been calculated to be elongated with parallel alpha helices within the subunit arranged along the axis of the pilus.

PHYSIOLOGY AND BIOCHEMISTRY OF PiLl

67

Brinton (1971) reported that treatment of F pili with sodium pyrophosphate at 55°C and pH 4.5 caused formation of fibres of about one-third the diameter of whole pili. Incubating pili under these conditions at pH 2.0 caused irreversible formation of vesicles. Tomoeda et af. (1975) also reported the splitting of pili into thinner fibres when treated with 0.05 M HC1 at 30°C. These results interpreted the structure of F pili as ribbon-like consisting of two parallel protein rods. However, diffraction data clearly suggest that pili are helical filaments, and these results can be explained as a rearrangement of the pilin subunits within the filament. X-Ray fibre diffraction studies have also been reported on pili from Ps. aeruginosu strains PAK and P A 0 (Folkhard et al., 1981). These bacteria have polar pili which are flexible filaments of about 6 nm diameter and 2.5 jim average length (Weiss, 1971; Bradley, 1972a). Native pili consist of a single subunit of M , 15,000 (Sastry et al., 1983). Previous publications (Frost and Paranchych, 1977; Paranchych et al., 1979) had suggested a molecular weight of 18,000 based on migration of pilin in sodium dodecyl sulphate-polyacrylamide gels (SDS-PAGE) and on its amino-acid composition. This led Folkhard et af.(198 1) to interpret from X-ray fibre diffraction data that native pili consist of 4.06-4.08 subunits in a 4.1 nm turn of helix. In light of more recent information (Sastry et al., 1983,1985a) indicating a 15,000 Da subunit, this was reinterpreted as 5.06-5.08 pilin subunits per turn (Watts et al., 1983a). Native pili have the overall appearance of a hollow cylinder of 5.2 nm diameter and a central channel of 1.2 nm diameter (Folkhard et al., 1981). A representation of a model for PAK or P A 0 pili from Ps. aeruginosa is shown in Fig. 2(b). Although X-ray fibre diffraction studies have not yet been reported for other NMePhe pili (i.e. N. gonorrhoeae, N. meningitidis, M . nonliquifaciens, M . bovis and B. nodosus),these pili have a similar structure to those from Ps. aeruginosa because they all have the same diameter and share a common Nterminal sequence (Paranchych et af., 1978; Sastry et af.,1985a; Elleman et af., 1986). Although numerous attempts were made in this laboratory (T. H. Watts and W. Paranchych, unpublished work) to prepare oriented fibres of gonococcal pili, we were unable to obtain diffraction patterns of good quality. However, Deal et al. (1985) claim to have grown protein crystals of gonococcal pilin aggregates. The crystals displayed 3.0 nm spacing in one direction with 20.0 nm spacing in the second; the third direction indicated a larger aggregate. Efforts are being made to use these crystals to determine the three-dimensional structure of the pilin subunit. These workers (Deal et al., 1985) have also suggested that the secondary structure of NMePhe types of pilin consist o f a series of four antiparallel a-helix domains in the analogy to that of tobacco mosaic virus coat protein (Bloomer et al., 1978). However, secondary structure predictions in this laboratory for pilin from Ps.

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aeruginosa (Sastry et al., 1983) suggest that the up and down 4-u-helix structural model (Weber and Salemme, 1980) may represent an oversimplification of NMePhe-type pilus structure. The structure of Bordetella perfussis pili was determined by optical density analysis of electron micrographs of paracrystalline bundles of purified pili. These pili are helical filaments with a 13 nm axial repeat containing five repeating units in two complete turns of a single start helix. The mass per unit length confirmed these results using a mass of 22,000 for serotype 2 and 21,500 for serotype 6 pili. Radial density profiles suggested that there was no axial channel and that the pilus diameter is 7.5 nm (Steven et al., 1986).

V. Organization and Expression of Pilin Genes A. CONJUGATIVE PILI

I . Plasmid-Encoded Elements Conjugative pili are specified by transfer regions of self-transmissible plasmids; the most detailed studies on bacterial conjugation have involved the IncF plasmids which have been extensively reviewed (Willetts and Skurray, 1980; Willetts and Wilkins, 1984; Clark, 1985; Silverman, 1986; Ippen-Ihler and Minkley, 1986). A transfer region generally encodes a plasmid-specific oriT sequence where the plasmid is nicked and transfer of one of the DNA strands into the recipient cell is initiated. There are several plasmid-specific transfer genes required to facilitate DNA transfer during this process. The role of the conjugative pilus is identifying a suitable recipient and bringing it into contact with the donor cell. This process is thought to involve retraction or disassembly of the pilus into the membrane. Thus, in addition to genes specifying synthesis and assembly of a pilus, there are, presumably, genes encoding pilus retraction. A third system encoded by the transfer region is surface exclusion or entry exclusion which prevents transfer of a given plasmid into cells already harbouring a related plasmid (Willetts, 1977a), and is a distinct process from that of incompatibility which prevents or slows replication of one of two plasmids within the same cell (Timmis, 1979). Surface exclusion has been shown to be a property of plasmids from the IncF complex (Willetts and Maule, 1986), P-1 (Olsen and Shipley, 1973; Barth and Grinter, 1977; Barth et al., 1978), N (Winans and Walker, 1985a), I (Meynell, 1969) and H (Taylor et al., 1985a) incompatibility groups. Other proteins encoded within the F transfer region have been identified but their function is not known. The current map of the F plasmid is reviewed by Ippen-Ihler and Minkley (1986).

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Although the functions encoded by the transfer regions of the various plasmids remains fairly constant, the organization of the genes can vary. The transfer regions of the IncF plasmids occur as one segment of the plasmid (Sharp et al., 1973; Willetts and Skurray, 1980; Finlay et al., 1983, 1984), whereas the transfer regions of other plasmids may be divided into as many as three segments, as with the IncN plasmid pKMlOI (Winans and Walker, 1985a). A partial list of plasmids whose transfer regions have been mapped includes the IncN plasmids R46 (Brown and Willetts, 1981) and pCUl (Konarska-Kozlowska and Iyer, 1981), RP4 (IncP-I) (Barth et al., 1978), RK2 (IncP- 1) (Meyer et al., 1977), R9 1-5 (IncP- 10) (Moore and Krishnapillai, 1982a, b), R27 (IncHI) (Taylor et al., 1985b) and the Ti plasmid (Holsters et a/., 1980). The location of the pilin gene is not known for these plasmids nor is the organization of RNA transcripts required for pilus expression. Genetic analysis of the F transfer region has shown that there are at least 14 genes, (traALEKBPVWCUFQHG), contiguous with each other within the large (33 kb) tra YZ transfer operon, that are required for synthesis of pilin and assembly into an intact pilus. The organization of cistrons and control elements within the F transfer region has been reviewed extensively (Willetts and Skurray, 1980, 1986; Ippen-Ihler and Minkley, 1986). The traA gene encodes propilin (Minkley et al., 1976), a polypeptide of molecular weight 13,200 (Kennedy et al., 1977; Moore et al., 1981; Frost et al., 1984), which is processed to pilin of molecular weight 7200 (Moore et al., 1981; Frost et al., 1984). The pilin molecule is acetylated (Frost et al., 1984) and may be further modified with glucose and phosphate moieties (Brinton, 1971;J. Armstrong et al., 1981). Processing of propilin to pilin requires the traQ product, but the function of true is unclear (Moore et al., 1982). Acetylation of the pilin protein may require the N-terminal portion of the traG gene product (Laine et al., 1985). The function of the other gene products required for pilus synthesis is unknown. However, the molecular weight and cellular location of these tra gene products has been identified by cloning the F transfer region onto multicopy plasmids or transducing phages and studying the gene products in minicells or by in uitro translation (Kennedy et al., 1977; Willetts and Skurray, 1980; Laine et a/., 1985). All of the gene products involved in pilus biosynthesis are associated with the cell envelope (Kennedy et al., 1977) and several fractionate with both the inner and outer membrane, suggesting that they may be localized at Bayer's junctions or sites of fusion of the two membranes (Bayer, 1975). While the IncF plasmids share many interchangeable gene products within their transfer regions, certain genes are plasmid-specific. These genes are involved in DNA metabolism during conjugation (traMY/),surface exclusion (traS7') and control of transfer-operon expression (traJandJinOP). The genes required for pilus biosynthesis appear to be interchangeable. This homology

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was evident through heteroduplex analysis of several IncF plasmids (Shfrp et al., 1973) and by genetic studies recently summarized by Willetts and Maule (1986). Expression of conjugative pili is usually very low in a bacterial population (0.1-1 Yo)because of the repressed state of the transfer region. This repression is alleviated for a few generations in a daughter cell until the concentration of a repressor is restored. This phenomenon is termed HFT for High Frequency of Transfer since these cells are fully piliated and are competent for conjugation (Meynell et al., 1968). Derepressed mutants of these plasmids can be isolated (Meynell and Datta, 1967; Bradley, 1984; Willetts, 1984) and these cells usually express 1-6 pili on each cell (Frost et al., 1985). This property of conjugative plasmids has been used extensively in identification of pili on cells. The ability of one plasmid to complement the mutation in a derepressed plasmid and rest-orerepression is termed fertility inhibition e n ' ) (Meynell et al., 1968). Two genes in F-like plasmids have been identified in repression of the transfer region, namely f i n 0 and JinP, which together form the FinOP repressor system (Willetts, 1977b). Derepressed plasmids usually carry a mutation within one of these two genes (Willetts and Maule, 1986); however, in at least two cases that have been studied, derepression is due to the presence of insertion elements. The F plasmid is a naturally occurring derepressed plasmid, and this is due to the presence of an IS3 element within thefin0 gene (Clark, 1985; Willetts and Skurray, 1986). The pED208 plasmid is the derepressed form of Folac and contains an IS2 element at the beginning of the transfer operon (B.B. Finlay, L.S. Frost and W. Paranchych, unpublished work). This particular mutation results in production of approximately 20 pili on each cell. The molecular basis for the FinOP repression system has been studied in some detail, and summarized in other reviews (Willetts and Skurray, 1980, 1986; Willetts and Maule, 1986). The large traYZ operon (33 kb) is positively regulated by the traJ gene product (about 25 kDa) which is transcribed on a separate operon and is, in turn, negatively regulated by thefin0 andJinP gene products. The sequence for the three alleles of traJof F-like plasmids (Willetts and Maule, 1986) are currently available (Fowler et al., 1983; Finlay et al., 1986a; L. S . Frost, B. B. Finlay and W. Paranchych, unpublished work). The traJ-encoded protein, TraJp, previously thought to be found in the outer membrane, is a cytoplasmic protein and is insoluble and co-purifies with the membrane fraction when overproduced on high copy-number plasmids (Cuozzo and Silverman, 1986). The traJ-encoded proteins from these three alleles are very different, but two regions are conserved and one region strongly resembles a DNA binding domain (Anderson et at., 1982; Takeda et al., 1983; L. S. Frost, B. B. Finlay and W. Paranchych, unpublished work). The protein TraJp stimulates transcription from the tra YZp promoter

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which is contained in the intergenic region between traJ and tray. There are three different known promoter sequences among F-like plasmids which match the three alleles of truJ. The exact location of this positively regulated promoter sequence has been mapped for F and R1-19 (Willetts, 19776 Gaffney et al., 1983; Fowler et al., 1983; Cram et al., 1984; Mullineaux and Willetts, 1985; Finlay et al., 1986b; Fowler and Thompson, 1986; Koronakis and Hogenauer, 1986; L. S. Frost, B. B. Finlay and W. Paranchych, unpublished results). ThefinP gene product appears to be a small anti-sense RNA molecule (Thompson and Taylor, 1982; Mullineaux and Willetts, 1985; Finlay et al., 1986b; Fee and Dempsey, 1986) which is complementary to the untranslated portion of the traJ mRNA. However, the f i n 0 gene product is absolutely required for repression of traJand, in turn, the transfer operon (Finnegan and Willetts, 1972). Thefin0 gene product may be a protein (Timmis et al., 1978; Dempsey and McIntire, 1983; Cheah et al., 1984; Willetts and Skurray, 1986) but the mechanism by which it interacts with finP and represses traJ transcription is unknown. ThefinP gene product is plasmid-specific, and six alleles among 12 plasmids studied have been identified (Willetts and Maule, 1986). The basis for this specificity is thought to reside in the loops of two stem-and-loop structures predicted from the sequence of thefinP gene (Finlay et al., 1986b). Thefin0 gene product is relatively non-specific, since thejinO mutation in the F plasmid can be complemented by a number of F-like plasmids and is the molecular mechanism of fertility inhibition (Egawa and Hirota, 1962; Meynell et al., 1968). Pilus expression can be affected by gene products from other F-like plasmids or plasmids from other incompatibility groups as well as chromosomally encoded gene products and small effector molecules such as cyclic AMP. 2. Chromosomally Encoded Control Elements At least four genetic loci have been implicated in the expression of the Ftransfer operon (Silverman, 1986). Mutations at these loci also affect the composition of the cell envelope (reviewed in Willetts and Skurray, 1986). The sfrA and cpxAB gene products affect the levels of TraJp in the cell (Beutin et al., 1981; Sambucetti et al., 1982; Silverman, 1986)although they do not act at the level of transcription initiation. The sfrA gene product may prevent premature termination (Beutin et al., 1981; Gaffney et al., 1983). They were originally thought to affect transport of TraJp to the outer membrane; however, TraJp is not an outer-membrane protein (see p. 70). The sfrB gene product is an 18,000 Da regulatory protein (Rehemtullah et al., 1986) which affects synthesis of the lipopolysaccharide core (Beutin er al., 1981;Sanderson and Stocker, 1981; Creeger et al., 1984). It also acts as a transcription

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antiterminator of the t r a y 2 operon (Beutin et al., 1981; Gaffney et al., 1983). These chromosomally encoded proteins seem to co-ordinate transfer operon expression and the structure and composition of the cell envelope. 3 . Plasmid Encoded Control Elements

F-Like plasmids are capable of inhibiting F transfer by supplying compatible fin0 orfinP gene products (see p. 71). Many apparently unrelated plasmids encode fertility-inhibition mechanisms other than the FinOP system for inhibiting F transfer (Willetts and Paranchych, 1974; Gasson and Willetts, 1975, 1977). These systems (FinQ,U,V,W,C) act at the level of premature termination at a number of sites within the traYZ or traM (FinW) transcript or inhibit the function of one or more of the transfer gene products (Gaffney et al., 1983). This could be a general phenomenon since, for instance, F and the IncN plasmid pKMlO1 inhibit transfer of RPl (IncP-1) (Olsen and Shipley, 1975; Tanimoto and Iino, 1983; Winans and Walker, 1985b).

4 . Small Effector Molecules One small molecule known to affect pilus expression is cyclic AMP, while little is known about control of piliation by other small molecules such as ppGpp. It has been known for many years that late stationary cultures of F+ or Hfr cells acquire an F- phenotype. Piliation is maximal during the logarithmic phase of growth, and decreases as cells enter the stationary phase under aerobic but not anaerobic conditions (Ippen and Valentine, 1967; Brinton and Beer, 1967; Biebricher and Duker, 1984). Piliation is dependent on the host strain and the number of pili on each cell in stationary-phase cultures can vary from none to almost the maximum. Harwood and Meynell (1975) observed that derepressed IncI plasmids in cya crp mutants of E. coli (Perlman and Pastan, 1969), which are defective in cyclic AMP synthesis (cya) or cyclic AMP receptor protein (crp), produce many more I pili than usual and that, in cya mutants, this effect can be reversed by addition of exogenous cyclic AMP. For cells carrying F-like pili, the results were less clear. F+ cells do not seem to have this sensitivity to cyclic AMP while the derepressed plasmids R538-1, R124rd, R1-19 and R136-1 (which is almost identical to R100- 1) gave 10-fold more pili in an E. cofi cyu strain. Lawn and Meynell (1972, 1975) found that addition of anti-pilus antibodies to E. cofi carrying an IncI plasmid, or subjecting the cells to vigorous washing, caused a dramatic rise in the extent of piliation. They suggested that multipiliation due to washing or the presence of antibodies was unrelated to the cyclic AMP effect and, indeed, may be connected to the equilibrium between outgrowth and retraction rather than transcriptional

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control by the cyclic AMP-CRP complex. Kumar and Srivastava (1983) found that Hfr(F+) strains carrying cya or crp mutations were transferdeficient, and that exogenous cyclic AMP, with the cya mutation alone, alleviated this Tra- phenotype. They speculated that transcription in this Hfr strain was controlled by cyclic AMP-CRP at the level of antitermination where this complex was required for synthesis of antiterminators which would prevent premature transcription of the long tra YZ operon or traJ gene. One interesting point in these experiments was the use as the wild-type control of E. coli CA8000 which contains the relA2 mutation and transfers at normal levels. Thus ppGpp may not be required for F pilus expression. Recently, the sequences of the promoter regions of the traJ and t r a y 2 operons for the three alleles of F-like plasmids have yielded information which helps to explain the previously noted effect of cyclic AMP-CRP (Fowler et al., 1983; Finlay et al., 1986a; L. S. Frost, B. B. Finlay and W. Paranchych, unpublished results). These two promoters are likely candidates as sites of action for the cyclic AMP-CRP complex. Using the consensus sequence for a CRP binding site (Ebright et al., 1984) which is defined as 5’-AA-TGTGA- T- - -TCA-ATA/T-3’, a potential CRP binding site was found within the promoter region of truJ in all three alleles. There was also a CRP-binding site within the R100-I tra YZpromoter region, while no strong site could be found within the F and R1-19 truYZ promoter regions. These CRP-binding sites could be involved in preventing transcription initiation at these promoters, and certainly a decrease in traJor t r a y 2 transcription would lower the level of piliation. Helmuth and Achtman (1978) reported that they added additional glucose to a culture as it entered the stationary phase of growth to encourage F-pilus production at high cell concentrations in order to facilitate F-pilus purification. This would support the idea that piliation increases under conditions where cyclic AMP production is suppressed (Adhya and Garges, 1982) for, as the concentration of glucose drops, the cyclic AMP concentration would rise and exert its effect at a number of sensitive promoters. Also, the sensitivity to arsenate of piliated cells grown in defined media containing sugars whose utilization is controlled by cyclic AMP (O’Callaghan et al., 1973b) could also involve cyclic AMP-CRP control where certain transfergene products are produced in lower amounts, emphasizing the sensitivity to the poison. Although cyclic AMP almost certainly affects expression of the various transfer operons, other factors are also involved. The effects of different oxygen concentrations and media composition on piliation cannot be wholly explained by the cyclic AMP-CRP effect, and a large number of unexplained observations concerning F pili production under various physiological conditions (for instance, temperature) are not understood.

14

W. PARANCHYCH AND L. S. FROST B. TYPE

1 PILI

The genes that encode production of Type 1 pili are located at 98 min on the E. coli linkage map (Swaney et al., 1977; Freitag and Eisenstein, 1983). Complementation studies of mutants defective in production of Type 1 pili indicated that at least three genes are required for production of these adhesins (Swaney et al., 1977).The gene encoding the structural component of Type 1 pili has now been cloned and sequenced at the nucleotide level. This gene was named JimA by Klemm (1984), and pilA by Orndorff and Falkow (1985). The two sequences were identical except that a Thr in position 140, reported by Orndorff and Falkow (1985), was absent from the sequence reported by Klemm (1984). The gene was found to specify a polypeptide 159 amino-acid residues long preceded by a 23 amino-acid residue signal peptide (Orndorff and Falkow, 1985). Klemm et al. (1985) found that the genes required for production of intact Type 1 pili are contained within a DNA segment of 8 kb. Four genes, designatedJimA, B, C and D, were found to be involved in synthesis of pili, and their order was shown to be (apparent molecular weight of the processed form of each gene product is shown in parentheses): fimB (23,000), JimA (16,50O),JimC (26,000) andJimD (89,000), organized in three transcriptional units. Orndorff and Falkow (1984a, b, 1985), Orndorff et al. (1985) and Maurer and Orndorff (1985) have also cloned the entire region responsible for Type 1 pili formation, and characterized the relevant genes. The designation and order of pili-related genes in their 9.4 kb fragment were: hyp (23,000), pilA (17,000), pilB (30,000),pilC (86,000), pilD (14,000), pilE (molecular weights not yet reported). The pilA gene encodes the structural component of Type 1 pili and corresponds to Klemm’sfimA gene product. The pilB and pilC genes correspond to Klemm’s fimC and fimD genes, respectively. Orndorff’s hyp gene is equivalent to Klemm’sJimB gene, while Klemm has not yet reported on’ the equivalent of the pilE gene. The pilE gene product has recently been shown to encode an adhesin which is distinct from the pilin subunit encoded by the pilA gene (Maurer and Orndorff, 1985). Mutants designated pilE possessed pili that were immunologically and morphologically indistinguishable from parental pili, but they were unable to cause agglutination of guinea-pig erythrocytes. It was suggested that the pilE locus specifies expression of an adhesin that becomes associated with the pilus to yield normal pili capable of haemagglutination (Maurer and Orndorff, 1985). It has been known for many years that Type 1 pili undergo “phase variation”, a metastable state in which bacteria switch back and forth between the piliated and non-piliated states at a rate of about to (Brinton, 1959, 1965; Eisenstein, 1981). The phase-switch function maps at 98 minutes,

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15

adjacent to the known pi1 (orjim) genes (Freitag and Eisenstein, 1983). Orndorff et al. (1985) and Abraham et al. (1985) have shown that it is the pilA gene that is subject to this metastable control. Orndorff et al. (1985) constructed a pi1A‘-lac2 fusion, and showed that it was subject to metastable transcriptional control. The rate of switching from the Lac+ to the Lacin the opposite per cell per generation and 6.2 x phenotype was 4 x direction. Abraham et al. (1985) subcloned the switch, sequenced the DNA, and determined the molecular basis for its activity. The switch is an invertible element different from that controlling flagella in Salmonella species; it is small, consisting of 3 14 bp bounded by 9 bp inverted repeats, and it is driven by a different recombinase. It is just upstream of the pili structural gene (pilA) and contains a consensus E. coli promoter when in the “on” orientation. Klemm (1986) reported that theJimB andJimE gene products direct the phase switch into the “on” and “off” positions, respectively. These two genes were sequenced and found to encode highly homologous proteins (around 23,000 Da) that were very basic and probably capable of interacting with the DNA. Orndorff and Falkow (1984b) and Orndorff et al. (1985) have also noted that, in addition to the metastable regulation of pilA, a second type of transcriptional regulation is effected by the product of the gene hyp, adjacent to pilA. The product of the hyp gene appears to be a repressor since it inhibits piliation (Orndorff and Falkow, 1984b). It was therefore suggested that hyp may effect the metastable expression of piliation (Orndorff and Falkow, 1984b). However, when the hyp gene was inactivated by Tn5 insertion, the E. coli strain exhibited a frequency of switching from Lac+ to Lac-, and vice versa, indistinguishable from that of the parental strain. Thus, hyp does not appear to affect the metastable variation but does affect the level of transcription of the pilA gene in the O N (transcribed) mode (Orndorff et al., 1985).The hyp gene is the equivalent ofjimE, reported by Klemm (1986), who has detected activator activity, not repressor activity, for this gene product. This suggests that there are two different systems of control of pilA transcription, namely an invertible element that controls phase variation and a second element that modulates the level of expression. C. PILl DESIGNATED PAP

A majority of uropathogenic E. coli strains carry mannose-resistant haemagglutination (MRHA) activity. This is in contrast to faecal strains where only 10% of the bacteria express MRHA (Hagberg et al., 1981; Van den Bosch et al., 1980; Minshew et al., 1978). Along with other virulence determinants, these strains usually produce Pap pili which are important in adherence to uroepithelial tissues (Hull et al., 1984; Kallenius et al., 1981; Korhonen et al., 1982). The Pap gene cluster has been mapped on the chromosomes of several

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clinical isolates of E. coli, and found to occur at a variety of locations. Also, more than one Pap gene cluster can occur on each chromosome although not all loci are necessarily expressed at once (Hull et al., 1981, 1986; Rhen et al., 1983b). The Pap gene cluster is often linked to other virulence determinants such as the chromosomal haemolysin determinant or the K1 capsular antigen. However, the intergenic region is not conserved (Low et al., 1984; Hull et al., 1986). It has been suggested that these virulence determinants are acquired by a transposition-like event since MRHA regions are bordered by a conserved sequence. This event may have occurred at some distant point in evolution since transposition of MRHA activity to a multicopy plasmid has not been possible, and the purported MRHA transposon may be degenerate (Goebel et al., 1973). The Pap gene cluster can be cloned on a plasmid vector as a single discrete entity of minimum size 10 kb, approximately, and the Pap pili, along with the associated properties of MRHA and adhesion, can be expressed by these chimeric plasmids (De Ree et al., 1985a, b; Hull et al., 1981; van Die et al., 1983, 1985; Clegg, 1982; Rhen, 1985; Rhen et al., 1983a, b, c). Hybrid chimeras containing DNA segments from several Pap-like gene clusters can express Pap pili and haemagglutination (Van Die et al., 1986a, b; Lund et al., 1985). Three Pap gene clusters cloned from separate E. coli isolates had very similar genetic organization and the genetically identifiable properties of pilin synthesis, pilin export. Pilus assembly and adhesion were trans complementable among the three gene clusters. The major differences in these gene clusters involved the central portion of the Pap pilin gene (papA) and one of the adhesin genes (papG) which, presumably, defines the antigenic variation within these proteins (Lund et al., 1985). Thus, the known gene clusters that express MRHA activity appear to be closely related to each other. This is reinforced by the protein-sequence homology among Pap pilins studied to date (Klemm, 1985; Baga et al., 1984; Rhen et al., 1985; Van Die et al., 1984a, b) as well as the highly conserved restriction maps for the Pap gene clusters currently available (Van Die et al., 1986a, b). The genetic organization of three Pap gene clusters (Pap, F71 and F72) has been described (Clegg and Pierce, 1983; Van Die et al., 1984a, b, 1985; Norgren et al., 1984; Baga et al., 1985). Synthesis of Pap pili can be separated from production of the adhesin as shown by transposon mutagenesis, isolation of mutants and recombinant-DNA manipulation. Thus, it is possible to isolate cells capable of expressing Pap pili but lacking adhesion function (Lindberg et al., 1984, 1986; Van Die et al., 1985, 1986b; Lund et al., 1985; Norgren et al., 1984; Normark et al., 1983). Normark and his coworkers have dissected the Pap system in some detail. They have identified nine genes (papZBAHCDEFG) involved in pilus

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synthesis, expression of the adhesin and control elements. The papA gene encodes the pilin subunit termed fimbrillin (16.5 kDa) and the DNA sequence reveals the presence of a 22 amino-acid residue signal peptide (Baga et al., 1984). The papA gene was preferentially transcribed on an 800 bp mRNA. This transcription was positively regulated by two cistrons (papB and papl) upstream ofpapA . ThepapB transcript also proceeded through the papA gene to give a 1300 bp mRNA. They speculated that thepapA mRNA (800 bp) is a processed form of the papBA transcript. The papl gene precedes papB and is weakly transcribed from the opposite strand (Baga et al., 1985; Norgren et al., 1984). The presence of glucose in the medium lowers pilus expression and adhesive properties of type 1 and K99 pili (Old and Duguid, 1970; Isaacson, 1980). Similarly, Pap pili production is affected by glucose concentration (SvanborgEden and Hansson, 1978), temperature (Goransson and Uhlin, 1984) and media composition. Baga et al. (1985) have shown that papB transcription is regulated by the cyclic AMP-CRP complex and have proposed a potential CRP-binding site upstream of the papB promoter. The papEFG genes, which are on a separate operon, are not required for pilus synthesis and specify the adhesin (Lindberg et al., 1986). Genes for papFG are associated with digalactoside-specific binding while papE mutants are capable of adherence to whole cells, but purified pili from this strain do not adhere (Lindberg et al., 1984). Mutations in papA, which destroy pilus production, do not prevent cell adherence to the digalactoside receptor, presumably because of expression of the adhesin (Lindberg et al., 1984; Norgren et al., 1984; Uhlin et al., 1985).ThepapE andpapFgenes encode Pap pilin-like proteins which may represent minor components of the pili or a subpopulation of pili which define the adhesin (Lindberg et al., 1986). The papG gene product is also involved in elaboration of the adhesin and is capable of antigenic variation. However, its sequence has not been determined and its precise function is unknown. The papC (81,000) and papD (28,500) gene products are required for pilus and adhesin assembly while the function of the remaining pap gene products is unknown. Pap pili are subject to phase variation which has been demonstrated for one strain containing four fimbrial A single colony was antigens, one of which was Type 1 pili (Rhen et al., 1983~). found to contain a number of variants expressing different fimbrial antigens suggesting that the rate of phase variation in this system is high. D. PILI DESIGNATED CFA/I

AND CFA/II

(csl, cs2 AND cs3)

Pili designated CFA/I are encoded by a group of related plasmids that are between 86-93 kbp in size and also carry genes for heat-stable enterotoxin (ST enterotoxin) (Smith et al., 1982a; Willshaw et al., 1982). Smith et al. (1982b)

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S . FROST

used transposon mutagenesis and complementation studies to map the genes responsible for expression of CFA/I pili. These genes were found in two regions of the plasmid separated by 37 kbp. Both regions were cloned onto compatible vectors, and it was shown that the ST genes are closely linked to one of these regions (designated region 1; Willshaw et al., 1983). Region 1 is contained within a DNA fragment of 6 kbp, while the second region (region 2) is within a 2.1 kbp fragment. Examination of the gene products of the two regions in minicells revealed that region 1 expressed the pilin subunit as well as proteins with molecular weights of 85,000, 40,000, 30,000, 28,000, 27,000, 25,000 and 15,000, whereas region 2 encoded polypeptides of 26,000, 15,000 and 14,000molecular weight (Willshaw et al., 1985). The functions of the nonpilin gene products are not yet known. The three CFA/II components (CS1, CS2 and CS3) can be distinguished serologically, and on the basis of pili morphology, pilin size and haemagglutination patterns (Smyth, 1982; Craviato et al., 1982). The genes responsible for production of CSl, CS2 and CS3 reside within the same plasmid of approximately 89 kbp size, which generally also codes for heat-labile (LT) and heat-stable (ST) enterotoxin (Penaranda et al., 1980; Mullany et al., 1983; Smith et al., 1983). Although CFA/II plasmids encode all three types of pili, expression is usually limited to one or two types. For example, the CFA/IIcontaining bacteria may produce components CS1 and CS3, CS2 and CS3, or CS2 or CS3 only. These patterns of expression appear to be related to the serotype or biotype of the host. Strains of serotype 0 6 : H16 have been shown to produce CSl or CS2. Moreover, CSl is produced only by 0 6 :H 1 6 of biotype A (rhamnose-negative), whereas CS2 is produced only by 0 6 :H 16 strains of biotype B, C and F (rhamnose-positive) (Craviato et al., 1982; Smyth, 1982).Component CS3 is produced independent of these biotypes and serotypes (Craviato et al., 1982). The mechanism underlying these phenotypic relationships is presently not understood. E. PILI DESIGNATED

K88 AND K99

Both K88 and K99 pili are encoded by plasmids that are approximately 75 kbp in size. Recombinant DNA studies on these two systems have revealed their genetic organization to be similar to that of the Pap and Type 1 pili systems, i.e. the genes are clustered, and five to nine gene products are required for pili expression (Klemm, 1985; Mooi and de Graaf, 1985). The K99 operon contains eight structural genes, at least seven of which appear to be required for K99 pilus formation. The K88 pilus system contains six structural genes, with at least five being located within a single transcriptional unit (Mooi and de Graaf, 1985). As already stated, pilin subunits encoded by the K88, K99, Pap and Type 1-related pilus systems show homology, indicating they are

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evolutionarily related. Details of the genetics and biochemistry of these systems have been reviewed extensively (Gaastra and de Graaf, 1982; Klemm, 1985; Mooi and de Graaf, 1985). G . PILI DESIGNATED

NMePhe

The genetic organization of NMePhe pili can tentatively be divided into two groups. One group, at present consisting of gonococcal (GC) pili alone, has developed a complex system of phase switching and antigenic variation involving gene rearrangement and classical control elements at the transcriptional and post-transcriptional levels. The other group, including pili from Ps. aeruginosa and Bacteroides nodosus, shows variation in terms of serogroup and serotype as well as pilin subunit size, but these are stably maintained and phase variation and gene rearrangement are not detectable in passaged laboratory strains. Expression of GC pili has been reviewed recently (So, 1986), and is briefly summarized to afford comparison with the other system.

I . The Gonococcal Pilus System Different states of piliation on gonococcal strains were first detected as differences in colony morphology (Swanson et al., 1971; Jephcott et al., 1971). A single colony was capable of generating several colony morphologies within a few generations. For instance, piliated cells generated non-piliated variants at a rate of one in 1000 while the reverse event occurred at a rate of one in lo4lo5. This high rate of switching was thought to be due to gene rearrangement and not simple mutation. Most of the information about organization of pilin genes in G C has been derived using a single isolate, MS11, originating at the Mount Sinai School of Medicine in New York (Swanson et al., 1985). Many regions of the chromosome contain pili-related sequences (Meyer et al., 1982; Segal et al., 1986; Swanson et al., 1986). These can be divided into silent (pilS) and expressed ( p i l E ) regions. The MS11 chromosome contains two functional expression sites, pilEl and pilE2 (Meyer et al., 1984) and two silent loci encoding the constant region of the pilin molecule (residues 1-30) (So, 1986; Segal et al., 1986) as well as other silent loci containing the SV and HV sequences, one of which, pilSI, has been studied in detail. The two pilE loci are about 20 kb apart and pilSl maps about 15 kb upstream of pilEl (Haas and Meyer, 1986). Haas and Meyer (1986) sequenced thepilSl silent region and showed that it contained six tandem pilin genes which lacked the constant N-terminal region and encoded the SV and HV regions of five different pilin proteins. These pilin sequences were flanked by 39 bp repeats also found in the expression site, and these authors postulated that these sequences represent cassettes of the SV and

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HV regions which are duplicated and transferred to the expression site(s) or other silent loci. They also suggested that the highly conserved residues interspersed within the HV and SV regions are the flanking regions of minicassettes that further alter pilin-protein sequence. However, these conserved residues may correspond to essential residues for pilus expression, mutation in which results in the P - rp + phenotype described by Bergstrom et al. (1986). The switch from the piliated ( P + ) to non-piliated (P-) phenotype can be caused by a deletion event in one or both of the expression sites, and this is thought to involve direct repeats found at these loci (Segal et af., 1985). However, this P + to P- switch is often not explainable by gene rearrangement; for instance, both pilE loci do not undergo this deletion process simultaneously. Thus, the presence of a phase switch-induced regulator which can act in trans on thepifElocus has been invoked (Segal et al., 1985; Hagblom et af.,1985; So, 1986). Other GC strains containing only one expression site, or a derivative of MSI 1 which has one expression site deleted, do not seem to express this regulator that is thought to influence transcription of functionally intact pilin genes (Bergstrom et af., 1986). Bergstrom et al. (1986) and Swanson et a f . (1986), using the primer extension technique which allows sequencing of mRNA transcripts (Hamlyn et al., 1981), have proposed that P- derivatives are the result of genetic rearrangements and mutations within the gene. They described the molecular basis of the reverting and non-reverting phenotype of P - strains described by Swanson et al. (1985). They used the GC strain with a single expression site already described, and found that non-reverting P- (Pn-) cells had a deletion at the 5’ end of the pilin gene resulting in loss of the single copy of the constant region of the pilin protein. This MS 1 1 strain had only one intact pilin gene, and the defect was not reversible by a recombination event from a second pilin gene. They then observed two types of reverting P- cells, P - rp + and P - rp - . These P-rp+ variants carried a sequence in the HV region that was identical to the copy 5 sequence from thepilSI locus described by Haas and Meyer (1986), suggesting that copy 5 encodes a non-functional pilin protein. The P-rp+ contained pilin in their membranes but did not elaborate a pilus. They found that the pilin in these P-rp+ cells was antigenically distinct from the original P cells and the sequence of the RNA transcripts showed multiple, discrete amino-acid substitutions, usually within the HV region. They concluded that these mutations affected assembly of pilin into pili since replacing the C-terminal portion of these pilin genes with DNA from P + cells restored piliation. The P - rp - cells encoded truncated pilin proteins which could be corrected by further mutation to give P + revertants. Gene rearrangement seems to depend on the general recombination pathway of the cell since the recA GC strain, constructed by M. Koomey

+

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(personal communication), is able to undergo the P + to P- transition at normal frequencies but has a much lower rate of reversion. If the P + to Ptransition was generated by pilin mutation (Bergstrom et al., 1986), by the presence of a repressor (So, 1986) or by a specific recombinase as suggested for the Type 1 pilus system (see p. 74), then perhaps the red-dependent event would affect antigenic variation, involving recombination between the silent and expression loci. This would be an essential step in the reversion from Pto P + cells. However, it has not been shown that antigenic variation requires the cell to pass through a P- stage (So, 1986). It should be emphasized that evidence of a switching mechanism involving an invertible DNA sequence has not been detected in GC. 2. Pilus Systems in Pseudomonas aeruginosa and Bacteroides nodosus

The genetics of pili in Ps. aeruginosa and B. nodosus are not well defined. However, some comparison to the GC system can be made. The number of copies of the pilin gene in Ps. aeruginosa appears to be one (Pasloske et al., 1985, and unpublished observations). The number of serogroups of pili in Ps. aeruginosa appears to be about 10 (W. Paranchych, F. Ehftekhar, D. P. Speert, K. Volpel and B. Pasloske, unpublished observations). Moreover, the pilus serotype remains stable in laboratory strains suggesting that phase variation and gene rearrangement are not common occurrences. However, Southern blot hybridizations of the DNA and immunoblots of the total cell protein of 64 clinical isolates from cystic fibrosis patients have revealed some interesting aspects of pilus expression (W. Paranchych, Q. Sun and D. P. Speert, unpublished observations). These isolates represent samples taken from a number of patients once a year for several years. All of the isolates had a single complete gene copy of the whole pilin gene, and the restriction enzyme digest patterns fell into approximately 10 groups (unrelated to serogroup) with many of the strains resembling PAK or PAO. When a 600 bp PstI fragment which contains the conserved N-terminal region of the pilin protein, was used as a probe, homology ranged from strong to very poor. If the 1.2 kb Hind11 fragment of PAK (Pasloske et al., 1985) was used as a probe, all of the digests hybridized strongly, and preliminary evidence suggested that a region downstream of the pilin gene, bordering one of the Hind11 sites, was highly conserved in all strains. The immunoblots of cellular proteins from these strains revealed that a majority of the strains reacted with either anti-PAK or anti-PA0 pili antiserum, to varying degrees, with only a few strains being totally nonreactive and representing either completely non-homologous serogroups or strains that did not produce pili. These immunoblots made it possible to distinguish strains that carried the major epitope of PAK or P A 0 pili and

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those that carried minor epitopes. The molecular weight of the pilin subunits of these strains was found to vary between 13,000 and 17,000, the molecular weight of PAK and P A 0 pilin being 15,000.Thus, the pilin gene seemed to be capable of undergoing changes in molecular weight reminiscent of the Streptococcuspyogenes M protein (Hollingshead et af., 1986)and was capable of phase variation and/or antigenic variation but not in the same time scale as gonococcal strains. Currently, sequence data on a number of interesting pilin genes from these isolates are being accumulated. However, the primer extension method cannot be used because there is no conserved sequence near the 3’ end of the gene. Similar results were also reported for pili of Bacteroides nodosus (Anderson et al., 1986). These pili were found to contain two proteins, the pilin subunit (17,000 Da) and a minor protein (80,000 Da) associated with the “cap-like” structure at the base of the pilus (Mattick et al., 1984). These basal structures are reminiscent of the knobs associated with the base of conjugative pili although no large protein has been reported in preparations of conjugative pili (see Table 1). Both of the B. nodosus pilus-associated proteins were powerful immunogens. Anderson et al. (1986) found that the electrophoretic mobility of both the fimbrial (1 6,000-1 9,000) and basal (77,00&80,000) proteins varied in molecular weight between serotypes of these antigens as well as the eight defined serogroups (A-H). One interesting serogroup, H, had two pilin subunits (6,000 and 10,000 Da) which proved to be the result of proteolytic cleavage of the encoded gene product (16,000 Da) (Elleman et af., 1986). While a number of NMePhe pilin clones have been expressed in E. coli, no cloned fragment has been capable of expressing a pilus. This could be due to, the fact that the accessory or assembly genes have not yet been identified and cloned. However, the assembly process may not function in E. coli even when all of the accessory proteins are provided, since the membranes of E. coli may not support the transport and processing of proteins from Ps. aeruginosa. Hence, work is in progress to construct an appropriate Pil- Pseudomonas mutant to provide a suitable background strain for these studies.

VI. Structure-Function Relationships of Pili Proteins Although pilus proteins are relatively small (in the range 7-28 kDa), they promote several important functions. Specific regions of the polypeptide chain are responsible for anchoring pilin to membranes, transporting pilin across membranes, interacting with various accessory proteins involved in pilus assembly, or subunit-subunit interactions. Moreover, surface-exposed regions of the pilus protein are often immunologically aWive, or function as

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recognition sites for pilus-specific bacteriophages or mammalian cell receptors. However, in some systems, pili attach to their mammalian cell receptor by means of an adhesin that is distinct from the major structural component of the pilus (Norgren et al., 1984). Both genetic and biochemical approaches are being used to identify and characterize these functional domains in various types of pili. A. CONJUGATIVE PILI

1. Chemical Composition of F Pili

F pili consist of a single, repeating subunit, pilin, of molecular weight 7200 (Moore et al., 1981a; Frost et al., 1984). The presence of an acetylatedterminus has been verified for F pilin (Frost et al., 1984),ColB2 pilin (Finlay et al., 1984)and pED208 pilin (Frost et al., 1983). The presence of carbohydrate and phosphate moieties on F-like pilin remains unresolved (Brinton, 1971; Tomoeda et al., 1975; Date et al., 1977; Willetts and Skurray, 1980). Brinton reported the presence of two phosphate groups which differed in acid lability and which were alkali-stable. One of the phosphate groups was associated with a phosphoglycopeptide with the same amino-acid composition as that of the first nine amino acids in F pilin. The presence of glucose and phosphate groups on F-like pili was investigated by G. D. Armstrong et al. (1981). They analysed the phosphate and carbohydrate content of pilin purified by gel-exclusion chromatography in the presence of sodium dodecyl sulphate (SDS). Both pED208 and ColB2 pilin contained three moles of phosphate per mole of pilin. With pED208 pilin, the phosphate was completely extracted with a chloroform-methanol mixture and was found to be derived from a mixture of phosphatidylglycerol and phosphatidylethanolamine (2 : 1 molar ratio, respectively). The composition of the E. coli K12 membrane was found to be 90% phosphatidylethanolamine and 10% phosphatidylglycerol, suggesting that pili have a greater affinity for phosphatidylglycerol or that F+ cells have an altered phospholipid composition. ColB2 pilin, with a molecular weight of 7000, is almost identical

to F pilin (see Fig. 3), and contajned 0.7-0,9rnoles ofphosphateper male of pilin after chloroform-methanol extraction. Analysis by 31P-NMR of pED208 and ColB2 pilin suggested that all pilus-associated phosphate could

be accounted for as phospholipids. However, the phosphoglycopeptidefrom F pilin reported by Brinton (1971), suggests that the phosphate residue associated with ColB2 pilin may represent a covalently bound phosphate on F-like pili which is involved in an unusual linkage not detected by NMR. Similarly, preparation of pure pilin by column chromatography in SDS removed all the carbohydrate from pED208 pilin and all but 1.O mole of D-

I

1 10 20 MET ASN ALA VAL LEU SER VAL GLN GLY ALA SER ALA PRO VAL LYS LYS LYS SER PHE PHE SER LYS PHE THR

11 I11 IV V

I I1 I11

IV V

TRP] ARG

a

40 50 LEU ALA ARG ALA VAL I L E PRO ALA ALA VAL LEU MET MET PHE PHE PRO GLN LEU ALA MET ALALEU ALA ARG ALA VAL I L E PRO ALA ALA VAL LEU MET MET PHE PHE PRO GLN LEU ALA MET ALALEU ALA ARG ALA VAL I L E PRO ALA ALA VAL LEU MET MET PHE PHE PRO GLN LEU ALA MET ALA-

30 LEU ASN MET LEU ARG LEU ASN MET LEU ARG LEU ASN MET LEU ARG LEU

20

10 I I1 I11

ARG

Ac ALA GLY SER SER GLY GLN ASP LEU MET ALA SER GLY ASN THR THR VAL LYS ALA THR PHE GLY LYS ASP

IV V

I I1 I11 IV V

I I1 I11 IV V

50

PHE PHE PHE PHE m L

LEU LEU LEU LEU E U

ALA ALA ALA VAL

40 VAL LEU VAL GLY ALA VAL MET VAL LEU VAL GLY ALA VAL MET VAL LEU VAL GLY ALA VAL MET VAL GLY ALA VAL MET U W V A L GLY[ALA]MET

LYS LYS LYS LYS MET

TRP TRP TRP TRP CYS

30 VAL VAL VAL VAL ILE

VAL VAL VAL VAL ILE

LEU ALA LEU ALA LEU ALA LE ALA IL!]ALA

GLU GLU GLU GLU G L

GLY PHE GLY PHE GLY PHE GLY PHE GLY ,LEU

ALA ALA ALA ALA VAL

ILE ILE LIE ILE VAL

ILE ILE LIE ILE VAL

SER VAL SER VAL SER VAL ER VAL SLE [VAL

60 70 PHE I L E ALA VAL GLY MET ALA VAL VAL GLY LEU PHE I L E ALA VAL GLY MET ALA VAL VAL GLY LEU PHE LIE ALA VAL GLY MET ALA VAL VAL GLY VAL GLY SER PHE I L E A LEU PHE b f ? V A L GLY ASP WPFL44)

SER SER VAL VAL SER SER VAL VAL SER SER VAL VAL SER SER I L E VAL SERWQlMET

TYR MET TYR MET TYR MET TYR MET TYR-ITHR

MET MET MET MET

m k 3

THR THR THR THR

LYS LYS LYS LYS LYS

ASN ASN ASN ASN A S

VAL YS VAL YS VAL YS VAL N E

3

LEU^

FIG. 3. The amino-acid sequence of the five types of conjugative pili including the sequence of the signal peptide. The prototype plasmids encodingeachpilus type are: I, F; 11, ColB2 or R538-1; 111, R1-19; IV, R100-1;V, pED208 (Willettsand Maule, 1986; Frost er al., 1985; B.B. Finlay, L.S. Frost and W.W. Paranchych, unpublished results). The non-homologous residues (with respect to the sequence of F propilin) are boxed.

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glucose per mole of protein from ColB2 pilin. Thus, ColB2, but not pED208 pilin, may also have a D-glucose moiety as reported for F pilin. An alternative interpretation of these observations is that the glucose and phosphate are noncovalently bound, and treatment with SDS and/or chloroform-methanol does not remove all of the phosphate and glucose from the F and ColB2 pilins. The nucleotide sequences of pilin genes from F-like pilin Types I-V showed that the major amino-acid differences occur at the N-terminus (Fig. 3). In the case of R1-19, the C-terminus contained an additional lysine residue. The sequence of the pED208 pilin gene revealed a number of conservative aminoacid substitutions in the central portion of the protein, and several substitutions involving a change in charge at the N- and C-termini. The leader sequence of pED208 propilin was significantly different from that of pilin Types I-IV, but the homology within the pilin proteins was striking. 2. The Antigenic Determinants of F-like Pili

The presence of five serotypes in F-like pili has been known for several years (Lawn and Meynell, 1970; Bradley and Meynell, 1978; Meynell, 1978). The prototypes in these studies were F, R538drd (R538-1 and ColB2 pilins are identical in sequence), R1-19drd (Rl-19), R100-ldrd (R100-1) and pED208, a derepressed derivative of FoIac (Falkow and Baron, 1962). Phenotypes R538-1 and RI-19 were stronglycross-reactive, F, R1-19, R538-1 and R100-1 were weakly cross-reactive and pED208 was serologically unique. These serotypes also correspond to the different types of F-like pili discerned by Willetts and Maule (1986) on the basis of phage-plating efficiency, which is a function of the phage attachment ability of a pilus type (Meynell, 1978). There are at least two antigenic determinants on F-like pili and the major epitope involves the N-terminus of the pilin molecule where approximately 80% of the antibodies raised against pili are directed (Worobec et al., 1983; Finlay et al., 1985). Using synthetic peptides which mimic the antigenic determinant, the acetyl group has been shown to be essential for the antigenicity of pED208 pili (Worobec et al., 1985) and F pili (Frost et al., 1986). With pED208 pili, the acetyl moiety and two leucine residues at positions 3 and 4 in the pilin protein define the major epitope of pED208 pili (Worobec et al., 1985). The nature of the secondary epitopes of F-like pili has not been chemically defined. However, the lysine residue at the C-terminus of R1-19 pilin allows these pili to be distinguished from the otherwise identical ColB2 pili, suggesting that this residue alters the secondary epitope in some way. Worobec et al. (1986) separated anti-pED208 pilus antiserum into two fractions by affinity chromatography using a synthetic peptide corresponding to the N-terminal region of pED208 pilin. Using immuno-gold labelling

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techniques, they found that the major epitope, located at the N-terminus, was not exposed on the sides of the pilus but was exposed on the surface of knobs at the base of the pilus and, perhaps, at the pilus tip. However, the pilus tip associates readily with knobs of other pili as previously noted by Brinton (1971), and it was difficult to resolve whether the observed binding of antibodies to the pilus tip was an artifact. Pili growing out from the cell did not appear to have antibodies at their tips. These results were repeated with the F pilus system using two monoclonal antibodies to F pili (Frost et al., 1986). These monoclonal antibodies reacted with two adjacent epitopes in the Nterminal region (residues 1-1 2) of F pilin and these epitopes were also exposed on the knobs at the base of the pilus although they did not appear to be at the pilus tip. Antibodies directed against other epitopes in pED208 pili were bound to the sides of the pilus. Since F and ColB2 anti-pilus antisera do not cross-react with heterologous synthetic peptides representing the N-terminal regions of these pilins, it was possible to show, by electron microscopy, that the common antigenic determinant, a secondary epitope, between these pilus types was also located on the sides of the pili. Furthermore, the anti-F pilus antiserum failed to react with the sides of R1-19 pili under the conditions employed to prepare electron-microscope specimens. However, prolonged incubation in a mild detergent uncovered this epitope (Frost et al., 1985) suggesting that the Cterminal lysine residue in RI-19 pilin masked, but did not destroy, the common epitope. These results correspond to the findings of Lawn et al. (1971) who observed “mixed” pili resulting from F and R1-19 residing in the same cell. F and R1-19 pili were unreactive with heterologous antisera, and the “mixed” pili, containing both types of subunit, reacted with the, homologous antiserum to give a patchy appearance in the electron microscope. 3. Phage Interactions with F-like Pili

Two classes of RNA phage attach to the sides of F-like pili at distinct sites. These are typified by R17 (or MS2, f2) and QB (Crawford and Gesteland, 1964) and have been extensively reviewed by Paranchych (1975). They are known to attach to the pilus by similar but distinct attachment proteins (A protein). These attachment proteins (39,000 Da) are highly insoluble and one copy is present per virion. A domain of the A protein is exposed on the phage surface and is presumably involved in phage attachment to the pilus. Based on data in Fig. 3, this involves residues 12-20 (approximately) and the last few residues at the C-terminus. In addition, there are several physical features on the pilus surface that may define the sites of phage attachment. These include the transverse grooves between each layer of subunits (1.28 nm) as well as the small longitudinal grooves between each adjacent subunit (see Fig. 2a).

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One of the amazing properties of F pili is their ability to cleave the R17A protein into two polypeptides to allow the phage RNA to leave the eclipsed phage particle and penetrate the cell (Krahn et al., 1972). The cleavage reaction does not occur in the cold or with free pili, suggesting that the process requires energy, presumably provided by the cell, which travels along the length of the pilus. This putative enzymic activity for F pilin is intriguing and leads to speculation as to the possible involvement of the elusive glucose and phosphate moieties (see Section VI.A.l). With regard to the interaction of filamentous DNA phage to conjugative pili, there are five copies of the attachment protein, pIII, at one end of the phage particle where the N-terminal portion forms a knob which is attached to the phage by the remainder of the molecule (Goldsmith and Konigsberg, 1977; J. Armstrong et al., 1981; Grant et al., 1981; Gray et al., 1981). If the F pilus contains five pilin subunits per turn of the helix, the five attachment proteins of f l phage could interact with the five pilin subunits at the tip and disassembly of the pilus and phage into the cell membrane may occur by similar mechanisms. For a review of fl phage structure and assembly, see Webster and Lopez (1985). 4 . F Pilus Interactions with Recipient Bacteria

During initiation of conjugation, the pilus tip recognizes a site on the recipient cell and a mating signal is transmitted to the donor cell which initiates DNA replication (Ou and Yura, 1982). The pilus is thought to retract into the donor cell, bringing the recipient cell surface into contact with the donor cell surface whereupon a fusion of the two cell envelopes is thought to occur, providing a conjugation bridge for the transfer of DNA between cells (Panicker and Minkley, 1985). Almost nothing is understood about the various steps in this process that involve the pilus, except that a functional pilus is an absolute requirement for conjugation. The F-pilus receptor on the recipient cell surface is the ompA-designated protein, a major outer membrane protein of E. coli (Manoil and Rosenbusch, 1982). This protein is also involved in structural maintenance of the cell envelope, and is a receptor for bacteriophages and colicins K and L (Lugtenberg and Van Alphen, 1983). It has a close association with lipopolysaccharide (LPS) such that mutations that affect the LPS often have a secondary effect on the stability of ompA in the outer membrane leading to conjugation-deficient (Con-) recipient cells (Schweizer and Henning, 1977; Schweizer et al., 1978; Manoil and Rosenbusch, 1982). The N-terminal half of the ompA protein traverses the outer membrane repeatedly in a cross-beta sheet conformation, exposing four regions on the cell surface (Morona et al., 1984). Two of these regions, around residues 25 and 154, are thought to be

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W. PARANCHYCH AND L. S. FROS?

involved in defining the conjugational proficiency of F- cells by acting as the receptor site for either the F pilus tip itself or another protein in the donor cell membrane (Morona et al., 1985). Although R1-19 (pilus Type 111) and F (Type I) appear to share the same R100-1 (Type IV) apparently receptor to varying degrees on F- cells (ompA), has a different receptor which involves LPS (Havekes et al., 1977a). IncI plasmids (R144) have yet another receptor involving LPS alone (Havekes et al., 1977b). 5. F-like Pili and Surface Exclusion

Another receptor for the pilus tip is an outer-membrane protein (TraTp) which is part of the surface (or entry) exclusion system encoded by conjugative plasmids (Lederberg et al., 1952; Willetts and Maule, 1974; Minkley and Ippen-Ihler, 1977; Achtman et al., 1977; 1980). This protein, encoded by IncF plasmids, is expressed by the traT gene in the transfer operon (Kennedy et al., 1977; Manning et al., 1980). It blocks mating-pair formation between related donor cells by preventing mating-pair stabilization (Achtman et al., 1977). Protein F TraTp is thought to have intimate contact with the ompA protein, blocking the site on OmpAp, which is the receptor for the F pilus. The traT protein also has sequence homology to the OmpAp-recognizing portion of the OmpAp-specific phages (Riede and Eschbach, 1986). Protein TraTp is a highly expressed lipoprotein (Perumal and Minkley, 1984)which may interact with the tip of the pilus and thus prevent donor-recipient cell recognition (Willetts and Maule, 1974, 1986; Minkley and Willetts, 1984). The second transfer gene involved in surface exclusion, traS, which maps immediately upstream from traT(Achtman et al., 1980), encodes a gene product which is associated with the inner membrane and appears to inhibit DNA transfer (Achtman et al., 1977; Manning and Achtman, 1979). F-like plasmids are capable of expressing surface-exclusion systems which are distinct from that of the F plasmid (Alfaro and Willetts, 1972; Willetts and Maule, 1974). Willetts and M a d e (1986) defined four surface-exclusion variants which, for the most part, correspond to the different alleles for F-like pili. A fifth surface-exclusion system would be that of pED208 (Finlay and Paranchych, 1986). The sequence of the traT genes of F, pED208 (Finlay and Paranchych, 1986; E. G. Minkley, personal communication) and R100-1 (Ogata et al., 1982) are available. The homology between the three proteins is striking, and the difference between F and R100-1 TraTp lies in one aminoacid substitution (Gly->Ala) in the middle of the protein. If the pilus tip recognizes TraTp, then it reacts to very subtle changes in sequence of F and R 100-1. Another possibility is that pilus-TraTp recognition may be rather non-specific and the stringency of the surface-exclusion phenomenon may be

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encoded by the traS gene product which appears to be unique in each case (Hansen et al., 1982; Ogata et al., 1982; Finlay and Paranchych, 1986; E. G . Minkley, personal communication). 6 . Surface Features of F-like Pili

The pilus is capable of interacting with a number of biological macromolecules. The tip of the pilus is the site of interaction with the cell surface of the recipient cell during mating-pair formation (Ou and Anderson, 1970; Achtman and Skurray, 1977) and other donor cells during the process of surface exclusion (Willetts and Maule, 1974). The tip is also the site of attachment of filamentous DNA phages M 13, fl ,fd (Caro and Schnoss, 1966) and the sides of the pilus are the sites of attachment for the spherical RNA phages R17 and QB (Crawford and Gesteland, 1964; Paranchych, 1975). In addition, antibodies are capable of recognizing epitopes on the pilus surface. One approach to elucidating pilus structure has been the identification of pilus variants which have an altered traA gene affecting phage attachment, donor ability or pili per cell. These include naturally occurring variants such as those specified by closely related IncF plasmids (Fig. 3; Paranchych, 1975; Frost et al., 1985; Willetts and Maule, 1986) and point mutants of the F plasmid itself (Silverman et al., 1967, 1968; Tomoeda et al., 1972; Orosz and Wootton, 1977; Burke et al., 1979; Willetts et al., 1980). Many F pilus variants have altered susceptibility to phages R17 or QB but retain fl sensitivity. No transfer-proficient, fl resistant variant has been isolated, suggesting that transfer, pilus outgrowth and fl attachment are closely linked. Figure 3 shows the amino-acid substitutions for four traA point mutants previously described (Willetts et ai., 1980) and summarized in Table 2. Taken together, the phage sensitivity patterns (Types I-V) and the information on the immunodominant regions of F-like pilin allow certain surface domains of the pilin subunit to be identified. The N-terminus may be exposed at the tip of F-like pili in a unique configuration and provide the specificity for pilus-related phenomena. However, electron microscopy has not clearly indicated this (see Section VI.A.2). Other types of conjugative pili have pointed tips which are clearly visible by electron microscopy and appear to be the site of attachment for a variety of filamentous phages (see Table I). These points appear to be conical arrangements of subunits instead of the tubular arrangement found in the pilus shaft. It is possible that F-like pili also have a unique configuration of pilin subunits at their tips, but that these are not clearly resolved by electron microscopy. Meynell et al. (1974) raised antisera to sheared pili in order to enrich for antibodies to the ends of pili. They found that these antibodies were unable to distinguish between F and Rl-19 pili, suggesting that a shared

W. PARANCHYCH AND

90

L. S. FROST

TABLE 2. Pilus types, efficiency of phage plating and effect of cyanide on levels of piliation of Escherichia coli strains with various F-like plasmids and their derivatives. From Willetts et al. (1980) and Frost et al. (1985). Efficiency of plating F-like plasmid

Pilus type

Inc group

F R538-1 COLB2 COLB4 RI-19 R100-I EDP208

I I1 I1 I1 111 IV V

FI F11 FII FII FII FII FV

fl

R17

QB

Pili per 100cells

100

100

90 1 3

70 86 120 60 8 0

100 95 115 117 3 3 0

143 212 360 164 180 108 1700

10

2 100

Efficiency of plating traA mutants

Donor ability

WPFL44 WPFL46 WPFL47 WPFL51

1I7

5 14 55

fl 79 R 74 1

R17

QB

Pili per 100cells

6 R R 20

R R 0.2 R

325 14 98 145

Treated with cyanide (20min) 14 37 374 80 40 22 1100 Treated with cyanide (20min) 121 6 7 0

Addition of cyanide indicates the retraction ability of that pilus type. R indicates resistance. Inc is the abbreviation for “incompatibility”.

epitope is exposed at the ends of broken pili. Experiments with two monoclonal antibodies to F pili support this hypothesis. F pili that had been sheared by sonication generated new binding sites for the monoclonal antibody (JEL92) that recognized the region near Met-9, whereas no new sites were created for the antibody (JEL93) that recognized the N-terminus (Frost et al., 1986). Jacobson (1972) made the observation that sheared pili are capable of attaching fl phage, suggesting that the N-terminal region (residues 1-12) may be exposed at the tip of an intact pilus, while the region around Met-9 is presumably exposed at the tip of broken pili and would be involved in f l attachment. In addition, the lysine residue at position 10 in R100-1 is probably partly responsible for the low phage-fl infectivity in bacteria carrying this plasmid. With the exception of pED208, a charged residue at the C-terminus (Rl-19 and WPFL44) also affects fl sensitivity suggesting that the N- and C-termini are in close apposition. If the glycophosphopeptide reported by Brinton (1971) is correct, then the glucose and phosphate moieties would be within the first nine residues of F pilin and would not be exposed on the sides of the pilus but would be exposed at the tip or buried within the structure. Since pED208 (which is glucose- and

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phosphate-free) is fully sensitive to f l ,the phosphate and glucose would not be important in fl infection, However, they could play a role in R17 or QB infection since pED208 is completely resistant to these phage. A region roughly corresponding to residues 12-22, as well as several residues near the C-terminus, may be exposed on the lateral surface of the pilus. Mutations in these regions affect R17 and QB attachment and efficiency ofplating (see Table 2 and Fig. 3). Charged residuesat the C-terminus (RI-19, pED208 and WPFL44) lower QB sensitivity more than that of phages R 17 or fl, whereas non-conservative substitutions within residues 11-22, usually involving a change of charge, greatly affect R17 phage sensitivity with lesser effects on QB and fl (R100-1, pED208, WPFL47 and WPFL46). The mutation in WPFL46 decreases pilus number and donor ability suggesting that pilus assembly has been affected and that the region after residue 20 may be involved in holding the subunits together. The central region of the pilin molecule is highly hydrophobic and is probably involved in the quarternary structure of the pilus where at least four surfaces of the molecule take part in subunit-subunit interactions. Minkley et al. (1976) observed that whole pili could not be iodinated with Chloramine-T, suggestingthat the tyrosine residue at position 42 is inaccessible. The low PI of pili (pH 4.15; Brinton, 1971) is puzzling since the three acidic residues are outnumbered by the five lysine residues. If the phospholipid content of pili is not contributing to its acidic nature (see Section VI.A.I), then the acidic residues must be on the pilus surface while the basic residues would be buried within the structure. Although there are several reports of trace amounts of other proteins that have been reported in pilus preparations (Date et af., 1977; Helmuth and Achtman, 1978), no other protein has routinely been found associated with pili either as a tip protein or a basal protein. Recently, a small amount of protein of molecular weight 8000 has been found in preparations of F pili examined by SDS-PAGE and stained with a silver staining reagent (K. IppenIhler, personal communication). If the pilus tip recognizes all three proteins, namely the N-terminal portion of the fl phage protein (pIII) (Hill and Petersen, 1982), the sequences near residues 25 and 154 in OmpA (Chen et al., 1980) and the surface-exclusion protein TraTp (Finlay and Paranchych, 1986), then some common aminoacid sequence among these proteins might be expected to serve as an F-pilus attachment site. A comparison of these sequences was undertaken to determine whether any sequence homology between these three proteins is detectable. No strong homology was found, suggesting that either different domains on the F pilus ate involved in these interactions or other features in an F + cell are the true receptors in these processes.

92

W. PARANCHYCH AND L. S. FROST

7 . F Pilus Assembly and Retraction

The F pilus contains only a single repeating subunit, and free pili do not appear to contain other proteins arranged in a basal structure such as is found on flagella (Iino, 1969). Recently, Silverman (1987) detected a small structure in sphaeroplasts of E. coli which spans the entire cell envelope. This agrees with the proposal by a number of workers that the F pilus originates from a plasmid-encoded organelle spanning a junction of the inner and outer membranes (Bayer, 1975; Achtman and Skurray, 1977; Laine et al., 1985). The steps involved in assembly of a pilus presumably involve expression of the pilin gene product, propilin, processing of propilin to pilin, modification of the pilin subunit (acetylation and, possibly, phosphorylation and glycosylation), formation of a pilin pool in the membrane at the site of pilus synthesis, and, presumably, assembly of the various transfer-gene products and host factors required for pilus outgrowth and retraction. There may also be a site at the base of the pilus for a membrane complex involving the plasmid DNA, awaiting the signal to begin transfer after establishment of a stable mating pair. All F-like pili have an extremely long leader sequence which is twice the average length for a prokaryotic leader sequence (Michaelis and Beckwith, 1982; Perlman and Halvorson, 1983). Ippen-Ihler and her coworkers have studied the steps in processing of the pilin precursor to mature pilin and have published a review of their findings (Laine et al., 1985; Ippen-Ihler and Minkley, 1986). In short, they suggest that the 14,000-Da pilin precursor protein is processed to the mature pilin subunit of 7000 Da in the presence of the traQ gene product. There may be a two-step processing mechanism with an 8000 dalton intermediate. The 7000 Da polypeptide (Ap7) does not react with anti-F pilus antisera but a slightly more slowly migrating polypeptide called Ap7* reacts with these antibodies. Conversion of Ap7 into AP7* is facilitated by traG and may correspond to the acetylation event required for formation of antigenically competent pilin. Processing of propilin to pilin occurs in the inner membrane as does the acetylation reaction since all four species of pilin can be found at this location (Ap 14,8,7,7*) while only Ap7* is found in the outer membrane suggesting that only the mature form is polymerized into pilin. The traQ gene product is required for efficient processing to occur, even though the cleavage site at the N-terminus strongly resembles the preferred substrate for the signal peptidase of E. coli Ala-X-Ala-Ala (Perlman and Halvorson, 1983).The requirement for electrochemical potential within the membrane for protein translocation (reviewed in Wickner and Lodish, 1985) is interesting in that pilus outgrowth is also energy-requiring and the two processes may be linked. The process of F-pilus outgrowth and retraction is controversial and

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evidence in favour of retraction is circumstantial. Several models for pilus function have been advanced (Brinton, 1971). The most widely accepted model involves pilus outgrowth and retraction being in equilibrium where energy is required for pilus assembly (Marvin and Hohn, 1969; Curtiss et al., 1969; Novotny and Fives-Taylor, 1974; O’Callaghan et al., 1978). Interfering with the energy source by addition of cyanide causes the pili to disappear. This was presumed to be evidence of pilus retraction since there was no increase in the numbers of F pili or pilin in the culture supernatant (Novotny and FivesTaylor, 1974). Similar results were obtained following addition of arsenate providing the cells were grown in minimal media on a carbon source other than glucose (OCallaghan et al., 1973b, 1978). Heating the cells to 50°C caused pilus retraction although this was thought to be by a different mechanism (Novotny and Fives-Taylor, 1978). Cooling cultures to 20°C caused pili to be shed into the medium while quick chilling to 0°C prevented this (Novotny and Lavin, 1971). Furthermore, addition of anti-pilus antibodies or RNA phages, which bind to the sides of the pilus, prevents disappearance of F pili presumably by physically blocking the retraction mechanism. These experiments would suggest that pilus outgrowth requires energy whereas retraction does not, and that conditions that block outgrowth may also cause retraction. Plasmid ColB2, an Inc FII plasmid closely related to the Inc F1 F plasmid, is insensitive to cyanide in that its pili do not retract (Frost et al., 1986). Similarly, retraction-minus mutants of F have been isolated (Burke et al., 1979), whereas ColB4 and pED208 have intermediate levels of retraction in the presence of cyanide and could represent natural mutants in the gene(s) responsible for retraction. The best evidence for F-pilus retnaction comes from experiments with filamentous bacteriophage where phage attachment to F pili has been followed by a visible shortening of the pili on the cells (Jacobson, 1972). However, Paranchych and his coworkers reported that RNA- and DNAphage infection was accompanied by an increase in pili fragments in the medium (Paranchych et al., 1971;OCallaghan et al., 1973a).Achtman and his coworkers have elaborated on the initial observations of a donor and recipient cell connected by a pilus (shown in Brinton, 1971; Ou and Anderson, 1970) and demonstrated that the mating pairs form large aggregates which are stabilized by the activity of traN and traG (Achtman et al., 1978a, b). Initial contacts between mating cells are easily disrupted by SDS which dissociates the pili while subsequent steps, when the mating cells have fused together, are SDS-resistant. Although no real evidence for pilus retraction can be proposed on the basis of these experiments, it is an attractive possibility. Whether pili retract into the donor cell or depolymerize into the recipient cell membrane is unknown. Preliminary efforts to identify transfer proteins in the recipient cell, especially pilin, were not successful (W. Paranchych, unpublished observations).

94

W. PARANCHYCH AND L. S. FROST

B. ADHESIVE PILI OF

Escherichia coli

Elucidation of the primary structures of the K99, Pap, and Type 1 pilins has revealed several common characteristics: (a) they are approximately 18 kDa in size, (b) the N-terminus is hydrophobic, (c) they show significant homology at the N- and C-termini, (d) there is a cysteine loop in the N-terminal half of the protein and (e) they have a penultimate tyrosine residue at the C-terminus (Mooi and de Graaf, 1985; Nonnark et al., 1985).Although homologies at the N- and C-termini are also present in CFA/I (14 kDa) and K88 (around 27 kDa) pilins, these proteins are significantly different in size and lack the cysteine loop (Mooi and de Graaf, 1985). It is worth noting that predictions concerning secondary structure and hydrophilicity in all of these pilins give similar profiles (Baga et al., 1984; Klemm, 1984; Roosendaal et al., 1984), suggesting that they contain analogous structural domains. Most of the above-mentioned pilins from E. coli contain highly hydrophobic amino-acid residues in the C-terminal part of the molecule, suggesting that these regions may be involved in subunit-subunit interactions (Klemm, 1985; Mooi and de Graaf, 1985). Three positions, in particular, contain highly conserved aromatic amino-acid residues. Such residues (particularly tyrosine), have been implicated in the maintenance of pilus structure in both Ps. aeruginosa (Watts et al., 1983a) and Type 1 (McMichael and Ou, 1979). Surface domains of pilus proteins in intact pili are presumably immunogenic towards the host’s immune system. Since these regions are not directly involved in maintenance of the pilus structure, they usually tolerate significant amino-acid changes, and this provides a means of evading the host’s immune system through antigenic variation. Examples of this are seen among K88 pili, which contain at least three serological subtypes, ab, ac and ad (Klemm and Mikkelsen, 1982; Dykes et af., 1985), Type 1 (Salit e l a[., 1983), and Pap pili (Nonnark et al., 1985; Mooi and de Graaf, 1985; Klemm, 1985). In all cases, the antigenic determinants were inferred on the basis of secondary structure and hydrophilicity predictions, and a comparison of amino-acid sequences in related types of pili. Only with Ps. aeruginosa (Watts et al., 1983b; Sastry et af., 1985a) and N. gonorrhoeae (Schoolnik et al., 1984, 1985) pilins have the proteins been mapped for antigenic determinants by means of proteolytic and chemical cleavage of pilin, and immunological characterization of the resulting peptides. Little is known about the localization of receptor-binding domains in adherence pili in E. coli. As already mentioned (Section III.C.2), Type 1 pili promote MS haemagglutination, whereas most other pathogenic strains promote MR haemagglutination. Pap pili bind to the a-D-Gal-(l+4))-fi-DGal unit of the P blood-group antigens of uroepithelial cells, while K88, K99, CFA/I, CFA/II and some less frequently occurring non-digalactoside-specific

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95

MR pili, termed X-adhesins (Vaisanen et al., 1981), are associated with intestinal epithelium. Interaction of these pili with eukaryotic cells has long been thought to be mediated directly by pilin, since purified pili have been shown to mediate agglutination of erythrocytes and other eukaryotic cells, and to inhibit competitively binding of intact bacteria. However, recent studies have demonstrated that, in the case of Pap and Type 1 pili, the adhesin (i.e. the structure that actually mediates bacterial cell binding) is distinct from the monomer making up the pilus (Norgren et al., 1984; Maurer and Orndorff, 1985; Normark et al., 1985; Minion et al., 1986). Although the location of the adhesin molecule in the pilus has not yet been determined, there is good reason to believe that one or more minor proteins promoting the adhesin function are positioned at the pilus tip (Mooi and de Graaf, 1985; Normark et al., 1985). Thus, it is possible that one of the roles of the major structural units is extension of the adhesin molecule some distance from the cell surface to facilitate interaction with the host receptor. Two different subunit genes have also been located within the operon in the case of K88 pili (Mooi and de Graaf, 1985). The large subunit (26 kDa) constitutes the major component of K88ab pili. The existence of the small subunit (17.6 kDa) was inferred from DNA sequence data and the analysis of K88ab mutants. Although it has never been detected as a minor component of K88ab pili, mutants lacking this protein were unable to agglutinate erythrocytes or bind to intestinal epithelial cells. One interpretation for this observation was that the 17.6 kDa protein may serve as an adhesin (Mooi and de Graaf, 1985). However, Jacobs et al. (1987) have shown that two tripeptides (Ser-Leu-Phe, Ala-Ile-Phe) from the major pili subunit (p26) of K88 pili are effective in inhibiting the haemagglutinating activity of purified K88 pili and the adherence of pili to intestinal epithelial cells. Of particular interest was the fact that the region surrounding the Ser-Leu-Phe sequence showed significant homology to gonococcal pilin, suggesting that this region may also promote adherence in the case of N. gonorrhoeae pili, and that it may represent the receptor binding domain in both K88 and gonococcal pili. Nucleotide sequencing of pilin genes from E. coli, and pilin-expression studies have shown that MS and MR pilins are exported across the bacterial cytoplasmic membrane, and that they are all endowed with N-terminal signal sequences that confer the membrane translocation function (Klemm, 1985; Mooi and De Graaf, 1985; Normark et al., 1985). In order for the subunits to be processed, translocated and assembled into intact pili, various auxiliary proteins are needed, and it has been found that three to six such helper proteins exist. However, the mode of pilin interaction with these accessory proteins has not yet been elucidated (Klemm, 1985; Mooi and de Graaf, 1985; Normark et al., 1985).

96

W. PARANCHYCH AND L. S . FROST

C. PILI DESIGNATED

NMePhe

NMePhe Pili are expressed by Ps. aeruginosa, N. gonorrhoeae, N . n z @ g & f d i s , M . nonliquifaciens, M . bovis, B. nodosus and V. cholera (see Section III.C.3). To date, structure-function studies have been performed primarily on pili from Ps. aeruginosa and N . gonorrhoeae. These studies are summarized below. 1 . Pilifrom Pseudomonas aeruginosa

Pili from Ps. aeruginosa are multi-functional organelles which mediate adherence of Ps. aeruginosa to human buccal epithelial cells (Woods et al., 1980; McEachran and Irvin, 1985; Paranchych et al., 1986), human tracheal epithelial cells (Ramphal et al., 1984; Palmer et al., 1986; R. T. Irvin, unpublished observations) and human cornea (Reichert et al., 1982). The nature of the pilus receptor on human epithelial cells is not yet understood. Paranchych et al. (1985) measured pilus binding to a variety of synthetic sugars representing many di-, tri- and tetra-saccharide structures found in mammalian glycoproteins and glycolipids, and failed to reveal any significant binding to any of the sugar moieties examined. However, a wide spectrum of binding activities was observed when a variety of proteins and enzymes were used as binding substrates. Of 30 proteins tested, phosphorylase b, pyruvate kinase and aldolase showed highest pilus-binding activity, whereas proteins such as cytochrome c, trypsin and band-3 protein bound pili very poorly. It was concluded that pili may have been recognizing a polypeptide domain that resembles the true pilus receptor, suggesting that it is perhaps a protein rather than oligosaccharide in Nature. Pili from Ps. aeruginosa also promote a phenomenon known as “twitching motility” (Bradley, 1980c), and they serve as receptors for a number of pilusspecific bacteriophages for Ps. aeruginosa including RNA-containing types (Feary et al., 1964; Bradley, 1972b), DNA-containing filamentous forms (Takeya and Amako, 1966; Bradley, 1973a) and forms containing heads and long non-contractile tails (Bradley, 1973b). Both twitching motility and phage sensitivity require that pili are retractile, i.e. able to undergo a process known as “retraction” (Bradley, 1972b, c, 1974a, b, 1980~).Mutants of Ps. aeruginosa which have non-retractile pili are multipiliated, phage-resistant and unable to promote twitching motility (Bradley, 1974b, 1980~).One such mutant of strain Ps. aeruginosa K, called PAK/2Pfs (Bradley, 1974b) was instrumental in facilitating the first purification and characterization of polar pili from Ps. aeruginosa (Frost and Paranchych, 1977; Frost et al., 1978; Paranchych et al., 1978). Recently, W. Paranchych and his colleagues obtained direct evidence that pili from Ps. aeruginosa are important virulence factors in pathogenesis

PHYSIOLOGY AND BIOCHEMISTRY OF PILI

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attributable to the bacterium. Using the thermal (Stieritz and Holder, 1975), neutropenic (Cryz et al., 1983) and CaClz (Tamura and Tanaka, 1985)mouse models, the LD50 values obtained with a Pil- strain of Ps. aeruginosa P A 0 were between lo2 to lo4 higher than those obtained with the isogenic Pil+ strain. Moreover, when mice were injected with 50 p1 of anti-pilus IgG at the time of bacterial challenge, the LDsOvalues were about lo4 higher than in control mice injected with non-immune IgG. It is therefore evident that polar pili from Ps. aeruginosa represent an important virulence factor in pathogenesis. The antigenic regions of the pilus protein of Ps. aeruginosa PAK were determined by Watts et al. (1983b) and Sastry et al. (1985a). Arginine-specific cleavage was used for the initial fragmentation of the pilus protein into four large peptide fragments called cTI (1-30), cTII (31-53), cTIII (54-120) and cTIV (121-144). Fragments cTIII and cTIV were further cleaved into a variety of subfragments. The various peptide fragments were subjected to immunoblot and direct ELISA studies, as well as competition ELISA experiments. Four distinct epitopes were identified: one weak (reacting with less than 5% of the anti-plus antibodies) cross-reactive epitope in the N-terminal region (130), a moderately immunogenic epitope (reacting with 20% of the anti-pilus antibodies)in fragment 70-81, a strongly immunogenic epitope (reacting with 60% of the anti-pilus antibodies) in fragment 82-1 10, and a weak (reacting with less than 5% of the anti-pilus antibodies) epitope in fragment 128-144. The cross-reactive epitope in the N-terminal region was detectable only in denatured pili, suggesting that this region of the protein is buried at subunit interfaces in intact pili. This conclusion was supported by spectral studies performed at PAK pili by Watts et al. (1983a), who showed that Tyr-24, Tyr27 and at least one Trp are involved in subunit-subunit interactions. The epitopes in cTIII and cTIV were considered to be type-specific, since they were not cross-reactive with pilus-specific antiserum raised against the closely related Pseudomonas strain PAO. Moreover, it was evident from competition ELISA studies that the immunodominant epitopes in cTIII were at least partially conformation dependent (Sastry et al., 1985a). The fact that the antigenic determinants in cTIII and cTIV are type-specific suggests that several different pili serotypes exist among naturally occurring strains of Ps. aeruginosa. However, the extent of this serological polymorphism is presently unknown. Bradley and Pitt (1975) and Woods et al. (1982) identified at least five distinct serotypes. W. Paranchych and his colleagues (unpublished observations) screened 64 clinical isolates for the PAK and P A 0 serotypes using whole-cell immunoblot assays, and found that about 34% were PAK-specific and 20% were PAO-specific. In addition, the amino-acid sequences of four different pilus types are now known (Sastry et al., 1985b; Johnson et al., 1986b; W. Paranchych and his colleagues, unpublished observations) and are shown in Fig. 4.

PAK PA0 CD4 PA103

Met Met Met Met

-5 Lys Lys Lys Lys

+1 Ala Ala Ala Ala

Gln Gln Gln Gln

Lys Lys Lys Lys

Glu Glu Glu Glu

Gly Ala Gly A l a Gly A l a Gly Ala

Ser Ser Ser Ser

45 Thr Thr Thr Thr

Thr Thr Thr Thr

Val Val Val Val

Glu Glu Glu Glu

Leu Leu Leu Leu

Lys Lys Lys Lys

Asn Asn Asn Asn

15 le le le le

PAK PA0 CD4 PA103

10 Leu Leu Leu Leu

Met Met Met Met

I I I I

l l l l

e e e e

Val Val Val Val

Val Val Val Val

e e e e

Pro Pro Pro Pro

Gln Gln Gln Gln

Tyr Tyr Tyr Tyr

25 Gln Gln Gln Gln

Ala Ala Ala Ala

Leu Leu Leu Leu

A l Ala Ala Ala

a [ Thr Thr Thr

k i Ile Ile Ile

G l u A l a Leu G l u e r Leu G l u Ser Leu G l u Ser Leu

Ser Ser Ser Ser

Arg Arg Arg Arg

Gly Gly Gly gly

Trp Ile Ile Ile

I I I I

l l l l

30

PAK [Ser V a l L y s S e r PA0 A l a Gly S e r L y s CD4 A l a Gly Ser L y s PA103 A l a G l y S e r L y s

I I I I

Ser Ser Ser Ser

Pro Pro Pro Pro

PAK PA0 CD4 PA103

l l l l

5 Glu Glu Glu Glu

Arg Arg Arg Arg

Ala Ala Ala Ala

I I I I

e e e e

20 Ala Ala Ala Ala

Val Val Val Val

Asn Asn Asn Asn

e e e e

l l l l

e e e e

Tyr Tyr Tyr Tyr

PAK PA0 CD4 PA103

l l l l

I I I I

Leu Leu Leu Leu

Ala Ala Ala Ala

I I I I

I I I I

Thr Thr Thr Thr

Ala Ala Ala Ala

Gly Gly Gly Gly

Ala Ala Ala Ala

Phe Phe Phe Phe

Leu Leu Leu Leu

e e e e

PAK PA0 CD4 PA103

Gly Gly Gly gly

l l l l

35

55

Q

60 G l y Thr Gly Thf G l u A Ile Ile A l a Ser T h r A l a Asp I l e Leu I l e G l y T h r T h r A l a S e r T h r A l a Asp

:G: y r Va Gly T h r T h r T y r V a l Gly T h r T h r T y r Val G l y

Val Val Ile Ile

(RlamqAs; G l u P r o As Asp G l u L y s Asp G l u L y s

90 PAK I l e A l a Leu Lys P r o PA0 ~ A l a ~ \ A s p CD4 V a l A l a Val T h r I l e L y s PA103 Val A l a V a l T h r I l e L y s

~

~

Asp P r o ~ + & Asp T h r Asp T h r

80 Ala Ala Ala Ala

95 A l a As G l y G l y Asp Gly Asp

Asn Asn Asn Asn

Lys Lys Lys Lys

Leu Leu Leu Leu

Gly G l Gly Gly

85 Thr y m Thr Thr

100 G l y M A l a Asp I l e ]ASP I l e + G l y T h r Val L v s Gly Thr Ls;

(Ilel

PAK T h r a T h r Phe PA0 &Phe T h r Phe C04 Phe T h r Phe A l a T h r G l y G l n S e r S e r P r o L y s Asn A l a Gly PA103 P h e m P h e A l a T h r Gly G i n S e r Ser P r o L y s Asn A l a Gly PAK Lys I l e PA0 l y s m CD4 L s Glu PA103 & G l u PAK

I I I I

l l l l

e e e e

Cys T h r Ser

Thr Thr Thr Thr

120 Leu[ThrlArg L e u Asn A r g L e u Asn A r g L e u Asn A r g

Thr Thr Thr Thr

Ala&Asp Ala Ala Glu Ala Glu

Phe P h T h r G l n G l u G l u M e t Phe T h r G l n G l u G l u M e t Phe

130 Lys p a Thr G l y Val T r p T h r

l y m T r p aGGly Val T r Gly Val T r p

140 I l e Pro e m P r o I l e Pro I l e Pro

Lys Lys Lys Lys

Gly Gly Gly Gly

Cys Cys Cys Asn L s P r o Cys A s n h P r o

FIG. 4. The amino-acid sequence of pilins from Pesudomonas aeruginosa strains PAK, PAO, CD4 and PA103. Boxed areas represent regions of non-homologous amino-acid sequence or spaces created to obtain maximum alignment. The PAK and PA0 sequences were obtained from Sastry et al. (1985a). The sequence of CD4 (a clinical isolate of Ps. aeruginosa obtained from a patient with cystic fibrosis) represents unpublished data of W. Paranchych and his colleagues. The sequence of PA103 (unpublished data) was provided by Kit Johnson and Steve Lory (personal communication).

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99

Three of the pilins described in Fig. 4 (PAO, CD4 and PA103) are serologically identical (PA0 type), whereas PAK is a unique serotype. There is about 70% homology between the PAO-specific pilins and PAK, about 80% homology between P A 0 and CD4, while CD4 and PA103 are almost identical (97% homology). The N-terminal domain (1-55) is very highly conserved and is referred to as the “constant region”. The greatest variation occurs in the central part of the molecule (56-1 10) which contains the major antigenic determinants in PAK pilin, and presumably in the P A 0 types as well. The Cterminal region (1 1 1-147) contains few significant amino-acid differences among the four proteins, and is therefore referred to as the semiconserved region. This domain contains the highly conserved cysteine loop within which is located a weak antigenic determinant (Sastry et af., 1985a). This region has also been implicated in recognition of the pilus receptor on human buccal epithelial cells (Paranchych et af.,1986; R . T. Irvin, P. Doig, W. Paranchych and P. A. Sastry, unpublished observations). The cysteine loop and the highly immunogenic epitopes in the central portion of the molecule are presumably surface-exposed regions of the intact pilus, and it is believed that the phageattachment sites of the pilus protein will also be located within these putative surface-exposed domains. Determination of the nucleotide sequence of pilin genes in Ps. aeruginosa has revealed a highly conserved, positively charged six-residue leader sequence (Pasloske et af., 1985; Sastry et af., 1985b). Although the mechanism of pilin processing is not yet understood, it is clear that it must involve removal of the six-residue leader and N-methylation of the resulting N-terminal Phe residue. Pilus assembly presumably occurs in the outer membrane or at sites where the inner and outer membrane are fused to form adhesion zones (Bayer, 1975). Watts et a f .(1982) observed approximately equal amounts of pilin in the inner and outer membranes of Ps. aeruginosa, although they did not consider the possibility that one or both of these pilin pools may be unprocessed. Finlay et al. (1986c) showed that a chimera containing the PAK pilin gene was expressed in E. coli minicells and that both unprocessed and apparently processed pilin was present in the inner membrane. More recent studies in this laboratory (B. L. Pasloske, B. B. Finlay, L. S. Frost and W. Paranchych, unpublished observation) have shown that the sixresidue leader of pilin in Ps. aeruginosa is not processed in E. coli, and that the C-terminal half of the protein interacts with the N-terminal half to stabilize the pilus protein in the inner membrane. Truncated PAK pilin mutants encoding amino-acid residues -6 to 30 (36-mer) and -6 to +59 (65-mer) were constructed using recombinant DNA techniques, and expression of these products was compared with that of normal pilin. Each of the three genes (encoding normal pilin, the 36-mer and the 65-mer) was inserted into a highexpression vector system utilizing the T7 polymerase arid its promoter (Tabor

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and Richardson, 1985). The results of these studies showed that the hydrophobic N-terminal region (36-mer) is expressed in E. coli and inserted as a stable entity into the inner membrane. The 65-mer was also expressed and inserted into the inner membrane, but this polypeptide was gradually degraded to a product similar in size to the 36-mer. Intact pilin, which was also expressed well in this system and inserted into the inner membrane, was extremely stable and not degraded at all. These observations suggested that only the first 36 amino acids of propilin are required for membrane integration and that the C-terminal region of propilin may associate with the N-terminal region to stabilize the region between residues 3&65 against degradative processes. It is not yet known whether pilin integration into the inner membrane behaves similarly in Ps. aeruginosa, nor whether processing of propilin occurs in the inner or outer membranes. Moreover, the number of accessory gene products required for pilus assembly in Ps. aeruginosa has not yet been established.

2. Pili from Neisseria gonorrhoeae Strains of Neisseria gonorrhoeae are notable for their ability to vary the antigenic properties of their pili. Gonococcal pilins from clinical isolates are extremely heterogeneous with regard to size (Lamden et al., 1979), antigenicity (Buchanan, 1975) and amino-acid sequence (Hagblom et al., 1985; Haas and Meyer, 1986). The relationship between gonococcal virulence and a piliated phenotype was first noted by Swanson et al. (1971). Gonococcal pili were subsequently shown to be involved in haernagglutination (Buchanan and Pearce, 1976), attachment to spermatozoa (James et al., 1976), human buccal epithelial cells (Punsalang and Sawyer, 1973), epithelial cells in culture (Swanson, 1973), human endocervical cells (Schoolnik et al., 1984) and human fallopian tubes in organ culture (Ward et al., 1974; McGee et al., 1981). However, since no animal model exists for gonorrhea, it has not been possible to test directly the degree of virulence conferred by gonococcal pili. Although the pilus is apparently required for initial colonization of the genitourinary tract, a second surface antigen, the opacity protein (Op or PII) is also involved in adhesion of N. gonorrhoeae to the human mucosa (Lambden et al., 1979; Blake and Gotschlich, 1983). Thus, the role of pili as a virulence factor is inferred from the observed correlation between virulence and piliation. Application of the primer-extension nucleotide-sequencing technique to mRNA for gonococcal pilin has resulted in the determination of a large number of pilin-DNA sequences from a variety of gonococcal strains (Hagblom et al., 1985; Haas and Meyer, 1986; Bergstrom et al., 1986).

101

PHYSIOLOGY AND BIOCHEMlSTRY OF PlLl

I - - - _ _. _ _ -. _ _ _ _ j-1_ _ ._ _ I, _ _ _ - 1MePhe

48

64 68

- - - - - - -. -

86 90

- _0 _ _ - _

104110 116

,

126 121

I

CYS

_.__

144

154 151

I

CYS

Constant region Semivarioble region Hypervariable region

FIG. 5. Schematic diagram showing distribution of the constant, semivariable and hypervariable regions in gonococcal pilus proteins. Based on information in Hagblom et al. (1 989, Haas and Meyer (1986) and Bergstrom et af. ( 1 986).

Comparison of these sequences has shown that the pilin gene can be divided into constant, semivariable and hypervariable regions (Fig. 5). Nucleotide changes within the semivariable regions usually involve single-codon changes, while the hypervariable region has been shown to undergo single-codon substitutions as well as in-frame insertions and deletions of one to four codons at a time. These changes, particularly within hypervariable regions, result in the appearance of unique epitopes on the pilus (Hagblom et al., 1985; Rothbard et af., 1985; Sparling er al., 1986). Several attempts have been made to delineate the antigenic determinants in gonococcal pilins. Schoolnik et al. (1984) determined the complete amino-acid sequence of pilin isolated from gonococcal strain MSI 1 and the partial sequence of pilin from strain R10. The proteins were cleaved with CNBr to yield three fragments: CNBr-I (1-8), CNBr-2 (9-92), CNBr-3 (93-159). The pilin structure was found to have several features similar to pilin from Ps. aeruginosa, notably a homologous Nxerminal region and a cysteine loop near the C-terminus. Unlike the pseudomonad pilin, the major antigenic determinants were located in the cysteine loop near the C-terminus, whereas the central part of the molecule was found to contain a conserved receptorbinding region and possibly an immunorecessive epitope common to all gonococcal pilins (Rothbard et al., 1984, 1985). Particularly interesting was a study in which antisera were raised against each of seven synthetic peptides corresponding to constant and variable sequences of the pilin from gonococcal strain MSl1, and then tested for their ability to cross-react with intact pili from both homologous and heterologous strains, as well as their ability to inhibit bacterial adhesion to a human endometrial carcinoma1 cell line (Rothbard et al., 1985). These experiments indicated that antibodies against a semiconserved central region (69-84) were the most efficient in binding to pili from all strains tested. These antibodies also successfully inhibited a heterologous gonococcal strain from binding to the endometrial carcinoma cells, as did antibodies specific for the region 41-50. These observations suggest that the pilin is the adhesin responsible for recognizing the eukaryotic

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receptor and that it is the conserved regions of the protein which mediate the binding function. However, Virji and Heckels (1984) reported that monoclonal antibodies directed against the variable, but not the conserved, regions of the pilus inhibit gonococcal binding to epithelial cells. One interpretation of these data is that both the conserved and variable regions of the pilus are involved in receptor recognition. Another possible interpretation is that there is another protein, distinct from the pilus structural component, which is responsible for interaction with the mammalian cell receptor. As already mentioned (Section V.F. I), antigenic variation can result from the mixing and matching of semivariable and hypervariable gene segments involving silent and expression sites on the chromosome. The total number of silent loci in the chromosome is unknown at present, but Haas and Meyer (1 986) have shown that at least one silent locus, pilS1, contains six tandem pilus-gene copies containing only the semivariable and hypervariable domains but lacking the common N-terminal regions. These copies differed from each other in the same way as the variant sequences determined by Hagblom et al. (1 985). Presumably, such minicassettes from silent regions are constantly being transferred to expression sites to generate new serologically distinct pilus types. VII. Acknowledgements We wish to thank David Bradley for help in constructing Table 1 and Don Marvin for his manuscript on F pilus structure. We also thank Karin IppenIhler, Ned Minkley and Loren Day for useful discussions. REFERENCES

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Tschape, H. and Tietze, E. (1981). Biologisches Zentralblatt 100, 353. Tschape, H. and Tietze, E. (1983). Zeitschrifi fur Allgemeine Mikrobiologie 23, 393. Uhlin, B., Norgren, M., Baga, M. and Normark, S. (1985). Proceedings of the National Academy of Sciences of the United States of America 82, 1800. Vaisanen, V., Elo, J., Tallgren, L.G., Siitonen, A., Makela, P.H., Svanborg-Eden, C., Kallenius, G., Svenson, S.B., Hultberg, H. and Korhonen, T.K. (1981). Lancet ii, 1366. Vanden Bosch, J.F., Verboom-Sohmer, U., Postma, P., de Graaf, J. and MacLaren, D.M. (1980). Infection and Immunity 29, 226. Van Die, I. and Bergmans, H. (1984). Gene 32,83. Van Die, I., van den Handel, C., Hamstra, H.J., Hoekstra, W. and Bergmans, H. (1983). FEMS Microbiology Letters 19, 77. Van Die, I., van Gefen, B., Hoekstra, W. and Bergmans, H. (1984a). Gene 34, 187. Van Die, I., van Megen, Hoekstra, W. and Bergmans, H. (1984b). Molecular General Genetics 194,528. Van Die, I., Spierings, G., van Megen, I., Zuidweg, E., Hoekstra, W. and Bergmans, H. (1985). FEMS Microbiology Letters 28,329. Van Die, I., van Megen, I., Zuidweg, E., Hoekstra, W., de Ree, H., van den Bosch, H. and Bergmans, H. (1986a). Journal of Bacteriology 167,407. Van Die, I., Zuidweg, E., Hoekstra, W. and Bergmans, H. (1986b). MicrobiulPathogenesis 1.51. Virji, M. and Heckels, J.E. (1984). Journal of General Microbiology 130, 1089. Ward, M.E., Watt, P.J. and Robertson, J.N. (1974). Journal of Infectious Diseases 129,650. Watts, T.H., Worobec, E.A. and Paranchych, W. (1982). Journal of Bacteriology 152,687. Watts, T.H., Kay, C.M. and Paranchych, W. (1983a). Biochemistry 22,3640. Watts, T.H., Sastry, P.A., Hodges, R.S. and Paranchych. W. (1983b). Infection andlmmunity 42, 113. Weber, P.C. and Salemme, F.R. (1980). Nature. London 287,82. Webster, R.E. and Lopez, J. (1985). In “Virus Structure and Assembly” (S. Casjens, ed.), pp. 236267. Jones and Bartelett, Boston. Weiss, R.L. (1971). Journal of General Microbiology 67, 135. Wickner, W. and Lodish, H. (1985). Science 230,400. Willetts, N.S. (1977a). In “R Factor: Drug Resistance Plasmid” (S. Mitsuhashi, ed.), pp. 89-107. University of Tokyo Press, Tokyo, Japan. Willetts, N.S. (1977b). Journal of Molecular Biology 112, 141. Willetts, N.S. (1984). Methods in Microbiology 17, 33. Willetts, N.S. and Maule, J. (1974). Genetical Research 24, 81. Willetts, N.S. and Maule, J. (1986). Genetical Research 47, 1. Willetts, N.S. and Paranchych, W. (1974). Journal of Bacteriology 120, 101. Willetts, N.S. and Skurray, R.S. (1980). Annual Review ofGenetics 14,41. Willetts, N.S. and Skurrary, R.S.(1986). Absrracts of the American Society for Microbiology, Washington, D.C., p. 151. Willetts, N.S. and Wilkins, B. (1984). Microbiological Reviews 48, 24. Willetts, N.S., Moore, P.M. and Paranchych, W. (1980). Journal of General Microbiology 117, 455. Williams-Smith, H. and Huggins, M.B. (1983). Journal of General Microbiology 129,2659. Willshaw, G.A., Smith, H.R., McConnell, M.M., Barclay, E.A., Kmjulac, J. and Rowe, B. (1982). Infection and Immunity 37, 858. Willshaw, G.A., Smith, H.R. and Rowe, B. (1983). FEMS Microbiology Letters 16, 101. Willshaw, G.A., Smith, H.R., McConnell, M.M. and Rowe, B. (1985). Plasmid 13, 8. Winans, S.C. and Walker, G.C. (1985a). Journal of Bacteriology 161,402. Winans, S.C. and Walker, G.C. (1985b). Journal of Bacteriology 161,425.

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Woods,D.E.,Strauss,D.C.,Johanson, W.G., Jr., Berry,V.K.andBass, J.A. (1980).Infectionand Immuniiy 29, 1146. Woods, D.E., Iglewski, B.H. and Johanson, W.G., Jr. (1982). In “Microbiology-1982” (D. Schlessinger, ed.), pp. 348-352. American Society of Microbiology, Washington, D.C. Worobec, E.A., Taneja, A.K., Hodges. R.S. and Paranchych, W. (1983). Journa/of Bacteriology 153, 955. Worobec, E.A., Paranchych, W., Parker, J.M.R., Taneja, A.K. and Hodges, R.S. (1985). Journal of Biological Chemistry 260,938. Worobec, E.A., Frost, L.S., Pieroni, P., Armstrong, G.D., Hodges, R.S., Parker, J.M.R., Finlay, B.B. and Paranchych, W. (1986). Journal of Bacteriology 167,660.

Carboxysomes and Ribulose Bisphosphate Carboxylase/Oxygenase GEOFFREY A . CODD Department of Biological Sciences. University of Dundee. Dundee DDI 4 H N . U K

I . Introduction . . . . . . . . . . . . . I1 . Distribution and structureofcarboxysomes . . . . . . . A . Chemolitho-autotrophic prokaryotes . . . . . . . . B. Photo-autotrophic prokaryotes . . . . . . . . . C . Cyanelles . . . . . . . . . . . . . 111. Carboxysome composition . . . . . . . . . . . A . Carboxysomeisolation and studies inuirro . . . . . . . B. Immuno-electronmicroscopy . . . . . . . . . IV. Ribulose 1, 5-bisphosphate carboxylase/oxygenase (RuBisCO) . . . . A . Purification and structure . . . . . . . . . . B. Activation and catalysis . . . . . . . . . . C . Specificity and regulation . . . . . . . . . . D . Genetics . . . . . . . . . . . . . V . Carboxysome function . . . . . . . . . . . A . Are carboxysomes sites of carbon dioxide fixation in uiuo? . . . . B. Do carboxysomes protect ribulose 1,5-bisphosphate carboxylase/oxygenase. C . Are carboxysomes storage bodies? . . . . . . . . VI . Further aspects of carboxysomes . . . . . . . . A . Ecological markers for autotrophy . . . . . . . . B. Man-made ribulose 1, 5-bisphosphate carboxylase/oxygenase inclusion . . . . . . . . . . . . . bodies? References . . . . . . . . . . . . . .

156 157

.

I Introduction Carboxysomes. formerly known as polyhedral bodies. occur in several groups of microbes which grow on carbon dioxide as the principle carbon source. Within these autotrophs. carboxysomes only occur among microbes that utilize the Calvin.. or reductive pentose phosphate. cycle. Carboxysomes are ADVANCES IN MICROBIAL PHYSIOLOGY. VOL. 29 ISBN 0- 12-027729-8

Copyright 0 1988. by Academic Press Limited All rights of reproduction in any form reserved

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apparently confined to prokaryotes, but not all Calvin cycle bacteria contain these inclusion bodies. Although carboxysomes were observed by electron microscopy over 25 years ago (Jensen and Bowen, 1961), insight into their biochemical nature has been more recent and has stemmed from the discovery that the organelles in the colourless sulphur-oxidizing bacterium Thiobacillus neapolitanus contain ribulose 1,5-bisphosphate carboxylase/oxygenase (RuBisCO), the COrassimilating enzyme of the Calvin cycle (Shively el al., 1973a; Shively, 1974). The Calvin cycle prokaryotes, including organisms that derive energy from the oxidation of inorganic compounds (e.g. the chemolitho-autotrophic hydrogen-, sulphur-, nitrite-, ammonia-, carbon monoxide- and iron-oxidizing bacteria) and those that do so from sunlight (e.g. the phototrophic purple bacteria, cyanobacteria and prochlorophytes), continue to attract considerable research interest from the fundamental and applied viewpoints. These organisms are responsible for the primary production of organic carbon in many aerobic and anaerobic environments on a global basis, and perform key roles in the geochemical cycling of matter (for reviews see Schlegel, 1976; Smith and Hoare, 1978; Pfennig, 1978; Stewart, 1980; Bowien and Schlegel, 1981; Kuenen and Buedeker, 1982; Gibson and Smith, 1982; Fogg, 1982; Kuenen et al., 1985; Colby et al., 1985). The RuBisCO enzymes in particular of these organisms are of interest. These enzymes facilitate autotrophic growth, and knowledge of the properties of microbial RuBisCO enzymes contributes to the understanding of their evolution, and offer prospects of modifying the RuBisCO enzymes of higher plants. In the Calvin cycle prokaryotes that contain carboxysomes, a variable and sometimes predominant proportion of the cellular RuBisCO pool is located in these inclusion bodies. For earlier reviews on microbial RuBisCO enzymes see McFadden (1973; 1980), McFadden and Tabita (1974), McFadden and Purohit (1978) and Codd (1984), and on carboxysomes see Shively (1974) and Codd and Marsden (1984). In this review, recent advances in knowledge of the occurrence, composition, properties and possible functions of carboxysomes are examined and discussed with reference to the rapidly expanding field of research on RuBisCO enzymes. It is of interest to compare the RuBisCO enzymes of prokaryotes that contain carboxysomes with those that do not. Insight into carboxysome function also may be gained by comparison with other prokaryotic inclusions and with the compartmentation of RuBisCO in chloroplasts, and these aspects are also considered. Finally, the use of carboxysomes as ecophysiological markers, and the prospects of producing RuBisCO-containing inclusion bodies in recombinant micro-organisms, are discussed.

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11. Distribution and Structure of Carboxysornes

Polyhedral bodies, usually surrounded by a membrane, although this is not always apparent, have been widely observed among the nitrifying bacteria, colourless sulphur-oxidizing bacteria and cyanobacteria. Their designation as carboxysomes depends on the presence of RuBisCO in the inclusion bodies. Until recently, the strict demonstration of RuBisCO in the organelles had been confined to one or two members of the nitrifiers, sulphur-oxidizers and cyanobacteria and was limited by practical problems of carboxysome isolation. Although these problems have to some extent been overcome (see Section 111), the in uitro approach to carboxysome identification and characterization is currently being complemented by the use of immunoelectronmicroscopy on sections of whole cells. This has enabled the polyhedral bodies of additional organisms to be examined and has supported earlier views (Shively, 1974; Stewart and Codd, 1975; Codd and Marsden, 1984) that in a wide range of Calvin cycle prokaryotes these organelles contain RuBisCO. A. CHEMOLITHO-AUTROPHIC PROKARYOTES

I . Nitrite- and Ammonia-Oxidizing Bacteria

The ultrastructure of these organisms has received intensive study and carboxysomes have been observed in seven out of nine strains of nitriteoxidizing bacteria (Table 1). The nitrite-oxidizing bacteria vary in their nutritional capabilities with respect to carbon compounds (Smith and Hoare, 1977) and include versatile members which can grow on organic carbon sources as well as specialist members which can only grow chemolithoautotrophically. All of the versatile strains examined possess carboxysomes and these organelles are also present in Nitrococcus mobilis, which is a specialist. They are lacking, however, from the specialists Nitrospina gracilis and Nitrospira marina (Table 1). Among the ammonia-oxidizing bacteria, the characteristic mode of carbon nutrition is specialist, and carboxysomes have not been observed in intensive studies of any of the six specialist species belonging to six different genera (Table 1). Carboxysomes have been found in only one ammonia-oxidizer to date, a marine Nitrosomonas sp. (Wullenweber et al., 1977), although whether this isolate is nutritionally versatile or a specialist has not been reported. Comparison of the presence of carboxysomes and nutritional versatility is of interest in considering possible functions for the carboxysomes. The carboxysomes of nitrifying bacteria are typically located in the central region of the cell and are not in contact with the intracellular membranes

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TABLE 1. Presence of carboxysomes in nutritionally versatile and specialist species of nitrite-oxidizing and ammonia-oxidizing bacteria

Organism

Nitriteoxidizing bacteria Nitrobacter winogradsky Nitrobacter agilis (three strains) Nitrobacter hamburgensis X 14 Nitrobacter hamburgensis Y Nitrococcus mobilis Nitrospina gracilis Nitrospira marina Ammonia-oxidizing bacteria Nitrocystis oceanus Nitrospira briensis Nitrosolobus multiformis Nitrosomonas europea Nitrosovibrio tenuis Nitrosococcus mobilis Nitrosomonas sp.

Presence of Nutritional carboxysomes capability References" V" V V V Sb

S S S S S S S S n.r.c

1-3 4 4 5 5

6

7 8 9 10 11 12 13

Versatile, i t . facultatively heterotrophic. Specialist, i.e. obligately autotrophic. n.r., not reported. References: 1, Pope et al. (1969); 2, van Gool et al. (1969); 3, Bock et al. (1974); 4, Bock et al. (1983); 5 , Watson and Waterbury (1971); 6, Watson et al. (1986); 7, Murray and Watson (1965); 8, Watson (1971); 9, Watson ct al. (1971); 10, van Gool (1972); 11, Harmser al. (1976); 12, Koops er al. (1976); 13, Wullenweber et al. (1977). (I

"

possessed by several of these organisms (e.g. Bock et al., 1983). The inclusion bodies are 100-120 nm in diameter and consist of a core surrounded by a membrane or shell (van Gool et al., 1969; Bock et al., 1974). The icosahedral shape (Peters, 1974) and appearance of the carboxysomes of Nitrobacter winogradsky and several Nitrobacter agilis isolates accounted for the organelles in nitrifying bacteria being initially described as 'phage-like particles (Bock et al., 1974; Westphal and Bock, 1974).As in sulphur-oxidizing bacteria and cyanobacteria, carboxysome numbers per cell in the nitrifying bacteria vary with cell age and environmental conditions. Between one and five carboxysomes per cell is usual (e.g. Wullenweber et al., 1977; Bock et al., 1983),although up to 200 have been observed (Watson and Waterbury, 1971). As with carboxysome composition, data of carboxysome frequency under defined growth conditions can contribute to the understanding of carboxysome function.

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2. Colourless Sulphur-Oxidizing Bacteria Carboxysomes have been observed in ten out of 17 species of strains of Thiobacillus.For example, among the nutritionally specialist species, they are present in T. neapolitanus, T. thioparus and T. thiooxidans (Shively et al., 1970; Holt et al., 1974), T. kakobis (Reynolds et al., 1981) and T. albertis (Bryant et al., 1983), but not in T. denitrificans (Shively et al., 1970). Among the nutritionally versatile Thiobacillusspecies, carboxysomes are present in T. intermedius (Holt et al., 1974) and T . perometabolis (Katayama-Fujimura et al., 1984a,b), but they have not been observed in Thiobacillus versutus (formerly ThiobacillusA2; Taylor and Hoare, 1969; Shively et al., 1970), or T. novellus (Shively et al., 1970). Carboxysomes are not present in the nutritionally versatile Thiosphaera pantotropha (Robertson and Kuenen, 1983). No correlation has emerged between the presence of carboxysomes and the versatility of carbon nutrition alone of the sulphur-oxidizing bacteria (Codd and Marsden, 1984), although the occurrence of carboxysomes in the genus Thiobacillusshows good agreement when viewed against the classification system of Katayama-Fujimura et al. (1984b) based on nutritional and physiological features, fatty acid composition, their respiratory chain ubiquinones and DNA base composition (Table 2). Carboxysomes may thus be adopted as a taxonomic tool for the heterogeneous genus Thiobacillus (Katayama-Fujimura et al., 1984b), but the significance of the presence or absence of these inclusions in individual strains requires further understanding of carboxysome composition and function. Thiobacillus carboxysomes are typically located in the central region of the cell and show a polygonal (mostly ,hexagonal) profile in cell sections. Carboxysome size in vitro varies from about 70 to 500 nm in diameter, with a mean of about 110 nm (Shively et al., 1973a,b; Holt et al., 1974; Bryant et al., 1983; Katayama-Fujimura et al., 1984b). Elongated carboxysomes have also been seen in T. neapolitanus (Shively et al., 1973b). In all cases, the inclusions are of medium to high electron density under the electron microscope and are surrounded by a shell, 3.5 nm thick in T. neapolitanus (Shively et al., 1973b; Holt et al., 1974). The carboxysome shell is a single-layered structure as apparent from studies of numerous cell-thin sections (see Codd and Marsden, 1984) and the freeze-etching analysis of Thiobacillus IV-85 (Murphy et al., 1974), which revealed smooth inner and outer surfaces of the carboxysome membrane without the studded particles characteristic of bilayer membranes. The three-dimensional structure of T. neapolitanus carboxysomes has recently been studied in detail using intact and broken isolated organelles (Holthuizen, 1986; Holthuizen et al., 1986a). Mean carboxysome size was I 17 & 7 nm with a range from 97 to 132 nm. The hexagonal profiles typically seen by negative staining, freeze hydration and freeze-etching techniques are

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TABLE 2. Presence of carboxysomes in Thiobacillus species and strains classified according to the system of Katayama-Fujmura et al. (1984b)

Organism

Source

Group 1-1 Thiobacillus versutus (A2) Thi~bucillusversutus Thiobacillus novellus Thiobacillus sp. Thiobacillus sp.

ATCC" 25364 ATCC 27793 ATCC 8093 IAMh 12816 IAM 12817

Group 1-2 Thiobacillus ucidophilus Thiobacillus acidophilus

ATCC 27807 ATCC 27977

Nutritional capability/ Respiratory Presence of optimal growth chain carboxyconditions ubiquinone somes

Versatile; prefer Or

slightly alkaline growthmedia

Q-10 Q-10 Q-I0 Q-10 Q-10

-

-

+

Versatile; prefer acidic media

Q- 10 Q-I0

Versatile

4-8 Q-8 Q-8

Specialist

4-8 Q-8

Group 111-2 Thiobacillus neapolitanus ATCC 23641

Specialist

Q-8

+

Group 111-3 Thiobacillusferrooxidans ATCC 19859

Specialist

Q-8

+

Group I1 Thiobacillus delicatus IAM 12624 Thiobacillusperometabolis ATCC 23370 Thiobacih intermedius ATCC I5466 Group 111-2 Thiobacillus denitrijicans ATCC 23642 Thiobacillus thioparus ATCC 8 158

+

+ + +

-

+

American Type Culture Collection, Rockville, Md, USA.

* Institute of Applied Microbiology Culture Collection, University of Tokyo, Japan consistent with a pentagonal dodecahedron, which was confirmed by stereomicrography. Thiobacillus carboxysomes contain arrays of 10 nm diameter particles which are often arranged in rows or circles (Shively et al., 1973a,b; Holthuizen, 1986; Holthuizen et al., 1986a,b). These doughnut-shaped particles are released when carboxysomes are ruptured, and are RuBisCO molecules.

3. Hydrogen-Oxidizing Bacteria Ultrastructural surveys of a wide range of mesophilic hydrogen-oxidizing Calvin cycle bacteria have not shown the presence of carboxysomes in

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members of the genera Alcaligenes, Aquaspirillum, Pseudomonas, Corynebacterium, Xanthobacter, Azospirillum, Flavobacterium, Arthrobacter, Bacillus, Nocardia or Paracococcus (Walther-Mauruschat et al., 1977; Aragno and Schlegel, 1981; Shively et al., 1978). Thermophilic hydrogen bacteria have received little attention to date and the sole representative examined ultrastructurally, Pseudomonas thermophila K2, contains hexagonal membrane-bound bodies, 100-150 nm in diameter (Kostrikina et al., 1981). The bodies contain particles of about the same size as RuBisCO molecules and are associated with up to 80% of the extractable RuBisCO activity in vitro (Romanova et al., 1982). This strongly suggests the presence of carboxysomes in Ps. thermophila, and further studies on these organelles in thermophilic hydrogen-oxidizing bacteria will be of interest. B. PHOTO-AUTOTROPHIC PROKARYOTES

I . Cyanobacteria Although individual sections of cyanobacterial cells may fail to reveal these organelles, since they may number only one or two per cell, carboxysomes are typically present in cyanobacteria, whether growing photo-autotrophically, photoheterotrophically or chemoheterotrophically. Numerous examples can be found in the literature (e.g. Jensen and Bowen, 1961; Peat and Whitton, 1967; Gantt and Conti, 1969; Lang and Whitton, 1973; Wolk, 1973; Stewart and Codd, 1975; Stanier and Cohen-Bazire, 1977). The carboxysomes of unicellular cyanobacteria typically resemble those of the chemolitho-autotrophic prokaryotes by occurring in the centre of the cell, not in contact with the thylakoids or other inclusions; by displaying a relatively uniform size (100130 nm diameter) with a common hexagonal profile; and by being surrounded by a 3 4 nm thick membrane (Gantt and Conti, 1969; Nierzwicki-Bauer et al., 1983; Jensen and Rachlin, 1984). By contrast, a wide variation in carboxysome size, shape, number and location occurs among the filamentous cyanobacteria. Carboxysome size among this diverse assemblage varies from about 100 to 900 nm diameter and hexagonal profiles are not more frequently seen than four- or five-sided sections of the carboxysomes (e.g. Peat and Whitton, 1967;Stewart and Codd, 1975; Lang, 1977; van Eykelenburg, 1979; Jensen and Ayala, 1976a). Carboxysomes in filamentous cyanobacteria may be in the centroplasm region (Stewart and Codd, 1975) or peripherally located between the thylakoids (Lang and Whitton, 1973). The membrane surrounding the carboxysomes of filamentous strains appears as a monolayer (e.g. Jensen and Ayala, 1976a;Jensen, 1979).Close association of the carboxysomes in several Anabaena strains with microtubules has been observed (Jensen and Ayala,

I22

G.A. CODD

1976a,b). The significance of microtubules in these prokaryotes remains unclear, although this alignment of arrays of microtubules, with their longitudinal axis in close contact with a facet of the carboxysome, may be related to carboxysome assembly or mobilization of carboxysome contents in these species. Carboxysomes occur in free-living and symbiotic cyanobacteria, including species such as the Nostoc cyanobionts of Cycas revoluta root nodules, the liverwort Blasia pusilla and the lichen Peltigera canina (Stewart and Codd, 1975). The aerobic nitrogen-fixing filamentous cyanobacteria have evolved a high degree of intercellular structural differentiation. The incompatible processes of photosynthetic oxygen evolution and aerobic nitrogen fixation in the heterocystous strains are permitted to occur simultaneously by the metabolic segregation of oxygenic photosynthesis to the vegetative cells and of nitrogen fixation to the adjoining heterocysts (Fogg et al., 1973; Wolk, 1973, 1982; Stewart, 1980). The RuBisCO required to fix CO2 photosynthetically is present in the vegetative cells but lacking from mature heterocysts (Codd and Stewart, 1977a; Codd et al., 1980). Electron microscopy of large numbers of cells from six heterocystous N2-fixing cyanobacteria showed that the carboxysomes are also segregated and are not present in heterocysts (Stewart and Codd, 1975). 2. Prochlorophyta

A new division of prokaryotes has been proposed, the Prochlorophyta, to accommodate oxygenic phototrophs which contain chlorophyll a , as do cyanobacteria, but which also contain chlorophyll h, which is characteristic of green algae and higher plants but not of cyanobacteria (Lewin, 1976, 1977, 1981). Prochlorophytes lack the phycobilin pigments characteristic of cyanobacteria, but contain RuBisCO (Berhow and McFadden, 1983; Codd, 1984), consistent with their ability to assimilate CO2 via the Calvin cycle (Akazawa et al., 1978). Until recently, the only prochlorophytes known were unicellular forms living in symbiosis with marine didemnid ascidians and were assigned to the genus Prochloron. Prochloron symbionts from Diplosoma uirens and Didemnum molle contain polyhedral bodies similar to the carboxysomes in filamentous cyanobacteria (Whatley, 1977; Fisher and Trench, 1980; Cox and Dwarte, 1981). The Prochloron polyhedral bodies typically occur between the thylakoids in the peripheral cytoplasm and may be closely adpressed to the photosynthetic membranes. Biochemical investigations on the Prochloron polyhedral bodies have been constrained by the inability to grow Prochloron species in laboratory culture. However, we have demonstrated by immunoelectronmicroscopy using heterologous RuBisCO antiserum that the D . uirens Prochloron polyhedral bodies contain RuBisCO (A.M. Hawthornthwaite and G.A. Codd, unpublished work).

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A free-living planktonic prochlorophyte has recently been found growing abundantly in Dutch freshwater lakes (Burger-Wiersma and Mur, 1985; Burger-Wiersma et al., 1986). This organism contains polyhedral bodies (Burger-Wiersma et al., 1986) and is well established in laboratory culture. Characterization of the RuBisCO and polyhedral bodies from this organism is in progress in this laboratory. C. CYANELLES

Cyanelles are blue-green prokaryotes which resemble free-living unicellular cyanobacteria morphologically, but are obligate endosymbionts in eukaryotic protists (Kies, 1980, 1984; Kies and Kremer, 1986). Differences exist between cyanelles and cyanobacteria: Cyanophora paradoxa cyanelles are surrounded by a thin lysozyme-sensitive layer, presumably prokaryotic peptidoglycan, but a complete cyanobacterial type of cell wall surrounding the cyanelle cannot be seen (Hall and Claus, 1963; Schenk, 1970). However, the genome size and copy number of the C . paradoxa cyanelle are typical of chloroplasts, rather than unicellular cyanobacteria, and this cyanelle can therefore be viewed as a photosynthetic organelle which has been derived from an endosymbiotic cyanobacterium (Herdman and Stanier, 1977). The rapid labelling kinetics of I4CO2 assimilation by cyanelles are consistent with the operation of the Calvin cycle (Kremer et al., 1979) and the RuBisCO enzymes of the cyanelles from C . paradoxa and Glaucosphaera vacuolata have been characterized (Codd and Stewart, 1977b; Codd, 1984). Polyhedral bodies have been observed in several cyanelles. Size and number of the organelles varies widely from the single 600-900 nm central polygonal body in Gloeochaete wittrockiana (Kies, 1976, 1980) to the seventy or more 130-160 nm diameter inclusions of Paulinella chromatophora (Kies, 1980, 1984). A membrane cannot be seen around the single central electron-dense body of the C . paradoxa cyanelle or the terminal, pyrenoid-like body of Glaucocystis nostochinearum, although a monolayer surrounds the bodies in Gloeochaete wittrockiana and Paulinella chromatophora (Kies, 1976, 1980, 1984). Polyhedral bodies have not been observed in the Calvin cycle cyanelles of Glaucosphaera vacuolata (Kies, 1980). The central polyhedral body of the cyanelles of C . paradoxa and the terminal body of the Glaucocystis nostochinearum cyanelles have recently been confirmed by immuno-electronmicroscopy to be carboxysomes (Mangeney and Gibbs, 1987; Mangeney et al., 1987). The transition status of cyanelles between cyanobacteria and chloroplasts confers particular interest in their carboxysomes in terms of the carboxysomes of free-living cyanobacteria and of the RuBisCO-containing pyrenoids in algal chloroplasts.

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111. Carboxysome Composition

A knowledge of carboxysome composition is necessary to provide understanding of the functions of the organelles and their contribution to the physiology of the cell. A. CARBOXYSOME ISOLATION AND STUDIES

in vitro

I . Isolation and Stability of Carboxysomes in vitro The rationale for carboxysome isolation and discussion of the various approaches used up to 1983 were detailed by Codd and Marsden (1984). Carboxysomes have been isolated from few organisms to date; from Thiobacillus neapolitanus (Shively et al., 1973a; Beudeker et al., 1980, 1981; Beudeker and Kuenen, 1981; Cannon, 1982; Cannon and Shively, 1983), Nitrobacter winogradsky (Peters, 1974; Westphal, 1977;Westphal et al., 1979); Nitrobacter agilis (KI) (Westphal and Bock, 1974; Shively et al., 1977; Westphal, 1977; Westphal et al., 1979; Biedermann and Westphal, 1979), Nitrobacter hamburgensis H XI^) (Ebert, 1982),Nirrosomonas sp. (Harms et al., 1981), Anabaena cylindrica (Codd and Stewart, 1976) and Chlorogloeopsis fritschii (Lanaras and Codd, 1981a). The stability of carboxysomes in vitro varies according to source: for example, those released from ultrasonically-disrupted cells of N . agilis and T. neapolitanus, and purified by differential or isopycnic sucrose density gradient centrifugation, or by preparative agarose electrophoresis, retain their hexagonal profile in vitro (Shively et al., 1973a, 1977; Cannon and Shively, 1983). The isolated T. neapolitanus carboxysomes can be stored for at least two months at 4°C without changing their appearance if maintained in the presence of the protease inhibitor phenylmethylsulphonyl fluoride (Holthuizen et al., 1986b). In contrast, the carboxysomes from the filamentous cyanobacteria are less stable in vitro, although carboxysome yields from Ch.fritschii were optimized by cell disruption in a French Pressure Cell and the destabilizing effect of sucrose during subsequent carboxysome purification was lowered by the use of the silicon polymer, Percoll (Lanaras and Codd, 1981a). 2. Carboxysomal Proteins In all cases, isolated carboxysomes consist mainly of protein. The presence of RuBisCO, which is the most abundant component in carboxysomes, was initially indicated by the ability of isolated carboxysomes to catalyse RuBPdependent COz assimilation (Shively et al., 1973a, 1977; Codd and Stewart,

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1976; Harms et af., 1981; Lanaras and Codd, 1981 a). This was confirmed by the finding of immunological cross-reactivity between isolated broken carboxysomes and antibodies to the respective RuBisCO enzymes (Codd and Stewart, 1976; Beudeker et al., 1981; Holthuizen et af.,1986b). These findings are consistent with the presence within isolated carboxysomes of 1&12 nm particles which have the same dimensions and shape as RuBisCO molecules (Shively et al., 1973a, 1977; Lanaras and Codd, 1981a; Holthuizen, 1986; Holthuizen et af., 1986a,b). Differencesin the estimates of the total number of polypeptides present in isolated sodium dodecyl sulphate (SDS)-dissociated carboxysomes have been reported after SDS-polyacrylamide-gel electrophoresis (SDS-PAGE), followed by gel staining with Coomassie Blue (Table 3). It is not clear whether these totally represent real differences in carboxysome composition between species or whether they merely reflect differences in methodology between laboratories. It is possible that additional carboxysomal polypeptide components may be detected by the use of more sensitive autoradiography of SDS-PAGE gels of SDS-dissociated carboxysomes of 35S-labelled cells. Whether carboxysome composition for an individual species varies with physiological conditions during growth does not appear to have been investigated. Despite the apparent interspecific differences, the isolated carboxysomes from the different physiological groups of autotrophs are similar in the possession of two polypeptides which coincide in terms of molecular mass with the large (L) subunits and small (S) subunits of their respective RuBisCO enzymes (Biedermann and Westphal, 1979; Lanaras and Codd, 1981a,b; Cannon and Shively, 1983;Snead and Shively, 1978; Holthuizen et af.,1986b). The L and S subunit peptides of the dissociated isolated carboxysomes are present in equimolar amounts, as in the purified RuBisCO enzymes which have an 8L8S quaternary structure. The T. neapolitanus carboxysomes have recently been examined for glycoproteins and four bands have been found with M , values of 120,000, 85,000, 54,000 and 10,000 (see Table 3). Those of 120,000 and 85,000 Da are present in trace amounts. The 54,000 glycoprotein migrates during SDSPAGE close to the RuBisCO L subunit (Holthuizen et al., 1986b) and may account for earlier indications of L subunit heterogeneity in T. neapolitanus carboxysomes (Cannon and Shively, 1983). The 10,000 Da species was the most abundant glycoprotein and this, plus the other three glycoproteins, are components of the T. neapolitanus carboxysome shell (Holthuizen et al., 1986b), a finding in support of earlier proposals for the shell of the inclusions from T. neapolitanus (Cannon and Shively, 1983) and N. agilis (Biedermann and Westphal, 1979). The lack of effect of ether and chloroform on the shells of N. agilis carboxysomes and analysis of chloroform-methanol extracts of

126

G.A. CODD

TABLE 3. Comparative polypeptide composition” of isolated carboxysomes Source o f carboxysomes Nitrobacter agilish

Chlorogloeopsis Thiobacillus fritschii“ neapolitanud Polypeptide identity

120

GP

97 89 85

GP

110

74 72 69

69 67 66

56

56 54 52

LSU GP LSU

49 47 43 39 36 32 15

14 13 12 Total

7

13

8

13

ssug

10

GP

12-15

Values refer to M,in kilodaltons. Data from Biedermann and Westphal (1979). Data from Lanaras and Codd (198la). Data from Cannon and Shively (1983) and Holthuizen et al. (1986b). ‘GP, glycoprotein. LSU, large subunit of RuBisCO. 8 SSU, small subunit of RuBisCO.

’

the T. neapolitanus organelles has indicated that lipids are not present (Biedermann and Westphal, 1979; Holthuizen et al., 1986b). Apart from RuBisCO, the identity of the remaining carboxysomal polypeptides is unknown. Activities of other enzymes of the Calvin cycle have been sought in isolated, intact and broken carboxysomes. Activities of

CARBOXYSOMES AND RIBULOSE BISPHOSPHATE CARBOXYLASE/OXYGENASE

127

phosphoribulokinase, ribose-5-phosphate isomerase, fructose 1,6-bisphosphatase, NAD glyceraldehyde 3-phosphate dehydrogenase, or sedoheptulose 1,7-bisphosphatase were not detected in highly purified T. neapolitanus carboxysomespreparations, in contrast to cytoplasmic supernatants (Cannon and Shively, 1983; Holthuizen et al., 1986b). The possibility that phosphoribulokinase, which is metabolically contiguous with RuBisCO, may be a component of cyanobacterial carboxysomes was examined since the organelles of Ch.fritschii contain a cryptic 39,000 Da polypeptide (Lanaras and Codd, 1981a), and purification of this enzyme from Ch.fritschii showed it to consist of 40,000 Da subunits (Marsden and Codd, 1984). Only about 5% of the extractable Ch.fritschii phosphoribulokinase activity is associated with a cell-free particulate fraction (Lanaras and Codd, 198la). Sucrose density gradient centrifugation of carboxysome-containing extracts of this cyanobacterium did not result in the localization of the particulate phosphoribulokinase in the purified carboxysomes but in a thylakoid fraction (Marsden et ai., 1984). These findings have been confirmed by immuno-electronmicroscopy (see Section III.B, p. 130). Carbonic anhydrase has also been considered as a possible carboxysomal protein, since this enzyme is present at high activity in cultures of several cyanobacteria, grown in low concentrations of COZ,where it may function in inorganic carbon concentration to support the carboxylase reaction, versus the oxygenase reaction, of RuBisCO (Aizawa and Miyachi, 1986; Codd and Kuenen, 1987). The possibility also exists that carboxysomes may act as a COz-concentrating mechanism (Codd and Marsden, 1984). Carbonic anhydrase was not detected in either carboxysomal or cytoplasmic fractions of T. neapolitanus (Cannon and Shively, 1983), though the enzyme is present in high amounts in Nitrosomonas europea and Rhodospirillum rubrum which do not contain carboxysomes (Jahnke et al., 1984; Gill et al., 1984). High carbonic anhydrase activities were detected in extracts of air-grown Ch. fritschii and 90% of the activity is particulate (Lanaras et al., 1985). Although the enzyme has not been purified from Ch.fritschii, the presence of a cryptic 43,000 Da peptide in the carboxysomes purified from this source (see Table 3), along with the findings of Yagawa et al. (1984) that carbonic anhydrase from the carboxysome-containing cyanobacterium Anabaena variabilis has a M,value of 42,000 & 5,000, suggested that the Ch.fritschii carbonic anhydrase may be a carboxysomal enzyme. However, centrifugation of crude particulate Ch. fritschii extracts through sucrose density gradients yielded a sharp band of carbonic anhydrase which was well separated from the carboxysomes and thylakoids (Lanaras et al., 1985). The carbonic anhydrase of Ch. fritschii is apparently on the cell surface since high enzyme activity is exhibited by the whole cells and this is not influenced by cell disruption by ultrasonication or lyzozyme treatment (Table +

128

G.A. CODD

TABLE 4. Carbonic anhydrase activities of whole cells, broken cells and fractions of Chloroglaeopsis,fritschii

Source of enzyme"

Treatments

Specific activityh (units (mgprotein)-I)

Whole cells Whole cells Whole cells

-

Ultrasonicated Lysozyme lysate

0.3 1 0.29 0.24

Whole cells Whole cells Whole cells

+Acetozolamide (0.2 mM) +Ethoxyzolamide (20 p ~ ) Sulphanylamide (0.2 mM)

+

0.00 0.00 0.00

40,OOOg x 1 h 40,OOOgx 1 h

Pellet of broken cells Supernatant of broken cells

0.33 0.02

Purified carboxysomes Purified carboxysomes

-

0.00 0.00

Ultrasonicated

Cultures grown photo-autotrophically to mid-exponential phase on air throughout. (For comparison, specific activity ofwhole cells grown in 5% COz in air was about 0.07.) Activity was measured electrometrically: one unit of activity = (ApH/ ApH, - 1) x 10, where ApH and ApH, are the rates of pH change of enzyme-catalysed and -uncatalysed reactions (Lanaras ei al., 1985; G.A. Codd and K. Okabe, unpublished work).

4). The established carbonic anhydrase inhibitors acetozolamide, ethoxyzolamide and sulphanylamide completely inhibit whole cell enzyme activity (as with broken-cell extracts) with acetozolamide being used at concentrations that are unlikely to enter the cell. Several examples of external carbonic anhydrases exist along the green algae (Aizawa and Miyachi, 1986), but Ch. fritschii appears to be the first example of a cyanobacterium possessing an external carbonic anhydrase (Table 4). The function of the 43,000 Da Ch. fritschii carboxysome peptide remains cryptic (see Table 3).

3. Carboxysomes and Deoxyribonucleic Acid The possibility that carboxysomes may contain DNA was raised by Bock and his colleagues, since the 'phage-like inclusions of Nitrobacter spp. are of high ) and have a high 260/280 nm absorption ratio of 1.2 (Bock density ( P * ~ 1.296) et al., 1974; Westphal and Bock, 1974). This concept has been supported by the finding of a deoxyribonuclease-sensitivedouble-stranded filament, 14 pm long, with a buoyant density ( p 2 7 of 1.701 in ethidium bromide+aesium chloride gradients and an absorption maximum of 263.5 nm, associated with

CARBOXYSOMES AND RIBULOSE BISPHOSPHATE CARBOXYLASE/OXYGENASE

129

osmotically-broken isolated N . agilis and N. winogradsky carboxysomes (Westphal, 1977; Westphal et al., 1979). Extrachromosomal DNA occurs commonly in autotrophic prokaryotes. Plasmids have been detected in purple photosynthetic bacteria, cyanobacteria, colourless sulphur-, nitrite-, ammonia-, iron- and hydrogen-oxidizing bacteria and carboxydotrophic bacteria (Herdman, 1982; Saunders, 1984; Schlegel, 1984; Friedrich, 1987). The only group in which progress has been achieved in identifying plasmid function is the hydrogenotrophic bacteria, where genes for HZoxidation, CO2 fixation, denitrification and heavy metal resistance have been located on plasmids (Schlegel, 1984; Friedrich, 1987). These organisms, with the probable exception of Pseudomonas thermophila, have not been found to contain carboxysomes (see Section 11. A). Cryptic plasmids have been reported in the following carboxysome-containing Thiobacillus spp.: T . ferrooxidans, T . neapolitanus, T . acidophilus and T . intermedius (Martin et al., 1981; Holmes et al., 1984; Toth et al., 1981; Davidson and Summers, 1983) and in Nitrobacter hamburgensis X14 (Kraft and Bock, 1984). Plasmids are also present in related strains that do not contain carboxysomes, for example Thiobacillus versutus (Gerstenberg et al., 1982) and Thiosphaera pantotropha (Chandra and Friedrich, 1986). No correlation has emerged between the possession of plasmids and of carboxysomes among the chemolitho-autotrophs. Carboxysomes are a consistent feature of cyanobacteria (Codd and Marsden, 1984) but extrachromosomal DNA has only been detected in about half of the strains examined (Herdman, 1982). We have examined three carboxysome-containing cyanobacteria for extrachromosomal DNA and specifically for the presence of DNA in the carboxysomes (Vakeria et al., 1984). Anabaena PCC7120 contains a 5 kb plasmid although this was not associated with the carboxysomes in this strain. No extrachromosomal DNA has been detected in Gloeobacter violaceus, or in Ch.fritschii using six different plasmid isolation methods. These findings preclude the possibility that Ch. fritschii and Gloeobacter violaceus carboxysomes contain extrachromosomal DNA. Gel electrophoresis of isolated intact and disrupted carboxysomes from these three cyanobacteria did not indicate the presence of plasmid DNA (Vakeria et al., 1984), although a trace of DNA was associated with carboxysomal pellets. Restriction enzyme digestion patterns of this DNA were characteristic of chromosomal DNA. We conclude that some association between the outer surface of the cyanobacterial carboxysomes and chromosomal DNA exists in broken cell preparations. Electron microscopic evidence exists for the attachment of DNA to the exterior of the putative carboxysomes of Ps. thermophila (Romanova et al., 1982)and the cyanelles of C. paradoxa (Bohnert et al., 1983). Holthuizen et al. (1986~)have recovered DNA associated with the exterior of isolated T.

130

G.A.

CODD

neapolitanus carboxysomes. This shows the same restriction enzyme digestion pattern as chromosomal DNA and hybridizes with the latter. A highly polar external surface of the carboxysome shell would facilitate the attachment of chromosomal DNA in vitro. Whether such association occurs in vivo is of interest. This has been observed in T. neapolitanus and apparently in Ps. thermophila (Holthuizen et al., 1986; Romanova et al., 1982) and raises the question of whether DNA has a functional role in carboxysome assembly. B. IMMUNO-ELECTRONMICROSCOPY

Immuno-electronmicroscopy has found valuable application in the localization of enzymes and other cell components. This approach has been used to investigate the subcellular distribution of RuBisCO using colloidal gold. Heavy labelling of the carboxysomes in cell sections of the cyanobacteria

FIG. 1. Immuno-electronmicroscopic localization of RuBisCO in Chlorogloeopsis fritschii. Cell sections were labelled with RuBisCO antibodies raised in rabbits,

followed by goat-antirabbit immunoglobulin conjugated to 20 nm colloidal particles, which may be seen covering the carboxysomesections (C). Bar marker represents 1 pm.

CARBOXYSOMES AND RIBULOSE BISPHOSPHATE CARBOXYLASE/OXYGENASE

13 1

Chlorogloeopsis fritschii, Anacystis nidulans R2 and Anabaena cy lindrica was obtained using RuBisCO antiserum confirming the presence of RuBisCO in the carboxysomes (Fig. 1; Hawthornthwaite et al., 1985; Cossar et al., 1985). RuBisCO antiserum labelling of the cytoplasm also occurred, confirming that some of the RuBisCO pool in uiuo in the cyanobacteria is also soluble (Lanaras and Codd, 1981a,b,1982). In a group of 76 Ch. fritschii cells examined for RuBisCO gold colloid particle distribution, the RuBisCO pool appeared to be approximately equally distributed between the carboxysomes and cytoplasm (Fig. 2a). Inclusion bodies other than carboxysomes did not contain RuBisCO according to the immunogold technique, although loose association of the cytoplasmic enzyme with the thylakoids was observed in A. cylindrica vegetative cells (Cossar et al., 1985). In contrast to the A. cylindrica vegetative cells, RuBisCO antiserum labelling of the heterocysts did not occur, confirming the absence of RuBisCO and carboxysomes from these cells (Stewart and Codd, 1975; Codd and Stewart, 1977a; Cossar et al., 1985). Phosphoribulokinase antiserum-labelling of Ch. fritschii cell sections occurred in the cytoplasm, with some association with the inner surface of the cytoplasmic membrane, but no significant labelling of the carboxysomes was obtained (Fig. 2b). Gold immuno-electronmicroscopy has also recently confirmed the presence

T

50c

CA

CY

M

0

N

CA

CY

M

0

N

-"

FIG. 2. Distribution of colloidal gold particles in the localization of RuBisCO (a) and phosphoribulokinase(b) in Chlorogfoeopsisfritschii by immuno-electronmicroscopy. (a) RuBisCO antiserum: 76 cells were examined and 9345 gold particles counted; (b) phosphoribulokinase antiserum: 45 cells were examined and 28,853 gold particles counted. Gold particles associated with cell sections treated with null serum varied between 0 and 0.6% of the numbers obtained with antisera. CA, carboxysomes; CY, cytoplasm; M, cytoplasmic membrane; 0, all other discernible inclusion bodies; N, labelling on outer layers external to cytoplasmic membrane. Marker bars represent SD. From Hawthornthwaite et af. (1985).

132

G.A. CODD

of RuBisCO in the central dense body of the cyanelles of C. paradoxa and the terminal electron-dense body of Glaucocystis sp. cyanelles (Mangeney and Gibbs, 1987). As with the cyanobacterial carboxysomes, the cyanelle inclusions were not labelled with phosphoribulokinase antiserum, in contrast to the surrounding cytoplasm (Mangeney et al., 1987). Similar data using RuBisCO and phosphoribulokinase antisera have been obtained in this laboratory using sections of the symbiotic unicellular Prochloron sp. and the filamentous prochlorophyte of Burger-Wiersma et al. (1986) by A. M. Hawthornthwaite (unpublished work). Further examination of carboxysome proteins using antibodies raised against individual carboxysome components will be of interest in the comparison of carboxysomes between the different physiological groups of autotrophs and in the study of the assembly and breakdown of the organelles.

IV. Ribulose 1,ibisphosphate carboxylase/oxygenase (RuBisCO) As the COZ-assimilating enzyme of numerous major groups of autotrophic

prokaryotes (Codd, 1984), in addition to algae and higher plants, RuBisCO attracts extensive and intensive study. Research on RuBisCO has accelerated over the past decade and merits entire symposia devoted to the enzyme (e.g. Siegelman and Hind, 1978; Ellis and Gray, 1986). Earlier reviews of the structural, mechanistic, functional and regulatory aspects of RuBisCO are available (Akazawa, 1979; McFadden, 1980; Lorimer, 1981; Lorimer and Andrews, 1981; Miziorko and Lorimer, 1983; Codd, 1984; Akazawa et al., 1984). In this article, aspects of the RuBisCO enzymes of prokaryotes are reviewed, with reference where appropriate to the enzymes of plants and algae, with a view to understanding the physiology of autotrophy and carboxysome function in prokaryotes. A. PURIFICATION AND STRUCTURE

Understanding of the structural and functional properties of RuBisCO is aided by the availability of purified enzyme and RuBisCO has now been purified from over 40 prokaryotes (Akazawa, 1979; McFadden, 1980; Codd, 1984). This has been facilitated by a combination of favourable factors: RuBisCO is among the most abundant proteins in the autotrophic cell, accounting for up to 17% of total protein in Th. neapolitanus (Beudeker et al., 1981) and about 50% in Rhodospirillum rubrum (Tabita et al., 1983);many but not all RuBisCO enzymes are of high M , value ( 2500,000); enzyme activity assays are straightforward; and most enzymes are relatively stable in uitro. The main purification procedure for microbial RuBisCO involves sedimentation

CARBOXYSOMES AND RIBULOSE BISPHOSPHATE CARBOXYLASE/OXYGENASE

133

of a high speed cell-free supernatant extract through sucrose density gradients as the principal step (e.g. Tabita and McFadden, 1974a,b; Bowien et al., 1976; Codd and Stewart, 1977~; Torres-Ruiz and McFadden, 1985). A rapid yield of purified spinach RuBisCO has recently been achieved by ion-exchange fast performance liquid chromatography (FPLC) (Salvucci et a/., 1986a). This procedure may be useful for microbial RuBisCO purification (Gutteridge et a[., 1986). All known RuBisCO enzymes from eukaryotes are proteins with high M , values (500,000-550,000), consisting of eight large subunits (L; ca. 50,00055,000) plus eight small subunits ( S ; ca. 11,000-15,000). The RuBisCO enzymes from most of the 40 or more autotrophs, examined from all of the major physiological groups of Calvin cycle prokaryotes, also have high M , values (500,000-600,000) with 8L8S quaternary structures (Table 5). Significant exceptions, however, occur among the photosynthetic purple nonsulphur bacteria. The 114,000-Da RuBisCO of R . rubrum is a 2L molecule (Tabita and McFadden, 1974b; Schloss et al., 1979).This enzyme has proved to be of outstanding value in the study of RuBisCO catalysis and function since it lacks S subunits. The RuBisCO of Rhodomicrobium uannielii may be a 6L6S enzyme (Taylor and Dow, 1980). The genus Rhodopseudomonas (Rhodobacter) is particularly interesting since out of three species examined, all have been shown to contain two RuBisCO enzymes i.e. a 6L enzyme in TABLE 5. Summary of occurrence of 8L8S ribulose 1,5-bisphosphate carboxylase/oxygenase(RuBisCO) in autotrophic prokaryotes Enzyme source Group Purple sulphur bacteria Purple non-sulphur bacteria Sulphur-oxidizing bacteria H ydrogen-oxidizing bacteria Nitrifying bacteria" Cyanobacteria Prochlorophytesd Cyanelles

No. genera No. species 3 3 1

5 1 10

1 2

Presence of 8L8S RuBisCOO

3

3

5 5 7

3h

4 I 2

2 I5 1

15 1

2

2

''

a Data condensed from Codd (1984). except for (Ebert, 1982) and (Berhow and McFadden, 1983). Absent from Rhodospirillum rubrum (2L enzyme: Tabita and McFadden, 1974a) and perhaps from Rhodomicrobium vannielii (6L6S enzyme: Taylor and Dow, 1980). RuBisCO heterogeneity exists in the remaining three species which contain 8L8S enzymes.

I34

G.A. CODD

TABLE 6. Occurrence of multiple forms of ribulose bisphosphate carboxylase/ oxygenase (RuBisCO) in autotrophic prokaryotes

Source

Designation of RuBisCO

Rhodopseudomonas sphaeroides

Form I Form I1

550,000 360,000

8L8S 6L

Gibson and Tabita (1 977a)

Rhodopseudomonas capsulata

Form I Form I1

550,000 360,000

8L8S 6L

Gibson and Tabita (1977b)

Rhodopseudomonas blastica

Form I Form I1

555,000 350,000

8L8S 6L

Sani (1985), Dow (1987)

Nitrobacter Form I hamburgensis X14 Form I1

520,000 480,000

8(L,L’)8S 8(L,L’)8S

Ebert (1982)

Molecular Quaternary mass structure

Reference

addition to the 8L8S form (Table 6). The 8L8S (Form I) and 6L (Form 11) enzymes of Rps. sphaeroides are regulated independently in response to metabolites and growth conditions. The L subunits of Forms I and I1 show considerable differences in peptide maps of tryptic digests and they are different immunologically (Gibson and Tabita, 1977a, 1985; Weaver and Tabita, 1983; Jouanneau and Tabita, 1986). Structural, immunological, kinetic and regulatory differences also occur between Forms I and 11 from Rps. capsulata (Gibson and Tabita, 1977b; Shively et al., 1984) and these findings have recently been supported by Dow ( I 987) using Rps. blastica. These differences strongly suggest that L subunits of RuBisCO enzymes Forms I and I1 are different gene products and Tabita and colleagues have shown this to be the case in Rps. sphaeroides (Gibson and Tabita, 1986;Tabita et al., 1987; Section 1V.D). L subunit heterogeneity of other microbial RuBisCO enzymes has occasionally been observed, e.g. from Rhizobiumjaponicum and N . agilis, although the possibility remains that this may be due to instability in vitro (Purohit et a/., 1982; Harrison et al., 1979). The presence of two RuBisCO forms in Nitrobacter hamburgensis soluble extracts, with different M , values (Table 6), and different L and S subunits and different responses to culture age and conditions suggests the presence of 2 RuBisCO’s in this organism. As reviewed in Section 11, 10-12 nm particles can be seen in negativelystained carboxysomes in whole cells and in vitro (Shively, 1974; Codd and Marsden, 1984). These are RuBisCO molecules of the 8L8S type, which have been most closely examined from Alcaligenes eutrophus (Bowien et al., 1976; Bowien and Mayer, 1978) which does not contain carboxysomes. The Alcaligenes model, constructed on the basis of M , values of the holoenzyme

CARBOXYSOMES AND RIBULOSE BISPHOSPHATE

CARBOXYLASE/OXYGENASE

135

and the 8L and 8s subunits plus electron microscopy and immunology, is a 13 x 13 x 10.5 nm structure with a fourfold symmetry. There are four layers perpendicular to a central hole; the top and bottom layers each contain four spherical S subunits and the central two layers each contain four L U-shaped subunits with the arms pointing outwards. The two central layers of L subunits are eclipsed. This model is supported by electron microscopic observations on the 8L8S holoenzyme from the cyanobacterium Synechococcus sp. (Andrews et af., 1981). The deduced arrangement of subunits in these prokaryotic RuBisCO enzymes differs from that of the tobacco RuBisCO barrel-shaped model which has a fourfold molecular axis running down a central pore concentric with the barrel. The barrel is 10.5 nm, 13.2 nm in diameter at its widest point, and the central aqueous pore decreases in diameter from 4.9 nm at the ends of the barrel to 0.6 nm at the mid-point. The precise arrangement of the L and S subunits within the overall smooth outline of the barrel has not been elucidated so far (Eisenberg et al., 1978; Chapman et al., 1986). Possible differences between the basic structures of microbial and plant RuBisCO enzymes with 8L8S quaternary structures would be of evolutionary significance and the unknown arrangements of the subunits in the 8L8S enzymes of the cyanelles and prochlorophytes are of particular interest in this context. The R. rubrum 2L molecule has the shape of a distorted ellipsoid about 5 x 7.2 x 10.5 nm (50 x 72 x 105 A) in size, with tight and extensive subunit interactions (Branden et af., 1986; Schneider et af.,1986). These workers have incorporated four 2L ellipsoid discs into a model, with about the same dimensions as the bacterial 8L8S enzyme (Bowien et af., 1976) which could provide the 8L basis of the microbial and plant molecule. Despite the low primary sequence homology between the R . rubrum and hexadecameric RuBisCO L subunits, high homologies exist in regions identified as catalytic and enzyme activation sites and the R . rubrum enzyme can be expected to have further value in understanding the origin and assembly of the predominantly extant 8L8S RuBisCO enzymes. B. ACTIVATION AND CATALYSIS

1 . Activation

Pioneering work by Pon et al. (1963) indicated that RuBisCO activity in vitro is enhanced by preincubation with COZ and Mg2+ before the addition of ribulose 1,5-bisphosphate (RuBP). The basis of these observations has become a fundamental principle in the understanding of RuBisCO function: namely that the enzyme, from all sources examined, can exist in vitro in an active and inactive form (Miziorko and Mildvan, 1974; Lorimer et af., 1976;

136

G.A. CODD

Laing and Christeller, 1976; Badger and Lorimer, 1976; Lorimer, 1981; Miziorko and Lorimer, 1983). RuBisCO activation is an ordered, two stage, pH-dependent process which requires the slow binding of an activating C02 molecule (AC02,not bicarbonate) followed by the rapid addition of Mg2+: slow fast E + AC02+E -AC02+ Mg2++E -AC02-Mg2' (active) (inactive)

The AC02molecule is distinct from that which is assimilated by the catalytic reaction and binding of one AC02per L subunit occurs at the activation site by reactivation with an E-lysyl residue to form a carbamate. This reaction is favoured by alkaline pH values and results in the production of a negatively charged carbonyl group from a positively charged or neutral amino group. The carbonyl group provides a binding site for a Mg2+ ion (Lorimer and Miziorko, 1980). The AC02-binding step is the rate-limiting reaction: E-lys-NH2+AC02+H (inactive)

+

+ E-IYs-NH-~CO~ +Mg2++E-lys-NH-C02-

.Mg2

+

(active)

The activation site was rigorously localized as lysine-201 of the L subunit of the spinach enzyme and lysine 191 of R . rubrum RuBisCO (Lorimer, 1981; Miziorko and Lorimer, 1983; Donnelly et al., 1983). This is most likely to be the activation site for microbial RuBisCO enzymes as a whole since the lysine201 domain is highly conserved and carbamate formation has been indicated during activation bf the enzyme from Chromatium and Thiobacillus species (see Codd, 1984). Activation of RuBisCO of Alcaligenes eutroghus resulted in a decrease in from 17.5s (inactive) to 14.3s (active) the sedimentation coefficient , S20,w1, suggesting that this enzyme may undergo conformational change during activation (Bowien and Gottschalk, 1982). However, no such changes were observed during the activation of spinach RuBisCO (Donnelly et al., 1984) and this problem is yet to be resolved.

2. Catalysis

In all autotrophs the RuBisCO is a bifunctional enzyme capable of catalysing the carboxylation or oxygenolytic cleavage of RuBP: Ribulose 1,5-bisphosphate+CO2

carboxylase .2 (3-phosphoglycerate)

or oxygenase Ribulose I ,S-bisphosphate +0 2 -3-phosphoglycerate

+2-phosphoglycollate

The oxygenase reaction, an apparently wasteful process, may impose considerable physiological and ecological constraints on all Calvin cycle

CARBOXYSOMES A N D RIEULOSE BISPHOSPHATE CARBOXYLASE/OXYGENASE

I37

autotrophs other than the anaerobes. Oxygen is a linear competitive inhibitor of the carboxylase reaction and vice versa (Lorimer, 1981; Codd, 1984). The two reactions that compete for the pentose phosphate acceptor, RuBP, occur at the same catalytic site and the oxygenase reaction requires activation to proceed via carbamate formation at L subunit lysine-201 as described for the activation of the carboxylase reaction. The carboxylase and oxygenase common catalytic site is on the L subunit. Affinity labelling studies and comparative amino acid-sequence analyses have indicated the existence of two lysine residues, distinct from that at the activation site, which are essential for catalysis. These are lysine- 166 and -329 in the R . rubrurn L subunit, and lysine-175 and -334 in the spinach L subunit (Hartman et al., 1984, 1986). Evidence for the proximity of the two essential catalytic lysine residues has been obtained using cross-linking agents. A purified peptide has been produced and sequenced by Hartman et al. (1986) and the cross-linked lysine-166 and lysine-329 residues found to be only 1.2 nm (12 A) apart. These residues occur in the same loop region of the R . rubrum L subunit according to the three-dimensional structure determined by X-ray crystallography at 0.29 nm (2.9 A) (Schneider et al., 1986). Increased knowledge of the three-dimensional structure of the active-site of RuBisCO is necessary to enable active-site mutagenesis to be applied to attempt to manipulate the relative specificities of the carboxylase and oxygenase reactions in favour of increased COZassimilation. Chemical and mechanistic details of the carboxylation and oxygenation of RuBP are emerging. A carboxylation scheme consisting of five steps has been formulated from the outstanding work of Lorimer and his coworkers (e.g. Miziorko and Lorimer, 1983; Schloss and Lorimer, 1982; Pierce ef af., 1986; Lorimer et af., 1986): (a) RuBP undergoes enolization involving deprotonation at the C-2 position to give a nucleophilic 2,3-enediol at C-2; (b) carboxylation at the 2,3-enediol produces the unstable C6 intermediate, 2’-carboxy-3-keto-~-arabinitol 1,5-bisphosphate (CKABP); (c) the C-3 ketone form of CKABP is hydrated to the gem-diol form; (d) carbon-carbon cleavage of CKABP occurs by deprotonation to produce one molecule of lower D-glycerate 3-phosphate (IPGA) plus one C-2 carbanion form of upper D-glycerate 3-phosphate (uPGA); (e) finally the C-2 carbanion derivative is stereospecifically protonated to produce uPGA. This proposed mechanism for carboxylation applies to the RuBisCO enzymes of spinach and probably Synechococcus. Understanding of the complete mechanisms of the carboxylation and oxygenation reactions (Miziorko and Lorimer, 1983; Lorimer et af., 1986) will combine with that of

138

G.A. CODD

the three-dimensional structure of plant and microbial RuBisCO enzymes to enable the site-directed mutagenesis approach to be attempted by design in the goal of selectively modifying RuBisCO enzymes to favour the carboxylase at the expense of the oxygenase reaction. 3. Small Subunit Function

The available data indicate the mechanisms and site of carboxylation and oxygenation reactions are common throughout the plant and microbial 8L8S RuBisCO enzymes, and the role of the L subunits is well established. Uncertainty exists, however, about the precise function of the S subunits. Clearly, they are not necessary for activation, or catalysis of carboxylation or oxygenation by the 2L RuBisCO of R . rubrum or the 6L enzymes of the Rhodopseudomonads. Advances in the understanding of the functional role of S subunits have arisen from studies of the RuBisCO enzymes of cyanobacteria and the purple bacteria. Differences in the catalytic and regulatory properties of the Form I (8L8S) and Form I1 (6L) enzymes of Rps. sphaeroides may be due to the presence of S subunits in Form 1. On this basis, the S subunits can be inferred to: enable the requirement for Mg2+(or Mn2+) for activation and catalysis to be replaced by Ni2+or Co2+;increase the rate of C02-plus-Mg2+activation; increase the inhibition of this activation if RuBP is present; and confer a higher affinity for C02 (Gibson and Tabita, 1979). However, deduction of the role(s) of the S subunits by this means is limited since the L subunits of the Forms I and I1 RuBisCO enzymes of Rps. sphaeroides, and probably those of Rps. capsulata and Rps. blastica, themselves differ in peptide composition, physiological properties and genetic origin (see Section 1V.A). The removal of S subunits from the 8L8S RuBisCO enzymes of several microbes yields an 8L “catalytic core” whose carboxylase and oxygenase catalytic capacities are reduced equally to very low levels. The loss of RuBisCO activity in Synechococcus sp. proceeds in proportion to the degree of S subunit depletion (Andrews and Ballment, 1983) and as a corollary, the decreased activities of the 8L catalytic cores of Aphanothece halophytica and Chromatium vinosum are increased in proportion to the amounts of S subunits added (Asami et al., 1983; Incharoensakdi et al., 1985; Akazawa et al., 1984). S subunit addition to the Ap. halophytica L8 catalytic core does not affect the K, values for C02 or RuBP but increases the V,,, value for carboxylation by at least 2-3-fold (Asami et al., 1983). Akazawa and co-workers have extended this approach by comparing the effects of homologous and heterologous subunit reconstitution. The requirement for S subunits for catalysis by the 8L cores of Synechococcus sp. and Ap. halophytica can be completely satisfied by homologous S subunit addition and

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partly fulfilled by heterologous hybridization with the reciprocal cyanobacterial S subunits and isolated subunits from the purple bacterium Cr. vinosum (Incharoensakdi et al., 1985). In some instances structural differences between the RuBisCO enzymes of individual organisms may preclude successful hybridizations: S subunits from Synechococcus sp. and Ap. halophytica cannot restore the activity of the depleted C r . vinosum L8 core. However, these findings, in sum, are highly promising since they show that catalytically active hybrid 8L8S RuBisCO enzymes can be constructed in vitro by the combination of subunits from different microbes. This approach may be useful as a strategy to “improve” the catalytic properties of RuBisCO if these are influenced by S subunits. Incharoensakdi et al. (1985) have attempted this strategy using the subunits of the spinach and Ap. halophytica RuBisCO enzymes, which differ in that the apparent K , (COZ)value of the cyanobacterial enzyme is about ten times greater than that of the plant enzyme. Hydridization between the Ap. halophytica 8L core plus spinach S subunits occurred with an increase in carboxylase Vmax,but the apparent K, (C02) value of the hybrid enzyme was not significantly lower than that of the cyanobacterial holoenzyme. Heterologous hybridization has also been performed between the L subunit core of RuBisCO of Synechococcus sp. and S subunits from Prochloron sp and spinach and restoration of catalytic activity obtained (Andrews et al., 1984; Andrews and Lorimer, 1985). The heterologous S subunits bound about ten times less tightly than the homologous S subunits of Synechococcus sp. and the apparent K m (CO2) value of the heterologous construct was about twice as high (Andrews and Lorimer, 1985). Although the hybrid 8L (Synechococcus):8 s (spinach) enzyme differed in apparent K , (C02) value from either holoenzyme, the specificity factor, a measure of carboxylase versus oxygenase activity (see below) was not affected (Andrews and Lorimer, 1985). This suggests that the enzymatic partitioning between the carboxylation and oxygenation of RuBP may be specified by the L subunits only. Indeed the role for the S subunits is still little understood: although necessary for carboxylation and oxygenation by the 8L8S enzymes, the binding of COz and stable complex formation with the transition state analogue 2-carboxyarabinitol 1,5-bisphosphate (CABP) to the enzyme of Synechococcus sp. still occurred when the enzyme was depleted by more than 90% of its S subunits (Andrews and Ballment, 1984). These findings have been supported by similar findings of RuBisCO Cr. vinosum (Jordan and Chollet, 1985). The S subunits are not therefore apparently essential for the activation process. At apparent variance with these findings is the ability of a second form of RuBisCO containing 8L subunits only, purified from Cr. vinosum, to exhibit similar carboxylase and oxygenase activities to the 8L8S form (TorresRuiz and McFadden, 1985). The catalytically competent Cr. vinosum 8L molecule was extracted and purified under protective conditions (in the

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presence of poly(ethy1ene glycol) in contrast to the acid or alkaline treatments which earlier studies have included for the removal of S subunits (Codd and Stewart, 1977c; Andrews and Ballment, 1983; Jordan and Chollet, 1985). If such methodological differences account for the disparate observations of the RuBisCO of Cr. vinosum, then it may be inferred that the S subunits are required to renature L subunits. Full understanding of the fundamental grounds for requirement of S subunits which are present in virtually all microbial RuBisCO enzymes and throughout the eukaryotes remains an important constraint in the cognition of CO2 assimilation via RuBisCO. C. SPECIFICITY AND REGULATION

2. Carbon DioxidelOxygen Specijicity

The inhibition of plant and microbial COZassimilation via the Calvin cycle by 02,as initially observed during photosynthesis by Chlorella sp. (Warburg, 1920), can be partly accounted for by the competitive inhibition of the carboxylase reaction of RuBisCO by 0 2 (Lorimer, 1981; Codd, 1984). Although the oxygenase reaction is a characteristic feature of all RuBisCO enzymes, considerable interest exists in the possibility of finding enzymes with decreased oxygenase activities ,or in the possibility of selectively reducing this activity compared with carboxylase activity. The possibility of obtaining a 50% increase in plant productivity by abolition of the oxygenase reaction is of obvious attraction to agriculture (Somerville et al., 1983). Although the oxygenase reaction may be an inevitable and unavoidable characteristic of RuBisCO (Lorimer, 1981; Codd, 1984), there are grounds for investigating selective manipulation. The carboxylase and oxygenase activities of RuBisCO from R . rubrum and Euglena gracilis are differentially affected by metal ions and temperature (Robison et al., 1979; Jordan and Ogren, 1984; Wildner and Henkel, 1978). Furthermore, a naturally occurring variation in the relative kinetic paramelers of the two competing reactions has emerged among the autotrophic microbes, algae and higher plants. This is expressed as variation in the C02/02 specificityvalues for the enzymes (Laing et al., 1974; Ogren, 1984). The interactions between photosynthesis and plant photorespiration can be expressed in terms of RuBisCO kinetics: Photosynthesis/photorespiration = u,/tuo = VcKoC/tV&O

are the carboxylation and oxygenation velocities; Vcand V, are the V,,, values of these activities; K , and KOare the K , values for the CO2 and O2 concentrations (C, 0).f is 0.5 which is the stoichiometry between the amount of O2 consumed by the oxygenase and the amount of CO2 released during photorespiration (Ogren, 1984). Absolute values for the C02/02 substrate - oc and v,

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14 1

specificity of RuBisCO can be obtained by plotting the ratio of the carboxylase and oxygenase activities as a function of the ratio of the two substrates. From the above equation, the slope of the line equals VcKo/VoKc, which is the C 0 2 / 0 2substrate specificity. Values have been determined for a considerable number of the RuBisCO enzymes from C3- and Cq- (Calvin cycle and Hatch-Slack) pathways of higher plants and from one intermediary species. The substrate specificities are particularly clustered in the case of the C3-plants (Fig. 3). Fewer microbial enzymes have been characterized in this manner, although if plotted in taxonomic groups and in the likely sequence for the evolution of autotrophy, a trend can be discerned. The C02/02 specificities of the photosynthetic bacterial RuBisCO enzymes lacking S subunits are the lowest known. The Rps. sphaeroides Form I (8L8S) enzyme substrate specificity is significantly higher and the values for the cyanobacterial and microalgal RuBisCO enzymes generally lie between the photosynthetic bacterial Form I1 enzymes and the enzymes from higher plants (Fig. 3). It is likely that the low CO2/O2substrate specificities of the enzymes of the photosynthetic bacteria would not have disadvantaged those organisms that perform anoxygenic photosynthesis and show an anaerobic mode of autotrophic growth. The increase of 0 2 in the atmosphere, largely due to the

C4-plonts

0

0

C3/ C4-plants

micro-olgae

:: 0

Ic

0 . .

0

0 C 0 c

.-

-0a cyanobocterio

0

0"

0

0)

photosynthetic a 0

bacteria

0

0

I

1

I

I

20

40

60

8

0

,

FIG. 3. Carbon dioxide specificity (VcKo/Vo&) values for microbial and plant RuBisCO enzymes. From: 0 (Ogren, 1984); 0 (Kent and Tomany, 1984).

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development of oxygenic photosynthesis by the cyanobacteria, would have imposed increasing pressure on the RuBisCO enzymes of the ancestral aerobic autotrophic prokaryotes and selected for the development of RuBisCO enzymes with higher C02/02 specificities. These can be found among the cyanobacterial, microalgal and, to a greater extent, among the higher plant enzymes. Other strategies have been developed to lessen the inhibitory effect of increasing 0 2 tensions in the environment on the cyanobacteria and algae, i.e. C02- concentrating mechanisms (Aizawa and Miyachi, 1986), and in the C4-plants, namely the Hatch-Slack, or Co-dicarboxylic acid pathway (Hatch, 1977). These developments may have decreased further evolutionary pressure on the RuBisCO enzymes in these organisms. The C3-pathway higher plants, which lack these mechanisms for reducing photorespiration, remain as the group most susceptible to the inhibitory effect of 0 2 on photosynthesis. Their RuBisCO enzymes generally show among the highest substrate specificities and little interspecies variation exists (Ogren, 1984; Fig. 3). However, the variations in C02/02 substrate specificity throughout the phototrophs and in K, within the RubisCO enzymes of C3-plants (Lorimer, 1981; Miziorko and Lorimer, 1983) support the case for further attempts to manipulate specificity by recombinant DNA technology and site-directed mutagenesis. Data on the substrate specificities of RuBisCO enzymes from the aerobic chemolitho-autotrophic bacteria are lacking and it would be of interest from an evolutionary viewpoint to compare them with the range shown in Fig. 3. Similarly, insufficient information is available to compare the substrate affinities of RuBisCO enzymes from prokaryotes with and without carboxysomes (Codd and Marsden, 1984). Within the individual carboxysomecontaining species Anabaena variabilis and Thiobacillus neapolitanus, the K,,, (COZ) values of the enzyme derived in vitro from the cytoplasmic and carboxysomal pools are closely similar, e.g. 252_+7 and 2 6 9 k 9 PM for Anabaena variabilis (Badger, 1980). 2. Phosphorylated Eflectors

Numerous reports exist on the positive and negative effects of sugar phosphate intermediates of the Calvin cycle, 6-phosphogluconate (6PGLU) and nucleotides on plant and microbial RuBisCO enzymes in vitro. For example, several microbial RuBisCO enzymes, activated by preincubation with COZand Mg2+ in vitro are inhibited by 6PGLU (Tabita and McFadden, 1972; Gibson and Tabita, 1977a; Codd and Stewart, 1977c; Snead and Shively, 1978; Tabita and Colletti, 1979). On the other hand, stimulation of inactive RuBisCO by 6PGLU plus bicarbonate has been obtained with the isolated enzymes of Pseudomonas oxalaticus, cyanobacteria and the 2L enzyme of R . rubrum (Lawlis et al., 1978; Tabita and Colletti, 1979; Whitman

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el al., 1979). The findings with R . rubrum, also obtained with fructose 1,6-

bisphosphate, 2-phosphoglycollate and NADPH, are consistent with results obtained with higher plant RuBisCO enzymes and with the conclusion that the metabolite effectors interact on the L subunit of the enzymes at the catalytic site (Lorimer, 1981). Evidence for the regulation of microbial RuBisCO in vivo by these effectors may be considered: Bassham and his colleagues found that transfer of photosynthesizing steady-state cultures of cyanobacteria from the light to the dark caused immediate cessation of I4CO2 fixation and the production of 6PGLU among the labelled sugar phosphates (Pelroy and Bassham, 1972; Pelroy et al., 1976). These workers, and Stanier and Cohen-Bazire (1977), reasoned that BPGLU, which is not a Calvin cycle intermediate but rather accumulates as an oxidative pentose phosphate pathway intermediate only in the dark in cyanobacteria, may inhibit RuBisCO in vivo. Tabita and Colletti (1979) extended this investigation by measuring cyanobacterial RuBisCO in situ using toluene-permeabilized cells. Brief treatment with toluene permits the entry of RuBP, other sugar phosphates and nucleotides. Assuming that the measurement of RuBP-dependent I4CO2incorporation into acid-stable material by toluene-treated cells gives an estimation of potential in vivo RuBisCO activity, than the inhibition of the activated enzyme by exogenous 6PGLU, fructose 6-phosphate, fructose 1,6-bisphosphate, NADPH and ATP can occur in cyanobacteria in vivo. Furthermore, stimulation of non-activated RuBisCO by these compounds was obtained with the permeabilized cells (Tabita and Colletti, 1979). The significance of 6PGLU and other metabolites on C02 fixation by plant and microbial RuBisCO enzymes in vivo is questioned, however, since the metabolites occupy the catalytic site, i.e. the RuBP-binding site (Lorimer, 1981). Badger and Lorimer (1981) have proposed that the binding of an effector to the enzyme in vitro at the catalytic site will result in a (RuBisCOCOrMg2+-effector)complex. If, on dilution of the complex in an activity assay mixture, the effector dissociates rapidly from the complex, then the enzyme would remain activated, its catalytic site would be vacant and the effector would appear in a positive mode. If the effector only dissociated slowly from the complex on dilution for assay, then the catalytic site would remain essentially occupied, RuBP binding for carboxylation would be delayed and the effector would thus appear to act negatively (Badger and Lorimer, I98 1). Although metabolites may thus increase the activation state in vitro and decrease the K,,, (CO2) value, their contribution as positive effectors in vivo is difficult to accept since they bind at the catalytic site and thus prevent substrate binding.

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3. Endogenous Ribulose 1,5-Bisphosphate CarhoxylaselOxygenase Inhibitors

The binding of 6PGLU at the catalytic site of RuBisCO would clearly permit this metabolite of dark (heterotrophic) metabolism and related compounds to act as an inhibitor. Diurnal changes in RuBisCO activity in a range of higher plants have been observed (Vu et al., 1983,1984; Servaites et al., 1984; Seeman et al., 1985; Servaites, 1985).These are not explained by changes in RuBisCO protein content but by the binding of a phosphorylated compound which acts as a non-competitive inhibitor with respect to RuBP. The naturally-occurring inhibitor accumulates in the dark in the chloroplast stroma and preferentially binds to the activated form of RuBisCO. A full restoration ofcatalytic activity is rapidly achieved upon illumination (within one hour). The inhibitor binds sufficiently tightly to RuBisCO to permit it to remain associated with the enzyme purified from dark-incubated plants. Tobacco RuBisCO activity is inhibited about 90% at a ratio of 10 moles inhibitor per mole of RuBisCO active sites (Servaites, 1987). The inhibitor has been identified from garden bean and potato leaves as the pentitol monophosphate, 2'-carboxyarabinitol1-phosphate (2CAlP) (Gutteridge et al., 1987; Berry et al., 1987). Whether this or related compounds with structural similarity to the C-6 carboxylation intermediate, function to regulate microbial RuBisCO enzymes in vivo is unknown. The 2CA 1P would have the advantage over 6PGLU that it would not be needed in such high excess over the RuBisCO active sites (Servaites, 1987).

4. Ribulose 1,5-Bisphosphate CarboxylaselOxygenase Activase Further exciting advances have recently been made in the understanding of the regulation of higher plant RuBisCO in vivo.Among these is the discovery of a soluble protein needed for RuBisCO activation in vivo. The discovery of this enzyme, termed RuBisCO activase, has further helped to resolve the differences between the requirements for maximal RuBisCO activity in vitro and the optimal provision of these requirements in the whole plant (for primary references, see Ogren et al., 1986). RuBisCO activation in whole leaves at high light intensities is a widespread phenomenon and the isolation of an Arabidopsis thaliana mutant by Somerville et al. (1982), which was unable to activate RuBisCO in the light, provided the opportunity for comparative studies versus the wild type which shows light activation of the enzyme. The light activation lesion could not be accounted for by differences in the RuBisCO enzymes. Two polypeptides, M , 40-50 kDa, were present in the chloroplast stromal proteins of the wild type, but were absent from the Ar. thaliana mutant (Salvucci et al., 1985). The requirement for soluble chloroplast protein was demonstrated in a reconstituted assay system, which

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additionally contained purified RuBisCO, thylakoids and RuBP and exhibited a light-stimulated increase in RuBisCO activity. The possibility that RuBisCO light activation may occur via the ferredoxin/ thioredoxin system, which accounts for the light activation of other Calvin cycle enzymes in plants and cyanobacteria, was discounted: the Ar. thaliana mutant was not deficient in the ability to show light activation of chloroplast fructose 1,6-bisphosphatase (FBPase) which is light-activated by the thioredoxin/ferredoxin-mediated transfer of electrons from the photosynthetic electron transport chain. Furthermore, the light activation of RuBisCO proceeds in the presence of methyl viologen which inhibits FBPase photoactivation (Salvucci et al., 1986b). The precise role of light in the activation process is not yet known. The COZrequirement for chloroplast RuBisCO activation with RuBisCO activase is about 4 PM which is significantly below the atmospheric CO2 concentration of about 10 PM (Portis et al., 1987). The two polypeptides of the RuBisCO activase of Ar. thaliana have recently been estimated to be of about 41 and 44 kDa and antibodies raised to the enzyme cross-react with leaf extracts from the homologous wild-type Arabidopsis sp., and at least 14 other diverse plants (Salvucci et al., 1987). RuBisCO activase thus appears to be widely distributed in higher plants and the immunological cross-reactivities obtained indicate that the polypeptides have been conserved. The constituent polypeptides also appear to be present in the green alga Chlamydomonas reinhardtii (Salvucci et a/., 1987). if RuBisCO activase is indeed commonly involved in the control of RuBisCO in the eukaryotic phototrophs, then a search for the origin(s) of this protein among the photosynthetic prokaryotes will be of obvious interest. D. GENETICS

Research on the genetics of microbial RuBisCO enzymes is advancing rapidly. Progress on the location, cloning, heterologous expression and regulation of the L and S subunit genes from the various physiological groups of autotrophs has been of value to microbiology. These advances are of no less interest to plant geneticists and agronomists since the prokaryotic RuBisCO genetic systems have several advantages for study and manipulation over those in eukaryotes. In almost all of the eukaryotes examined to date, the genes for the L and S subunits of RuBisCO have been located in the chloroplast and nuclear chromosomes respectively (Coen er al., 1977; Kawashima and Wildman, 1972; Akazawa et al., 1984). Detailed consideration of the genetics of RuBisCO in eukaryotes is beyond the scope of this review and recent advances can be found elsewhere (Ellis and Gray, 1986). In prokaryotes that contain 8L8S RuBisCO enzymes, the genes for both types of subunit are located on the chromosome, as are the genes of the 2L and 6L prokaryotic RuBisCOs.

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Among the photosynthetic bacteria, the RuBisCO genes have been located and cloned from R . rubrum (Nargang et al., 1984; Somerville and Somerville, 1984), Rps. sphaeroides (Fornari and Kaplan, 1983; Quivey and Tabita, 1984; Gibson and Tabita, 1986; Tabita et al., 1987) and Cr. uinosum (Viale et al., 1985; Kobayashi et al., 1987). The L and S subunit genes for the Rps. sphaeroides Form I RuBisCO are located on a single 4 kb chromosomal fragment (Gibson and Tabita, 1986). Among the cyanobacteria, the L and S subunit genes have been shown to be closely linked and have been cloned from Anacystis nidulans (Synechococcus) 630 1 (Shinozaki and Sugiura, 1983,1985; Shinozaki et al., 1983; Christeller et al., 1985; Tabita and Small, 1985; Gatenby et al., 1985; Bradley et al., 1986; Gutteridge et al., 1986), Anabaena 7120 (Nierzwicki-Bauer et al., 1984; Gurevitz et al., 1985), Spirulina platensis (Tiboni et al., 1984) and Ch.fritschii (Vakeria et al., 1986). Location of the L and S subunit genes in the cyanelles of C .paradoxa was of particular interest since these organelles are incapable of living outside of their eukaryotic host and the possibility existed that the S subunit gene may have been among the genes thought to have been transferred from the cyanelle to the host nucleus. However, as in the free-living autotrophic prokaryotes, the L and S subunit genes of C . paradoxa are both on the cyanelle chromosome (Heinhorst and Shively, 1983; Bohnert et al., 1983; Mucke et al., 1984). They are closely linked: the L subunit gene is situated 105 nucleotides upstream from the S subunit gene and the spacer sequence does not indicate the presence of a promoter sequence. Indeed, cotranscription of the L and S subunit genes from the 2500 nucleotide sequence has been demonstrated by Northern blot analysis (Starnes et al., 1985). These findings are of evolutionary interest and raise the question of when and where the segregation of the L and S subunit genes occurred in the development of eukaryotic phototrophs. Until recently, RuBisCO gene research in the eukaryotes had centred almost entirely on chlorophytic (chlorophyll a plus b ) organisms, in which, as discussed earlier, the L and S subunit genes have been invariably found to be segregated to the chloroplast and nucleus. Reith and Cattolico (1986) have chosen to examine the DNA of the non-green chromophyte alga Olisthodiscus luteus and found that the L and S subunit genes are linked on the chloroplast DNA. This may also be so in the red algae Cyanidium caldarium and Porphyridium cruentum, whose plastids are though to have originated from cyanobacterial endosymbionts (Steinmuller et al., 1983). The eukaryotic RuBisCO enzymes with linked genes may be of practical interest, in addition to evolutionary interest, since it should be possible to obtain heterologous expression and assembly of the cloned non-chlorophyte 8L8S enzyme, in contrast to the enzyme from chlorophytic higher plants (c.f. Bradley et al., 1986). Complete nucleotide sequence data for the gene coding for the R . rubrum L subunit and for the L subunit genes for the cyanobacteria Anacystis nidulans

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6301 and Anabaena 7120 enzymes are available (Nargang et al., 1984; Shinozaki et al., 1983; Curtis and Haselkorn, 1983). The amino acid sequence ofthe R. rubrum L subunit, deduced from analysis of the enzyme purified from the organism itself (Hartman et al., 1984) and from Escherichia coli into which the L subunit gene was cloned (Somerville and Somerville, 1984; Nargang ef al., 1984) shows only 31-33% homology with the spinach L subunit. This is in contrast to the > 80% homologies between the spinach L subunit amino-acid sequence and those of other higher plants, green algae and cyanobacteria (see Hartman et al., 1984). Regions of nucleotide sequence homology are likely to be functionally important and have already been useful in confirming the activation and catalytic sites on the L subunits (Section 1V.B). Other regions sequences of high homology may be likely targets for attempts to modify RuBisCO by site-directed mutagenesis. Gutteridge et al. (1986) have investigated the effects of substituting a conserved aspartic acid residue at position 198 on the R . rubrum enzyme by glutamic acid. In this case, the mutant showed a 30% decrease in carboxylase and oxygenase activities although the apparent K,,,values for C02 and 0 2 and the activation rates with C02 and Mg2+ were unaffected. A reduction in turnover, perhaps due to a spatial need to accommodate the extra methylene group of glutamate, appears to be the sole effect in this initial attempt at site-directed mutagenesis, but systematic substitution of amino acids in the highly conserved regions of the subunits is likely to provide further mechanistic information and, it is hoped, to provide prospects for the selective manipulation of substrate specificities in favour of the carboxylase. Unlike plants that contain multiple copies of the L and S subunit genes, only one copy of the L subunit gene is present in R . rubrum (Somerville and Somerville, 1984) and single copies of each of the L and S subunit genes are present in cyanobacteria and the C . paradoxa cyanelles (Shinozaki and Sugiura, 1983; Nierzwicki-Bauer et al., 1984; Vakeria et al., 1986; Heinhorst and Shively, 1983). This indicates that the cytoplasmic and carboxysomal pools of RuBisCO are products of the same genes within each organism. Chlorogloeopsis fritschii contains only one set of L and S subunit genes (Vakeria et al., 1986) in agreement with findings with other cyanobacteria (Shinozaki et al., 1983; Nierzwicki-Bauer et al., 1984). However, Ch.fritschii, according to six plasmid isolation methods, does not appear to contain extrachromosomal DNA (Vakeria et al., 1984) and we wished to know whether extrachromosomal RuBisCO genes may be additionally present in a plasmid-containing strain. Microcystis 7820 contains 3 4 cryptic plasmids (Vakeria et al., 1985)but when these were probed under a variety of stringency conditions with a range of heterologous RuBisCO genes, no hybridization was detected. This was in contrast to positive hydridization between Microcystis chromosomal DNA and the L subunit probe from Anacystis

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nidulans (Vakeria et al., 1986). These findings indicate that if the carboxysomes of Microcystis sp. did contain extrachromosomal DNA, and there is no evidence for this in cyanobacteria (see Section III.A), then it is unlikely that such carboxysomal DNA would contain RuBisCO genes. Evidence for the presence of RuBisCO genes on extrachromosomal DNA exists, however, in the purple non-sulphur bacteria and hydrogen bacteria. The single copy of the Form I1 (6L) RuBisCO gene of Rps. sphaeroides is present on the chromosome (Fornari and Kaplan, 1983; Quivey and Tabita, 1984) but the single copy L and S genes of the Form I enzyme has been localized on an endogenous plasmid (Gibson and Tabita, 1986). The separate identity and location of the L subunit genes of the Form I and I1 enzymes confirm the observed differences in kinetic and physiological properties of the enzymes (Tabita et al., 1987) and open the possibility for unambiguous investigation of the physiological regulation of these enzymes at the genetic level. Duplicate L and S subunit genes exist in the nutritionally versatile hydrogen bacterium Alcaligenes eutrophus. The role of megaplasmids in lithoautotrophic metabolism is well established since hydrogenase genes are plasmid-encoded in several strains (Schlegel, 1984; Friedrich, 1987). For example, in Al. eutrophus HI 6 the self-transmissible 450 kb megaplasmid encodes for the two hydrogenases of this autotroph and the ability to grow chemolitho-autotrophically has been abolished by plasmid-curing of this strain (Friedrich et al., 1981). Since plasmid-cured strains are still able to grow organo-autotrophically on formate, it is clear that the essential genes of the Calvin cycle and any essential COZfixation regulatory genes must be on the chromosome (Bowien et al., 1984). However, the involvement of megaplasmids was indicated in the partial derepression of RuBisCO and phosphoribulokinase proteins upon transfer from chemoheterotrophic to autotrophic conditions. These findings were confirmed by the first demonstration of multiple phosphoribulokinase genes on the strain H 16 chromosome and megaplasmid (Klintworth et al., 1985). More recently, Bowien and colleagues have used the R . rubrum RuBisCO gene to probe for the L subunit gene and a synthetic DNA sequence constructed on the basis of the amino-terminal amino acid sequence of the homologous S subunit, to localize the S subunit gene in Al. eutrophus H 16. Both genes are present on the chromosome and the megaplasmid. In each case, the RuBisCO genes constitute an operon as evidenced by Northern blot hybridizations which indicate a 2.2 kb transcript size. In the same direction of transcription as the L and S subunit genes, but about 3.5 kb downstream and under separate transcriptional control in the chromosome and megaplasmid is the phosphoribulokinase gene (Bowien et al., 1987). Additional evidence exists for the presence of functional RuBisCO genes on the chromosome and megaplasmid of Al. eutrophus strain ATCC 17707 (Andersen and Wilke-Douglas, 1984; Andersen et al., 1986). The high

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degree of homology between the RuBisCO gene regions on the megaplasmid and chromosome and the expression of active enzyme in E. coli from the cloned plasmid genes (Bowien et al., 1987) suggest that the latter are functional and the role of the duplicate sets of Calvin cycle genes in these hydrogen bacteria requires investigation. Some preliminary generalizations concerning expression of RuBisCO genes in the prokaryotes may be ventured: although there is variation in the spacer region between the Land S subunits, cotranscription of the genes occurs in the cyanobacteria Anacystis nidulans 6301 (Shinozaki and Sugiura, 1985) and Anabaena 7120 (Nierzwicki-Bauer et al., 1984), and Al. eutrophus strains (Andersen et al., 1986; Bowien et al., 1987), as with the Cyanophora cyanelle genes (Starnes et al., 1985). Cotranscription with single promoter control is clearly one factor in the success widely achieved in the heterologous expression of structurally accurate and catalytically competent 8L8S RuBisCO from several prokaryotic sources. This is in contrast to the problems encountered in attempts to obtain expression of catalytically active RuBisCO of higher plants (see Bradley et al., 1986). Expression of microbial RuBisCO enzymes in E. coli to give at least wild-type activities has been obtained with the genes from R. rubrum (e.g. Somerville and Somerville, 1984) Rps. acidophila Forms I and I1 enzymes (Tabita et al., 1987), Cr. uinosum (Viale et al., 1985), An. nidulans (Tabita and Small, 1985; Gatenby et al., 1985), Spirulinaplatensis (Tiboni et al., 1984) and Ch.fritschii (Vakeria et al., 1986). Alcaligenes eutrophus RuBisCO genes have been expressed to produce active enzyme in Pseudomonas aeruginosa (Andersen et al., 1986) in addition to E. coli (Bowien et al., 1987). A further factor in the faithful production of the prokaryotic 8L8S enzymes in these bacterial expression systems is that the microbial RuBisCO enzymes do not require additional assembly factors to be cloned from the autotrophs. The cloned fragments containing the An. nidulans 6301 and Al. eutrophus ATCC 17707 RuBisCO genes did not code for other polypeptides (Shinozaki and Sugiura, 1983,1985; Tabita and Small, 1985; Andersen et al., 1986). The production of large quantities of microbial RuBisCO by recombinant DNA techniques will greatly facilitate study of enzyme structure and manipulation by site-directed mutagenesis and post-translational modification. This approach will be additionally useful for the production of greater quantities of potentially interesting RuBisCO enzymes from slower-growing autotrophs.

V. Carboxysome Function

Knowledge of the occurrence, structure and composition of carboxysomes has increased considerably over recent years but understanding of the

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function(s) of these organelles remains inadequate. The hypotheses proposed have all sought, justifiably, to examine carboxysome function in terms of the properties and role of RuBisCO, the most abundant carboxysomal protein component (Shively, 1974; Stewart and Codd, 1975; Beudeker et al., 1981; Lanaras and Codd, 1982; Codd and Marsden, 1984). Evidence for and against these hypotheses exist. A. ARE CARBOXYSOMES SITES OF CARBON DIOXIDE FIXATION

In Vivo?

The possibility that carboxysomes are active sites of C02 fixation in vivo is attractive since all carboxysomes contain RuBisCO. If so, then carboxysomal RuBisCO must be catalytically competent. In all cases examined, the RuBisCO extracted from carboxysomal fractions has been found to be closely similar to, or identical with, in quaternary structure, immunological, activation and catalytic properties, the enzyme from the soluble (cytoplasmic) fraction of the cell (Badger, 1980; Lanaras and Codd, 198la,b; Beudeker et al., 1981; Leadbeater, 1981; Cannon and Shively, 1983). These studies with the enzymes from several cyanobacteria and T. neapolitanus show that carboxysoma1 RuBisCO enzymes are capable of COZfixation at rates similar, on an enzyme protein basis, to their counterparts in the cytoplasm. The ability of carboxysomal RuBisCO to be activated in vivo has been suggested from the data of Cannon (1982). The transition state analogue 2-carboxyarabinitol 1,s-bisphosphate (CABP) binds irreversibly at the catalytic site of the RuBisCO L subunit and locks the activating C02 and Mg2+ onto the activation site, thereby stabilizing the otherwise labile carbamate (see Miziorko and Lorimer, 1983). When ['4C]CABP was supplied to chloroform-permeabilized T. neapolitanus cells, labelling of the cytoplasmic and carboxysomal RuBisCO was obtained (Cannon, 1982; Cannon and Shively, 1982). The possible action of chloroform in influencing the permeability of the carboxysome membrane in vivo may not have influenced these findings since the membrane lacks lipids. If the carboxysome membrane was unaffected by the chloroform permeabilization procedure then these findings indicate that carboxysomal RuBisCO is capable of being activated in vivo, and that the membrane is permeable to C02, Mg2+ and the pentose CABP. Further investigations on the possibility that carboxysomes are sites of C02 fixation have centred on measurements of carboxysome abundance and the distribution of RuBisCO between carboxysomes and cytoplasm in whole-cell studies of cultures maintained under different physiological conditions. Continuous culture of T. neapolitanus under C02-limitation resulted in a 3.5fold increase in total extractable RuBisCO activity compared with cultures grown on excess COZ (i.e. thiosulphate limitation) (Beudeker et al., 1980). Changes in total and carboxysomal RuBisCO activities in these cultures

CARBOXYSOMES AND RIBULOSE BISPHOSPHATE CARBOXYLASE/OXYGENASE

15 I

correlated with variations in carboxysome volume density and number per cell. However, maximal COz-fixation capacity by whole cells, obtained during growth on thiosulphate plus 5% C02, varied inversely with carboxysome numbers and carboxysomal RuBisCO activities. Rates of RuBisCO activity per unit of RuBisCO protein in carboxysomal extracts from steady-state chemostat cultures of T . neupolitunus were about the same as those of the cytoplasmic enzyme in extracts of thiosulphate- or nitrogen (ammonia)limited cells. In extracts of C02-limited cells the carboxysomal RuBisCO was 4.5 times more active per unit RuBisCO protein than the cytoplasmic enzyme (Beudekeret al., 1981).These findings confirm the potential of the carboxysoma1 RuBisCO to contribute to whole-cell C02 fixation. This possibility also exists in Thiobacillus intermedius, which produces carboxysomes when RuBisCO activity enables or supports cell growth (i.e. under chemolithoautotrophic and mixotrophic growth conditions respectively) and does not produce the organelles, or RuBisCO, during chemoheterotrophic growth (Purohit et al., 1976). Carboxysomes and RuBisCO are produced in nutritionally versatile cyanobacteria irrespective of the sources of carbon and energy (Codd and Marsden, 1984) and the possibility that the organelles are active in C 0 2 fixation is open. Carboxysome numbers and levels of carboxysomal compared to cytoplasmic RuBisCO are minimal during the exponential growth phase of photo-autotrophic batch cultures of Ch. fritschii, in contrast to stationary phase cultures when almost all of the cell RuBisCO complement is located in

TABLE 7. Carboxysome abundance and photosynthetic characteristicsof Synechococcus leopoliensis under different nutrient limitations in continuous chemostat culture. From Turpin et a). (1984) Growth rate

Nutrient limitation

No. carboxysomes per cell section Reactor DIC K; DIC' pmax p (day-') % p max" ( k SE) (PM) ~

DICb Phosphate Nitrate

0.25 1.71 0.27 1.56 0.27 1.56

13 85 13 80 13 80

3.40f0.30 0.57 f0.09 0.41fO.10 0.34f0.10 0.40f0.10 0.40 f0.10

~~~

4.4 1700 6000 4090 5500 2800

109 308 485 399 ndd nd

Approximate estimate of per cent of maximum specific growth rate 01). Dissolved inorganic carbon concentration in reactor vessel. Half-saturation constant for photosynthesis for dissolved inorganic carbon (DIC). Not determined.

1.9 1600 1250 1500 nd nd

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the carboxysomes (Lanaras and Codd, 1982). These findings suggest that the carboxysomes have a principal role other than in C02 fixation in these cultures but the latter possibility is not excluded. Turpin et al. (1984) have studied carboxysome abundance versus photosynthetic characteristics of the cyanobacterium Synechococcus leopoliensis in chemostat culture (Table 7). This valuable study shows that carboxysome number per cell section under carbon limitation at 85% of maximum specific growth rate is marginally, but significantly, less than in phosphate- and nitrogen-limited cells. Under severe inorganic carbon limitation, at only 15% of pmax,carboxysome sections per cell section increased to exceed carboxysome abundance in phosphate- and nitrogen-limited cells by between six and ten times (Table 7). This increase was accompanied by a decrease of three orders of magnitude in the half-saturation constant of photosynthesis for dissolved inorganic carbon. These findings suggest strongly that under air-saturated levels of 02, inorganic carbon supply has a major effect on carboxysome numbers in S. leopoliensis and that the carboxysomes are functional in this organism under severe inorganic carbon limitation. The ability of the carboxysomes to protect RuBisCO from inhibition by 0 2 under these conditions is considered in Section V.B. If carboxysomal RuBisCO is active in C02 fixation in uiuo, then it is essential that the carboxysome membrane is permeable to RuBP. Phosphoribulokinase, which produces RuBP by phosphorylating ribulose 5-phosphate, is not present in the carboxysomes of Ch. fritschii, Synechococcus sp., T. neapolitanus or the Cyanophora or Glaucocystis cyanelles (Lanaras and Codd, 1981a; Cannon and Shively, 1983; Hawthornthwaite et al., 1985; Holthuizen et al., 1986b; Mangeney et al., 1987). Carboxysomes d o not contain Calvin cycle enzymes other than RuBisCO (Cannon and Shively, 1983; Holthuizen et a f . , 1986b), with the further implication that if the carboxysomal enzyme fixes C02 in vivo, then the carboxysome membrane must be permeable to the products of RuBisCO, 3-phosphoglyceric acid and 2-phosphoglycolate if carboxysomal RuBP oxygenation occurs, in addition to RuBP. If carboxysomes are a site of C02 fixation in viuo, then it is further possible that they act as a CO2-concentrating mechanism to favour the carboxylase reaction and help to account for the lack of photorespiration in cyanobacteria. This prospect has been discussed in detail previously and the existence of a inorganic carbon transport and concentration system at the cyanobacterial cell membrane is well documented. Carbonic anhydrase is also involved in inorganic carbon concentration (Aizawa and Miyachi, 1986), although the enzyme is lacking from the carboxysomes of T. neapolitanus and Ch.fritschii (Cannon and Shively, 1983; Lanaras et al., 1985; Holthuizen et al., 1986b).

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B. DO CARBOXYSOMES PROTECT RIBULOSE 1,5-BISPHOSPHATE CARBOXYLASE/ OXYGENASE

Ribulose 1,5-bisphosphate carboxylase/oxygenases are inhibited by several chemical and physical agents. Although the physiological significance of the inhibitory effects of metabolic intermediates is questionable, the inhibitory effect of 0 2 on CO2 fixation is an undoubted and indeed universal feature of RuBisCO enzymes (see Sections IV. B and IV. C). If carboxysomes served principally to protect RuBisCO from inhibition by 0 2 , then this would imply that the carboxysomal enzyme was active in COZfixation in vivo, and that carboxysomes were a characteristic feature of aerobic autotrophs. Speculation about carboxysomal functions in the different groups of autotrophs should not overlook the possibility that the organelles may play different roles in different physiological groups. However, in the context of protection from 02,it is worthy of note that carboxysomes do not occur in the anaerobic purple sulphur- and purple non-sulphur bacteria. Within the aerobic autotrophs, however, the only group in which they consistently occur are the oxygenic cyanobacteria. Although the organelles are also present in the three prochlorophytes examined, they d o not occur in all cyanelles (see Sections 11. B and 11. C). A protective role against 0 2 would also be consistent with the presence of carboxysomes in the aerobic chemolitho-autotrophs. Although carboxysomes commonly occur among the sulphur-oxidizers and nitrifiers, their distribution is variable (see Tables 1 and 2) and of the many aerobic hydrogen-oxidizing bacteria examined, carboxysomes have apparently only been found in one (thermophilic) strain, Pseudomonas thermophila K2 (Kostrikina et al., 1981; Romanova et al., 1982). If carboxysomes do function to protect RuBisCO from inhibition by 0 2 , then variations in carboxysomes abundance and in the subcellular distribution of RuBisCO between the organelles and the cytoplasm may be observed under changing 0 2 tensions. The anthropocentric rationale supposes that the carboxysome-producing prokaryote would respond by forming more of the organelles to contain a greater proportion of the cell’s RuBisCO complement under adverse high O~/lowCOZconditions, further assuming that it possessed the ability to do so. The distribution of RuBisCO protein between the cytoplasm and carboxysomes in continuous cultures of T . neapolitanus was not influenced by varying 0 2 tensions, providing no support for the 0 2 protection hypothesis (Beudeker et al., 198I). The potential contribution of an O2 protection mechanism for RuBisCO of S. leopoliensis has been estimated by Turpin et al. (1984) at various ratios of internal to external dissolved inorganic carbon (DIC) ratios. The protection of RuBisCO from 0 2 may increase net photosynthesis by 100% and 18% at cyanobacterial DIC ratios of 100 and 1000 respectively. These observations leave unanswered the question

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G.A. CODD

of whether the carboxysomes confer 0 2 protection to RuBisCO. However, with a major increase in carboxysome numbers in S. leopoliensis in response to carbon limitation (Table 7), an active role of carboxysomes in cyanobacterial photosynthesis appears to be likely. Other agents that inhibit higher plant RuBisCO enzymes include sulphite, sulphate and hydrogen fluoride gas at concentrations present in industriallypolluted environments (Khan and Malhotra, 1982; Parry and Whittingham, 1983; Buckenham et al., 1982). The effect of these compounds on microbial RuBisCO enzymes has not received attention. However, the success of Calvin cycle prokaryotes in sulphur-containing environments (van Gemerden and de Wit, 1986)merits study of the effects of toxic sulphur compounds on microbial RuBisCO enzymes. The RuBisCO enzymes from freshwater and halophilic cyanobacteria are inhibited by NaCl in vitro (Cook, 1980; Incharoensakdi et al., 1986). Although intracellular osmotica, particularly glycine betaine, relieve the inhibition by NaCl, the relative susceptibilities of the carboxysomal RuBisCO pool compared to the cytoplasmic enzyme to in uitro and in uiuo treatments has not been determined. The possibility that carboxysomes may confer protection on RuBisCO in uiuo against adverse environmental parameters is under current study in this laboratory.

c. ARE CARBOXYSOMES STORAGE BODIES? The possibility that carboxysomes may serve as deposits for the storage of RuBisCO was proposed by Shively (1974) and Stewart and Codd (1975). Several observations support, but do not prove, this possibility. Carboxysome numbers per cell increase in old stationary phase cultures of Nitrobacter winogradskyi and decrease during reactivation of the cells by the supply of fresh nutrients (Bock and Heinrich, 1971). In nutritionally versatile cyanobacteria, RuBisCO and carboxysomes continue to be produced during chemoheterotrophic growth, although the enzyme is not required catalytically under these conditions. Levels of RuBisCO protein in photo-autotrophic and chemoheterotrophic Nostoc 6720 cultures are similar. In nitrate-grown chemoheterotrophic mid-exponential phase cells, most of the enzyme is present in the carboxysomes; in mid log-phase photo-autotrophic cells most of the enzyme is cytoplasmic (Leadbeater, 1981; Codd and Marsden, 1984). Most of the RuBisCO in this versatile autotroph is thus sequestered in the carboxysomes when it is not active, or required, for growth and a storage function for the organelles under these conditions can be inferred. However, in other versatile autotrophs, neither carboxysomes nor RuBisCO are produced during chemoheterotrophic growth, e.g. Thiobacillus intermedius (Purohit et al., 1976). Chemostat cultures have also been used to address the storage hypothesis.

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Carboxysomal RuBisCO protein levels were minimal during the growth of T.neapolitanus under nitrogen limitation, consistent with this hypothesis (Beudeker et al., 1981). In addition, a rapid doubling of carboxysomal RuBisCO protein levels was obtained following the addition of ammonia to nitrogen-limited cultures. However, the transfer of nitrogen-limited steadystate cultures to nitrogen starvation conditions did not result in a breakdown ofcarboxysomal or cytoplasmic RuBisCO, suggesting that the T. neapolitanus carboxysomes do not act as general protein reserves (Beudeker et al., 1981). The lack of effect of growth of S. leopoliensis under nitrogen (nitrate) limitation (Table 7) does not support the storage hypothesis in this nutritionally specialist cyanobacterium (Turpin et a/., 1984). The likelihood that carboxysomes serve as general protein reserves in cyanobacteria should be viewed against the presence of two other major nitrogen reserves in these organisms: the phycobilisomes, which serve as protein reserves in addition to accommodating the accessory pigments for photosynthesis, and the cyanophycin granules (Stanier and Cohen-Bazire, 1977). The cyanophycin polypeptide of the latter bodies consists of a copolymer of equimolar quantities of arginine and aspartic acid. Cyanophycin is not synthesized on ribosomes and presents simpler requirements as a general nitrogen reserve than the transcription and translation of RuBisCO genes and the assembly of their products into the 8L8S enzyme for its sequestration into membrane-bound carboxysomes. This argument does not militate against a role for carboxysomes in the specific storage of the RuBisCO enzyme under specific conditions.

VI. Further Aspects of Carboxysomes A. ECOLOGICAL MARKERS FOR AUTOTROPHY

Since carboxysomes only occur in prokaryotes that contain RuBisCO and are capable of assimilating CO2 via the Calvin cycle, then their easily recognized appearance may be useful as a structural marker for autotrophy in microbial ecology (Codd and Marsden, 1984). Unfortunately for this purpose, carboxysomes are not a feature of all Calvin cycle autotrophs although they are confined to autotrophs. The remarkable discovery of the dense growths of microbes and animals around deep-sea hydrothermal vents has been among the most exciting microbial ecophysiological advances of recent years (see Jannasch and Mottl, 1985). Primary production in these totally dark environments is performed by chemolitho-autotrophic bacteria, including free-living species of the sulphur-oxidizers Thiobacillus and Thiomicrospira and a range of symbiotic bacteria, living in vestimentiferan and pogarophoran

156

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CODD

worms and in molluscs (Cavanaugh, 1983; 1985). Several of these bacterial symbionts contain Calvin cycle enzymes but apparently no carboxysomes. However, other bacterial symbionts present in the gills of clams from sulphide vent environments have not been characterized enzymically, but they contain polyhedral electron-dense bodies which may be carboxysomes (Giere, 1985). Until recently, the marine non-heterocystous filamentous Nz-fixing cyanobacterium Oscillatoria (Trichodesmium) erythraea could not be grown in laboratory culture. This has been a constraint on research into the mechanism of N2-fixation in this major bloom-forming species and in particular on how the organism may enable the physiologically-incompatible processes of N2 fixation and oxygenic photosynthesis to occur in the same filament. A clear morphological differentiation at the subcellular level may, however, be seen along the filaments. Carboxysomes, indicating the presence of RuBisCO and photosynthetic capacity, are abundant in the terminal cells a t each end of the filaments and numbers of the organelles decrease toward the central carboxysome-free cells in the central region (Bryceson and Fay, 1981). The central cells are thought, from the absence of carboxysomes, not to perform photosynthesis. If these cells lack Photosystem I1 in addition to carboxysomes and RuBisCO, then the central cells may enable the 02-sensitive nitrogenase, if present, to function in an environment of reduced 0 2 tension, as in heterocystous cyanobacteria. B. MAN-MADE RIBULOSE 1 ,j-BISPHOSPHATE CARBOXYLASE/OXYGENASE INCLUSION BODIES?

The production of large amounts of microbial RuBisCO in foreign recombinant heterotrophic bacteria is now feasible (see Section 1V.D). For example, photosynthetic bacterial and cyanobacterial RuBisCO protein, when expressed in the presence of the gratuitous inducer isopropyl P-D-thiogalactopyranoside by E. coli, may account for up to 3-15% ofcell protein (Somerville and Somerville, 1984; Viale et al., 1985; Gatenby et al., 1985; Tabita and Small, 1985). It is likely that considerably higher levels of RuBisCO will be made by recombinant DNA techniques. At least 20 eukaryotic polypeptides, expressed in E. coli as fusion proteins or directly, accumulate as inclusion bodies (see Harris, 1983; Marston, 1986). Examples include insulin A and B chains and calf prochymosin. When produced in the original eukaryotic cells, these proteins are in soluble form. The man-made inclusion bodies are amorphous aggregates and are not surrounded by, or are in close contact with, membranes (Schoemaker et al., 1985; Schoener et al., 1985). Examples of normal E. coli proteins expressed to high levels using recombinant DNA methods that accumulate as inclusion bodies are also known (Cheng, 1983). The bacterial 8L8S RuBisCO is one of

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1 57

themost complex proteins to have been cloned and expressed in foreign hosts. Results so far indicate that most of the cloned RuBisCO is soluble; it will be of interest and value for comparison with authentic carboxysomes to see if inclusion bodies consisting of RuBisCO are produced in recombinant hosts. REFERENCES

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Archaebacteria: The Comparative Enzymology of Their Central Metabolic Pathways MICHAEL J . DANSON Department of Biochemistry. University of Bath. Clauerton Down. Bath BA2 7 A Y . England

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MICHAEL J . DANSON

I. Introduction A. ARCHAEBACTERIA

I . The Archaebacteria The majority of microbiologists, and many biochemists and molecular biologists, will be familiar with the concept of the archaebacteria. To such readers, recognition of the archaebacteria as a phylogenetically distinct group of organisms, that are as distinct from the eubacteria as they are from the eukaryotes, needs neither explanation nor justification. However, to those unfamiliar with the idea that there are at least three major groups (Kingdoms) of living organisms, a brief discussion of the archaebacteria might be helpful and may set in context the value of the comparative enzymology with which this review is primarily concerned. This discussion is not intended to provide a comprehensive review of the archaebacteria; for such information the reader is directed towards a number of excellent articles and books recently published (Fewson, 1986; Woese and Olsen, 1986; Kandler and Zillig, 1986; Woese and Wolfe, 1985; Kandler, 1982; Woese, 1981, 1987). The long-held view that all organisms can be divided into two Kingdoms, the prokaryotes and the eukaryotes, has been seriously questioned in recent years. Carle Woese and George Fox have demonstrated that 16s and 18s ribosomal RNA (rRNA) molecules are excellent molecular chronometers for measuring phylogenetic relationships among organisms: these rRNA species are constant in function, universal in distribution, easily isolated, are relatively large molecules and have both moderately and highly conserved parts of their sequence across large phylogenetic distances (Woese, 1985, 1987). Therefore, it is argued that rRNA sequence comparisons can accurately measure both small and large phylogenetic distances. The method of comparison is that of oligonucleotide cataloguing (Sanger et al., 1965; Uchida et al., 1974; Woese et al., 1976) whereby 16S/18S rRNA is digested with ribonuclease T1 and the resulting oligonucleotides are separated by twodimensional paper electrophoresis and sequenced. The catalogue so generated is characteristic of an organism, and can be quantitatively compared with that of any other organism by means of a binary association coefficient which takes into account oligonucleotide sequences of six bases or longer (Fox et al., 1977). The coefficient measures the fraction of bases in any two catalogues found in oligonucleotides common to the two catalogues, and is related to the actual number of nucleotide differences between the rRNA sequences, but in an unknown and non-linear way. Phylogenetic analysis by oligonucleotide cataloguing (Woese and Fox, 1977; Fox et al., 1980; Woese, 1981) has led to the proposal that there are at

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least three Kingdoms of living organisms: (a) the Eubacteria (the true bacteria); (b) the Eukaryotes; and (c) the Archaebacteria. Although eubacteria and archaebacteria may represent primary evolutionary lineages, the eukaryotic cell is now thought to be a genetic chimera; that is, the cytoplasm and organelles may be of different descents, evidence having been presented for the eubacterial origin of chloroplasts and mitochondria (Gray, 1982; Gray and Doolittle, 1982; Spencer et al., 1984). Archaebacteria encompass three basic phenotypes, namely methanogenic, halophilic and sulphur-dependent (reviewed by Woese and Olsen, 1986; Fewson, 1986; Woese, 1987). The methanogens are obligate anaerobes that reduce carbon dioxide to methane. Methanogenesis is obligatory and no secondary energy sources have been identified (Balch et al., 1979; Whitman, 1985). Halophiles need high concentrations of sodium chloride with some members growing in saturated (5.2 M) salt solutions and others having the additional requirement of high pH optima (pH 9-10) (Kushner, 1985). Sulphur-dependent archaebacteria (Stetter and Zillig, 1985) are found in thermophilic environments (55-lOO0C)and can either reduce or oxidize sulphur to produce energy. Aerobic and anaerobic species are known, so too are autotrophs and heterotrophs. Many sulphur-dependent archaebacteria are thermoacidophiles, growing at acidities as low as pH 1.0 and, for this reason, the thermoacidophilic archaebacterium Thermoplasma acidophilum is often grouped phenotypically with these organisms even though it is an obligate heterotroph with no dependence on sulphur. These extreme conditions in which archaebacteria are found impose phenotypes apparently well suited to the type of environment thought to exist during early life on earth, 3-4- lo9 years ago. From such observations the name archaebacteria was tentatively suggested (Woese and Fox, 1977), although the question of their “primitive nature” is still a matter of debate.

2. Archaebacterial Phylogeny The concept of the archaebacteria as a phylogenetically distinct group, originally formulated from the rRNA oligonucleotide catalogues, has been strengthened by analysis of the complete sequences. The 16S/18S rRNA sequences from various methanogenic, halophilic, thermoacidophilic and sulphur-dependent archaebacteria and from a number of eubacteria and eukaryotes have been determined (referenced in Woese and Olsen, 1986) and the data support a clear distinction between the archaebacterial kingdom and those of the eukaryotes and the eubacteria (Fig. I). Similarly, the complete sequence data are consistent with the archaebacteria comprising two main divisions (Fig. 2); namely, ( I ) the thermophilic sulphur-dependent archaebac-

ARC HAEBACTERIA sulldobw solfatwicus

m e r w o t w s tmax 0.1

Hdobacterium volcmii Mathmspiriiium hunqalai &thanobacterium formicicum Mathamcoccus vannieilii ?kmococcus cder

EUBACTERIA

EUKARYOTES

FIG. 1. Unrooted phylogenetic tree constructed from 16s (or 18s) rRNA sequences. Complete rRNA sequences were aligned and estimates of sequence divergence (mutations fixed per sequence position) were calculated and used by Woese and Olsen (1986) to infer the phylogenetic tree. The scale bar corresponds to a tree branch length of 0.1 mutations fixed per sequence position. Reproduced with the permission of Woese and Olsen (1986).

0.01

Methanococcus vannielii

-

Methanobacterium formicicum Thermoplasme acidophilum Methanospirillum hungatei Halococcus morrhuae Halobacterium cutirubrum lhrmococcus celer

Sulfoobus solfataricus Thermproteus tenax

FIG. 2. Unrooted phylogenetic tree for the archaebacteria based on 16s rRNA sequences. The tree was constructed by Woese and Olsen (1986) from alignment of complete rRNA sequences as described in the legend to Fig. 1. The scale bar corresponds to a tree branch length of 0.01 mutations fixed per sequence position. Reproduced with the permission of Woese and Olsen (1986).

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teria, but not including Thermococcus celer and Tp. acidophilum; and (2) the methanogens and extreme halophiles, plus Tc. celer and Tp. acidophilum. This conclusion has recently been supported by measurements of total rRNA hybridization homologies (Klenk et al., 1986). The rRNAs of 17 species of archaebacteria were hybridized to corresponding and noncorresponding nitrocellulose-bound DNAs. Hybridization homologies were calculated from hybridization yields, corrected for different genome lengths and numbers of rRNA operons per genome, and from them a phylogenetic tree was constructed (Fig. 3). This tree resembles that obtained by comparison of the total sequences of 16s rRNAs (Fig. 2), except in a few details of the precise branching orders. Most significantly, it suggests that Thermococcales represent a third branch of the archaebacterial kingdom beside the branch of the methanogens with halophiles and that of the Sulfolobales with Thermoproteales. The thermophilic phenotype appears in all three of the major branches of archaebacteria, and Woese and Olsen (1986) have therefore raised the possibility of this being the ancestral archaebacterial phenotype. Criteria other than the rRNA sequences have also been used to deduce phylogenetic relationships. These include 5 s rRNA secondary structural features (Fox et al., 1982; Wolters and Erdmann, 1986), the structure of DNAdependent RNA polymerases (Zillig et al., 1985) and ribosome morphology THERMOCOCCALES Cdduplex woesei METHANOMICROBIALES THERMOPLASMALES

Methonolobus findorius

Thermaplosmo ocidophilum

SULFOLOBALES

METHANOCOCCALES Methonococcus vannielii

hfethanofhermus fervidus METHANOBACTERIALES Mefhombacleriumfhermooufotrophicum

fenax THERMOPROTEALES Dewlfurococcus mucows

FIG. 3. Phylogenetic tree of the archaebacteria based on hybridization homologies. The tree was constructed by Klenk et al. (1 986) from hybridization homologies between rRNAs and nitrocellulose-bound DNAs (see text for details). The organisms on both sides of the tree have the same mean distance to the vortex at the centre of the tree. Reproduced with the permission of Klenk et al. (1986).

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MICHAEL J . DANSON

(Lake et al., 1984, 1985). In all three cases, a specific relationship between the sulphur-dependent archaebacteria and the eukaryotes is inferred, with the eubacteria arising from the halophilic-methanogenic branch. Moreover, Lake et al. (1984, 1985) and Lake (1986) propose that sulphur-dependent archaebacteria form a kingdom called “eocytes”, that halophilic archaebacteria and the eubacteria form another called the “photocytes” and that the term “archaebacteria” is reserved to cover only the methanogens. These other proposed branching orders have been critically reviewed by Woese and Olsen (1986) and by Woese (1987). Whatever one’s position, however, it is clear that archaebacterial phylogeny is still a matter of contention, and it is a priority to make further comparable measurements of sequence homologies.

3. Biochemistry of the Archaebacteria Many biochemical features of the archaebacteria serve to reinforce their distinct phylogenetic position (reviewed in Fewson, 1986) and a number of these are outlined in this section. In contrast to the straight-chain fatty acyl ester-linked glycerolipids (with sn-1,2-glycerol) of eubacterial and eukaryotic membranes, archaebacterial lipids are isopranyl ether-linked glycerolipids with an sn-2,3-glycerol configuration (Langworthy, 1985). The thermoacidophilic and some methanogenic glycerolipids contain tetra-ethers, allowing formation of lipid “monolayer” membranes, with the additional feature in thermoacidophiles that the biphytanyl chains may contain from one to four cyclopentyl rings (Langworthy, 1985). In addition, the archaebacterial cell-envelope is distinctive in its lack of the characteristic eubacterial murein (Kandler and Konig, 1985).In its place are found residues of N-acetylated glucosamine or galactosamine, Llysine, L-glutamate, L-alanine or L-threonine and N-acetyl-L-talosaminuronic acid. Finally, the base modifications of transfer RNA (tRNA) are characteristic of archaebacterial species (Gupta, 1985). Interestingly, although the above features are unique to archaebacteria, certain others may be typically eubacterial or eukaryotic. Thus, on the eubacterial side, archaebacteria are distinctly prokaryotic in organization and morphology. Also, the rRNA components of the archaebacterial ribosome have a number of eubacterial features (Matheson, 1985); the 70s ribosome contains one molecule of 5S, 16s and 23s rRNA (Viscentin et al., 1972) and the organization of these rRNA genes is similar to that in eubacteria (Hofman et al., 1979). Furthermore, a putative Shine-Dalgarno sequence has been found in a halophilic 16s rRNA (Kagramanova et al., 1982) and its secondary structure more closely resembles that of eubacterial 16s rRNA than that of the eukaryotic 18s molecule (Kagramanova et al., 1982; Gupta et al., 1983).

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Further morphological aspects of the ribosome are eubacterial in nature although, at the level of the individual ribosomal components, the structural features are significantly closer to eukaryotic molecules than to those of the eubacteria (Matheson, 1985). However, as indicated previously, Lake et al. (1984, 1985) consider the morphology of the archaebacterial ribosome to be distinctive and have used it to deduce phylogenetic relationships. Archaebacteria also have many typically eukaryotic characteristics. (a) Introns have been found in the genes for tRNAL""and tRNASerof Suvolobus solfataricus (Kaine et al., 1983), for tRNA'F of Halobacterium volcanii (Daniels et al., 1985) and for 23s rRNA of Desulfurococcus mobilis (Kjems and Garrett, 1985). (b) Basic histone-like proteins associated with DNA have been found in thermoacidophilic (Searcy and Stein, 1980; Green et al., 1983) and methanogenic (Thomm et al., 1982) archaebacteria. Partial sequence analysis of the HTa protein from Tp. acidophilum shows it is more homologous to eukaryotic nuclear DNA-binding proteins than it is to counterparts from eubacteria (Searcy and DeLange, 1980). (c) Some mRNA molecules have long polyadenylated sequences at the 3'-termini, similar to those in eukaryotes (Ohba and Oshima, 1983; Oshima el al., 1984) and initiation of protein synthesis from mRNA seems to occur in archaebacteria with methionyl-tRNA as in eukaryotes rather than with N-formylmethionyltRNA as in eubacteria (Gupta, 1985). (d) All archaebacteria possess an elongation factor (EF-2) which is ADP-ribosylated by diphtheria toxin, as are eukaryotic EF-2s (Klink, 1985). (e) The transcriptional apparatus of archaebacteria shows a uniform and "eukaryotic-type" antibiotic sensitivity although that of the translational system is heterogeneous (Bock and Kandler, 1985).

4 . Enzymology of the Archaebacteria It is clear that, for the purposes of establishing the phylogenetic status of the archaebacteria, much emphasis has been placed on the molecular biology of these organisms and on the chemical nature of their cell walls and membranes. However, it is also becoming clear that the pathways of metabolism in archaebacteria and their constituent enzymes are equally fruitful areas for investigation. It is the purpose of this review to concentrate on the enzymology of archaebacteria, not in isolation but in comparison with that of eubacteria and eukaryotes. Therefore, the enzymes of the central metabolic pathways have been chosen for review because, as explained in the following section, not only are these pathways thought to be some of the first cellularly established metabolic routes but also because they are the most studied and well-characterized systems in non-archaebacterial species.

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MICHAEL J . DANSON

B. CENTRAL METABOLISM OF EUBACTERIA A N D EUKARYOTES

It has been argued that the universal occurrence of sugars and the use of their derivatives for biosynthesis might suggest that development of a sugar-based biochemistry was an early evolutionary innovation (Gest and Schopf, 1983). Indeed, D-glucose is now the most abundant compound in the biosphere and its catabolism may have been one of the first energy-conversion processes to have been successfully exploited by living cells. Such reasoning has led to an investigation of the pathways of hexose metabolism in archaebacteria but, before this work is described, it would seem pertinent to discuss briefly those catabolic routes in eubacteria and eukaryotes. I . Pathways of Glucose to Pyruvate

The pathways of sugar catabolism in eubacteria have been comprehensively reviewed by Payton and Haddock (1985) and by Cooper (1986) and are summarized, with those of eukaryotic organisms, in Fig. 4. The following summary indicates the salient features of these pathways from which a comparison with those in archaebacteria can best be appreciated. The Embden-Meyerhof glycolytic pathway is characteristic of eukaryotic cells and a large number of anaerobic and facultatively anaerobic eubacteria. In glycolysis, glucose is phosphorylated to fructose 1,6-bisphosphate which in turn undergoes aldol cleavage to two interconvertible three-carbon sugars, dihydroxyacetone phosphate and glyceraldehyde 3-phosphate. The two trioses enter a common route of catabolism to pyruvate (Fig. 4). A key enzyme of the glycolytic sequence, 6-phosphofructokinase, is absent from many strictly aerobic eubacteria, implying that this pathway is inoperative in these organisms. Instead, glucose can be catabolized via the Entner-Doudoroff pathway (Entner and Doudoroff, 1952); glucose 6-phosphate is oxidized to 6-phosphogluconate before being dehydrated to 2-keto-3-deoxy-6-phosphogluconateand undergoing subsequent aldol cleavage to yield glyceraldehyde 3-phosphate and pyruvate (Fig. 4). Glyceraldehyde 3-phosphate is metabolized to give a second molecule of pyruvate via the same sequence of reactions as in the glycolytic sequence. Consequently, only one molecule of ATP is generated for each molecule of glucose fermented, as opposed to two ATP molecules via glycolysis. There is a third route for the catabolism of glucose (Fig. 4), namely the hexose-monophosphate pathway (pentose-phosphate pathway; see Racker, 1948, 1957). It is probable that the reactions of this pathway do not operate generally as a cycle for glucose oxidation, but serve to provide the cell with NADPH and pentose and tetrose sugars. Further oxidation of these sugars to glyceraldehyde 3-phosphate is possible, the metabolic fate of which is the same as in the Embden-Meyerhof and Entner-Duodoroff pathways.

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FIG. 4. Pathways of glucose catabolismin eubacteria and eukaryotes.The three major metabolic routes for catabolism of glucose are the Embden-Meyerhof glycolytic sequence (+), the Entner-Doudoroff pathway (+) and the pentose-phosphate pathway (- - +). The sequence of reactions from glyceraldehyde 3-phosphate to pyruvate is common to all three pa€hways.

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MICHAEL J. DANSON

A few species of eubacteria possess the hexose monophosphate pathway reactions coupled to the phosphorolytic cleavage of pentose phosphates or hexose phosphates, as found in the pentose-phosphate phosphoketolase pathway (Heath et al., 1956) and the hexose phosphate-pentose phosphate phosphoketolase pathway (Scardovi and Trovatelli, 1965; de Vries et al., 1967) (reviewed by Cooper, 1986). Again, glyceraldehyde 3-phosphate is produced, and is metabolized to pyruvate as for the other pathways discussed. Finally, mention should be made of the methylglyoxal pathway (Cooper and Anderson, 1970) which has been found in many members of the Enterobacteriaceae and in some clostridia and pseudomonads (Cooper, 1984, 1986). In this pathway, dihydroxyacetone phosphate, which is formed from glyceraldehyde 3-phosphate by the enzyme triose-phosphate isomerase, is converted into methylglyoxal and then into pyruvate, but without generation of ATP by substrate-level phosphorylation. Thus, this pathway is always accompanied by the common pathway for conversion of glyceraldehyde 3-phosphate into pyruvate. In summary, it is this common trunk sequence from triose phosphate to pyruvate,, utilizing phosphorylated intermediates, that can be identified as constituting the true central pathway for sugar catabolism (Cooper, 1986)and it is a priority to determine if this ubiquity in the eubacterial and eukaryotic kingdoms is maintained in archaebacteria. 2. Enzymes of Glucose Catabolism

It is inappropriate in a review on archaebacterial enzymes to discuss in detail enzymes of glucose catabolism in eubacterial and eukaryotic species. However, it is important to note that the enzymes of the Embden-Meyerhof glycolytic pathway, and therefore of the common pathway converting glyceraldehyde 3-phosphate into pyruvate, have been the subject of detailed molecular studies. Complete amino-acid sequences are available for all of the glycolytic enzymes, except for phosphoglucose isomerase, and high-resolution crystal structures have been derived for all but aldolase and enolase (reviewed by Fothergill-Gilmore, 1986). From a consideration and comparison of these structures, Fothergill-Gilmore (1986) concludes that the glycolytic pathway appears to have arisen primarily by random association of enzymes converging on similar structures. A clear exception to this is the divergent evolution of bisphosphoglycerate mutase and monophosphoglycerate mutase, with other possible examples in glyceraldehyde 3-phosphate dehydrogenase and phosphoglycerate kinase. On the other hand, is is thought that gene duplication and divergence have given rise to multiple isoenzymes within glycolysis and that this may have been a crucial means of adapting to the various metabolic needs of different tissues and organisms.

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Clearly, this enormous data-base on glycolytic enzymes provides a firm basis on which to begin a study of archaebacterial enzymes. Indeed, it is probable that a structural investigation of these proteins will make significant contributions to answering questions currently posed concerning evolution of the glycolytic pathway. It is unfortunate that there is not yet a similar level of information for enzymes of the Entner-Doudoroff and hexose monophosphate pathways, that is for the reactions leading to glyceraldehyde 3phosphate; only when we have this information can we begin to speculate on evolutionary relationships of the three main catabolic routes of sugar catabolism.

3. Metabolic Fate of Pyruvate In eubacteria growing under anerobic conditions, pyruvate is metabolized fermentatively, that is it serves as a sink for reducing equivalents generated in its formation from glucose. This contrasts with oxidative metabolism described below, where redox carriers in the electron-transport chain act a: electron sinks, resulting ultimately in reduction of oxygen. Micro-organisms exhibit a diverse array of fermentative reactions from pyruvate, all of which can be classified into six main types (reviewed by Morris, 1985). These are lactic, ethanolic, butyric, mixed acid, propionic and homoacetic fermentations. In eukaryotes and eubacteria growing under aerobic conditions, the most common fate of pyruvate is its oxidative decarboxylation to acetyl-CoA. This reaction is catalysed by the pyruvate dehydrogenase multienzyme complex, the mechanism of which will be described in detail later in this review. Complete oxidation of the acetyl-CoA then proceeds via the citric acid cycle (Fig. 5 ) , yielding reducing equivalents in the form of NADH and FADH2, the reoxidation of which is coupled to synthesis of ATP via an electron-transport chain. In addition, the citric acid cycle also provides key intermediates for a variety of biosyntheses and, consequently, replenishment is required via a number of anaplerotic reactions such as those catalysed by pyruvate carboxylase, phosphoenolpyruvate (PEP) carboxylase and PEP carboxykinase (Kornberg, 1966). A survey of a wide variety of organisms, especially amongst eubacterial species, reveals that the citric acid cycle exibits considerable diversity, both in the portions of the cycle selected for use (Weitzman, 1985) and in the patterns of catalytic activity, regulation and molecular structure of its constituent enzymes (Weitzman and Danson, 1976;Weitzman, 1981). Therefore, in a manner similar to that described for catabolism of glucose to pyruvate, the enzymology of the subsequent oxidative reactions provide a rich source of comparison with counterparts in archaebacteria.

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MICHAEL J . DANSON

Pyruvote

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Citratet CoA

cis -Acan;ote

Fumarate FA0

lsocitrate

1

2-Oxoglutarate N NDPtPi T P q Succinyl-CoA

NAD' + CoA

FIG. 5. The citric acid cycle.

11. Archaebacterial Pathways of Central Metabolism

In order to compare the enzymes in archaebacteria with those of eubacteria and eukaryotes, it is first necessary to establish the pathways of hexose catabolism in the three archaebacterial phenotypes. Although many investigations of the biochemistry of archaebacteria are still in their infancy, one principle is quite clear and this is the fact that one cannot assume that the situation found in eubacteria is necessarily present also in archaebacteria. This is as true for pathways of sugar metabolism as it is for the molecular biological characteristics and for membrane and cell-wall structures. Therefore, this section will be concerned with a review of the central metabolism of archaebacteria, and from the pathways discovered enzymes can then be selected for comparison with their counterparts in other organisms. A. HEXOSE CATABOLISM

1. Halophiles

Many of the extreme halophiles utilize proteins and amino acids rather than carbohydrates as a source of carbon (Larsen, 1981). However, Tomlinson and Hochstein (1972a,b) isolated several carbohydrate-metabolizing strains and

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one of these, initially designated Mg but later classified as Halobacterium saccharovorum (Tomlinson and Hochstein, 1976), has been shown to catabolize glucose and galactose via a modified Entner-Doudoroff pathway (Fig. 6; Tomlinson et al., 1974). Glucose is initially oxidized to gluconate by an NAD+-dependent glucose dehydrogenase and is then converted stoicheiometrically into 2-keto-3-deoxygluconate via gluconate dehydratase. 2-Keto-3deoxygluconate is phosphorylated by ATP and the 2-keto-3-deoxy-6phosphogluconate so generated undergoes aldol cleavage to equimolar amounts of pyruvate and glyceraldehyde 3-phosphate. This sequence of reactions has been termed the “modified” Entner-Doudoroff pathway (Andreesen and Gottschalk, 1969) and differs from the Entner-Doudoroff pathway (see Section 1.B) in that the latter involves the phosphorylation of glucose and therefore operates entirely with phosphorylated metabolites. Enzymes metabolizing the glyceraldehyde 3-phosphate to a second molecule of pyruvate via the common trunk sequence discussed in Section 1.B have been detected in H. saccharovorum (Tomlinson et al., 1974). It is therefore in the initial reactions of glucose catabolism that this halophile is unusual. It should be noted that acetate is produced by H. saccharovorum when grown on glucose (Tomlinson and Hochstein, 1972b, 1976). However, the enzymology of the conversion of pyruvate into acetate has not been reported, except that it presumably procedes via acetyl-CoA production by the pyruvate:ferredoxin oxidoreductase (see Section 1II.B). Two additional points can be made concerning this modified EntnerDoudoroff pathway in H . saccharovorum. First, cell extracts also reduced NAD+ in the presence of glucose although at a rate only 8% of that observed with NADP+. This may be significant as thermoacidophilic archaebacteria possess a glucose dehydrogenase which can utilize both cofactors and, as discussed later, the enzyme has a much higher K,,, value for NAD+ than for NADP+; thus, the low activities observed by Tomlinson et al. (1974) may be a consequence of a dual-specificity glucose dehydrogenase, with a similar ratio of K,,, values for its cofactors. Secondly, Tomlinson et at. (1974) found that pyruvate production from glucose did not have an absolute requirement for ATP. Again, this may be an important factor in that species of SulJblobus and Thermoplasma appear to utilize a non-phosphorylated pathway for glucose catabolism; thus H. saccharovorum may also possess such a metabolic route, perhaps in addition to the modified Entner-Doudoroff pathway already described. Oxidation of glucose 6-phosphate or 6-phosphogluconate by cell extracts of H. saccharovorum was not observed, suggestingthe absence of both the Embden-Meyerhof glycolytic sequence and the conventional EntnerDoudoroff pathway. However, a hexokinase activity was detected although its metabolic function remains unclear. At present, there are few data on pathways of glucose catabolism in other

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FIG. 6. Pathways of glucose catabolism in halophilic and thermoacidophilic archaebacteria.The modified Entner-Doudoroff pathway of halophiles (+) and the non-phosphorylated Entner-Doudoroffpathway of Sulfolobus solfataricus and Thermoplasma acidophilum (- - +) are shown alongside the classical Entner-Doudoroff pathway of eubacteria (-) from Fig. 4. Conversion of glyceraldehyde into pyruvate via glycerate (- - +) has been demonstrated only in Thermoplasma acidophilum (see text for details).

CENTRAL METABOLIC PATHWAYS OF ARCHAEBACTERIA

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species of halophilic archaebacteria. Hochstein (1978) reported the presence of the requisite enzymes for a modified Entner-Doudoroff pathway in a number of halophilic strains (M4, M7, Gt-1, L-1, U-14 and U-18). D'Souza and Altekar (1983) could not detect the presence of 6-phosphofructokinase in Halobacterium halobium, nor could they detect conversion of glucose, fructose, glucose 6-phosphate or fructose 6-phosphate into triose phosphates in the presence of ATP and MgC12. These observations would seem to rule out the presence of an Embden-Meyerhof sequence for catabolism of glucose, but would not argue against an Entner-Doudoroff pathway as NAD(P)+ was not included in the incubation mixtures. The modified Entner-Doudoroff pathway of H . saccharovorum is unusual but is not unique to the archaebacteria; it has been reported in species of Clostridium (Andreesen and Gottschalk, 1969; Bender et al., 1971), Alcaligines, Achromobacter (Kersters and De Ley, 1968) and Rhodopseudomonas spheroides (Szymona and Doudoroff, 1960). However, with the exception of Rh. spheroides, the pathway has been shown to operate in gluconate-grown cells, with no reports on whether glucose can be catabolized. 2. Thermoacidophiles

Interestingly, the thermoacidophilic archaebacteria metabolize glucose through a further modification of the Entner-Doudoroff pathway, in which the phosphorylation step is omitted altogether in formation of the first molecule of pyruvate. This sequence of reactions was elucidated by De Rosa et al. (1984) in Sulfalobus solfataricus, a species that is able to grow on glucose as the sole carbon source. The absence of 6-phosphofructokinase from S. solfataricus and preliminary investigations with differentially labelled ['4C]glucoses(De Rosa et al., 1983) indicated that glucose was not catabolized via the glycolytic pathway. Instead, De Rosa et al. (1984) demonstrated that glucose is converted into gluconate in an NAD(P)+-dependent dehydrogenation step and that this is subsequently dehydrated to 2-keto-3-deoxygluconate. So far this is the same as the modified Entner-Doudoroff pathway found in H . saccharovorum, but, at this point in S. solfataricus, 2-keto-3deoxygluconate is not phosphorylated; rather it is cleaved direct to pyruvate and glyceraldehyde. This non-phosphorylated version of the Entner-Doudoroff pathway is outlined in Fig. 6. Each step has been tested in uitro at 70"C, which is within the growth-temperature range of the organism, and intermediates have been identified by chromatography, spectroscopy, synthetic evidence and/or by enzymic assays. The metabolic fate of the glyceraldehydein S. solfataricus was not defined by De Rosa et al. (1984); conversion into glyceraldehyde 3phosphate was considered to be a possibility as incubation with ATP and with

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MICHAEL J. DANSON

glyceraldehyde resulted in formation of ADP. However, production of glyceraldehyde 3-phosphate could not be detected by enzymic assay. Using radiorespirometric assays of glucose oxidation, Wood et al. (1987) have provided evidence for the in vivo operation of this pathway in Sul$olobus brierleyi and Sulfolobus strain LM (similar to Sulfolobus acidocaldarius). Release of I4CO2 from glucose specifically labelled in carbon atoms 1,2, 3,4 and 6 was measured using suspensions of bacteria previously grown in tetrathionate medium supplemented with glucose. Substantial release of carbon atoms 1 and 4 by both strains was consistent with the nonphosphorylated Entner-Doudoroff pathway, the excess of C-1 release over C-4 indicating that an oxidative pentose-phosphate pathway may also be operative. In S. brierleyi, the very low release of C-2 suggests that this pathway may be non-cyclic, but in strain LM the high release of C-2, coupled with significant C-3 and C-6 release, indicates a conventional oxidative cycle. My colleagues and I have found that the non-phosphorylated version of the Entner-Doudoroff pathway is also operative in Thermoplasma acidophilum (Budgen and Danson, 1986a). Intermediates of this pathway were identified by enzymic analysis or by thin-layer chromatography, and individual enzymes involved were assayed and their kinetic parameters determined. Interestingly, we could find no evidence for conversion of glyceraldehyde into glyceraldehyde 3-phosphate; rather glyceraldehyde is oxidized to glycerate by an NADP+-dependent dehydrogenase and glycerate is then converted into 2phosphoglycerate by glycerate kinase (Fig. 6). Enolase and pyruvate kinase complete the generation of a second molecule of pyruvate. Up to this point in the metabolic scheme, there is no net yield of ATP in Tp. acidophilum nor are there reactions for regeneration of NAD+ and NADP+. Searcy and Whatley (1984) found that cultures of this organism produced and excreted significant amounts of acetic acid when grown on glucose. We have recently shown (N. Budgen and M. J. Danson, unpublished observations) that acetyl-CoA, generated from pyruvate by pyruvate:ferredoxin oxidoreductase (see Section III.B), can be converted into acetate by cell extracts of Tp. acidophilum in the following reaction:

+

+

+

Acetyl-CoA ADP PiaAcetate CoA + ATP We have demonstrated the stoicheiometry of this reaction and can find no evidence either for formation of acetyl-phosphate from acetyl-CoA (via a phosphotransacetylase) or for synthesis of ATP and acetate from acetylphosphate (via an acetate kinase). Thus, the enzyme concerned is thought to be acetyl-CoA synthetase (ADP-forming) (EC 6.2.1.13). This activity was first detected in the human eukaryotic parasite, Entamoeba histolytica, where it is suggested that conversion of acetyl-CoA into acetate is an important energyconserving step (Reeves et al., 1977). In Tp. acidophilum, AMP and

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pyrophosphate cannot substitute for ADP and inorganic phosphate, and the synthetase is thus distinguished from the more commonly distributed AMPutilizing acetyl-CoA synthetase (EC 6.2.1.1) which has been found in both eubacterial and eukaryotic organisms (Londesborough and Webster, 1974) and in the archaebacteria Methanobacterium thermoautotrophicum (Oberlies et al., 1980) and Thermoproteus neutrophilus (Schafer et al., 1986). In both of these archaebacteria, the enzyme is thought to serve an acetate-assimilatory role. Using radiorespirometric analyses similar to those reported by Wood et al. (1987) for species of Sulfolobus, my colleagues and I have provided confirmatory evidence for the operation of the Entner-Doudoroff pathway in Tp.acidophilum (N. Budgen and M. J. Danson, unpublished observations). In addition, the radioactivity recovered in the excreted acetate is consistent with its formation from pyruvate and acetyl-CoA as already suggested. It is probable that some acetyl-CoA will enter the citric acid cycle, generating further energy and more NAD(P)H. The mechanism for regeneration of reduced cofactor(s) has not yet been fully defined. The respiratory chain of Tp. acidophilum appears to consist of only a b-type cytochrome (Hollander, 1978; Searcy et al., 1978; Searcy and Whatley, 1982)and a quinone (Hollander et al., 1977). No proton-translocating adenosine triphosphatase (ATPase) has been identified although a membrane-bound ATPase, proposed to function as a sulphate-exporting translocase, has been found (Searcy and Whatley, 1982; Searcy, 1986). Thus, it has been suggested that energy is produced in Tp. acidophilum by substrate-level phosphorylation (Searcy and Whatley, 1982), formation of acetate providing one such mechanism. In contrast, S. acidocaldarius seems to depend energetically on respiration-coupled phosphorylation (Anemuller et al., 1985) and a plasma-membrane associated ATPase has been identified as a proton-translocating candidate for this oxidative phosphorylation (Wagaki and Oshima, 1986; Lubben and Schafer, 1987). Characterization of the purified enzyme suggests a close relationship with the group of eubacterial and eukaryotic Fo F1-ATPases (Lubben et al. 1987). In my investigations of Tp. acidophilum (Budgen and Danson, 1986a) the activities of the following glycolytic enzymes could not be detected: 6phosphofructokinase, fructose 1,6-bisphosphatase, fructose 1,6-bisphosphate aldolase, glyceraldehyde 3-phosphate dehydrogenase and phosphoglycerate mutase. However, on the basis of respiratory activities, Searcy and Whatley (1984) conclude that glycolysis is operative in addition to the pentose phosphate pathway. At present, the apparently contradictory data on the absence or presence of glycolysis remain unresolved, although the observed respiratory activities on gluconate and glyceraldehyde and conversion of glucose into gluconolactone are consistent with the operation of the nonphosphorylated Entner-Doudoroff pathway.

182

MICHAEL J . DANSON

3 . Methanogens

Most methanogenic archaebacteria are autotrophs, gaining their energy by formation of methane from COz and Hz. Some methanogens are able to' disproportionate a variety of CIcompounds and acetate to methane and carbon dioxide and, in addition, to reduce the carbon dioxide to methane. Some species are restricted to methanol or acetate degradation. The literature on assimilation of carbon in methanogens is extensive and, for comprehensive reviews, the reader is referred to Fuchs and Stupperich (1984, 1986), Kirsop (1984), Whitman (1985), Zeikus et al. (1985) and Jones et al. (1987). What is important to the present discussion is that the carbon is eventually fixed into acetyl-CoA and therefore hexose metabolism is mainly in the direction of carbohydrate synthesis rather than degradation. Thus, the gluconeogenic route is the pathway that has been most intensively studied and this will be described below; it will be seen that a reverse of the Embden-Meyerhof pathway effects the synthesis of glucose from acetyl-CoA. However, there will be some carbohydrate turnover, albeit small, and using I3C-NMR spectroscopy Evans et al. (1985, 1986) have shown that, even though glucose is not an energy or carbon source for Methanobacterium thermoautotrophicurn, this organism will take up and catabolize this carbohydrate. [ I-13C]Glucoseand [6-13C]glucosewere individually fed to growing cells and both led to formation of triose phosphates exclusively labelled in the C-3 position. It is argued that this provides strong evidence for catabolism of glucose via the Embden-Meyerhof pathway, the reverse of which is the gluconeogenic route in this organism. However, two observations must be taken into account. First, as will be discussed later, although enzymes for gluconeogenesis have been found in M. thermoautotrophicum, there is no report of the presence of 6-phosphofructokinase, a key enzyme of the Embden-Meyerhof pathway. This activity must be present for glucose to be catabolized as proposed by Evans et al. (1985,1986). Secondly, Fuchs et al. (1983) claim that glucose synthesis is unidirectional, the activity of fructose 1,6-bisphosphate aldolase being detectable only in the direction of aldol condensation. A low level of the reverse reaction may have been found although there were enzymic activities in cell extracts which interfered with estimation of triose phosphate formation. Confirmation of the pathway of glucose degradation in this methanogen therefore awaits analysis of the individual steps at the enzymic level. Such an investigation is all the more important given the apparent absence of the Embden-Meyerhof pathway from halophilic and thermoacidophilic archaebacteria.

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183

B. GLUCONEOGENESIS

As noted previously, halophilic archaebacteria can use amino acids and proteins as carbon sources. Similarly, species of Sulfolobus and Thermoplasma can grow heterotrophically on yeast extracts (Brock, 1978). Species of Sulfolobus can also grow autotrophically (Wood et al., 1987 and references therein) as can other thermophilic archaebacteria and the methanogens. Therefore, all of the archaebacteria will possess the capacity to carry out gluconeogenesis from C3 or Cq compounds. The pathways of gluconeogenesis have not been established in species of Sulfolobus or Thermoplasma. There are no reports of the absence or presence of necessary enzymes in any species of Sulfolobus, whereas, in Thermoplasma acidophilum, fructose 1,6-bisphosphatase, fructose 1,6-bisphosphate aldolase, glyceraldehyde 3-phosphate dehydrogenase and phosphoglycerate mutase were searched for but were not detected (Budgen and Danson, 1986a). It is possible that a reversal of the non-phosphorylated Entner-Doudoroff pathway could occur in these organisms, but such a suggestion awaits proper investigation. Interestingly, we have found high activities of triose phosphate isomerase in both S. acidocaldarius and Tp. acidophilum. The function of this enzyme is at present unclear in that it appears to be metabolically “isolated” unless, in the case of Sulfolobus, glyceraldehydecan indeed be phosphorylated to glyceraldehyde 3-phosphate (De Rosa et al., 1984). In contrast to the situation in thermoacidophiles, the routes effecting synthesis of glucose in halophilic and methanogenic archaebacteria have received considerable attention. It appears that gluconeogenesis proceeds via a reversal of the Embden-Meyerhof glycolytic pathway in organisms of both of these phenotypes, despite the possibility discussed previously that they use different routes for glucose catabolism. D’Souza and Altekar (1983) reported formation of pyruvate from glyceraldehyde 3-phosphate and the production of triose phosphates from phosphoenolpyruvate, 2-phosphoglycerate and 3-phosphoglycerate in unfractionated cell extracts of Halobacterium halobium grown on a medium containing salts, peptone and citrate. In these extracts, the enzymes fructose 1,dbisphosphate aldolase, fructose 1,6-bisphosphatase and phosphoglucose isomerase were detected, thereby indicating the potential for functional reversal of glycolysis in H. halobium. Further work on these enzymes (D’Souza and Altekar, 1982; Dhar and Altekar, 1986a,b) has demonstrated that both class I (Schiff baseforming) and class I1 (metal-requiring) fructose 1,6-bisphosphate aldolases can be found among the halophiles although only one type is found in any one species. In non-archaebacterial species, class I aldolases are usually found in higher eukaryotes whereas the class I1 enzyme is characteristic of fungi and eubacteria (reviewed in Morse and Horecker, 1968; Horecker et al., 1972).

184

MICHAEL J. DANSON

However, class I aldolases have also been found in Peptococcus aerogenes (Lebherz and Rutter, 1973), Lactobacillus casei (Kaklij and Nadkarni, 1974), Escherichia coli (Stribling and Perham, 1973), staphylococci (Gotz et d., 1979) and mycobacteria (Bai et al., 1975). It will be interesting to compare the halophilic class I aldolases with their eukaryotic and eubacterial counterparts, particularly as this same enzyme type has been found in another archaebacterium, namely M. thermoautotrophicum (Fuchs et al., 1984). It has been noted previously that in M. thermoautotrophicum the synthesis of glucose from acetyl-CoA is essentially via a reversal of the EmbdenMeyerhof glycolytic pathway. Autotrophic growth involves synthesis of acetyl-CoA from two molecules of C 0 2 and reductive carboxylation then leads to production of pyruvate. Pulse-labelling with I4CO2and ['4C]pyruvate long-term labelling studies (Jansen et al., 1982) suggested that glucose synthesis proceeds via 3-phosphoglycerate and head-to-head aldol condensation of two triose phosphate molecules to form fructose 1,6-bisphosphate. Enzymic studies indicated direct synthesis of phosphoenolpyruvate from pyruvate and ATP (Eyzaguirre et al., 1982):

+

Pyruvate + ATP H20=Phosphoenolpyruvate+ AMP + Pi and subsequent enzymes of the Embden-Meyerhof pathway have been detected in cell extracts with specific activities sufficiently high to account for the in vivo rate of carbohydrate synthesis (Jansen et al., 1982; Fuchs et al., 1983). I3C-NMR spectroscopy (Evans et al., 1985, 1986) has confirmed this gluconeogenic sequence and in vivo 3'P-NMR spectroscopic studies (Kanodia and Roberts, 1983; Seely and Fahrney, 1983) identified a unique metabolite, namely 2,3-cyclopyrophosphoglycerate,in M. thermoautotrophicum (Fig. 7). This cyclic pyrophosphate occurs in high intracellular concentrations (up to 20 mM)in cells grown in the presence of excess phosphate. Its function remains chase experiments (Evans et af.,1986) to be established. although '3C02/12C02

0

/CH2

-

CH

\

0

FIG. 7. Structure of 2,3-~yclopyrophosphoglyceratefound in Methanobacterium thermoautotrophicum.

CENTRAL METABOLIC PATHWAYS OF ARCHAEBACTERIA

185

suggest that it is an important gluconeogenic metabolite whose turnover is directly linked to carbohydrate synthesis. The precise mechanism of synthesis is also still undefined; it is formed after phosphoenolpyruvate but is succeeded by 2,3-bisphosphoglycerate.Studies on gluconeogenesis in other methanogenic archaebacteria have not been reported. C . GLYCEROL SYNTHESIS

All archaebacteria have lipids based on ether linkages, and all such glycerol ethers contain an sn-2,3-glycerol configuration which is in direct contrast to the sn-1,2-glycerolin ester-linked glycerolipids of eubacteria and eukaryotes. Synthesisof archaebacterial glycerolipids is under active investigation and has been reviewed by Langworthy (1985) and De Rosa et al. (1986). Having discussed routes of glucose catabolism and anabolism in these organisms, my concern in this short section is to look at the supply of glycerol from these pathways. Kates and his coworkers have investigated the origin of the glycerol residues in lipids of the halophilic archaebacteria (Kates et al., 1970; Kates and Kushwaha, 1978). Halobacterium cutirubrum was grown on variously labelled [''C]- and [3H]glycerols;retention of 3H in C-I and C-3 was observed, but 3H from C-2 was almost completely lost. Although the authors regarded this loss as a significant feature of the etherification step, De Rosa et al. (1986) interpret it as possibly being due to interconversion of glycerol (or glycerol phosphate) and dihydroxyacetone (or dihydroxyacetone phosphate). Retention of 3H at C-1 and C-3 further suggests that, when the organism is grown on glycerol, there is no aldo-keto isomerization between dihydroxyacetone and glyceraldeyde. However, this may not be so when it is grown with carbohydrate as the sole carbon source. It has already been noted that glyceraldehyde 3-phosphate is produced from glucose and thus production of glycerol via dihydroxyacetone phosphate is a distinct possibility. Glycerol phosphate dehydrogenase activity has been detected in cell extracts of H . cutirubrum (Wassef et al., 1970), although this would produce sn-glycerol 3phosphate (and not the required sn-glycerol I-phosphate) as its stereochemical specificity is the same as that of the eubacterial and eukaryotic enzymes. Glycerol could be produced from glycerol phosphate or from reduction of dihydroxyacetone via glycerol dehydrogenase which has also been found in this organism (Baxter and Gibbons, 1954). If glycerol is itself alkylated then the configuration of the chiral centre in the glycerol residue of the lipid could be determined by stereospecificity of the alkylation step. De Rosa et al. (1982, 1986) have shown glycerol to be incorporated into ether lipids of S. acidocaldarius without change in the hydrocarbon skeleton. They therefore suggest that the ether-forming step in biosynthesis of these

186

MICHAEL J . DANSON

lipids occurs without any loss of hydrogen from any of the glycerol carbon atoms, which in turn could be directly alkylated by geranylgeranylpyrophosphate. We have suggested (Budgen and Danson, 1986b) that glycerol may be provided by the activity of a glycerol: NADP+ oxidoreductase (EC 1.1.1.72) which we have found in both S. acidocaldarius and Tp. acidophilum. This enzyme catalyses reduction of glyceraldehyde (formed by the non-phosphorylated Entner-Doudoroff pathway) according to: DL-Glyceraldehyde + NADPH

+ H +=>NADP++Glycerol

In cell extracts of these two thermoacidophiles we have not been able to detect sn-glycerol3-phosphate dehydrogenase, glycerol kinase or DL-glyceraldehyde kinase, suggesting a non-phosphorylated glycerol as the glycerolipid precursor. Nothing is known of the origin of the glycerol residue of methanogenic lipids although the presence in the Embden-Meyerhof glycolytic sequence of dihydroxyacetone-phosphate and glyceraldehyde 3-phosphate might suggest that these are the precursors as in eubacterial and eukaryotic organisms. D. THE CITRIC ACID CYCLE All archaebacteria can convert pyruvate into acetyl-CoA. The enzyme involved is pyruvate oxidoreductase, the structure and catalytic mechanism of which are described and compared with those of the eubacterial and eukaryotic pyruvate dehydrogenase multienzyme complexes in Section 1II.B. In non-archaebacterial species, acetyl-CoA can enter either the pathway of fatty-acid biosynthesis or the citric acid cycle. The question then is, do the archaebacteria possess this cycle and if so, is it complete and do they use it in an oxidative or reductive capacity? As with glucose catabolism to pyruvate, we shall see that each of the three archaebacterial phenotypes possesses its own distinctive variation of this pathway, which in turn is adapted to the environmentally determined growth conditions. 1. Halophiles

It has already been noted that halophilic archaebacteria are chemoorganotrophic organisms that can fulfil their energy requirements by metabolism of amino acids and other nitrogenous compounds. Some strains can utilize carbohydrates but they are not a requirement and therefore it is probable that halophiles will possess an oxidative citric acid cycle. Aitkin and Brown (1969) reported the presence of enzymes of this and the glyoxylate cycle in H. halobium, and my colleagues and I have found the key enzymes citrate synthase and succinate thiokinase in a range of both classical and alkaliphilic

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halophiles (Danson et al., 1985). Not all citric acid-cycle enzymes have been searched for in all of these organisms, but the nutrients able to support growth are all consistent with halophiles possessing an oxidative aerobic pathway. 2. Thermophiles The situation in the thermoacidophilic and thermophilic archaebacteria is more complex and may depend on whether growth is heterotrophic or autotrophic, aerobic or anaerobic. Indeed, the species diversity within this phenotype is probably a reflection of their phylogenetic differences, as discussed in Section I.A, and one should perhaps not look for common features in their citric acid cycles. The species whose pathways have been investigated are Thermoplasma, Sulfolobus and Thermoproteus, organisms which cover the range of growth requirements and abilities; their citric acid cycles will be discussed in turn. Thermoplasma acidophilum is an aerobic obligate heterotroph, and it is probable that it possesses an oxidative citric acid cycle. Although radioactive tracer studies have not been reported, the following enzymes have been found: citrate synthase (Danson et al., 1985; Grossebuter and Gorisch, 1985), isocitrate dehydrogenase (M . J. Danson, unpublished observations), 2oxog1utarate:ferredoxin oxidoreductase (Kerscher et al., 1982), succinate thiokinase (Danson et al., 1985) and malate dehydrogenase (Gorisch et al., 1985; Grossebuter et al., 1986). Many strains of Sulfolobus species are facultatively autotrophic: that is, they can grow autotrophically on COz as the sole carbon source (with energy being produced aerobically by oxidation of elemental sulphur to sulphuric acid, or, in some species, anaerobically by hydrogen-dependent reduction of sulphur), or heterotrophically on yeast extract and various other carbon sources including glucose and sucrose (for reviews see Stetter and Zillig, 1985; Stetter, 1986; Stetter et al., 1986; Wood et al., 1987). Using I4CO2 pulselabelling techniques, Kandler and Stetter (1981) found that early products of heterotrophic and autotrophic I4CO2 assimilation in S. brierleyi were intermediates of the citric acid cycle. The authors interpreted the data as being inconsistent with a Calvin cycle and suggested a reductive carboxylic (citric) acid pathway the exact nature of which is yet to be defined. Wood et al. (1987) studied autotrophic, mixotrophic and heterotrophic growth of several Sulfolobus strains. Enzyme assays confirmed the absence of a Calvin cycle and demonstrated that a small proportion of COz fixation could occur through carboxylation of pyruvate and phosphoenolpyruvate; no further evidence for the proposed reductive citric-acid pathway was obtained. However, these authors suggest that simultaneous use of acetate or glucose and COz for cellular biosyntheses indicates that the COz-fixing cycle must operate simultaneously with both assimilatory and oxidative pathways.

( a ) With carbon dioxide

1

Oxaloacetate

Fumarate

i

& Succinate

Phosphoenolpyruvate

t

Pyruvate

Citrate

2-Oxog lutarate

(b) With carbon dioxide and acetate

Phosphoenolpyruvate

Fumarate

Succinate

Acetyl-CoA

co2

FIG. 8. Proposed carbon-assimilation pathways in Thermoproteus neutrophilus. 4utotrophic carbon dioxide assimilation via a reductive citric acid cycle. (b) Heterotrophic acetate assimilation via an incomplete citric acid cycle in which fumarate reductase activity is repressed. Reproduced with permission of Schafer et al. (1 986). (P’

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189

Wood e f af. ( 1 987) also report that there is little evidence from glucose radiorespirometry for large-scale use by Sulfolobus of an oxidative citric acid cycle for terminal oxidation of acetate produced from pyruvate. A number of citric acid-cycle enzymes have been detected in heterotrophically grown S. acidocaldarius, namely citrate synthase (Danson et al., 1985;Grossebuter and Gorisch, 1985), isocitrate dehydrogenase (Danson and Wood, 1984), 2oxog1utarate:ferredoxin oxidoreductase (Kerscher et al., 1982) and malate dehydrogenase (Gorisch et al., 1985). Thermoproteus neutrophilus is a strictly anaerobic archaebacterium and, although once thought to be an obligate autotroph, has been shown to be a facultative autotroph. Carbon assimilation into T. neutrophilus has been studied by Schafer et al. (1986). Autotrophically grown cells appeared not to contain ribulose 1,5-bisphosphatecarboxylase or carbon monoxide dehydrogenase; rather, all of the enzymes of a reductive citric acid cycle could be detected, except for a citrate-cleaving enzyme. Incorporation of [1,4-’4C]succinate was consistent with the operation of such a pathway. Acetate was a preferred carbon source and suppressed autotrophic COz fixation. Acetategrown T. neutrophilus exhibited an incomplete citric acid cycle in which fumarate reductase activity was “repressed”. These two proposed pathways are illustrated in Fig. 8. It is thus interesting to note that both lines of the sulphur-associated archaebacteria, represented by S . brierleyi and T, neutrophilus, appear to fix CO;!by a reductive citric acid cycle. The exact nature of such a pathway is yet to be defined although it is possibly similar to the reductive citric acid cycle in Chlorobium thiosulfatophilum (Evans et al., 1966).

3. Methanogens Methanogenic archaebacteria exhibit a third version of the citric acid cycle in addition to the oxidative pathway found in halophiles and the reductive cycle of sulphur-dependent thermoacidophiles. Two anabolic variations of an incomplete citric acid cycle are found in the methanogens. From incorporations into glutamate, aspartate and alanine of [U-’4C] acetate (Fuchs et al., 1978a), [U-’4C]succinate (Fuchs and Stupperich, 1978) and of [U-’4C]fumarate(Fuchs et al., 1978b), it appears that M . thermoautotrophicum possesses an incomplete, reductive citric acid cycle leading from oxaloacetate to 2-oxoglutarate via succinyl-CoA (Fig. 9). The enzymes required for such a pathway, namely phosphoenolpyruvate carboxylase (Daniels, 1974; Kenealy and Zeikus, 1982), malate dehydrogenase, fumarate reductase, succinate thiokinase and 2-oxoglutarate synthase (Fuchs and Stupperich, 1982 and references therein), have been found in cell extracts of this methanogen and with sufficient activities to account for this proposed function. Evidence for an incomplete reverse citric acid cycle has also been

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MICHAEL J . DANSON

ATP

C02\

.

2H+,'

*+-

c

Acetyl-CoA

2c02

_--.__ r

Oxaloacetate

Citrate +CoA

NAD+ Malate

+.

cis -Acon,itate

, ,

*

I

Succinate CaA ATP ADPtPi

k' 2-Oxoglutorate

2H

co2 FIG. 9. The citric acid cycle of methanogenic archaebacteria. The pathways shown are those proposed for Methanobacterium thermoautotrophicum (+) and Methanosarcina barkeri (- - +).

found in Methanospirillum hungatei (Ekiel et al., 1983) and Methanococcus uoltae (Ekiel et af., 1985). In contrast, Methanosarcina barkeri appears to have an incomplete but oxidative citric acid cycle, with 2-oxoglutarate being synthesized via citrate synthase, aconitase and isocitrate dehydrogenase (Daniels and Zeikus, 1978; Weimer and Zeikus, 1979) (Fig. 9). Thus, no methanogen has yet been found with a complete cycle, whether oxidative or reductive. E. PATTERNS OF THE ARCHAEBACTERIAL CENTRAL METABOLIC PATHWAYS

Now that the central metabolic pathways are being established in representatives of the three archaebacterial phenotypes, halophiles, thermoacidophiles and methanogens, it is tempting to try to discern patterns of similarity and diversity within these organisms. However, in doing so it must be remembered that, in some cases, only a few species of each phenotype have been studied and that these may not be representative of other member organisms as yet uninvestigated. Moreover, the general phenotypes do not necessarily correspond to genotypic relationships and therefore any emerging pattern may reflect adaptation of phylogenetically distinct organisms to similar environ-

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191

ments. Nevertheless, phenotypic patterns of these metabolic pathways are emerging and they are outlined below. Halophilic archaebacteria catabolize glucose via a modified EntnerDoudoroff pathway in which phosphorylation precedes the aldol-cleavage step leading to glyceraldehyde 3-phosphate and pyruvate. Gluconeogenesis is probably via reversal of an Embden-Meyerhof sequence and there is thought to be an oxidative citric acid cycle. The thermoacidophiles, namely species of Thermoplasmaand Sulfolobus, catabolize glucose via a further modification of the Entner-Doudoroff sequence in which glyceraldehyde and pyruvate are produced from glucose without phosphorylation. Data are not yet available on their gluconeogenic pathways, but the heterotrophic Tp. acidophilum appears to have an oxidative citric acid cycle whereas species of Sulfolobus growing autotrophically may use a reductive reversal of this pathway for COZ fixation. Autotrophically, Thermoproteus neutrophilus is similar to species of Sulfolobus but heterotrophic growth on acetate may use an incomplete oxidative cycle. Methanogens may possibly use an Embden-Meyerhof sequence of reactions to effect both catabolism and anabolism of glucose. Autotrophically they fix COZvia an activated acetic-acid pathway and then 2oxoglutarate synthesis (for production of glutamate) is via an incomplete citric acid cycle. Some methanogens have a reductive half of the pathway from oxaloacetate to 2-oxoglutarate, whereas others proceed oxidatively via citrate. No methanogen has yet been found with a complete cycle. These phenotypic patterns are summarized in Table 1. It is imperative to study these pathways in more archaebacterial species to see if this phenotypically associated diversity holds or whether the situation is more complicated. It is premature to speculate on the “primitive” pathways until we have a more definitive idea of which genera are primitive from a genotypic point of view. However, a number of points concerning possible evolutionary origins of the central metabolic pathways are worthy of comment in the light of archaebacterial metabolism. First, it has often been argued that the Embden-Meyerhof pathway is the ancient energy-conserving route of hexose catabolism. Gest and Schopf (1983) point out that it is a simple type of energy-conversion process, only one kind of hydrogen (electron) carrier is required (NAD+), fermentative ATP generation occurs via substrate-level phosphorylation and the net yield of ATP is low, as would be expected of a primitive energyconversion mechanism. In these arguments one must now take into account that, in halophiles and in species of Sulfolobus and Thermoplasma, organisms from the major archaebacterial branches (see Figs. 2 and 3), it is an EntnerDoudoroff-type pathway through which glucose appears to be catabolized. Yields of ATP are lower than those from the glycolytic pathway, and there is no reason why this metabolic route could not be used anaerobically if suitable reactions were available for re-oxidation of reduced cofactors.

TABLE 1. Proposed central metabolic pathways of archaebacteria Organism

Glucose catabolism

Gluconeogenesis

Citric acid cycle

Modified Entner-Doudoroff pathway

Reverse Embden-Meyerhof pathway

Complete oxidative cycle

Non-phosphorylated Entner-Doudoroff pathway

Unknown

Complete reductive cycle (autotrophic growth) Complete oxidative cycle (heterotrophic growth)

(1) Halophiles Halobacterium saccharovorurn Halobacterium halobiurn Halophiles M4, M7, Gt-1, Ll, U-14, U-18

(2) Thermophiles Sulfolobus species

Thermoplasrnaacidophilurn

Non-phosphorylated Entner-Doudoroff pathway

Unknown

Complete oxidative cycle

Thermoproteus neutrophilus

Unknown

Unknown

Complete reductive cycle (autotrophic growth) Incomplete oxidative cycle (heterotropbic growth)

Embden-Meyerhof pathway

Reverse Embden-Meyerhof pathway Unknown

Incomplete reductive cycle

(3) Methanogens Methanobacterium thermoautotrophicurn Methanosarcina barkeri

Unknown

Incomplete oxidative cycle

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A second point concerns the evolution of the citric acid cycle (reviewed by Weitzman, 1985). One scenario (Gest, I98 I ) envisages reductive conversion of oxaloacetate into succinate via malate and fumarate as having emerged in association with hexose fermentation in order to regenerate NAD+ from NADH. Thus, of the two molecules of pyruvate formed from glucose, one might be used biosynthetically while the other could be carboxylated to oxaloacetate which then supplies an electron sink in its conversion into succinate. Succinate itself is also a biosynthetic precursor in formation of porphyrin and amino acids via succinyl-CoA. As discussed by Weitzman (1985), formation of acetyl-CoA from pyruvate via an oxidoreductase is likely to have evolved in early anaerobic organisms, and this may have led to evolution of the other half of the citric acid cycle whereby 2-oxoglutarate is synthesized from oxaloacetate and acetyl-CoA via citrate and isocitrate. A primitive cell might thereby possess two arms of the cycle. Anaerobic photosynthesis is generally accepted as an early evolutionary innovation after that of fermentation. Thus, pyruvate oxidation could be driven in the reverse direction by photoreduced ferredoxin, and evolution of a 2-oxoglutarate synthase from ferredoxin-linked pyruvate synthase would result in the complete citric acid cycle operative in the anaerobic reductive capacity. Indeed, the reductive citric acid cycle may have been the primitive complete pathway, the oxidative, bioenergeticcycleawaitingavailabilityof02 as a suitable electron acceptor. It may be significant, therefore, that the reductive cycle is thought to be present in species of Sulfolobus, Therrnoproteus and, incompletely, of Methanobacterium, Methanospirillurn and Methanococcus, organisms that again are representative of the major archaebacterial branches (see Figs. 2 and 3). Also, as discussed in detail in Section III.B, all archaebacteria investigated possess oxidoreductase enzymes for interconversion of pyruvate and acetyl-CoA, and 2-oxoglutarate and succinyl-CoA, the analogous 2-0x0 acid dehydrogenase complexes probably appearing in an oxidative atmosphere after divergence of the eubacteria. Evolutionary considerations are necessarily speculative, but it is felt that these studies on archaebacterial metabolic pathways are laying the foundation for proper comparisons to be made in the future between archaebacteria, eubacteria and eukaryotes. One thing is clear, and that is that all three kingdoms possess variations on a small number of metabolic routes, routes that must have been established before the separation into the three evolutionary lineages, and which have subsequently been adapted to particular metabolic needs and environmental conditions. A more informative evaluation of metabolic evolutionary relationships will come through investigations of the enzymes catalysing these chemical interconversions; the beginnings of such studies in the archaebacteria are the subject of the next section on “enzymic diversity”.

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111. Archaebacterial Enzyme Diversity

Now that some of the pathways of glucose catabolism have been established in representatives of the halophilic, thermoacidophilic and methanogenic archaebacteria, we are in a position to begin enzymological comparisons with the counterparts in eubacterial and eukaryotic species. However, it will be obvious from the previous section that these archaebacterial metabolic routes have only recently been defined and, consequently, studies of the enzymes involved are still at a preliminary stage. Therefore, the enzymes and enzyme systems that I have chosen to discuss in this section are those with which I have some personal involvement and which are either unique to archaebacteria or have well-studied parallel systems in eubacteria and eukaryotes. Thus, the survey is necessarily incomplete and focuses on archaebacterial enzymes, the study of which will allow comparisons to be made across all three proposed evolutionary kingdoms. Indeed, it is the species-dependent diversity of these enzymes that is the emphasis of this review.

A. DEHYDROGENASES WITH DUAL COFACTOR SPECIFICITY

The nicotinamide nucleotide-dependent dehydrogenases of eubacterial and eukaryotic organisms are characteristically specific for either NAD+ or NADP+, and it is uncommon to find such enzymes which are catalytically active with both cofactors. However, it is becoming clear that this is not the situation in the thermoacidophilic archaebacteria where a number of dehydrogenases have been found which accept both NAD+ and NADP+ on the same protein molecule. These are described in turn, with particular attention paid to whether or not the separate NAD+- and NADP+-dependent dehydrogenases occur in other organisms. 1. Zsocitrate Dehydrogenase of Sulfolobus acidocaldarius

Isocitrate dehydrogenase catalyses the nicotinamide nucleotide-dependent oxidative decarboxylation of isocitrate to 2-oxoglutarate in the citric acid cycle:

+

Isocitrate NAD(P)+*2-Oxoglutarate+ NAD(P)H

+H + +COZ

Eukaryotes possess both NAD+- and NADP+-specific isocitrate dehydrogenases (reviewed by Colman, 1983 and by Plaut and Gabriel, 1983). The NAD+-linked enzyme is confined to mitochondria and is allosterically regulated in a manner consistent with a controlling role in the energy-yielding function of the cycle. The NADP+-dependent isocitrate dehydrogenase is

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found in both the mitochondrion and the cytoplasm but, unlike the NAD+linked enzyme, is not subject to allosteric control. Mechanistically, the NAD+- and NADP+-isocitrate dehydrogenases share common features; for example, both enzymes use the same side of the nicotinamide ring (Chen and Plaut, 1963a),in both enzymic reactions a proton from the solvent replaces the carboxyl group of isocitrate with retention of configuration (Lienard and Rose, 1964; Englard and Listowsky, 1963; Rose, 1966) and chemical modification studies indicate the possibility of similarities in the active-site regions (Colman, 1983). However, the two enzymes differ markedly in physicochemical properties; the NADP+ 4socitrate dehydrogenase is a monomeric protein (Mr 60,000) which may dimerize in the presence of substrate, whereas the NAD+-linked enzyme possesses three different subunits [a(Mr 39,000), b(Mr4 1,000) and y(Mr41,OOO)l in a stoicheiometry of 2: 1:1 (Colman, 1983; Plaut and Gabriel, 1983; and references therein). This NAD+-isocitrate dehydrogenase also appears to form reversibly associated aggregates, an octameric species having been detected, with the possibility of additional, more complex structures. Eubacterial isocitrate dehydrogenases have been reviewed by Weitzman (1981). The majority of eubacteria appear to contain only an NADP+-linked enzyme (Mr100,000) which has been shown to be a dimeric protein. A few eubacteria have been found to contain an NAD+-isocitrate dehydrogenase ( c g . Streptococcus bouis, Acetobacter suboxydans and Thiobacillus thiooxiduns), whereas others (e.g. Acetobacter peroxydans, Xanthomonas pruni and Hydrogenomonas eutropha) contain both NAD+- and NADP+-linked enzymes (see Weitzman, 1981, and references therein). Clearly there is a diversity of isocitrate dehydrogenase enzymes in eukaryotic and eubacterial organisms, but the NAD+- and NADP+-specific enzymes are separate and distinct proteins. The one possible exception is the isocitrate dehydrogenase of Rhodomicrobium vannielii (Morgan et al., 1986) which may be able to use both NAD+ and NADP+ (D. J. Kelly, personal communication). This general eubacterial and eukaryotic situation contrasts markedly with isocitrate dehydrogenase from Sulfolobus acidocaldarius (Danson and Wood, 1984). Cell extracts of this archaebacterium were found to possess both NAD+- and NADP+-dependent activities, both of which showed hyperbolic dependence of enzymic velocity on cofactor and substrate concentrations. While K , values for the two cofactors differed considerably [KmaPP3.4 ( f 0.5) mM for NAD+ and 30 (k0.2) p~ for NADP+], the Vma, values [0.10-0.13 ] pmol min-'(mg protein)-'] and K , values for isocitrate [lo (f0.5)p ~ were similar for both NAD+- and NADP+-dependent isocitrate dehydrogenase activities. We presented evidence that both enzymic activities are functions of the same protein; NAD+ and NADP+ competed with each other for the enzyme and did so with K; values equal to their K , values; thermal

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inactivation resulted in loss of both activities at the same rate, and copurification was observed through a number of different procedures (Danson and Wood, 1984).The enzyme has an M , value of 96,000 ( f4,000); it has recently been purified to homogeneity and is probably a dimeric protein (T. Oshima, personal communication). The pure enzyme retains both NAD+ and NADP+ activities. It is premature to consider the evolutionary significance of this dual specificityisocitrate dehydrogenase although it is tempting to compare it with the two (NAD+ and NADP+) enzymes found in eukaryotic species. For instance, for the archaebacterial enzyme, the K,,,value for NAD+ is approximately a hundred times greater than that for NADP+. A similar ratio has been found for the separate eukaryotic isocitrate dehydrogenases, although the values for the constants are at least 10-fold higher in the enzyme from S. acidocaldarius. However, the archaebacterial isocitrate dehydrogenase was assayed at Sac,and it has been reported (Dalziel, 1975) that an increase in temperature produces a decrease in nucleotide-binding affinity for some dehydrogenases. Interestingly, Chen and Plaut (1963b) found that the pig-heart NAD+-linked isocitrate dehydrogenase also binds NADPH but probably at a different site from NAD+. Thus no catalytic activity is detectable with NADP+ but, like the enzyme from S . acidocaldarius, it does bind both cofactors. The reciprocal situation of NAD(H) binding to the NADP+-specific enzyme has not been reported. Finally, it will be interesting to see if other genera of thermoacidophilic archaebacteria possess dual-specificity isocitrate dehydrogenases; at present, we have preliminary kinetic data to suggest that the enzyme from Thermoplasma acidophiliumis also active with both NAD+ and NADP+ (L. Smith, M. J. Danson and D. W. Hough, unpublished work). Whatever the distribution, it is felt that molecular characterization of the isocitrate dehydrogenase from S. acidocaldarius will yield important evolutionary and mechanistic information on cofactor specificity.

2.Glucose Dehydrogenase from Species of Sulfolobus and Thermoplasma It has already been noted in the discussion of glucose metabolism in Sulfolobus solfataricus (De Rosa et al., 1984) and Tp. acidophilum (Budgen and Danson, 1986a) that cell extracts of these two thermoacidophilic archaebacteria possess both NAD+- and NADP+-linked glucose dehydrogenase activities. Evidence strongly suggests that in both organisms it is one enzyme that utilizes both cofactors. The glucose dehydrogenase from S. solfataricus has been purified to homogeneity (Giardina et al., 1986)and shown to be a tetrameric protein with a native M , value of 124,000. The pure enzyme is indeed catalytically active

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with both nicotinamide nucleotide cofactors, maximum activity being found at 77°C and in the presence of Mg2+, Mn2+ or Ca2+. With NAD+ as coenzyme, among a number of monosaccharides examined only D-glucose, Didose and D-xylose were oxidized at a high rate and it has therefore been proposed that the dehydrogenase specifically oxidizes sugars presenting equatorial orientation of hydroxyl groups at C-2, C-3 and C-4. A broader substrate specificity was found with NADP+ although all of the sustrates were oxidized at a lower rate than that of glucose in the presence of NAD+. However, the KmaPPvalue for NADP+ was 40 times lower than the KmaPPvalue for NAD+ and the KmaPPvalue for glucose with NADP+ as cofactor was 20 times lower than with NAD+. Given that it is K,,,/K,,,that determines enzymic specificity in the sense of discrimination between two competing substrates (Fersht, 1985), then glucose may preferentially be oxidized by the glucose dehydrogenase from S. solfataricus with NADP+ as electron acceptor. The enzyme has not yet been purified from Thermoplasma acidophilum but we have kinetic evidence (N. Budgen and M. J. Danson, unpublished work) that it is also a single enzyme which utilizes both NAD+ and NADP+ and oxidizes both glucose and galactose. In a manner similar to the enzyme from S. solfataricus, the glucose dehydrogenase from Tp. acidophilum has a much higher KmaPPvalue for NAD+ than for NADP+, although the affinities for glucose and galactose were not significantly different. Almost all glucose dehydrogenases from non-archaebacterial species are either NADi- or NADP+-specific. However, notable exceptions are found in various Bacillus species. The enzyme has been purified from B. subtilis (Fujita et al., 1977) and catalyses the oxidation of glucose with NAD+ and NADP+, the two cofactors binding with approximately equal affinities. Similarly, glucose dehydrogenase from B. cereus is active with both cofactors (Sadoff, 1966). Both Bacillus species are spore formers, and Giardina et al. (1986) noted that heat-resistant endospores and thermophilic archaebacteria have evolved a similar enzymic protein; however, the significanceof this correlation remains unclear, particularly in view of the observation that the mesophilic Halobacterium saccharouorum may also have a dual-specificity glucose dehydrogenase (Tomlinson et al., 1974). 3 . Malate Dehydrogenase

Malate dehydrogenase catalyses the interconversion of malate and oxaloacetate in the citric acid cycle: Malate + NAD(P)+=>Oxaloacetate+ NAD(P)H

+H +

Most eubacterial and eukaryotic species possess an NAD+-specific malate dehydrogenase (EC 1.1.3.37) although many higher plants also have an

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MICHAEL J. DANSON

NADP+-dependent enzyme (EC 1.1.1.82) (Hatch and Slack, 1969; Johnson, 1971; Johnson and Hatch, 1970). In two archaebacterial phenotypes a malate dehydrogenase with dual cofactor specificity has been found. The enzyme from M . hungatei is catalytically active with NADH and NADPH (Sprott et al., 1979). It is reported that the catalytic rate with NADPH is only 10% of that with NADH although, without a knowledge of K , values, the preferred cofactor cannot be identified. In contrast, NADPH is not an effective electron donor for malate dehydrogenase from M . thermoautotrophicum (Zeikus et al., 1977). Malate dehydrogenases from Tp. acidophilum and S. acidocaldarius also accept both NAD+ and NADP+ (Grossebuter et al., 1986).The enzymes have been purified to homogeneity and shown to be tetrameric proteins with subunits of M , N 36,000. In non-archaebacterial species, both dimeric and tetrameric malate dehydrogenases have been found although the subunit Mr values are all in the range 32,000 to 36,000 (Murphey et al., 1967a,b; Sundaram et al., 1980; Weitzman, 1981). The dimeric form is the more widely distributed enzyme, the larger enzyme having been found in some species of Bacillus, Corynebacterium, Butyribacterium, Sarcina, Pediococcus and Neisseria. The dual-specificity malate dehydrogenase from M . hungatei has a native M , value of 62,000 as judged by gel filtration and is therefore probably a dimeric enzyme. This is also the case for the enzyme from H . halobium (native M,84,000) but which is NAD+-specific (Mevarech et af., 1977). 4 . Concluding Remarks

These few examples serve to illustrate the fact that dual cofactor specificity is not an uncommon feature of dehydrogenase enzymes in the thermoacidophilic archaebacteria. Indeed, the phenomenon may turn out to be far more prevalent than the current observations suggest. In the case of isocitrate dehydrogenase from S. acidocaldarius, for example, the K , value for NAD+ is two orders of magnitude greater than that for NADP+ and catalytic activity with the former could well have been overlooked if low concentrations of both cofactors had been used. Thus, it may well be worth investigating a number of apparently cofactor-specific archaebacterial dehydrogenases for observable activity with higher concentrations of the other nicotinamide nucleotide coenzyme. Even so, the dual-cofactor specificity of the enzymes described above is a notable feature of the thermophilic archaebacteria. It has been suggested that the wider specificity may be a consequence of adaptation to the high environmental temperatures in which the organisms grow, although these extremes are not required for the dual activities; thus isocitrate dehydrogenase activities with NAD+ and NADP+ are in the same ratio at 30°C as they are at 55°C (Danson and Wood, 1984). There have also been

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suggestions that thermophilicity is an ancient phenotype and the lack of absolute cofactor specificity may reflect this antiquity and may have been preserved only in organisms that still grow in the specialized ecological niches characteristic of these archaebacteria. Whatever the evolutionary reason for selection of dehydrogenases which are active with both NAD+ and NADP+, it is clear that a detailed study of their structures may provide an insight into the nature of how other dehydrogenases distinguish between the two coenzymes. Although a number of dehydrogenases from non-archaebacterial species have been crystallized and their structures elucidated, there are very few comparisons possible between NAD+ and NADP+ binding to the same or similar enzymes. The first direct comparison between two related enzymes that are NADP+-linked (human glutathione reductase) and NAD+-linked (E. coli dihydrolipoamide dehydrogenase) was made by Rice et al. (1984). These two proteins are highly homologous in primary structure and the amino-acid sequence of the dihydrolipoamide dehydrogenase has been compared with the sequence and three-dimensional structure of glutathione reductase. It was shown in the latter protein that the 2’-phosphate of NADP+ is bound by multiple interactions with Arg-218, His-219 and Arg-224; in the dihydrolipoamide dehydrogenase these three residues are replaced by Met, Pro and Phe, consistent with its requirement for NAD+. Similarly, the structures of glucose dehydrogenases from Bacillus megaterium and B. subtilis (both of which can utilize NAD+ and NADP+) have been predicted by comparison with the known structure of lactate dehydrogenase (NAD+-specific) (Hones et al., 1987). An aspartate residue binding the 2’-0H of the adenosine ribose ring of NAD+ in lactate dehydrogenase is replaced by an asparagine residue in glucose dehydrogenase; the authors suggest that this replacement permits both NAD+ and NADP+ binding in glucose dehydrogenase and further explains why NADP+ would bind more tightly than the non-phosphorylated coenzyme. B.

2-OX0 ACID : FERREDOXIN OXIDOREDUCTASES

Conversion of the 2-0x0 acids pyruvate and 2-oxoglutarate into their corresponding acyl-CoA thioesters are crucial reactions in intermediary metabolism. The nature of the enzymic systems catalysing such reactions depends on the organism concerned and the conditions under which it is growing. Representatives of these different systems have been characterized kinetically, mechanistically and structurally, and they therefore provide one of the best sources for detailed enzymic comparisons between eukaryotes, eubacteria and archaebacteria.

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1. Eukaryotes and Eubacteria

In aerobic environments, eukaryotes and many respiratory eubacteria oxidatively decarboxylate the 2-0x0 acids via the pyruvate and 2-oxoglutarate dehydrogenase multienzyme complexes, both of which use NAD+ as the electron acceptor (Reed, 1974; Perham, 1975). The mechanism of the complexes is shown in Fig. 10(a). Each complex molecule is composed of multiple copies of three separate enzymic activities, namely a 2-0x0 acid decarboxylase (El), a dihydrolipoyl acyltransferase (E2) and dihydrolipoamide dehydrogenase (E3). Enzyme E 1 catalyses the thiamin pyrophosphate (TPP)-dependent decarboxylation of the 2-0x0 acid and the enamine so formed is then transferred to lipoic acid, a cofactor coval4ntly bound to the ENH2 group of a lysine residue of enzyme E2. The acyl mbiety is passed from the lipoyl-lysine to CoA and the dihydrolipoyl-lysine so generated is reoxidized by enzyme E3, the catalytic mechanism of which involves FAD and alternate oxidation and reduction of an intrachain disulphide bond (Williams, 1976). The reducing equivalents are finally accepted by NAD+ to form NADH. The central feature of this catalytic mechanism is therefore lipoic acid which acts as a swinging-arm to connect the active sites of these multienzyme complexes and so channels the substrate through the three component enzymes. The E2 component constitutes the structural core of the 2-0x0 acid dehydrogenase complexes to which the E l and E3 components are noncovalently bound. In both eukaryotic and eubacterial 2-oxoglutarate dehydrogenase complexes, this core possesses octahedral symmetry, being composed of 24 E2 polypeptide chains (DeRosier and Oliver, 1972; Reed, 1974). The same symmetry and number of polypeptides are found in the E2 core of the pyruvate complex from Gram-negative eubacteria (Reed, 1974; Danson et al., 1979a), whereas in Gram-positive eubacteria and eukaryotes there are 60 E2 polypeptide chains arranged with icosahedral symmetry (Reed, 1974; Henderson et al., 1979). This species-dependent diversity of structure has been discussed, along with that observed in other citric acid cycle enzymes, by Weitzman (1981). An interesting connection between the multimeric structure of the 2-0x0 acid dehydrogenase complexes and their catalytic mechanism has been discovered. This is the phenomenon of “active-site coupling”, whereby lipoic acid not only serves to connect E l , E2 and E3 but can effect rapid transfer of acyl groups between neighbouring E2 polypeptide chains within one complex molecule. This active-site coupling has been observed in both pyruvate and 2oxoglutarate dehydrogenase complexes of eubacteria and eukaryotes (Bates et al., 1977; Collins and Reed, 1977; Danson et al., 1978a; Cate and Roche, 1979; Stanley et al., 1981).We have suggested that this mechanism ofcoupling

W

[TPP-H]

R-C-COOH

R-E-SCOA

+H+ Net reaction

R R-C-COOH + CoASH+ NAD'

-

NAD'

FI R-C-SCOA+CO~

+

NADH+H*

(b)

FeS

red

FeS

FeS

FeS

OX

red

ox

FIG. 10. Reaction mechanisms of the 2-0x0 acid dehydrogenase multienzyme complexes and the 2-0x0 acid:ferredoxin oxidoreductases. (a) The 2-0x0 acid dehydrogenase multienzyme complexes El (a 2-0x0 acid decarboxylase), E2 (a dihydrolipoyl acyltransferase) and E3 (dihydrolipoamide dehydrogenase) represent the three enzymic activities that make up the complexes; TPP-H, thiamin pyrophosphate; Lip, lipoic acid; B is a base on the dihydrolipoamide dehydrogenase component. (b) The 2-0x0 acid:ferredoxin oxidoreductases of the archaebacteria. TPP-H indicates thiamin pyrophosphate; Fd ferredoxin and FeS an enzyme-bound iron-sulphur cluster.

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the decarboxylation of a 2-0x0 acid on a particular E l component with the acylation of CoA at a physically removed E2 enzyme may permit a rate enhancement at low substrate concentrations (Danson et al., 1978b). Thus, a rationale for the multimeric structure of these complexes has been offered. Under anaerobic conditions, NAD+ would act as an unacceptable electron trap and therefore, in strictly and facultatively anaerobic eubacteria, two possible alternatives exist for the NAD+-linked 2-0x0 acid dehydrogenase complexes. The first retains the oxidative process but, in this case, the electrons are transferred to ferredoxin or flavodoxin (acceptors with a more negative redox potential than NAD +) through 2-0x0 acid oxidoreductases. These enzymes are far simpler than the dehydrogenase complexes, being smaller, oligomeric proteins with a M , value below 300,000 (Uyeda and Rabinowitz, 1971; Gehring and Arnon, 1972; Blaschkowski et al., 1982; Kerscher and Oesterhelt, 1982). A great advantage to photosynthetic and chemolithotrophic organisms is the opportunity for the oxidoreductase reaction to be driven in the reverse direction, effecting carbon dioxide fixation via a reductive carboxylic acid cycle (Fuchs et al., 1980). The second alternative for the dehydrogenase complexes is to avoid oxidation of pyruvate by converting it into acetyl-CoA and formate via action of pyruvate formate lyase (Knappe et al., 1969; Payton and Haddock, 1985).

2. Archaebacteria In archaebacteria, the conversion of pyruvate and 2-oxoglutarate into their respective acyl-CoA esters is catalysed by 2-0x0 acid oxidoreductases, with no alternative enzyme systems yet detected. The oxidoreductases have been found in H . halobium (Kerscher and Oesterhelt, 1981a,b), Tp. acidophilum, S. acidocaldarius, D . mobilis (Kerscher et al., 1982)and M . thermoautotrophicum (Zeikus et al., 1977). Therefore the pattern of enzymes is constant throughout the archaebacteria, whether aerobic or anaerobic, autotrophic or heterotrophic. However, there is a diversity of electron acceptors in the reactions, the halophiles and thermoacidophiles using ferredoxin and the methanogens the deazaflavin derivative F42,,. Pyruvate and 2-oxoglutarate:ferredoxin oxidoreductases have been purified from H . halobium (Kerscher and Oesterhelt, 1981a). Their respective M , values were found to be 256,000 and 248,000, respectively and both have an a$2 subunit structure with, respectively, polypeptide Mt values of 86,000 and 42,000 for the pyruvate enzyme and 88,000 and 36,000 for the 2-oxoglutarate protein. Electron paramagnetic resonance signals and absorption spectra of the purified proteins indicated the presence of [4Fe-4S]2+(2+.i+) clusters. Thiamin pyrophosphate (TPP) was also found to be present although neither lipoic acid nor flavin nucleotides could be detected. Thus the archaebacterial

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enzymes appear similar to the 2-0x0 acid oxidoreductases of eubacteria but are fundamentally different from the 2-0x0 acid dehydrogenase multienzyme complexes. The proposed catalytic mechanism of the archaebacterial2-0x0 acid:ferredoxin oxidoreductases (Kerscher and Oesterhelt, 1981b) is shown in Fig. 10(b) for comparison with that of the 2-0x0 acid dehydrogenase complexes. The 20x0 acid is decarboxylated in a TPP-dependent reaction to give hydroxyalkylTPP;therefore it is proposed that one electron is abstracted and transferred to the enzyme-bound iron-sulphur cluster (and subsequently to ferredoxin) leaving a free radical-TPP species. This intermediate then reacts direct with CoA to form acyl-CoA with concommitant transfer of a second electron to the iron-sulphur cluster. Again, ferredoxin serves to reoxidize the enzyme’s redox centre.

3. A Unique Catalytic Mechanism for Oxidoreductases? The fundamental differences between the catalytic mechanisms of the archaebacterial 2-0x0 acid oxidoreductases and of the eubacterial and eukaryotic 2-0x0 acid dehydrogenase complexes have been stressed (Kerscher and Oesterhelt, 1981b, 1982; Oesterhelt et al., 1983). As a balance to this view, it may be constructive, for evolutionary considerations, to stress some of the similarities of the catalytic processes. First, in both enzyme systems, the 2-0x0 acids are decarboxylated via the cofactor TPP to give initially a hydroxyalkyl derivative. Next, in the oxidoreductases, an electron is abstracted from this intermediate before transfer of the acetyl group to CoA and abstraction of the second electron, i.e. there are separate redox and group-transfer steps. Recent evidence suggests the possibility of similar discrete steps in the pyruvate dehydrogenase complex even though the acetyl group is first transferred to lipoic acid and then to CoA (Flourney and Frey, 1986). Flourney and Frey (1986) detected formation of acetyl-TPP from pyruvate in the pyruvate dehydrogenase complex of Escherichia coli and, furthermore, demonstrated that this acetyl group can be transferred to dihydrolipoamide on E2 with an efficiency of approximately 95%. Thus, acetyl-TPP appears to be chemically competent in this catalytic mechanism, suggesting transfer of reducing equivalents from hydroxyethyl-TPP to the lipoyl groups before transfer of the acetyl moiety. However, it still remains to be determined whether this aspect of the catalytic mechanism is the kinetically competent route. If it is, then there may be a closer similarity with the mechanism of the 2-0x0 acid oxidoreductases than initially proposed. Reoxidations of the enzyme systems after formation of acyl-CoA are obviously quite distinct. In the 2-0x0 acid dehydrogenase complexes of eubacteria and eukaryotes, there is therefore the FAD-containing dihydroli-

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poamide dehydrogenase component whose function is to reoxidize the E2dihydrolipoamide groups. This enzyme is not part of the archaebacterial oxidoreductases as these do not contain lipoic-acid residues. It is therefore extremely interesting to note that we have discovered the enzyme dihydrolipoamide dehydrogenase in archaebacteria; this discovery, and its possible functional and evolutionary significance, are discussed in the next section. In the present context it is sufficient to note that, even though it represents a major difference between the 2-0x0 acid dehydrogenase complexes and the oxidoreductases, the enzyme is present in archaebacteria. As a final point of comparison, whereas halophiles and thermoacidophiles possess ferredoxins as electron acceptors, in methanogenic archaebacteria the reducing equivalents from pyruvate oxidation are accepted by the deazaflavin derivative F420. In 2-0x0 acid dehydrogenase complexes, electrons are also transferred to a flavin (FAD) on the E3 component, dihydrolipoamide dehydrogenase, before acceptance by NAD+ . Thus, even in redox aspects, the two enzyme systems are not totally dissimilar. 4 . Evolutionary Considerations

Kerscher and Oesterhelt (1982) have pointed out that all species of archaebacteria so far investigated contain either a 2-0x0 acid:ferredoxin oxidoreductase of the type found in anaerobic eubacteria, or an analogous enzyme using F420 as electron acceptor. Thus, they suggest that these enzymes existed before divergence of the archaebacteria, eubacteria and the ancestral eukaryotic line of descent, and that it seems likely that the 2-0x0 acid dehydrogenase complexes evolved soon after the development of oxidative phosphorylation (the complexes have only been detected in respiratory organisms). Organisms that have retained their anaerobic niches still use 20x0 acid:ferredoxin oxidoreductases. However, both enzyme systems are found in cyanobacteria (Bothe and Nolteersting, 1975),so too in E. coli, which also possesses a pyruvate-formate lyase (Blaschkowski et al., 1982), although in the latter species the oxidoreductase is expressed at a level 0.1-1 % of the other two pyruvate-metabolizing enzyme systems. Evolution of the 2-0x0 acid dehydrogenase complexes remains an intriguing subject, especially as archaebacteria lack them but nevertheless contain a TPP-dependent decarboxylase activity (in the oxidoreductases), an enzymic activity to transfer the CZmoiety of hydroxyethyl-TPP to CoA (also in the oxidoreductases) and a dihydrolipoamide dehydrogenase enzyme. Possible evolutionary relationships of these to components of the 2-0x0 acid dehydrogenase complexes remain speculative in the absence of detailed protein-chemical information, but it would seem an area worthy of investigation.

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Finally, mention should be made of the ferredoxin electron acceptors of the archaebacterial2-0x0 acid oxidoreductases as they too have been considered from an evolutionary point of view (Kerscher and Oesterhelt, 1982; Kerscher et al., 1982; Oesterhelt et al., 1983). In eubacteria and eukaryotes, two major types of ferredoxin have been identified. One is widely distributed among eubacteria, especially the photosynthetic and fermentative anaerobes, and contains [4Fe-4S] clusters as redox centres; this ferredoxin is termed the “bacterial-type”. The second, the “chloroplast-type”, contains [2Fe-2S] clusters, and is found in chloroplasts and in just one eubacterial group, the cyanobacteria. The cyanobacterial origin of eukaryotic chloroplasts is discussed by Gray and Doolittle (1982). Thermoacidophilic archaebacteria probably contain [4Fe-4S] ferredoxins (Kerscher et al., 1982), the sequences of which have been obtained for the proteins from Tp. acidophilum (Wakabayashi et al., 1983) and S. acidocaldarius (Minami et al., 1985). Preliminary X-ray diffraction studies have been reported for the ferredoxin from Tp. acidophilum (Tsukihara et al., 1985). Alignment of the protein sequences indicates that the ferredoxins from the two thermoacidophiles are homologous, and the overall distribution of cysteine residues suggests that they resemble other bacterial ferredoxins. However, archaebacterial proteins show more complicated structures than their eubacterial counterparts and it has been proposed (Minami et al., 1985) that this may reflect adaptation of the former to extreme environments. Surprisingly, halobacteria possess [2Fe-2S] ferredoxins. Protein sequences from two species of halophilic archaebacteria have been determined (Hase et al., 1978, 1980) and they both show remarkable homology with the ferredoxins of green plants and cyanobacteria. In fact, halobacteria are the only organisms known to contain ferredoxins homologous to the “chloroplast-type’’ proteins and, as discussed by Kerscher and Oesterhelt (1982), this poses some evolutionary problems. Occurrence of this type of ferredoxin in one eubacterial branch argues against its presence before divergence of archaebacteria and eubacteria. Thus, it has been speculated (Kerscher and Oesterhelt, 1982; Oesterhelt et al., 1982) that halobacteria may have adopted this and other proteins by the exchange or acquisition of genetic material from another organism before the high degree of genetic specialization was reached to prevent such processes from taking place. Furthermore, with the sequences of ferredoxins from thermoacidophiles showing homology with the eubacterial proteins and that from Methanosarcina barkeri (Hausinger et al., 1982) being similar to clostridial ferredoxin, Minami et al. (1985) suggest that these data support the existence of multiple phyletic lines in the origin of the archaebacteria. Clearly, as with 2-0x0 acid oxidoreductases, further information on the sequence of ferredoxins from more archaebacterial species will throw additional light on these intriguing observations and suggestions.

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MICHAEL J. DANSON

C. DIHYDROLIPOAMIDE DEHYDROGENASE

As described in the previous section (see Fig. 10a), the enzyme dihydrolipoamide dehydrogenase (EC 1.8.1.4) catalyses NAD+-dependent oxidation of dihydrolipoamide (reviewed by Williams, 1976). This is the only known function of the enzyme and one which it carries out as an integral component of the pyruvate, 2-oxoglutarate and branched-chain 2-0x0 acid dehydrogenase multienzyme complexes (Reed, 1974; Perham, 1975; Pettit et al., 1978). As also described, these complexes have never been found in archaebacterial species (Aitken and Brown, 1969; Kerscher and Oesterhelt, 1982; Danson, 1982) and my colleagues and I were therefore surprised to find the presence of dihydrolipoamide dehydrogenase activity in the halophilic archaebacteria (Danson et al., 1984). Having found this enzymic activity in both classical (Halobacterium halobium and Halobacterium volcanii) and alkaliphilic (Natronobacterium pharaonis, Natronobacterium gregoryi and Natronococcus occultus) halopiles and shown the expected stoicheiometry of the reaction to be one mole of NADH produced per mole of dihydrolipoamide oxidized, we have followed up the unexpected discovery of its presence by attempting to answer three fundamental questions that immediately come to mind. First, is this a true dihydrolipoamide dehydrogenase as found in eukaryotic and eubacterial species and, if so, how similar are its enzymological and structural properties to the non-archaebacterial counterparts? Secondly, are halophiles the only group to possess this enzyme in the absence of 2-0x0 acid dehydrogenase complexes or is a similar situation found in other archaebacteria or even in eukaryotes or eubacteria? Thirdly, what is the function of a dihydrolipoamide dehydrogenase in the absence of 2-0x0 acid dehydrogenase complexes? 1 . Halophilic Dihydrolipoamide Dehydrogenase As might be expected, the catalytic activity of dihydrolipoamide dehydrogenase is optimal in the presence of 2~ NaC1. The enzyme exhibits a hyperbolic dependence of velocity on both dihydrolipoamide and NAD+ concentrations, is specific for NAD+ and is noticeably resistant to thermal denaturation (Danson et al., 1984). We have purified to homogeneity the enzyme from H . halobium, and have shown it to be a dimeric flavoprotein with a polypeptide chain with an M , value of 58,000 (k3,OOO) (Danson et al., 1986). These properties are remarkably similar to those of dihydrolipoamide dehydrogenases from 2-0x0 acid dehydrogenase complexes of eukaryotes and eubacteria. The similarity extends further to the catalytic mechanism of the nonarchaebacterial enzyme; this involves alternate oxidation and reduction of an intrachain disulphide bond (Fig. lOa), the polypeptide sequences around

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which have been determined for the proteins from E. coli and pig heart (Stephens et al., 1983). At present, we have no sequence information for the halophilic dihydrolipoamide dehydrogenase, but we have provided strong evidence from chemical modification experiments that it too possesses an active-site disulphide bridge. Thus, the enzyme is only susceptible to inactivation by the thiol reagent, iodoacetic acid, in the presence of its reducing ligands, dihydrolipoamide or NADH (Danson et al., 1984). That this modification is taking place at a reduced disulphide bond (i.e. the dithiol form of the enzyme) has been confirmed by similar experiments with the reagent (p-aminopheny1)dichloroarsine; this trivalent arsenical reacts specifically with vicinal thiol groups leaving single thiols unmodified. As predicted, the reagent is a powerful inactivator of substrate-reduced dihydrolipoamide dehydrogenase from H . halobium (Danson et al., 1986). On the question of how similar in structure it is to the non-archaebacterial enzymes, a comparison of amino-acid compositions as a measure of sequence similarities (Cornish-Bowden, 1977) gave no detectable homology between the enzyme from H . halobium and those from E. coli and pig heart (Danson et al., 1986). The same analysis estimated a homology of approximately 70% between eubacterial and eukaryotic dihydrolipoamide dehydrogenases, in good agreement with the value of 57% identity on the basis of partialsequence comparisons (Stephens et al., 1983). The apparent lack of homology with the archaebacterial enzyme is possibly a consequence of adaptation of the halophilic dihydrolipoamide dehydrogenase to conditions of high concentrations of salt and therefore true estimates of homology must await full sequence data. Attempts to obtain an N-terminal sequence for the H . halobium enzyme have been made (Danson et al., 1986). No sequence was obtained, strongly suggesting that the N-terminal amino acid is blocked; support for this conclusion has been gained from the observed resistance of the enzyme to the exopeptidase, aminopeptidase M. Nevertheless, we can conclude that halophilic archaebacteria do possess a true dihydrolipoamide dehydrogenase, although ,,its function in the absence of the 2-0x0 acid dehydrogenase complexes remains to be elucidated. 2. Dihydrolipoamide Dehydrogenase in Other Archaebacteria

My colleagues and I have discovered the presence of dihydrolipoamide dehydrogenase in the thermoacidophile Tp. acidophilum (Smith et at., 1987) and possibly in the thermophiles Desulfurococcus mucosa and Thermococcus celer and in the methanogen Ms. barkeri (M. J. Danson, D. W. Hough and N. Walker, unpublished work). Thus, its presence in the absence of 2-0x0 acid dehydrogenase complexes may be of general occurrence in archaebacteria. The dihydrolipoamide dehydrogenase from Tp. acidophilum has an M , value

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MICHAEL J . DANSON

of 118,000 (k11,000), is specific for NAD+ and is inactivated by (paminopheny1)dicloroarsine in the presence of reducing ligands (Smith er al., 1987). In these respects, it is similar to other dihydrolipoamide dehydrogenases, but we have preliminary evidence that the enzyme may be membraneassociated, the possible significance of which is discussed later in this review. We have extended our studies outside the archaebacteria to see if similar situations occur in eubacterial and/or eukaryotic species. Our first choice of organism was the eukaryotic parasite, Trypanosoma brucei, the causative agent of African Sleeping Sickness. The bloodstream form of this organism is totally dependent on glycolysis for production of ATP as it has no carbohydrate or lipid reserves and its mitochondrion does not possess a competent citric acid cycle. Consequently, it does not possess 2-0x0 acid dehydrogenase complexes and, under aerobic conditions, glucose is metabolized almost completely to pyruvate which is then excreted (Fairlamb, 1982). Thus, T. brucei provided us with a suitable eukaryotic organism in which to look for dihydrolipoamide dehydrogenase, and we have recently discovered the presence of the enzyme in the bloodstream form (Danson et al., 1987). As with all other dihydrolipoamide dehydrogenases, it specifically catalysed stoicheiometric oxidation of dihydrolipoamide by NAD+, and modification with trivalent arsenicals indicates the operation of a reversibly reducible disulphide bond in catalysis. In addition to these expected properties, we have found that the enzyme is specifically located in the plasma membranes of the organism; the dihydrolipoamide dehydrogenase copurifies with these membranes and, having extracted it from them with detergent, it can be reconstituted in an active form into phospholipid vesicles. The membrane association of the dihydrolipoamide dehydrogenase in T. brucei and possibly in Tp. acidophilum may represent the first steps in elucidation of its function in the absence of 2-0x0 acid dehydrogenase complexes. As yet, we have not looked for the enzyme in those anaerobic eubacteria that metabolize 2-0x0 acids via oxidoreductase enzymes, but it would not be surprising to find the dihydrolipoamide dehydrogenase in this kingdom too. 3. Metabolic Function of Dihydrolipoamide Dehydrogenase

Studies on archaebacteria, and latterly on T . brucei, have raised the distinct possibility that the enzyme dihydrolipoamide dehydrogenase has a metabolic function in addition to its established role in 2-0x0 acid dehydrogenase complexes (Danson, 1987).One should, however, keep in mind the possibility that functional pyruvate and 2-oxoglutarate dehydrogenase complexes are present in archaebacteria but have, for reasons of instability, for example, remained undetected.

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Data from T. hrucei may suggest a function specific to a membrane location. Indeed, evidence has been presented (Richarme, 1985; Richarme and Heine, 1986) that lipoic acid and dihydrolipoamide dehydrogenase may be involved in the binding protein-dependent transport of galactose and maltose in E. coli, and a membrane-bound form of the enzyme has been found in this organism (Owen et al., 1980). Additionally it has been observed that the dithiol-specific trivalent arsenical, phenylarsine oxide, inactivates insulinstimulated hexose transport by 3T3-LI adipocytes (Frost and Lane, 1985), but the site of modification has not yet been defined. Also, a mechanism by which lipoic acid or its dehydrogenase might be involved in transport phenomena has not yet been suggested, although the cofactor’s lipid solubility and its ability to carry protons may be important factors. We have no information relating to the role of dihydrolipoamide dehydrogenase in archaebacteria nor, in fact, on whether the presumed substrate, lipoic acid, is present in these organisms. The search for lipoic acid is thus a priority and, to this end, we have developed a method using gas chromatography-mass spectrometry to detect lipoic acid as its oxidized lipoate methyl ester (Pratt et al., 1986). Whatever the additional role(s) of dihydrolipoamide dehydrogenase, it is clear that further studies of the enzyme in archaebacteria and T. brucei, where 2-0x0 acid dehydrogenase complexes are not found, will be valuable in identifying any new function(s). D. CITRATE SYNTHASE AND SUCCINATE THIOKINASE

It has already been noted in this review that enzymes of the citric acid cycle may provide a valuable source of comparison between different groups of organisms. Thus, even though there is a fundamental unity associated with the citric acid cycle in that basically the same cellular reactions occur throughout a diverse range of species, it is becoming increasingly clear that enzymes catalysing these reactions possess a very real diversity at the molecular level. This diversity is reflected in their subunit structure, catalytic activity and modes of allosteric regulation and it shows a strong correlation with the taxonomic status of each organism. This has already been illustrated with the pyruvate dehydrogenase complex where, in the few species studied so far, the enzyme from Gram-positive eubacteria closely resembles the complex from eukaryotic mitochondria in arrangement and symmetry of its constituent subunits, but contrasts significantly with the enzyme from Gram-negative eubacteria. Archaebacteria appear not to possess this enzyme complex, and the comparison could not therefore be fully extended to these organisms, although I have compared it with the alternative 2-0x0 acid oxidoreductase system instead. The molecular diversity of the citric acid-cycle enzymes, citrate synthase

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and succinate thiokinase, has been studied in considerable detail in numerous species and, with these enzymes, the comparisons can be extended to archaebacteria. Correlation of structural and functional diversity and its relationship to the taxonomic status of the source organism in eubacteria and eukaryotes are quite remarkable and have been comprehensively reviewed by Weitzman and Danson (1976) and by Weitzman (1981). The details are summarized here so that a more informed comparison can be made with the features found in archaebacterial enzymes.

I . Citrate Synthase and Succinate Thiokinase in Eukaryotes and Eubacteria Citrate synthase catalyses condensation of acetyl-CoA with oxaloacetate to form citrate and so effects entry of carbon into the citric acid cycle. It may therefore be considered as the first enzyme of the cycle and, with the conclusion that it may be the rate-limiting step on the pathway (Krebs and Lowenstein, 1960), its regulatory characteristics have been the subject of extensive examination. A survey of members of eubacterial genera (Weitzman and Jones, 1968) revealed that citrate synthases from Gram-negative species are powerfully and specifically inhibited by NADH, no effect being produced by NAD+, NADP+ or NADPH. Reduced nicotinamide adenine dinucleotide is a primary end product of the citric acid cycle and so the observed inhibitory effect of this metabolite may serve to regulate the flux through the pathway. Within this group of organisms, NADH inhibition of citrate synthases from strictly aerobic bacteria is overcome by the low-energy signal AMP, whereas reactivation by AMP is not observed with the enzyme from facultative anaerobes. Weitzman and Jones (1 968) rationalized this difference on the basis that strict aerobes are absolutely dependent on the cycle for production of energy and, therefore, might require regulation of the pathway’s initial enzyme according to their energetic states. On the other hand, facultatively anaerobic organisms produce energy by fermentation and, accordingly, they control enzymes of those metabolic routes. Inhibition by NADH in both groups of Gram-negative eubacteria is thought to be allosteric in nature, evidence coming from kinetic studies (Harford and Weitzman, 1979, desensitization by chemical modification (e.g. Weitzman, 1966; Senior and Dawes, 1971; Danson and Weitzman, 1973, 1977; Weitzman and Danson, 1976; Talgoy et al., 1979), immunodesensitization (Weitzman, 1981) and desensitization through in uivo mutagenesis (Danson et al., 1979b; Weitzman et al., 1978). In contrast to these effects, citrate synthases from Gram-positive eubacteria and eukaryotes are not allosterically inhibited by NADH but are isosterically inhibited by ATP (Weitzman and Danson, 1976; Weitzman, 1981 and

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21 I

references therein). The inhibition is competitive with the substrate, acetylCoA, and probably arises from their common structural features. Accordingly, other nicotinamide nucleotides inhibit these citrate synthases, the order of effectiveness being NADPH > NADH > NADP+ > NAD+. Direct kinetic evidence for the isosteric nature of ATP inhibition has been obtained (Harford and Weitzman, 1975), although measurements in situ have thrown doubt on its significance in vivo (Weitzman and Hewson, 1973; Matlib et al., 1978). Although allosteric inhibition by NADH is restricted to citrate synthases from Gram-negative eubacteria, one would expect all citrate synthases to exhibit at least some sensitivity to ATP if it merely binds as a consequence of its structural homology to acetyl-CoA. My colleagues and I therefore proposed (Weitzman and Danson, 1976) that the sensitivity of a citrate synthase to ATP might be directly related to the affinity of the enzyme for acetyl-CoA. If one assumes that the K,,, value for acetyl-CoA is a measure of the enzyme’s affinity for this substrate, then our hypothesis is supported by the finding that, for diverse citrate synthases, a linear plot of Ki for ATP versus K,,, for acetyl-CoA is obtained (M. J. Danson, S . Harford and P. D. J. Weitzman, unpublished work). These energetically related controls of citrate synthase are mirrored by a structural diversity at the level of their oligomeric nature. Weitzman and Dunmore (1 969a) revealed the existence of two groups of citrate synthases, ‘‘large’’ and “small”. The ‘‘large’’ enzymes are, without exception, from Gram-negative eubacteria whereas those from the Gram-positive eubacteria and eukaryotes are all of the “small” type. Purification of a number of citrate synthases representative of these groups suggests that the “large” citrate synthases are hexameric (Tong and Duckworth, 1975; Mitchell and Weitzman, 1983) whereas the “small” enzymes from both the eubacteria and eukaryotes are dimeric (Wu and Yang, 1970; Singh et al., 1970; Moriyama and Srere, 1971; Robinson et al., 1983). Despite differences in the sizes of the native citrate synthases, the subunit M , values are relatively constant (40,OOC 50,000). Thus it is thought to be the oligomeric state of the citrate synthase that determines its regulatory sensitivities; this conclusion is supported by our finding that in viuo mutagenesis of the wild-type hexameric, NADH-sensitive enzyme of E. coli can cause it to dissociate into a dimeric form with resultant loss of its allosteric regulations and the gain of isosteric ATP inhibition (Danson et al., 1979b). Mention should also be made of the biosynthetic controls operating on citrate synthases. The enzyme from the facultatively anaerobic Gramnegative eubacteria is inhibited by 2-oxoglutarate (Wright et al., 1967; Weitzman and Dunmore, 1969b). The regulation is probably allosteric, and its physiological significance may result from lack of a 2-oxoglutarate

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dehydrogenase complex in these organisms when they grow anaerobically (Amarasingham and Davis, 1965), resulting in 2-oxoglutarate as the end product of the citrate-synthase arm of the citric acid cycle. Support for this rationale comes from other organisms, some of which are Gram-positive eubacteria, that also lack the 2-0x0 acid complex and whose citrate synthaseis inhibited by 2-oxoglutarate (see Weitzman and Danson, 1976 and references therein). The regulatory and molecular characteristics of citrate synthases are summarized in Table 2. Although the correlation of these parameters with taxonomic grouping of the source organism holds for most of the species investigated, there are exceptions to the pattern. These are discussed by Weitzman (1981) and, in the majority of cases, the deviation from the norm can be interpreted on the basis of the particular metabolic modes of the organisms in question. However, three of the “exceptions” are species of Halobacterium and, as discussed below, they are in fact not exceptions but merely examples of a more extensive pattern which now has to include archaebacteria. Succinate thiokinase catalyses the reaction:

+

+

+

Succinyl-CoA NDP Pi = Succinate NTP+ CoA where NDP and NTP are nucleoside di- and tri-phosphates, respectively. Like citrate synthase it therefore uses the high free energy of hydrolysis of a

TABLE 2. Properties of citrate synthases and succinate thiokinases of eubacteria and eukaryotes Eubacteria Gram-negative Strict aerobes Citrnte synthase Native Mr value Oligomeric nature Inhibitors (a) allosteric

-

290,000 6n

Eukaryotes Gram-positive

Facultative anaerobes

-

290,000 6n

NADH (Reactivation (No reactivation by AMP) by AMP) 2-oxoglutarate (allosteric?) (b) isosteric ATP (Ki > 5mM) ATP (Ki= 1mM) ATP (Ki < 1mM)

Succinate thiokinase Native M,value Oligomeric nature

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thioester acyl-CoA bond but, in this case, for substrate level phosphorylation rather than to synthesize a carbonsarbon bond. This enzyme also exhibits a pattern of molecular sizes (Weitzman and Kinghorn, 1978; Weitzman, 1981) and one which mirrors that of citrate synthase. Gram-negative eubacteria produce a “large” succinate thiokinase ( M r = 155,000) which in E. coli has been found to be an a& structure (Bridger, 1974). In contrast to this, Grampositive eubacteria and eukaryotes possess a “small” enzyme ( M r= 75,000) which in pig heart is an a j dimer (Bridger, 1974). Exceptions to this taxonomic pattern have not been found. The data are summarized along with those for citrate synthase in Table 2.

2. Citrate Synthase and Succinate Thiokinase in Archaebacteria So remarkable and consistent are the patterns of enzymic diversity in eubacteria and eukaryotes that a survey of the properties of citrate synthases and succinate thiokinases was prompted in the archaebacteria (Danson et al., 1985). Molecular sizes were determined by gel filtration on Sephacryl S-200, the column being calibrated with standard globular proteins of known sequence and therefore precisely determined molecular masses. Unfractionated archaebacterial cell extracts were analysed. The previously reported observations (Cazzulo, 1973; Weitzman and Kinghorn, 1983) that species of Halobacterium possesses a ‘‘small” type citrate synthase but a “large” succinate thiokinase were reproduced for both classical (H. halobium, H. volcanii and H. vallismortis) and alkaliphilic (Natronobacterium pharaonis, Nb. gregoryi and Natronococcus occultus) halophilic archaebacteria. The M , values determined (citrate synthase: 105,00&112,000; succinate thiokinase: 170,00&184,000) are both slightly larger than values for the non-archaebacterial counterparts (see Table 2). However, these may not be significant differences as it must be remembered that their determination and calibration of the Sephacryl column with standard proteins were necessarily performed in the respective presence and absence of 4~ NaCl and it is difficult to be sure that the two filtrations are absolutely equivalent. The halophilic citrate synthase has only been partially purified (Higa and Cazzulo, 1975) but it is likely that it will turn out to be a dimeric protein. The finding of a “small” citrate synthase and a “large” succinate thiokinase appears to be a pattern that is characteristic of halophilic archaebacteria and not of any other organism so far investigated except for Thermus aquaticus (Weitzman and Kinghorn, 1983). On the basis of oligonucleotide cataloguing, Thermus aquaticus is not an archaebacterium but groups with Gram-negative eubacteria; its pattern of citrate synthase and succinate thiokinase is therefore a notable exception. In contrast to halophiles, the thermoacidophilic archaebacteria Tp. acido-

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philum and S. acidocaldarius, possess citrate synthases and succinate thiokinases which are both of the “small” type. We have purified the citrate synthases from Tp. acidophilum and S . acidocaldarius to homogeneity and, by sodium dodecyl sulphate (SDS)-polyacrylamide-gel electrophoresis, we have shown them to be dimeric enzymes of subunit Mr=42,000 (M. J. Danson, D. W. Hough, T. Pun, L. Smith and S. Wearne, unpublished work). This combination is therefore precisely that found in Gram-positive eubacteria and eukaryotes and is a similarity supported by regulatory characteristics (see below). We have detected the presence of citrate synthase in Ms. barkeri(Danson et a[.,1985) but the levels were too low to determine an M , value. Succinate thiokinase activity could not be found, consistent with the previous report that these two enzymes have not been found in the same species of methanogen (Weimer and Zeikus, 1979). In addition to determination of molecular sizes, the effects of ATP, NADH and 2-oxoglutarate on the archaebacterial citrate synthases were also investigated (Danson et al., 1985). In contrast to the “small” enzyme from Gram-positive eubacteria and eukaryotes, the halophilic citrate synthases are ; expected, there was no allosteric only weakly sensitive to ATP (K, > 4 m ~ )as inhibition by either NADH or 2-oxoglutarate (see also Cazzulo, 1973, 1978). On the other hand, the “small” citrate synthases from thermoacidophilic archaebacteria were sensitive to ATP in a manner competitive with acetylCoA, results that have been confirmed by Grossebuter and Gorisch (1985). The regulatory behaviour of the methanogenic citrate synthase was not easy to investigate as it is present in low activity. Also, the presence of malate dehydrogenase, at a level 30 times higher than that of citrate synthase, made assays of NADH inhibition difficult to interpret on account of the rapid depletion of the substrate oxaloacetate. Despite these difficulties,we examined the citrate synthase from Ms. barkeri (Danson et al., 1985) and found it unusual in being sensitive to all three metabolites, namely ATP, NADH and 2oxoglutarate. The enzyme is also inhibited by ADP and AMP in the order ATP > ADP > AMP. However, the enyzme was not inhibited by NADPH, suggesting that the effect of NADH is unlike the less specific, isosteric nucleotide inhibition found in “small” citrate synthases. On account of difficulties in the assay referred to above, these data must be viewed with caution and there is still the need to repeat the investigations when the methanogenic citrate synthase can be partially purified. An inspection of the kinetic parameters determined for the archaebacterial citrate synthases reveals the same trend as that reported for the enzyme from eubacterial and eukaryotic organisms, namely that the K, value for ATP is higher the greater the K,,, value for the substrate acetyl-CoA. These data add further support to our hypothesis (Weitzman and Danson, 1976) that the

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general nucleotide inhibition of this enzyme reflects the structural similarity between substrate and inhibitor, the affinity of one being a consequence of the affinity of the other. Succinate thiokinases from all halophiles and thermophiles tested (Danson et al., 1985) and from M . thermoautotrophicum (Fuchs and Stupperich, 1982) were ADP-specific, no activity being detectable with GDP. Weitzman (1 981) reviewed the nucleotide specificity of diverse succinate thiokinases from nonarchaebacterial species and observed that only the enzymes from Grampositive eubacteria and plants are similarly ADP-specific. However, further investigations (P. D. J. Weitzman and T. M. Jenkins, personal communication) reveal a more complex situation. In addition to ADP-linked succinate thiokinase, GDP- and IDP-linked succinate thiokinase activities have also been detected in Gram-positive eubacteria, and preliminary evidence suggests that these activities may reside on enzymes distinct from the ADP-linked thiokinase. This brief survey of citrate synthases and succinate thiokinases in the archaebacteria is summarized in Table 3. It would seem that each of the three phenotypic groups, the halophiles, thermoacidophiles and methanogens, possesses its own distinctive pattern of subunit arrangements and regulatory sensitivities of these enzymes. The patterns found in halophiles and methanogens appear to be unique, whereas those from species of Thermoplasma and Sulfolohus closely resemble the situation found in Gram-positive eubacteria and eukaryotes. There is considerable information on the structure of citrate TABLE 3. Properties of citrate synthases and succinate thiokinases of archaebacteria Halophiles Thermoacidophiles Methanogen Halobacterium species Natronobaclerium species Methanosarcina Natronococcus species Suljblobus Thermoplasma barkeri

Citrate synthase Native M I value Oligomeric nature Inhibitors (K,values) Ki-NADH (mM) Ki-2-oxoglutarate(mM) Ki-ATP( m ~ )

-

Succinate thiokinase Native M, value Oligomeric nature

-

110,000 (2n ?) > 4

> 20 > 4

180,000 ?

83,000 2n

85,000 2n

? ?

4.6 2.2 0.9

5.4 5.4 2.2


ND

-68,000 ?

ND

N D indicates that enzyme activity was not detected. Data in this table were taken from Danson et al. (1985). See also Grossebuter and Gorish (1985) for comparable values for the citrate synthases from the thermoacidophilic archaebacteria.

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synthases from non-archaebacterial organisms, and it would therefore seem a priority to obtain amino-acid sequences of archaebacterial citrate synthases for comparison with the already sequenced enzymes of E. coli (Ner et al., 1983; Bhayana and Duckworth, 1984), Acinetohacter anitratum (H.W. Duckworth, personal communication), Saccharomyces cervisiae (Suissa et al., 1984) and pig heart (Bloxham et al., 198I). Alignment of these sequences shows regions of significant homology, especially near residues thought to be involved in catalysis, the similarity between the eukaryotic enzymes being more extensive than between either of them and the bacterial protein. A citrate synthase from a Gram-positive eubacterium has yet to be sequenced, but immunochemical comparisons (Pullen et al., 1985) revealed it to share structural similarities with the eukaryotic enzyme. In addition to the sequence data, an X-ray structure of citrate synthase from pig heart has been obtained (Remington et al., 1982; Wiegand et al., 1984). Clearly, with this detailed information on citrate synthases and with species-dependent diversity of the enzyme at a well-established stage in eubacteria and eukaryotes, a detailed study of archaebacterial citrate synthases might provide us with an excellent enzymic comparison across all three evolutionary lineages. E. COMPARATIVE ENZYMOLOGY

It was the stated intention of this section on “enzyme diversity” to make comparisons between the three kingdoms. This has necessitated a detailed discussion, not only of archaebacterial enzymes but also of the corresponding enzymes in eubacteria and eukaryotes. As each enzyme has been described, comparisons have been made and it is hoped that the correlations observed will stimulate similar studies in a wider selection of archaebacterial species. Only then can meaningful contributions be made from comparative enzymology to the study of taxonomic relationships of these organisms with those of the other two kingdoms. However, even with the limited information on archaebacterial enzymes, one cannot help but notice the similarity between the enzymology of species of Sulfolohus and Thermoplasma, genera which have been distinctly separated on a phylogenetic basis (see Section LA). These two genera of thermoacidophiles catabolize glucose via a non-phosphorylated Entner-Doudoroff pathway, they possess dual cofactor-specific isocitrate, glucose and malate dehydrogenases, they have 2-0x0 acid oxidoreductases with homologous [4Fe-4S] ferredoxins as electron acceptors and their citrate synthases are indistinguishable on basic catalytic, regulatory and oligomeric criteria. These enzymological features contrast with those of the halophilic and methanogenic counterparts and, given that such characteristics have been used as an aid to eubacterial taxonomy (e.g. see Weitzman, 1981), they

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perhaps should be considered in future discussions on evolutionary relationships within the archaebacteria.

IV. Structure of Archaebacterial Enzymes Archaebacteria grow in extreme environments and therefore their macromolecules will be structurally adapted to function under such conditions. In extreme halophiles, the intracellular concentration of salt is at least equal to that in theenvironment (between 3.5 M and saturation) but is mainly KCI with lower concentrations of NaCl (Larsen, 1981). Thermoacidophilic archaebacteria grow at temperatures between 55 and 110°C and in pH values as low as pH 1-2. Although there can be no temperature differential between the outside and inside of a cell, it is thought that cytoplasmic pH values are approximately neutral. Thus, in a study of archaebacterial enzymes, we are interested in adaptation to high concentrations of salt or extremes of temperatures, factors which, in most other organisms, would lead to denaturation of all classes of biological macromolecule and to cell death. Such an interest is not only of academic or evolutionary importance; with the increased use of enzymes in organic syntheses and other industrial processes (e.g. Battersby, 1985), not only might archaebacterial enzymes be of immense potential in themselves, but a knowledge of the factors conferring their stability might lead to the possibility of engineering such features into “normal” enzymes and proteins. Structural studies on archaebacterial proteins are not at an advanced stage, and of these only a few relate directly to enzymes of central metabolic pathways. In this section the general features conferring halophilicity and thermophilicity will be briefly described and, where possible, illustrated with examples of enzymes to which reference has already been made in this review. A. HALOPHILIC ENZYMES

Halophilic enzymes are not only stable in high concentrations of salt, but actually require them for maintainance of their native structure and for their activity. The structural nature of their adaptation to such extremes of salinity has been reviewed by Lanyi (1974, 1979), Jaenicke (1981), Kushner (1986) Werber et al. (1986) and Eisenberg and Wachtel (1987), and the reader is referred to these publications for detailed accounts and experimental evidence. A number of general features emerge from this extensive literature. In non-halophilic proteins and enzymes, high concentrations of salt disrupt native structures and the effect often follows the order of the Hofmeister series for anions (Hatefi and Hanstein, 1969; Herskovitz et al., 1977). Thus, it has

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been suggested that salt withdraws water from proteins and makes hydrophobic bonds particularly strong, causing polypeptide chains to aggregate and collapse. Halophilic proteins seem to have overcome these potential effects by increasing the frequency of polar amino acids and decreasing that of non-polar residues. This was initially observed with bulk protein of halophilic archaebacteria (Reistad, 1970; Lanyi, 1974), but has since been confirmed with a number of purified individual enzymes (Rao and Argos, 1981) including two of the central metabolic pathways, dihydrolipoamide dehydrogenase (Danson et al., 1986) and malate dehydrogenase (Mevarech et al., 1977). The increased frequency of polar amino-acid residues in halophilic proteins is due to a greater percentage of those with negatively charged side chains, that is, glutamate and asparate. These acidic amino-acid residues increase stabilization of the protein by effectively competing with salt for available water and, thereby, they retain an appropriate hydration layer even at high ionic strengths. Acidic amino-acid residues are the most effective with respect to hydration capacity, as they bind 6.0-7.5 mol water per mol residue, compared with 3.0-4.5 mol per mol for basic residues (Kuntz, 1971).Thus, the increased frequency of glutamate and aspartate residues, rather than arginine and lysine residues, is rationalized. Lysine residue contents of halophilic proteins are characteristically lower than their non-halophilic counterparts and, given that the codons for glutamate are GAA and GAG and for lysine are AAA and AAG, Bayley (1966) speculated that a mutational error in the first adenine base of the lysine codons might lead to higher glutamate and concomitantly lower lysine contents in adaptation to high concentrations of salt. Reistad (1970) points out that alanine residue content of bulk halophilic protein is also consistently low, and envisages similar mutational events in the transition from this amino-acid residue (codons: GCA, GCG, GCC and GCU) to residues of glutamate (GAA and GAG) and aspartate (GAC and GAU). Of course, if archaebacterial phenotypes are more primitive than eubacterial, it is the adaptation to low concentrations of salt that we should be considering. As in all proteins, hydrophobic bonds are crucial for maintenance of the native structure. The lower frequency of non-polar amino-acid residues in halophilic proteins therefore means that high concentrations of salt are required to produce the necessary hydrophobic bonding. Lanyi (1974) suggested that the new hydrophobic bonds created at such ionic strengths involve marginally hydrophobic groups which are normally not well suited for non-polar interaction. Hence, it has been found (e.g. Louis and Fitt, 1971; Mevarech et al., 1977; Danson et al., 1986) that halophilic proteins contain significantly more borderline hydrophobic residues (e.g. alanine and threonine) than the non-halophilic counterparts. It should be noted that the

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increased frequency of alanine residues in these specific proteins is in contrast to the data of Reistad (1970) from bulk proteins. Although these changes in primary sequence in response to high concentrations of salt are obviously of significance to the stablity of halophilic proteins, there will undoubtedly be adaptations in their secondary, tertiary and quaternary structures. Testing this assumption will come with determination of the three-dimensional structures of proteins from extreme halophiles. As yet such information is limited, but the available data do suggest specific conformational adaptations to the salt concentrations (reviewed by Eisenberg and Wachtel, 1987). One of the best characterized halophilic proteins is the citric acid-cycle enzyme, malate dehydrogenase, from Halobacterium marismortui. It has been purified to homogeneity, and by equilibrium centrifugation and SDSpolyacrylamide-gel electrophoresis it has been shown to be a dimeric protein with a native M , value of 84,000 (k4000) (Mevarech et al., 1977). As mentioned previously, amino-acid analysis showed an excess of 10.4 mol% of acidic residues. The enzyme is active at high concentrations of salt ( > 2 M NaCl) (Mevarech and Neumann, 1977) and, from measurements of density increments, it was found that native malate deh,drogenase bound unusually large amounts of water (0.87 g water (g protein)-') and salt (0.35 g NaCl (g protein)-'), far in excess of those usually encountered in non-halophilic macromolecules (Pundak and Eisenberg, 1981). Also, as discussed previously, the water binding, may, in part, be due to the large number of glutamate and aspartate residues in the protein; however, using the data of Kuntz (1971), a maximal hydration of 0.35-0.40 g water (g protein)-' was calculated for malate dehydrogenase on the basis of its amino-acid composition. Thus, it was suggested that special structural features of the enzyme, in addition to its high content of glutamate and aspartate residues, may contribute to the binding of water. This suggestion was supported by the observation that decreasing the concentration of salt causes dissociation of the dimeric enzyme and unfolding of the polypeptide chains, with concomitant loss of water- and salt-binding properties (Pundak et al., 1981). That is, these properties seem to be associated with the intact structure of the enzyme and, when the protein is denatured, the water-binding properties become as expected from the charged nature of its amino-acid composition (Zaccai et al., 1986a). Data from small-angle X-ray and neutron scattering and from ultracentrifugation experiments on the malate dehydrogenase from H. marismortui have been analysed together to yield a model for the enzyme particle formed by the protein and its interactions with water and salt (Zaccai et al., 1986b). The hypothetical model is based on these data and the assumption that the structure around the active site is similar to that of cytoplasmic malate dehydrogenase from pig heart, for which the crystal structure has been solved

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(Hill ef al., 1972; Tsernoglou et al., 1972). The halophilic enzyme particle hasa 1982) and a Stokes’ radiusof radius of gyration of 3.2 nm (32 A) (Reich et d., 4.3nm (43 A) suggesting an involuted surface. By combining the data from the X-ray and neutron-scattering studies, the radius of gyration of the protein moiety alone was calculated to be about 2.8 nm (28 A) and of the associated water and salt distribution to be about 4.0 nm (40 A). Thus, most (but not all) of the protein component must lie at smaller radii and it is proposed (Zaccaiet al., 1986b) that the halophilic malate dehydrogenase has a core similar to that of the eukaryotic enzyme, with about 20% of the protein extending loosely out of the core, forming a large interface with solvent. The calculated relative mass of protein in the core of the halophilic enzyme (80%) is in good agreement with the size differences between the archaebacterial ( M , 84,000) and eukaryotic ( M r 72,000) malate dehydrogenases. One might reasonably speculate that the acidic residues could be concentrated within the extended loops of the enzyme. The primary sequence of the enzyme is not yet known, but such a situation is found in halobacterial ferredoxins (Werber et al., 1986; Eisenberg and Wachtel, 1987). The aminoacid sequences of ferredoxins from H . marismortui and H . halobium have been determined (Hase et al., 1978, 1980) and a comparison with the protein from Spirulina platensis shows that the most striking difference between the sequences is the addition of 22 residues to the N-terminus of the halophilic ferredoxins. This region is rich in aspartate and glutamate residues but lacks lysine and arginine residues. X-Ray crystallographic studies of the ferredoxin from H . marismortui (Sussman et al., 1986) indicate that the N-terminal region may form a separate domain, and low-angle X-ray scattering data support the concept of an extended N-terminus (Eisenberg and Wachtel, 1987).

B. THERMOPHILIC ENZYMES

In mesophilic organisms, macromolecules such as proteins and nucleic acids are denatured by heat. In thermophiles, growing at temperatures in excess of 5 5 ° C such molecules are stable, and the temperature optimum of their enzymes is frequently above the optimum for growth of the organism. However, contrary to original expectations, thermophilic enzymes do not have extensive and pronounced structural features that confer thermostability; rather the differencesbetween them and mesophilic proteins are in general subtle and often difficult to detect (for reviews see Ljungdahl and Sherod, 1976; Singleton, 1976; Perutz, 1978; Zuber, 1979, 1981; Brock, 1985). At present, these differences have been studied only in proteins of nonarchaebacterial thermophiles, but it is highly likely that the principles elucidated are equally applicable to archaebacterial enzymes.

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From comparisons of amino-acid sequences and three-dimensional structures of ferredoxins, haemoglobins and glyceraldehyde 3-phosphate dehydrogenases from mesophilic and thermophilic organisms, Perutz (1978) argues that extra salt bridges on the protein surface are the sources of their greater heat stability. Furthermore, a consideration of relative rates of thermal inactivation reveals that the energy of a few salt bridges is indeed sufficient to protect thermostable enzymes from denaturation by heat. For example, a comparison of the three-dimensional structure of glyceraldehyde 3-phosphate dehydrogenase from the thermophile Bacillus stearothermophilus (Biesecker et af.,1977)with that of the lobster enzyme (Moras et al., 1975) shows that the thermophilic enzyme has salt bridges between the four subunits, but that these interactions are lacking in the mesophilic protein. In kanamycin nucleotidyltransferase, one extra salt bridge can significantly affect the themostability of the enzyme (Matsumura et al., 1984). The gene for the enzyme encoded by a plasmid in B. stearothermophilus was determined and compared with that from Staphylococcus aureus. Only one base difference was found between the two, resulting in substitution of a threonine residue in the mesophilic protein with a lysine residue in the thermophilic enzyme. This lysine residue is involved in increased electrostatic bridging without any significant change in the three-dimensional structure, and is therefore thought to be the cause of the increased thermal stability of the Bacillus enzyme. Brock (1 985) points o u t that salt bridges may not fully explain thermostability and he refers to the work of Yutani et af.(1977) who found that an increase in hydrophobicity in the alpha subunit of tryptophan synthetase from E. cofi greatly increased the stability of the enzyme. A similar point is made by Zuber (1981) from a study of lactate dehydrogenases from a series of thermophilic, mesophilic and psychrophilic bacilli. In addition to a higher proportion of ionic bonds in the thermophilic enzyme, the internal hydrophobicity of the protein is increased by substitution of polar residues (especially serine and threonine) by hydrophobic amino-acid residues. From the point of view of archaebacterial enzymes and proteins, there is the need for determination of their three-dimensional structures from thermophilic species. Malate dehydrogenase from Tp. acidophilum has been crystallized (Grossebuter et al., 1986) and preliminary X-ray diffraction studies have been carried out on ferredoxin (Tsukihara et af., 1985) and superoxide dismutase (Morris et al., 1985) from this same organism. Thus we can expect detailed structures to be available in the near future. In terms of thermophily, archaebacteria span the range from Thermoplasma, which grows optimally at around 55°C to Pyrodictium (Stetter, 1982; Stetter ef al., 1983)and Pyrococcus (Fiala and Stetter, 1986)which can grow at temperatures up to 110°C and 103"C, respectively. The upper temperature limit of life is still undefined but probably lies in the range between 110 and

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150°C (Stetter, 1986). Thermophilic marine bacteria, isolated from the vicinity of waters at 350°C from sulphide chimneys (“black smokers”) along the East Pacific Rise, have been reported to grow at a pressure of 26.85 kPa (265 atm) and a temperature of 250°C (Baross and Deming, 1983). However, Trent et al. (1984) concluded that the evidence indicating bacterial growth at 250°C could be due to artefacts produced in the medium used and to contaminants introduced during sample processing. Baross and Demhg (1984) dispute this claim but it has been demonstrated that, under “black smoker” conditions, biological macromolecules are hydrolysed rapidly and the consequent hydrolysis products such as amino acids are themselves unstable (White, 1984; Bernhardt et al., 1984). Proteins, DNA and ATP all have half-lives of less than one second at these temperatures and pressures, rendering impossible life as we know it. If any organism is to live at temperatures greater than 200”C, then it will have to be composed of some unique macromolecules; archaebacteria have provided us with many surprises, but even they could not be expected to meet this challenge.

V. Concluding Remarks

The concept of the archaebacteria as a phylogenetically distinct group is now established and widely accepted. However, the debate will continue as to their evolutionary relationships to eubacteria and eukaryotes, and to whether they reflect primitive organisms. This article has not attempted to question the phylogenetic position of archaebacteria but has accepted the view that any consideration of evolutionary events must take into account this group of organisms on an equal footing with eubacteria and eukaryotes. Although the majority of biochemical comparisons between these kingdoms has concentrated on molecular biological characteristics, this review has attempted to redress this imbalance by showing that metabolic pathways and their constituent enzymes provide a valuable source of comparative biochemistry. Furthermore, in primitive organisms, it would have been the establishment of such pathways for energy production and biosynthesis that would have been the basis for survival and successful competition during early life on earth. Therefore, a continued and intensive investigation of archaebacterial enzymology is essential to our understanding of cellular evolution.

VI. Acknowledgements I am indebted to Professor David Weitzman (University of Bath) who introduced me to the concept of enzyme diversity and has enthusiastically

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supported my work in this area. Special thanks are also due to my colleagues Dr.David Hough (University of Bath) and Professor Kenneth Stevenson (University of Calgary, Canada) with whom I enjoy close research collaborations, and to Miss Kathy Pratt (Calgary). Financial support for the work of the author from SERC (UK), NSERC (Canada), The Royal Society, The British Council and NATO (in the form of a Grant for International Collaboration in Research with Professor Stevenson) are gratefully acknowledged. Finally, I thank my wife, Janet, for the typing and proof reading of the entire manuscript. REFERENCES

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Physiology of Lipoteichoic Acids in Bacteria W . FISCHER

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I . Introduction . . . . . . . . . . . . . I1 . Occurrence and structure . . . . . . . . . . . A . Poly(g1ycerophosphate) lipoteichoic acids . . . . . . . B. Poly(digalactosy1, galactosylglycerophosphate) lipoteichoic acid . . . C . Glycerophosphate-containinglipoglycan . . . . . . . D . Succinylated lipomannan . . . . . . . . . . E . “Lipoteichoic acid” from Sirepioroccus pneumoniae . . . . . F. Quantitative aspects . . . . . . . . . . . 111. Metabolism . . . . . . . . . . . . . A . Biosynthesis of poly(g1ycerophosphate) lipoteichoic acids . . . . B. Biosynthesis of related macroamphiphiles . . . . . . . C . Influence of lipoteichoic acid biosynthesis on the turnover of membrane lipids . . . . . . . . D . Addition of chain substituents . E . Turnover of o-alanine-ester residues in lipoteichoic acid and transfer to wall teichoic acid . . . . . . . . . . . . F . Conditions affecting the cellular content and synthesis of lipoteichoic acid . G . Conditions affecting chain substitution . . . . . . . H . Degradation and excretion of lipoteichoic acids . . . . . . IV . Cellular location . . . . . . . . . . . . V . Biological activities . . . . . . . . . . . . A . Lipoteichoic acid carrier . . . . . . . . . . B. Interaction with cell-wall lytic enzymes (autolysins) . . . . . C . Interaction with divalent cations . . . . . . . . VI . Concluding remarks . . . . . . . . . . . . VII . Acknowledgements . . . . . . . . . . . . References . . . . . . . . . . . . .

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I. Introduction Teichoic acids (Greek: teichos. wall) were detected in 1958 by Baddiley and his coworkers (Armstrong et al., 1958. 1959) in a search for a role of CDP-

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glycerol and CDP-ribitol which they had earlier discovered in Gram-positive bacteria. The first representatives were isolated from cell walls and shown to be polymers of ribitol phosphate or glycerol phosphate usually bearing glycosyl residues or alanine ester or both. In view of the structural diversity which was recognized in subsequent work, the term teichoic acids was redefined to include all bacterial wall, membrane or capsular polymers containing glycerophosphate or ribitol phosphate residues (Baddiley, 1972). A group of poly(g1ycerophosphate) teichoic acids was differentiated and named “intracellular” teichoic acids because they were extracted from whole cells but, unlike previously studied teichoic acids, absent from cell walls (McCarty, 1959; Critchley et al., 1962). Later on, intracellular teichoic acids were located in the cytoplasmic membrane and were therefore renamed “membrane teichoic acids” (Shockman and Slade, 1964). A third change of name, to lipoteichoic acids, was finally suggested when eight years later “teichoic acid lipid complexes” were detected (Wicken and Knox, 1970) and subsequently recognized as amphiphiles in which the poly(g1ycerophosphate) is covalently linked to a membrane glycolipid by a phosphodiester bond (Toon et al., 1972). Thus, lipoteichoic acids are, as amphiphiles, anchored in the cytoplasmic membrane by hydrophobic interaction, whereas teichoic acids are covalently linked through a particular linkage unit to the peptidoglycan of the cell wall (Coley et al., 1978; Ward, 1981; Hancock and Baddiley, 1985). Many Gram-positive bacteria contain both polymers, but lipoteichoic acids are more widespread and their synthesis has been considered to be less dependent on growth conditions (Ellwood and Tempest, 1972a; Wicken and Knox, 1975a). Teichoic acids and lipoteichoic acid are structurally and metabolically unrelated: Staphylococcus aureus for example contains a poly(D-alanyl,cr,fi-Nacetyl-D-glucosaminyl ribitol phosphate) teichoic acid and a poly(D-alanyl glycerophosphate) lipoteichoic acid. Even if both polymers, such as in Bacillus subtilis Marburg, possess a poly(g1ycerophosphate) chain, their biosynthetic pathways are different. The glycerophosphate residues of lipoteichoic acids are derived from phosphatidylglycerol, those of teichoic acids from CDPglycerol, and accordingly the glycerophosphate residues of these polymers have enantiomeric stereochemical configurations (Fig. 1). More recently “lipoteichoic acids” have been discovered that possess poly(hexosylg1ycerophosphate) and glycerophosphate-carrying lipoglycan structures. However, in spite of these structural variances, these amphiphiles are related to the classical poly(g1ycerophosphate) lipoteichoic acids in so far as they share with them the same biosynthetic origin and stereochemical configuration of their glycerophosphate residues. Reviews on several aspects of teichoic acids and lipoteichoic acids have appeared (Archibald and Baddiley, 1966; Baddiley, 1972; Knox and Wicken,

235

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

I

H b CIOH

8 H 2 C-O--F;-OH I

0-

(a1

B

CH2OH

CH2OH ~

I

I

0 H~ II

HZC-0-P-OH I

0-

~

R

CH20H

I QC - O b Y2-o-CHmCdOH R- 4 C4H I

?

H 2 C-0-P-0-C I

(b)

II

H2

0-

(C)

FIG. 1. Stereochemical configuration of glycerophosphates. sn-Glycero-I-phosphate (a) is present in lipoteichoic acids and glycerophosphoglycolipids (Fischer, I98 1 ), sn-glycero-3-phosphate(b) in CDP-glycerol, wall teichoic acids, their linkage units (for review, see Ward, 1981) and phosphatidylglycolipids (Wilkinson and Bell, 1971; Fischer et al., 1973b). Phosphatidylglycerol (c) contains both enantiomers, the sn-3-isomer in acylated and the sn-I-isomer in non-acylated form. It can therefore serve in biosyntheses as a donor of either phosphatidyl or sn-glycero- 1 -phosphate residues.

1973; Archibald, 1974; Wicken and Knox, 1975a, 1980; Lambert et al., 1977a; Ward, 1981; Fischer, 1987b). Methods used in the preparation and structural analysis of teichoic acids have been detailed in reviews by Archibald and Baddiley (1966) and Fischer (1988). For immunological properties and activities of lipoteichoic acids in mammals the reader is referred to the reviews by Wicken and Knox (1975a, 1980) and Fischer (1988). 11. Occurrence and Structure A. POLY(GLYCEROPHOSPHATE) LIPOTEICHOIC ACIDS

The predominant type of lipoteichoic acids contains a 1,3-1inked poly(g1ycerophosphate) chain attached by a phosphodiester bond to a glycolipid or phosphatidylglycolipid which, in the free state, occur as characteristic membrane lipids in Gram-positive bacteria (Fischer, 198I; Ishizuka and Yamakawa, 1985). Likewise characteristic of Gram-positive bacteria are glycerophosphoglycolipids which are short-chain homologues of lipoteichoic acids carrying a sn-glycero-1 -phosphate residue on the glycolipid in place of the poly(g1ycerophosphate) chain. They are usually minor membrane components and may be considered intermediates in biosynthesis of lipoteichoic acids. The structural relationship between glycolipids, glycerophosphoglycolipids and lipoteichoic acids is illustrated in Fig. 2, using Staph. aureus as an example. A comprehensive survey of the distribution of poly(g1ycerophosphate) lipoteichoic acids among Gram-positive bacteria has in retrospect been

H2COH

I I

HbC4OH

(c)

9-

n

T2 HC+-C-R

I

H2-CC-R

FIG. 2. Structural relationship between glycolipids (a), glycerophosphoglycolipid (b) and lipoteichoic acid (c) in Staphylococcus aureus (X, 70% malanine ester, 30% H; n = 25-29). The same set of compounds has been found in Bacillus licheniforrnis and Bacillus subtilis. For references see Fischer (198 1).

provided by McCarty (1959) in a serological search for extractable poly(glycerophosphate) antigen. This study also revealed that pneumococci, clostridia, corynebacteria, group-0 streptococci and Gram-negative bacteria lack the antigen consistently. Among the large group of non-haemolytic and viridans streptococci, both positive and negative strains were encountered. Since then, the occurrence of poly(g1ycerophosphate) lipoteichoic acids has been confirmed for many taxa by isolation and characterization (Knox and Wicken, 1973; Wicken and Knox, 1975a; Fischer et a]., 1980a,b; Fischer, 1981). 1, Lipid Anchor

The lipid moiety renders lipoteichoic acids amphiphilic and serves to anchor them in the cytoplasmic membrane by hydrophobic interaction. Since glycolipids differ in structure among Gram-positive bacteria, usually in a genus-specific manner (Shaw and Baddiley, 1968; Shaw, 1974), the glycoiipid moieties of lipoteichoic acids which are derived from them vary accordingly. Their structures and occurrence are listed in Table 1. As can be seen,

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

231

TABLE I . Lipid anchors of lipoteichoic acids Lipid anchor Ia

Ia+Ib

Ia+Ic

Occurrence Glc(a 1-2)Glc(a 1-3)acylzGro Streptococcus pyogenes (Streptococcus , mesenterhemolyticus D-58)"Leuconostoc oide8 Glc(al-2)Glc(a 1-3)acyl2Gro Luctococcus lactif , Lactococcus garvieae Glc(a 1-2)Glc(a 1-3)acylzGro ("Streptococcus lactis Kiel 421 72")', Lac~ O ~ O C C UrafinoIactiP,k, S Lactococcus pfan6 tarumd.' Lacy1 Glc(a 1-2)Glc(a 1-3)acyl2Gro Enterococcusfaecalisb,eg

Glc(al-2)Glc(al-3)acyl2Gro 6

I1

+

IIIa IIIb

IIIa + IIIc

+

Ptd Glc(B1-6)Glc(B1-3)acylzGro Bacillus licheniformid'.'. Bacillus subtilis',', Bacillus pumilus', Micrococcus variansb, Staphylococcus aureuf,', Staphylococcus xylosusk Gal(a I-Z)Glc(a 1-3)acyhGro Lactobacillus fermentum"'. "Streptococcus Gal(al-2)Glc(al-3)acyl~Gro tactis motile" d,k 6 acyl Gal(aI -2)Glc(al-3)acyl2Gro Listeria monocytogenes and other Listeria Gal(a 1-2)Glc(a 1-3)acylzGro species".",P 6 Ptd

IVa IVb

V

Glc(~I-6)Gal(aI-2)Glc(a1-3)acylzGro Lactobacillus caseF, Lactobacillus helvetiGlc(B1-6)Gal(a1-2)Glc(a 1-3)acylzGro cus', Lactobacillus plantarum' 6 Lacy1 acyl2Gro Bacillus coagulans', Bacillus megaterium'. mutant strain of Bacillus licheni$ormis'

References: "Fischer el al. (1980a); hFischer et al. (1981); 'Koch and Fischer (1978); dSchleifer et al. (1985); 'Toon et al. (1972); 'Ganfield and Pieringer (1975); ZFischer et al. (1983); "Button and Hemmings (l976a); 'Iwasaki et al. (1986); W . Fischer (unpublished observations); 'Duckworth et al. (1975); W . Fischer and T. Fleischmann-Sperber (unpublished work); "Hether and Jackson (1983); "Ruhland and Fiedler (1987); Wchikawa et al. (1986); qNakano and Fischer (1978); 'Fischer et al. (1980b); ,'Button and Hemmings (1976b).

'

dihexosyldiacylglycerolsare prevalent, trihexosyldiacylglycerolsoccur occasionally, whereas monohexosyldiacylglycerols have not yet been detected as lipoteichoic acid constituents, although they are consistently present in membranes as the biosynthetic precursor of dihexosyldiacylglycerols. In a number of lipoteichoic acids in addition to the glycolipid a derivative of it has been found (Table 1) that carries either a third fatty acid ester (Ib, IIIb, IVb) or a sn-3-phosphatidyl residue (Ic, IIIc). Third fatty-acid and phosphatidyl

238

W. FISCHER

FIG. 3. Space-filling model of GroP-6Glc(~1-6)Gal(a1-2),acyl-6Glc(a I-3)acy12Gro from Lactobacillus casei showing a glycolipid moiety with three fatty-acyl chains.

residues are generally bonded to C-6 of the inner diacylglycerol-linked hexosyl residue which, as illustrated in Fig. 3, leads to a coherent hydrophobic moiety. Molecular species with three or four fatty-acyl residues usually constitute less than 50% of the total lipoteichoic acid. The third and fourth fatty-acyl residue may be expected to anchor these species more firmly in the membrane because the critical micellar concentration of lipid amphiphiles decreases with increasing number of aliphatic carbon atoms (Tanford, 1980; Marsh and King, 1986). A non-glycosylated diacylglycerol residue as the lipid anchor was for the first time found in the lipoteichoic acid of a mutant strain of Bacillus licheniformis which, lacking phosphoglucomutase, was unable to synthesize UDP-glucose and accordingly glycolipids (Button and Hemmings, 1976b). Recently a diacylglycerol residue as the lipid anchor of lipoteichoic acid has also been observed in wild-type strains of Bacillus coagulans and Bacillus megaterium (Iwasaki et al., 1986). It is of interest that B. megaterium does not naturally contain glycolipids in its membrane (W. Fischer, unpublished observation), while the membrane lipids of B. coagulans remain to be studied

239

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

in this respect. The presence of fatty-acyl residues on glycerol residues of the poly(g1ycerophosphate)chain as suggested earlier (Wicken and Knox, 1975a) has not been confirmed. Variable amounts of deacylated lipoteichoic acid have often been reported to accompany fully acylated lipoteichoic acids in phenol-water extracts from bacteria. This may, however, be an artifact produced during preparation by the action of endogenous lipases (Kessler and Shockman, 1979b; Kessler et al., 1979). Phenol-water extraction of lipoteichoic acids from a range of Gram-positive bacteria in my laboratory usually yielded none or negligible amounts of partially or fully deacylated lipoteichoic acid, if all operational steps prior to extraction were done immediately after harvesting at low pH values and low temperatures (Fischer et al., 1980b). As shown in Table 2, the fatty-acyl composition of lipoteichoic acids is usually quite similar to that of total cellular lipids, thus reflecting the origin of the lipid moiety from membrane lipids. If residues of cis- 1 1,12-methylenoctoTABLE 2. Percentage fatty-acyl composition of Iipoteichoic acids and membrane lipids in various Gram-positive bacteria Fatty-acyl composition (per cent of total) c140 cl4 I c l 6 0 cl6 I CIS0 CIS I Cl9c; Enterococcus faecalis NCIB 8 191" Lipoteichoic acid Total lipids Lactococcus lacfis NCDO 712 Lipoteichoic acidb Total lipids' Lactococcus garvieae Kiel42172 Lipoteichoic acidb Total lipids' Micrococcus varians ATCC 29750h Lipoteichoic acid Total lipids Bijidobacterium bijidum DMS 20239d Macroamphiphile Total lipids

5.7 3.4

-

37.7 37.7

8.7 7.5

-

40.0'

12.6 7.7

-

0.4

30.3 21.7

3.3 3.9

1.9 2.3

35.1' 16.9 24.V 40.9

11.4 4.5

0.5 0.5

33.5 28.5

4.5 2.8

2.9 4.7

43.2f 2.1 4 8 . 9 10.6

26.9 25.6

1.6 1.9

15.7 12.5 7.1 22.4

-

43.@ 42.5g

6.5 4.6

-

33.5 38.7

0.9 1.2

7.9

35.w 15.5

13.6 12.7

References: "Toon et al. (1972); hW. Fischer (unpublished work); 'Schleifer "Fischer (1987a). "Cis-II ,12-methylene-octadeanoic acid. 'Predominantly cis-A1 1-0ctadecenoic acid. goctadecenoic acid with unusual, unidentified location of the double bond. Predominantly cis-A9-octadecenoic acid.

ct

a!. (1985);

240

W.FISCHER

decanoic acid are present, their content is usually higher in membrane lipids than in lipoteichoic acids at the cost of cis-All-octadecenoic acid, its biosynthetic precursor (Law, 1971; Golfine, 1972). Since cyclopropanization of unsaturated fatty-acyl residues occurs at the level of completed lipids (Thomas and Law, 1966; Cronan, 1968) and towards the end of logarithmic growth (Jungkind and Wood, I974a, b; Obermeier, 1978), the observed differences suggest that lipoteichoic acid might be less accessible to cyclopropane synthetase, possibly due to its location in the outer layer of the membrane (see Section IV). 2. Chain Structure

Poly(g1ycerophosphate) lipoteichoic acids are generally believed to contain a single unbranched chain linked by a phosphodiester bond to the glycolipid moiety, although these structural details have been definitely proved in a few structures only. Structural studies on glycerophosphoglycolipids, their shortchain homologues, have directly shown the presence of a phosphodiester linkage and further suggested that the poly(g1ycerophosphate) chain may be linked consistently to C-6 of the non-reducing hexosyl terminus distal to the diacylglycerol moiety (Fischer et al., 1973a, 1978c; Fischer and Landgraf, 1975; Laine and Fischer, 1978). The sn-glycero-1-phosphate configuration of the glycerophosphate residues (see Fig. 1) suggested by their biosynthetic origin from phosphatidylglycerol (Section 111. A), has been chemically proved for glycerophosphoglycolipids (Fischer et al., 1973a, 1978c) and the terminal glycolipid-linked glycerophosphate residue of lipoteichoic acids (Fischer, 1981). In addition to variations in glycolipid structure, species and genus variations may concern the length of the poly(g1ycerophosphate) chain and the nature of substitution on C-2 of its glycerol residues, as is shown in Table 3. With a single unbranched chain on the glycolipid, the chain length varies between about 16 and 40 glycerophosphate units, being fairly constant for different strains of a given species. According to chain substitution four groups of lipoteichoic acids can be distinguished (Table 3). The first group lacks any substituent, the second and third groups carry D-alanyl ester or glycosyl residues, and the fourth group contains both of these substituents. So far, no amino-acid residue other than D-alanine has been detected, while the glycosyl substituents comprise a narrow range of common monosaccharide residues. Most lipoteichoic acids carry monohexosyl substituents. Mono-, di-, tri-, and tetrahexosyl residues may occur concomitantly on lipoteichoic acids of Enterococcusfaecalis strains. As can be seen from Table 3, among these strains there are remarkable differences in the degree of glycosylation of the poly(g1ycerophosphate) chain as well as in the number of glucosyl residues of

TABLE 3. Substitution and chain length of poly(g1ycerophosphatc) lipotcichoic adds

Molar ratios to phosphorus Lipoteichoic acid from: Micrococcus varians, ATCC 29750 Bacillus megaterium, various strains

D-Alanine none none

Glycosyl substituents” none 0.05

32k4 20-25

1, 5

a-GlcNAc6

none none none none none none 0.03-O.10

40 24 25 29 27 28 25-30

3.4, 5 6 7,8 9 9 5 7, 10, I 1

a-Gal a-Glc14c

0.4M.43 0.82

17 20

1, 12

a-Gal

Lactobacillus casei, 7469,DSM 20021 Lactobacillus helveticus, DSM 20075

0.56-0.63 0.67 Streptococci group A 0.47-0.56 Lactococcusplantarum, NCDO 1869 0.45 Lactococcus raflnolactis, NCDO 617 0.52 Staphylococcus xylosus, DSM 20266 0.294.40 Stuphylococcus aureus H,DSM 20233 0.30-0.87

(GroP). References

2

~

Bacillus coagulans, various strains Enterococcusfmcalis, ATCC 9790

none none

Enterococcusfaecalis. Kiel 27738 0.48 a-Glcf 0.47 0.45 Enterococcusfaecalis, DSM 20478 0.23 a-Glc$ Enterococcusfaecalis, NCIB 8191 a-Glc2,~‘ > 0.90 0.88 Enterococcusjaecalis, NCIB 39 0.41 a-Glcl.2‘ > 0.90 Lactococcus lactis, various strains 0.10-0.50 a-Gal 0.174.60 Listeria monozytogenes, ATCC 153 I3 0.17 0.31 a-Gal Listeria, various species 0.21-0.36 a-Gal 0.114.34 Bacillus lichenijormis, DSM 13 0.51 a-Gal, a-GlcNAc 0.02,0.18 Bacillus lichenformis, AHU 1371 0.69 a-GlcNAc 0.15 Bacillus subtilis, W 23 0.40 a-Glc, a-GlcNAc 0.20,0.21 Bacillus subtilis, various strains 0.35-0.55 a,fi-Glc, a-GlcNAd‘ 0.04,0.214.43

19k 1 19k I n.d.‘ n.d. 19-26 23 16-33 27 28 24 25-33

2 1, 5

5 13 13 I , 6.9 14, 15

14 5

2 16 2

“Glycosyl residues, as so far studied, belong to the D-series and are in the pyranose form. *Absent from Staphylococcusaureus H gol-’ aR(W. Fischer, unpublished work), a mutant lacking the usual Nacetylglucosaminylsubstituents on the wall teichoic acid (Heckels et al., 1975). CStructures: a-Glc(1-; a-Glc( I-Z)a-Glc(I-; a-Glc(l-2)a-Glc(l-2)a-Glc( I-; a-Glc(l-2)a-Glc(l-2)a-Glc(l-2)aGlc( 1-. dNot present in all strains, while the anomeric form is strain-specific. References: I , Fischer et al. (1981);2, lwasaki et ul. (1986);3 , Kelemen and Baddiley (1961);4. Nakano and Fischer (1978);5, W. Fischer (unpublished work); 6,Fischer et a / . (1980b); 7, Fischer et a / . (1980a);8, McCarty (1964);9.Schleiferet al. (1985);10, Rajbhandari and Baddiley (1963);I I . Fischer and Rose1(1980);12,Cabacungdn and Pieringer (1985);13. Wicken and Baddiley. (1963);14,Ruhland and Fiedler (1987);15, Uchikawa el u/. (1986); 16. Fischer and Koch (1981).

242

W. FISCHER

each substituent. In a few bacteria the glycosyl substituents are group antigens (for a review see Wicken and Knox, 1975a). The presence of non-substituted glycerol residues in almost all lipoteichoic acids (Tables 3 and 4) raises the question as to whether these lipoteichoic acids are mixtures of substituted and non-substituted chains or whether all chains are partially substituted. Anion-exchange chromatography on columns of DEAE-Sephacel separates molecular species of lipoteichoic acid in the order of increasing negative charge, i.e. in the order of decreasing alanine-ester content (Fischer and Rosel, 1980). Using this procedure, it could be shown that unsubstituted species are absent from the lipoteichoic acid of Staph. aureus and that all molecular species are alanylated within a narrow range (Fischer and Rosel, 1980). Lectin-affinity chromatography specific for a-D-galactopyranosyl residues likewise demonstrated the absence of nongalactosylated species from the lipoteichoic acid of Lactococcus lactis (Wicken and Knox, 1975b). A means to study the distribution of alanyl residues along the chain was provided by the discovery of a phosphodiesterase (Schneider and Kennedy, 1978) which, along with a phosphomonoesterase, degrades Dalanyl lipoteichoic acid stepwise from the terminus distal to the lipid moiety (Childs and Neuhaus, 1980; Fischer et al., 1980b). Using this procedure with the lipoteichoic acid from Staph. aureus, it was shown that each third of the chain had the same D-alanine-phosphate ratio. This revealed a homogeneous distribution of alanyl ester substituents along the chain suggesting either a regular or a random arrangement (Fischer et al., 1980b). "P NMR Spectroscopy which, as recently discovered, provides a considerable amount of information on chain substitution (Batley et al., 1987) indicates a random rather than a regular distribution of alanine ester substituents in the lipoteichoic acid of Staph. aureus (W. Fischer and W. Bauer, unpublished TABLE 4. Chain composition of substituted poly(g1ycerophosphate) lipoteichoic acids".From Fischer and Koch (1981) Species: Bacillus subtilis

Grob AlaGro GlcGro GlcNAcGro

Lactococcus lactis

Enterococcus faecalis

W-23

Marburg

NCDO 712

Kiel27738

0.23 0.42 0.20 0.21

0.28 0.38 0.17 0.18

0.31 Grob 0.21 AlaGro GalGro 0.39 AlaGalGro 0.09

Grob AlaGro GlczGro AlaGlc2Gro

0.23 0.29 0.29 0.19

'Measurement after hydrolysis with 40% (w/w) aqueous hydrogen fluoride (Fischer et al. 1980b, 1981); values quoted are molar ratios to phosphorus. 'Non-substituted glycerol.

PHYSIOLOGY OF LIPOTElCHOIC ACIDS IN BACTERIA

243

observations) and Lactobacillus fermentum (Batley et al., 1987). The glycosyl substituents in the lipoteichoic acid of Ent. faecalis are also randomly distributed. Two or three different substituents on one lipoteichoic acid (Table 3) may be linked, as shown in Table 4, either individually to separate glycerol residues or, as with alanine ester residues, be in part attached to glycosyl substituents. The high degree of glycosylation of the lipoteichoic acids from Ent. faecalis strains NCIB 8191 and NCIB 39 suggests that most of the alanine ester is bonded to glycosyl substituents (Table 3). That hexosyl and alanyl substituents occur on the same rather than on separate chains was demonstrated with the lipoteichoic acid from L. lactis. It did not separate on DEAESephacel into species containing either D-alanyl ester or galactosyl residues and, on rechromatography, after removal of the alanyl substituents, unsubstituted species did not appear (W. Fischer, unpublished observations). B. POLY(DIGALACTOSYL,GALACTOSYLGLYCEROPHOSPHATE)LIPOTEICHOIC ACID

An unusual structure was detected in the lipoteichoic acid from Lactococcus garvieae (Fig. 4). Digalactosyl residues are intercalated between the glycerophosphate residues, while the glycerophosphate residues are consistently substituted at C-2 with monogalactosyl residues (Koch and Fischer, 1978; Schleifer et al., 1985). As in other species of Lactococcus the lipid anchor is Glc(cr 1-2)Glc(a1-3)acyl2Gro and the 6-0-acylated derivative thereof (see Table 1). The chain structure is reminiscent of poly(hexosy1 glycerophosphate) wall teichoic acids (Archibald, 1974) but differs from them by containing sn-glycero-1-phosphate residues (Koch and Fischer, 1978). C. GLYCEROPHOSPHATE-CONTAINING LIPOGLYCAN

The lipoteichoic acid from Bijidobacterium bifidum was originally reported to consist of two 1,2-linked poly(g1ycerophosphate) species, one carrying a glucan, the other a galactofuranan on the glycerol terminus of the poly(g1ycerophosphate) chain distal to the lipid moiety (Op den Kamp et al., 1984). Recent studies suggested the structure depicted in Fig. 5 (Fischer, 1987; Fischer et al., 1987). In this structure, the amphiphilic chain is a linear lipoglucogalactofuranan and the glycerophosphate residues are not intercalated into the chain but are attached as monomeric side branches to the galactofuranosyl residues. As in poly(g1ycerophosphate) lipoteichoic acids, the glycerophosphate residues have the sn- 1-configuration and are in part substituted by alanine esters which, however, have the L-configuration in contrast to the D-alanyl residues of poly(g1ycerophosphate) lipoteichoic acids. The lipid anchor was tentatively identified as /?-~-Galp(1-3)acylzGro and may

245

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

CH20-X

I

HbCIOH

I

9

H 2 C-0-7-0-

- X = Ala ( 2 0 - S o 0 / ~ )- ,H

[

‘i

0-H- -0-C-H uH2] ~ - ~ - - @ 3 H 2

3 -0-C-R H2 -0-5-R

0 FIG. 5. Structure of a lipoglycan from Bijidobacterium bijidurn DSM 20239 bearing sn-glycero-1-phosphate side chains partially substituted with L-alanyl ester. n, 7-1 0; m,8-1 5.

therefore be derived from one of the membrane galactoglycolipids from this organism (Veerkamp, 1972). In contrast to poly(g1ycerophosphate) lipoteichoic acids, in this novel amphiphile the glycerophosphate residues are kept a certain distance from the membrane by the glucan moiety. Actinomycetes and Streptococcus sanguis biotype B lack serologically detectable poly(g1ycerophosphate) lipoteichoic acids (Hamada et al., 1976, 1980). In place of them they seem to contain amphiphilic heteropolysaccharides whose structures have not yet been unravelled (Wicken et al., 1978; Yamamoto et al., 1985). Small amounts of possibly monomeric glycerophosphate residues may be present. D. SUCCINYLATED LIPOMANNAN

Micrococcus luteus, MicrococcusJlavus and Micrococcus sodonensis also lack serologically detectable lipoteichoic acids, but instead possess a succinylated lipomannan (Powell et al., 1974, 1975; Owen and Salton, 1975a; Pless el al., 1975; Fig. 6). The hydrophilic moiety contains 50-70 (1-2)-, (1-3)-, and (1 -6)linked a-D-mannopyranosyl residues and two 2,4-substituted branch points. Between 10 and 25% of the mannosyl residues are substituted with ester-linked succinate. A neutral lipomannan lacking succinyl residues has been isolated from Micrococcus agilis (Lim and Salton, 1985). As shown with the lipomannan from M . luteus, the lipid moiety is diacylglycerol (Powell et al., 1975), possibly as part of the sequence Man(crl-3)Man(crI-3)acyl~Gro which is the major membrane glycolipid in M . luteus (Lennarz and Talamo, 1966). A functional analogy of lipomannan to lipoteichoic acids has been proposed in view of their amphiphilic nature, membrane localization, net negative charge and similar Mg2+-binding properties (Powell et al., 1975; Wicken and Knox, 1980). The glycerophosphate-containing lipoglycan of BiJidobacterium bifidum (Section I1.C) may be considered a structural link between lipomannans and lipoteichoic acids.

246

( k)

Ma nnosylfmannosyl A I

T

W . FISCHER

(

H

CH20H

w d 0 - C H 2

/\ 0 0- -10

I HC-0-CO- R

I

H,C-0-CO-R

FIG. 6. Structure of the succinylated lipomannan from Micrococcus luteus. E. “LIPOTEICHOIC ACID” FROM

Streptococcus pneumoniae

Streptococcus pneumoniae strains possess in place of a poly(g1ycerophosphate)-type lipoteichoic acid a unique macroamphiphile, the pneumococcal Forssman antigen, also called pneumococcal lipoteichoic acid. Although known since 1943 as “lipocarbohydrate” (Goebel et al., 1943) and during the last decade intensively studied for biological activities (Section V.B), the structure of this polymer has not been unravelled. It contains fatty-acyl residues (5.7-6.5%), ribitol phosphate, galactosamine, glucose and choline phosphate (Goebel et al., 1943; Fujiwara, 1967; Brundish and Baddiley, 1968; Briles and Tomasz, 1973). With the exception of the fatty-acyl residues the composition is similar to that of the pneumococcal wall teichoic acid whose structure has been established as shown in Fig. 7. In spite of the similarity in

iH A C

FIG. 7. Structure of the repeating unit of the wall teichoic acid from Sfreptococcus pneumoniae (Jennings et al., 1980).The pneumococcal Forssman antigen (pneumococcal “lipoteichoic acid”) has a similar composition but, in addition, contains fatty-acyl esters on a non-identified lipid moiety.

PHYSIOLOGY OF LlPOTElCHOlC ACIDS IN BACTERIA

241

composition, lipoteichoic acid is not a precursor of this wall teichoic acid as was deduced from pulse-chase experiments using radiolabelled choline (Briles and Tomasz, 1975). F. QUANTITATIVE ASPECTS

Earlier estimates of the cellular content of lipoteichoic acids may have yielded too low values, because difficulties in quantitative extraction of lipoteichoic acids from bacteria have not been recognized until recently (Fischer and Koch, 1981; Huff, 1982; Fischer et al., 1983). Values between 1 and 3% of the cell dry weight have been reported, and about 2-3% may be calculated from the estimate of 120 pmol lipoteichoic acid phosphorus or glycerol per gram dry weight (Wicken et al., 1973; Fischer et al., 1983). For certain bacteria the content may vary depending on growth conditions as will be discussed in Section III.F.5. In Staph. aureus, grown in batch culture, lipoteichoic acid, teichoic acid and nucleic acids contribute 13,29 and 58%, respectively, to total polymer phosphorus (Fischer et al., 1983). Taking the different chain lengths of teichoic acid and lipoteichoic acid into account, their molar ratio is approximately 1.5. Estimates of lipoteichoic acids and membrane lipids suggest that, in the membrane of L. lactis and Staph. aureus, lipoteichoic acids represent every tenth and twentieth lipid amphiphile molecule, respectively (Fischer, 1981; Koch et al., 1984). Having long chains, lipoteichoic acids nevertheless represent approximately 50% of the membrane amphiphile glycerol in Staph. aureus (Koch et al., 1984), 40% in spores of B. megaterium (Johnstone et al., 1982) and 30% in Ent. faecalis (Carson et al., 1979; Shungu et al., 1980).

111. Metabolism A. BIOSYNTHESISOF POLY(GLYCEROPHOSPHATE) LIPOTEICHOICACIDS

In 1974, in vivo pulse-chase experiments by Glaser and Lindsay and by Emdur and Chiu provided suggestive evidence that, in Staph. aureus and Strep. sanguis, the glycerophosphate required for lipoteichoic acid biosynthesis is derived from phosphatidylglycerol. Support for the donor function of phosphatidylglycerol was obtained by in vitro experiments using membrane preparations of Strep. sanguis (Emdur and Chiu, 1975; Mancuso et al., 1979), Ent. faecalis (Ganfield and Pieringer, 1980) and toluene-treated cells of Lactobacillus casei (Childs and Neuhaus, 1980). CDP-Glycerol containing snglycero-3-phosphate could not substitute for phosphatidylglycerol, as was shown with particulate enzyme preparations from Enr. faecalis (Pieringer et

248

W. FISCHER

al., 1981). In experiments with B. subtilis, inhibition of biosynthesis of phosphatidylglycerol with 3,4-dihydroxy-butyl- 1-phosphonate, an analogue of sn-glycero-3-phosphate, resulted, as expected, in a block in lipoteichoic acid synthesis (Deutsch et al., 1980). In pulse-chase experiments in which growing cells of Staph.aureus (Koch et al., 1984)and B. subtilis (Koga et al., 1984)were labelled with [3H]glycerol, phosphatidylglycerol was isolated and, after hydrolysis with moist acetic acid or phospholipase C, the labelling pattern of the diacylglycerol and glycerophosphate moieties was studied separately (Fig. 8). A rapid and virtually complete turnover of the non-acylated glycerol moiety into lipoteichoic acid was observed and, when the phosphate group was labelled, it showed an analogous behaviour (Koga et al., 1984). A promising novel system for studying details of lipoteichoic acid synthesis is afforded by membrane vesicles released from L. casei by treatment with penicillin (Ntamere and Neuhaus, 1987). These vesicles catalyse incorporation of label from ['4C]glycero-3-phosphate and UDP-['4C]glucose into poly(g1ycerophosphate) and glycolipid. The mode of chain elongation was elucidated by differential radioisotope

FIG. 8. Turnover of the non-acylated glycerophosphate moiety of phosphatidylglycerol into lipoteichoic acid in growing Staphy[ococcus aureus in a pulse-chase experiment using [2-3H]glycerol; the arrow indicates the onset of the chase. Symbols: 0, lipoteichoic acid; A , glycerophosphate moiety and 0 , diacylglycerol moiety of phosphatidylglycerol released by treatment with phospholipase C and separated by phase partition (Koch et al., 1984).

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

249

labelling techniques. In Ent.faecalis, lipoteichoic acid was pre-labelled in vivo with ['4C]glycerol and subsequently elongated in vitro using particulate (Cabacunenzyme preparations and [I (3)-gly~erol-~H]phosphatidylglycerol gan and Pieringer, 198I). In Lb. casei, [2-gly~erol-~H]lipoteichoic acid, formed in growing bacteria, was elongated in toluene-treated cells using ['4C]glycero3-phosphate as the second label (Taron et al., 1983). The lipoteichoic acids were isolated and degraded stepwise from the glycerol terminus distal to the lipid moiety by the joint action of phosphodiesterase and phosphomonoesterase from Aspergillus niger. In both polymers the ratio of the two isotopes in the glycerol released indicated chain growth distal to the lipid anchor (Fig. 9). This mode of chain growth is identical with that for wall teichoic acids (Burger and Glaser, 1964; Kennedy and Shaw, 1968) and differs from chain-extension of peptidoglycan which is elongated by transfer of the growing chain to the next repeating unit nearest to the lipid carrier (Ward and Perkins, 1973). Growth distal to the lipid carrier, as in lipoteichoic acid synthesis, allows the whole chain to be synthesized linked to the definitive lipid anchor. It requires,

U

100

200

300

LOO

500

T I M E (h) FIG. 9. Data showing mode of lipoteichoic acid chain extension in Lactobacillus casei. Lipoteichoic acid in bacteria was labelled in succession with [2-3H]glycerol and ['4C]glycerol,isolated, and from the glycerol terminus degraded stepwise by the joint action of phosphodiesterase and phosphomonoesterase from Aspergillus niger. At time intervals the glycerol released was analysed for 14C-and 3H-radioactivity. From Taron et al. (1983).

250

W. FISCHER

on the other hand, that during the synthesis either the chain or the growing terminus of the chain remains in contact with the membrane. Synthesis of glycolipids has been studied with crude enzyme preparations from various Gram-positive bacteria. Generally, 1,2-di-O-acyl-sn-glycerolis the initial lipid substrate to which the monosaccharide residues are sequentially transferred from nucleotide-linked hexosyl substrates (Kaufman et al., 1965; Lennarz and Talamo, 1966; Pieringer, 1968; Veerkamp, 1974). The diacylglycerol residue had been suggested to originate from phosphatidic acid through the action of a phosphatidic acid phosphatase (Krag et al., 1974) before it became evident that amounts of diacylglycerol, far greater than needed for glycolipid synthesis, result from lipoteichoic acid synthesis (see Section 1II.C). In lactococci (Fischer et al., 1978a; Schleifer et a!., 1985), enterococci (Fischer et al., 1973b, 1978b) and lactobacilli (Fischer et al., 1978b; Nakano and Fischer, 1977,1978) phosphatidyl and fatty-acyl diglycosyldiacylglycerolipids that occur as lipid moieties in lipoteichoic acids are frequently also present in the free state among membrane lipids. These glycolipid derivatives may therefore be used as acceptor substrates in lipoteichoic acid synthesis rather than be formed by acylation or phosphatidylation of the completed polymer. The additional finding of sn-glycero-1-phosphate-bearing derivatives of these acyl- and phosphatidylglycosyldiacylglycerolipids supports this idea and leads, in Ent. faecalis for example, to the putative biosynthetic sequence depicted in Fig. 10. Biosynthesis of phosphatidylglucosyldiacylglycerol has been demonstrated with particulate enzyme preparations from Ent. faecalis which catalysed phosphatidyl transfer from bisphosphatidylglycerol and phosphatidylglycerol to the glycolipid. Since these membrane preparations also catalysed conversion of phosphatidylglycerol to bisphosphatidylglycerol and the latter was more active in phosphoglycolipid synthesis, the following sequence of reactions has been suggested to occur (Pieringer, 1972): Glc(a1 -2)Glc(a1 -3)acyl2Gro PtdlGro Gro

‘PtdGro

With particulate membrane preparations from the same bacterium, it could further be demonstrated that radiolabelled phosphatidyldiglucosyldiacylglycerol was used as an acceptor substrate in lipoteichoic acid synthesis which proceeded from endogenous phosphatidylglycerol (Ganfield and Pieringer, 1980). The radiolabel appeared in a water-soluble product which seemed to increase in length on longer incubation times as indicated by increasing

FIG. 10. Structural and possible biosynthetic relationships between glycolipid, phosphoglycolipids and the two molecular species of lipoteichoic acid in Enterococcus fueculis. For structures of phosphoglycolipids (Fischer et ul., 1973a, b; Fischer and Landgraf, 1975) and lipoteichoic acid (Toon et ul., 1972; Ganfield and Pieringer, 1975; Fischer et ul., 1981) see the references given; for chain substituents of completed lipoteichoic acids (X)see Table 3.

252

W. FISCHER

T I M E (h) FIG. 1 1. Time course of incorporation of [g~uc~se-’~C]phosphatidyldiglucosyldiacylglycerol (nmol (mg protein)-’) into saline extractable polymer from a chlorofommethanol-water (1 : 1 :0.125, by vol.) supernatant ( 0 ) and an insoluble proteincontaining pellet fraction (0)by membrane preparations of Streptococcus fuecalis. From Ganfield and Pieringer (1980).

insolubility in chloroform-methanol-water (Fig. 1 1). The water-soluble product was characterized as micellar non-substituted poly(g1ycerophosphate) by column chromatography and analysis. Short-chain homologues of lipoteichoic acids were also synthesized in toluene-treated cells of Lb. casei and, under appropriate conditions, elongated to water-soluble polymers (Brautigan et al., 1981; Taron et al., 1983). On Bligh-Dyer phase partition, these short-chain homologues separated into the chloroform layer and might therefore be earlier intermediates in assembly of lipoteichoic acid than are the short-chain products obtained from Ent.faecalis which partitioned into the aqueous layer (Fig. 11). In pulseechase experiments in which the fatty-acyl residues of growing Staph. aureus were labelled with [I4C]acetate, the I4C-label appeared in and liposuccession in Glc(B1-3)acy12Gro, G l c ( ~ l - 6 ) G l c1(-3)acylzGro ~ teichoic acid (Fig. 12). The glycerophosphoglycolipid that had been isolated and characterized earlier from this organism (Fischer et al., 197%) has been shown to be an intermediate in this sequence by the pulse-chase experiment with [3H]glycerolin Fig. 13 (Koch et al., 1984): Glc2acyl2Gro + PtdGro-+ GroP-Glc2acyl2Gro+ acylzGro +24 acyl2Gro. GroP-Glc2acyl2Gro+ 24 PtdGro+(GroP)2~-Glc~acyl2Gro

As shown in Figure 13, the glycerophosphate moiety of the glycerophosphoglycolipid gained and lost radioactivity very rapidly, which is expected if the small pool of glycerophosphoglycolipid turns over to the large pool of lipoteichoic acid (see Table 5). On the other hand, the radiolabel of the

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

253

-a lo5

-U I

45

75

105 135 165 195

t T I M E imin) FIG. 12. Pulse-chase kinetics of lipid amphiphiles in growing Staphylococcus aureus on labelling of fatty acids with [14C]acetate.The arrow indicates onset of the chase. Symbols: A , phosphatidylglycerol; 0 , diacylglycerol; 0 , lipoteichoic acid; 0 , diglucosyldiacylglycerol; 0 , glucosyldiacylglycerol; 0 , phosphatidic acid. From Koch et al. (1984).

glycerol residue in the glycolipid moiety increased continuously throughout the chase period (Fig. 13), gaining radioactivity through glycolipid synthesis from the long-lasting label in the diacylglycerol pool (Fig. 12). Two chemically different linkages are formed in lipoteichoic acid synthesis; the first is the linkage of glycerophosphate to the glycolipid, the second is that of glycerophosphate units to each other. The discovery of glycerophosphoglycolipids in a wide range of Gram-positive bacteria, together with the usual absence of higher homologues (Fischer et al., 1978b) suggests that two glycerophosphate transferases might be involved. One would recognize the glycolipid substrate, thereby forming the glycerophosphoglycolipid intermediate, and a second would polymerize the chain. The initial transferase seems to be highly specific since, in some bacteria, such as in Lb. cusei, certain glycolipids are strictly selected (Fig. 14), and in all instances studied the glycerophosphate residue is exclusively linked to C-6 of the non-reducing hexosyl terminus of the glycolipid moiety. Non-glycosylated diacylglycerol

254

W.FlSCHER

T I M E (rnin) FIG. 13. Pulse-chase kinetics of the two glycerol moieties of GroP-+6Glc(B1-6) GIc(B1-3)acylzGro,the glycerophosphoglycolipidof Staphylococcus aureus, on labelling growing cells with [2-3H]glycerol. Symbols: diglucosyldiacylglycerol moiety; A , glycerophosphate moiety; A, glycerophosphate-glycolipid which escaped conversion into lipoteichoic acid. From Koch e f al. (1984) where details may be found.

.,

found as the lipid moiety in the lipoteichoicacid of certain bacteria (see Table 1) suggests that in these bacteria diacylglycerol or phosphatidylglycerolserves as the initial acceptor substrate. B. BIOSYNTHESIS OF RELATED MACROAMPHIPHILES

Although biosynthesis of the poly(glycosylg1ycerophosphate) lipoteichoic acid from Lactococcusgarvieae has not been studied, the sn-1-configuration of its glycerophosphate residues (Koch and Fischer, 1978; Fischer et al., 1982) provides strong evidence for phosphatidylglycerol being the glycerophosphate donor. A set of galactosylated sn-glycero-1-phosphoglycolipids, detected in this organism (Fischer et al., 1979), can be incorporated into a putative biosynthetic sequence leading to a compound that carries Gal(a16)Gal(al-3),Gal(a 1-2)Gro-1-phosphate, the complete repeating unit of the

H

V

W

-

R

y

2

HCUCO-R

1

H2C-0-CO-R

E

H -KO-R H2 -0-CO-R

'Q

w ~ I ~ - C O - R

H -0-CO-R

FIG. 14. Structures of glycolipids, glycerophosphoglycolipids and lipoteichoic acid from Lactobacillus casei. For details, see Fischer et a[.(1978~)and Nakano and Fischer (1977, 1978), for substitution of lipoteichoic acid (X) see Table 3.

258

W. FISCHER

lipoteichoic acid, on the definitive glycolipid moiety (Fig. 15). One might therefore suggest an assembly of the repeating unit on the growing chain by successive transfer of glycerophosphate and individual galactosyl residues. The sn-glycero- 1-phosphate side chains of the lipoglycan from Bifidobacterium bijidum (for its structure see Fig. 5) have been shown to be derived from phosphatidylglycerol. Membrane preparations catalysed transfer of radiobut not from CDP-[“’C]glycerolto a activity from ph~sphatidyl[’~C]glycerol product having the properties of the macroamphiphile of this organism (Op den Kamp et al., 1985a). Since, in these experiments, incorporation of radioactivity into the polymer from UDP-[14C]glucoseand UDP-[’4C]galactose was not observed, the glycerophosphate was apparently transferred to preformed lipoglycan. That galactofuranosyl and glycerophosphate residues may be transferred separately to the completed lipoglucan is inferred from the finding of the molecular species in the native amphiphile that have a smaller number of galactofuranosyl residues incompletely substituted with glycerophosphate (Fischer, 1987). Whether synthesis of the glucogalactofuranan moiety requires hexosyl derivatives of phosphoryl undecaprenol as glycosyl donors remains to be studied. In Micrococcus luteus, cr-D-mannopyranosyl-I -phosphoryl undecaprenol is the substrate for synthesis of the lipomannan (Scher et al., 1968; Scher and Lennarz, 1969). It is formed from GDP-a-D-mannose which, on the other hand, is directly used for synthesis of mono- and dimannosyldiacylglycerol in this organism (Lennarz and Talamo, 1966). If the synthesis of lipomannan starts from dimannosyldiacylglycerol,both activated mannosyl donors will be required for total synthesis. C . INFLUENCE OF LIPOTEICHOIC ACID BIOSYNTHESIS ON THE TURNOVER OF MEMBRANE LIPIDS

In Gram-positive bacteria a rapid turnover of membrane lipids, particularly of phosphatidylglycerol, was recognized I5 years ago (Short and White, 1970, 1971), but it was not until recently that the driving force for this process was detected in lipoteichoic acid biosynthesis (Koga et al., 1984; Koch et al., 1984). The dynamics involved for example in Staph. aureus can be deduced from the data in Table 5 . Lipoteichoic acid, although present at not more than 6 mol% contains approximately 50% of the total amphiphile glycerol and three times the amount of the non-acylated glycerol moiety of phosphatidylglycerol. The total membrane phosphatidylglycerol must therefore turn over three times for biosynthesis of lipoteichoic acid in one bacterial doubling. The diacylglycerol formed concomitantly is six times the amount present in the diacylglycerol pool. If, in the chase after labelling with [2-3H]glycerol,radioactivity in the diacylglycerol moieties of all of the lipid amphiphiles is balanced, it becomes

259

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

TABLE 5. Composition of lipid amphiphiles in logarithmically growing Stuphylococcus aureus. From Koch et al. (1 984) Composition Amphiphile

(per cent of amphiphile glycerol)

(mol Yo)

33.0 6.5 1.1

Glc(~l-6)Glc(B1-3)acyl~Gro GroP-6Glc(~1-6)Glc(~1-3)acyl~Gro

50.4 9.9 1.1 <0.1 23.6 1.5 7.6 0.1

(GroP)2~6Glc(B 1-6)Glc(B1-3)acylzGro (lipoteichoic acid)

5.7

48.6

Phosphatidylglycerol Lysylphosphatidylglycerol Bisphosphatidylglycerol Phosphatidic acid Diacylglycerol Glc(B1-3)acy12Gro

-

7.7 0.5 2.5

evident that the diacylglycerol is completely preserved (Table 6). Since, however, only a minor fraction is required for the synthesis of glycolipids and the glycolipid moiety of lipoteichoic acid (Table 5), the major part must recycle through phosphatidic acid to phosphatidylglycerol (Fig. 16). Evidence for this recycling was provided by the different labelling kinetics of the two glycerol moieties of phosphatidylglycerol in pulsechase experiments (Fig. 8). During the pulse, the diacylglycerol moiety was labelled more slowly than the TABLE 6. Balance of radioactivity in the diacylglycerol moieties of lipid amphiphiles in growing Staphylococcus aureus during a chase after labelling with [2-3H]glycerol.From Koch et al. (1984) Time after chase (min)

0

10

60

90

120

150

180

168 70 53 53 40 384

[3H]Glycerol( lo3dpm)

Diacylglycerol moiety of: Phosphatidylglycerol Lysylphosphatidylglycerol Diacylglycerol Diglucosyldiacylglycerol Lipoteichoic acid

301 38 33 5 3

260 50 57 9 6

204 70 57 32 24

194 74 50 41 25

172 74 50 33

162 70 51 52 37

Sum

380

382

387

384

377

372

48

2 60

W. FISCHER

cjtyc e r o p h o s p h o t e

CUDPGtc

GlycoLipid*

\

1

LUDP

v Glc(i)l-3)oc y L2Gro

non-acylated glycerol moiety suggesting that it is derived from a slowly labelled precursor pool rather than being formed by de nouo synthesis from glycerophosphate. Accordingly, after the chase, the diacylglycerol moiety lost the label more slowly than the non-acylated glycerol moiety. Further support for the recycling scheme comes from the synchronous behaviour, after the chase, of the radioactivity in the fatty-acyl residues of the diacylglycerol moiety of phosphatidylglycerol, free diacylglycerol and phosphatidic acid (see Fig. 12). Particularly remarkable is the persistence of the radioactivity in phosphatidic acid which, due to the small pool size and key position in lipid synthesis, would rapidly lose label after the chase if it were only formed by de nouo synthesis via glycerophosphate from unlabelled glycerol (Fig. 16). The same difference in turnover of the two glycerol moieties of phosphatidylglycerol was also observed in other Gram-positive bacteria that synthesize lipoteichoic acid (Lombardi et al., 1980; Card and Finn, 1983; Koga et al., 1984) whereas, in a Bacillus strain lacking lipoteichoic acid, both glycerol moieties turned over synchronously (Koga et al., 1984). A prerequisite for recycling of diacylglycerol is the existence of a diacylglycerol kinase. This

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

26 1

enzyme was demonstrated in toluene-treated cells and membranes of Lh. casei (Taron et al., 1983) and in membranes of Staph. aureus (H. U. Koch, unpublished observations). In Bacillus megaterium, two distinct pools of phosphatidylglycerol have been observed (Lombardi et al., 1980). The major and rapidly metabolizing pool is the precursor of diacylglycerol and, therefore, presumably involved in lipoteichoic acid synthesis. D. ADDITION OF CHAIN SUBSTITUENTS

1. Glycosylation

Although native lipoteichoic acid from En?.faecalis is highly substituted at C-2 of the glycerol moieties with mono-, di-, tri- and, eventually, tetraglucosyl residues (Wicken and Baddiley, 1963; Toon et al., 1972; Cabacungan and Pieringer, 1985), synthesis in vitro of the poly(g1ycerophosphate) chain by particulate enzyme preparations proceeds in the absence of UDP-glucose, non-substituted lipoteichoic acid being formed (Ganfield and Pieringer, 1980). Conversely, glycosylation is independent on poly(g1ycerophosphate) synthesis. Radioactivity was incorporated into preformed lipoteichoic acid when particulate enzyme preparations were incubated in the presence of UDP-[3H]glucose. After hydrolysis with 40% (w/w) aqueous hydrogen fluoride the radioactivity was detected in mono-, di-, tri- and tetraglucosyl residues, whereby elongation of diglucosyl substituents appeared to be prevailing (Cabacungan and Pieringer, 1985). In the living cell, glycosylation may also occur on growing lipoteichoic acid, as is suggested by the existence of a galactosylated di(g1ycerophospho)glycolipid which was extracted with membrane lipids from L. lactis strains and carried, like the lipoteichoic acid of this organism, an a-D-galactopyranosyl residue on one of the glycerophosphate moieties (Laine and Fischer, 1978). A particulate enzyme from Strep. sanguis ATCC 10556 incorporated glucose from UDP-['4C]glucose into a poly(g1ycerophosphate) which was tentatively characterized as lipoteichoic acid (Mancuso et al., 1979). Incorporation into this polymer was preceded by formation of labelled glucosyl lipid which was soluble in chloroform-methanol and characterized as glucosyl-1-phosphorylundecaprenol. Glucosylation of lipoteichoic acid was therefore formulated as a two-step reaction (Mancuso and Chiu, 1982): UDP-Glc +undecaprenylphosphate+Glc- 1-phosphorylundecaprenol+ UDP Glc- 1-phosphorylundecaprenol+ LTA +Glc-LTA + undecaprenylphosphate

If the second reaction, the existence of which is so far based on indirect evidence, will be established, the requirement for hexosyl- 1-phosphorylunde-

262

W. FISCHER

caprenol as donor substrate would be noteworthy because glycosylation of wall teichoic acids, which occurs in a similar membrane-associated reaction, proceeds directly from hexosyl- 1-diphosphoryl nucleosides (for review, see Ward, 1981). 2. Addition of D-Alanyl Residues

Alanylation of lipoteichoic acid has been established by Neuhaus and his coworkers using membranes and toluene-treated cells of Lb. casei (Neuhaus et af.,1974; Childs and Neuhaus, 1980). The reaction requires both membrane preparation and supernatant fraction. The supernatant fraction contains high activity of a D-alanine-activating enzyme (Baddiley and Neuhaus, 1960): Enzyme +D-alanine+ A T P e D-Alanyl-AMP-enzyme

+PPi.

Following gel-permeation chromatography of the supernatant fraction, another protein component was detected which is required for incorporation of D-alanine into membranes in the presence of D-alanine-activating enzyme (Linzer and Neuhaus, 1973). It was named D-alanine: membrane acceptor ligase, and is thought to be involved in D-alanyl transfer from the activated intermediate to the membrane: D-Alanyl-AMP-enzyme + membrane acceptor~D-Alanyl-membraneacceptor + AMP.

The ultimate membrane acceptor was identified as the poly(g1ycerophosphate) lipoteichoic acid present in membrane preparations (Childs and Neuhaus, 1980). It is not yet known whether the ligase acts as a D-alanyl transferase or is, prior to alanyl incorporation, required for binding of the Dalanyl-AMP-enzyme complex to the membrane. Additional membrane components seem to be necessary as recently suggested by the discovery of mutant strains of Lb. casei, two of which were almost completely lacking D-alanyl ester in their lipoteichoic acids (Ntamere et al., 1987). In spite of the presence of a normal or increased lipoteichoic acid content, membrane fragments of these mutant strains were unable to incorporate D-alanine when incubated with supernatant proteins from the parent strain, whereas the reverse combination showed activity. When toluene-treated cells or membranes together with supernatant proteins from Lb. casei were incubated under appropriate conditions with ~-['~C]alanine, labelled lipophilic compounds were detected (Brautigan ef al., 1981). In contrast to lipoteichoic acid, they were extractable with a monophasic chloroforn-methanol-water mixture and, after phase partition, were recovered in the organic layer. These compounds contained ~-['~C]alanine in ester linkage, showed chromatographic properties similar to the glycerophosphoglycolipids from Lb. casei and were therefore considered

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

263

alanylated short-chain homologues of lipoteichoic acid. A D-alaninecontaining short-chain homologue was isolated from B. licheniformis, and 1'-phospho-6Glc its structure was identified as 3'(2')-O-~-alanyl-sn-glycerol(/31-6)-Glc(/31-3)acylzGro(Fischer, 1982). Incorporation of D-alanine seems therefore to be independent of chain length and may occur in vivo concomitantly with poly(g1ycerophosphate) synthesis. Accordingly, with toluene-treated cells of Lb. casei, evidence for elongation of alanylated lipophilic compounds has been obtained (Brautigan et al., 198 I ; Taron et al., 1983). In an attempt to establish the site of lipoteichoic acid at which D-alanine is incorporated, the lipoteichoic acid of toluene-treated cells of Lb. casei was labelled in sequence with ~-[~H]alanine and ~-['~C]alanine (Childs et al., 1985). Stepwise enzymic hydrolysis of the double-labelled chain from the glycerol terminus revealed a constant ratio of 3Hand I4Cradioactivities. There are two explanations for this observation. One of them is that D-alanine is randomly esterified to any available glycerophosphate residue on the chain; the other that D-alanine is added to a particular site, e.g. near the lipid moiety, and then redistributed along the chain by intra- and inter-chain D-alanyl migration (Neuhaus, 1985). Evidence has been reported for a non-enzymic inter-chain transfer of alanyl-ester residues from ['4C]alanyl lipophilic lipoteichoic acid to hydrophilic long-chain lipoteichoic acid (Childs et al., 1985). Further experiments are, however, necessary to evaluate the efficiency of non-enzymic trans-esterification between non-adjacent hydroxyl groups. On the other hand, random incorporation of alanyl-ester residues along the chain would require parallel orientation or bending back of the lipoteichoic acid to the membrane where one might expect esterification to take place. E. TURNOVER

OF D-ALANINE-ESTER RESIDUES IN LIPOTEICHOIC ACID AND TRANSFER TO WALL TElCHOlC ACID

D-Alanine-ester residues of lipoteichoic acids, like those of wall teichoic acids are, even at pH 7, susceptible to spontaneous base-catalysed hydrolysis (Archibald and Baddiley, 1966; Childs and Neuhaus, 1980; Fischer et al., 1980b; Fischer and Koch, 1981), suggesting that a continuous loss of esterbound D-alanine occurs in viuo. On the other hand, D-alanyl/glycerol molar ratios as high as 0.7 are normally found in the lipoteichoic acid of Staph. aureus (Fischer and Rosel, 1980). In pulsechase experiments with ~['~Clalanine, using growing Staph. aureus cells in which lipoteichoic acid had been prelabelled with [3H]glycerol,the I4C label of lipoteichoic acid rapidly decreased after the chase, whereas the 3H label remained constant (Haas et al., 1984). In view of the stability of the 3Hlabelled poly(g1ycerophosphate) chain, the decrease of I4C label indicated the

264

W. FISCHER

expected loss of D-alanine-esterresidues from lipoteichoicacid. In addition to base-catalysed hydrolysis, an enzyme-catalysed process seemed, however, to be involved because the velocity of alanyl-residue turnover turned out to be 20 times faster than that of base-catalysed hydrolysis at the same pH value. from lipoteichoic acid was a gain of Correlated with loss of ~-['~C]alanine radioactivity in wall-linked D-alanyl-ester, suggesting a transfer of D-alanyl residues from lipoteichoic acid to teichoic acid. Strong support for this transfer was obtained by the chase kinetics of ~-['~C]alanylglyceroland ~-['~C]alanylribitolwhich, by treatment of whole cells with 40% (w/w) aqueous hydrogen fluoride, were released from lipoteichoic acid and teichoic acid, respectively, and separated in an amino-acid analyser. As shown in Fig. 17, loss of radioactivity from D-alanylglycerol was accompanied by an increase of label in glycosylated and total ribitol from teichoic acid. Whether this transfer is the only pathway for incorporation of D-alanine into teichoic acid remains to be studied. The rapid and almost complete loss after the chase of ~-['~C]alanine-ester residues from lipoteichoic acid in the experiment described in Fig. 17 would

I

30

60

90

120

150

180

T I M E AFTER C H A S E (rnin)

FIG. 17. Turnover of ~-['~C]alanyl residues from lipoteichoic acid to wall teichoic acid in growing cells of Staphylococcus aureus. Symbols: 0,alanyl lipoteichoic acid; 0 , alanylglycerol from lipoteichoic acid; A , total ribitol-linked alanine residues and A , alanyl N-acetyl glucosaminyl ribitol from wall teichoic acid. For explanation, see the text. From Haas et al. (1984) where details may be found.

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

265

rapidly produce substantial amounts of D-alanine-free lipoteichoic acid unless the vacant positions were re-esterified. Re-esterification studies were conducted using toluene-treated cells of Staph. aureus which, under appropriate conditions, do not synthesize lipoteichoic acid but retain the capacity to incorporate D-alanine into lipoteichoic acid and teichoic acid (Koch et al., 1985). As can be seen in Fig. 18, the lipoteichoic acid of toluene-treated Staph. aureus cells lost D-alanine-ester residues and, as expected, the rate of loss increased with increasing pH value. On incubation in the presence of ATP and Mg2+,the moderate loss of D-alanine-ester residues ~-['~C]alanine, at pH 6 was completely compensated for by incorporation of new I4C-labelled D-alanine and, therefore, the D-alanine/glycerol molar ratio of the lipoteichoic acid remained constant. The accelerated loss of alanine-ester residues at pH 7.5 apparently induced faster re-esterification so that the D-alanine/glycerol ratio decreased by less than 10% although loss of D-alanine-ester residues over the same time period amounted to 60%. A still more accelerated incorporation, leading to a transient increase in the alanine/glycerol ratio was observed when the experiment at pH 6 was done with Staph. aureus cells that, due to growth in the presence of high concentrations of salt, contained lowalanylated lipoteichoic acid (Koch er al., 1985). Like the D-alanyl-ester residues incorporated into lipoteichoic acid in the double-labelling experiresidues incorporated into ment described above (p. 263), ~-['~C]alanine-ester lipoteichoic acid by re-esterification were homogeneously distributed along the chain. With toluene-treated cells of Staph. aureus, as with living cells, a precursorproduct relationship suggested D-alanyl transfer from lipoteichoic acid to teichoic acid. In contrast to growing Staph. aureus, the acceptor teichoic acid in toluene-treated cells could not have been growing membrane-associated polymer because teichoic acid synthesis did not take place. Transfer must therefore have occurred to completed wall-linked teichoic acid. Re-alanylation of completed teichoic acid may also play a role in living Staph. aureus, as is suggested from the observation that, after sublethal heating, a 55% loss of ester-bound D-alanine from injured cells was repairable during recovery (Hurst et al., 1975). F. CONDITIONS AFFECTING THE CELLULAR CONTENT AND SYNTHESIS OF LIPOTEICHOIC ACID

For a range of bacteria a number of growth conditions have been reported to affect the amount of cellular lipoteichoic acid. Some of these results require further confirmation because, in the meantime, it has been observed that extraction of lipoteichoic acid from non-disintegrated bacteria may be incomplete, and variable extractability of lipoteichoic acid from bacteria

f""-" 1

0 I-

A l o / Gro

4

$1

(z

0.6

0.6

I

0.6

-

-

0

0

C"C 3 Ala /A La

I-

Q 0.L

W

4

0. L

z

L"C1 Ala/Ala

t

t-

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0. 21

O

L l

2

TIME AFTER

3

L

CHASE [h)

O

1

2

3

L

INCUBATION T I M E

lh)

FIG. 18. Loss ofmalanine-ester residues from lipoteichoic acid (a) and re-esterification of the vacant positions with new D-alanine (b, c) in toluene-treated cells of Staphylococcusaureus. In (a) lipoteichoic acid was prelabelled with ~-['~C]alanine and chased at the indicated pH values in the presence of non-labelled D-alanine, ATP and Mg2+ions. In (b) and (c) unlabelled cells were incubated under otherwise identical conditions in the presence of ~-['~C]alanine at the pH values indicated. From Koch et al. (1985) where details may be found.

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

261

grown under different conditions can a priori not be precluded (Fischer and Koch, 1981; Huff, 1982; Fischer et al., 1983). Results also have frequently been reported as amounts of cellular lipoteichoic acid relative to cellular mass. This does not necessarily reflect changes in cellular lipoteichoic acid content because changes in cell size may occur concomitantly as is, for example, observed if growth rate is varied (Sud and Schaechter, 1964; Ballesta and Schaechter, 1972). 1. Growth Stage and Growth Rate

During logarithmic growth of Ent. faecalis (Kessler and Shockman, 1979a) and B. rnegateriurn(Johnstone et al., 1982)in medium containing [3H]glycerol, radioactivity incorporated into lipoteichoic acid roughly paralleled the growth curve and remained in Ent. faeculis a constant fraction of the total glycerol incorporated. A constant cellular content of lipoteichoic acid was also found on a weight basis during all stages of growth for Ent. faecalis and Staph. aureus (Huff,1982). In certain bacteria, the amount of cellular lipoteichoic acid decreases in the stationary phase due to loss of the polymer into the culture fluid (see p. 272). Variability in the amount of cellular lipoteichoic acid could be demonstrated by changing the growth rate in chemostat culture. With Ent. faecalis, an increase in doubling time from 30 to 200 min resulted in a tenfold increase in total cellular glycerol, and the distribution of glycerol residues between lipoteichoic acid and lipids appeared to remain essentially unchanged (Carson et al., 1975). Streptococcus rnutans strains Ingbritt and BHT showed a relatively constant amount of cellular lipoteichoic acid at generation times between 1 and 10 h in glucose-containing media at pH 6 (Jacques et al., 1979a,b). A fourfold increase at the shorter generation times was, however, observed for Strep. mutuns Ingbritt when glucose in the growth medium was replaced by sucrose (Jacques et al., 1979a). Cells of Lactobacillus casei and Lactobacillus fermenturn contained a maximum amount of cellular lipoteichoic acid at intermediate generation times between 2 and 3 h (Wicken et al., 1982). 2. Efect of p H Value and Carbohydrate Source

The pH value of growth media can also affect the amount of cellular lipoteichoic acid as was shown by studies on Strep. rnutans strains BHT (Jacques et al., 1979b) and Ingbritt (Jacques et al., 1979a; Hardy et al., 1981). With strain Ingbritt, for example, the cellular amount increased fourfold when the pH value was increased from 5.5 to 7.0 (Fig. 19). Similar but less pronounced was the effect of pH value with Lb. fermenturn and Lb. casei (Wicken et al., 1982).

268

W. FlSCHER

n -

6.0 r

0 5.0

L a V 4.0 I--

= ? 3.0

= V

0 -

I

. . -

1.0 . 2.0

< O

-

Q

J

0

JI II

5.5

6.0

6.5

7.0

7.5

p H VALUE FIG. 19. Effect of pH value o n the amount of cellular lipoteichoic acid detected by rocket immunoelectrophoresis against lipoteichoic acid antiserum for Streptococcus mutans Ingbritt grown in medium containing 0.5% fructose at a dilution rate of 0.1 h-'. Amount of Iipoteichoic acid is given as rocket height (cm) standardized on 50 pg (dry weight) of cells. From Hardy et al. (1981) where details may be found.

In continuous and batch culture, the cellular lipoteichoic acid may further be influenced by the carbohydrate source in the growth medium. As shown in Fig. 20, with Strep. mutans Ingbritt the highest levels were obtained with D-fructose at a high dilution rate, intermediate levels with sorbitol at intermediate dilution rates, and the lowest level was seen with D-glucose. With Strep. mutans BHT, however, it was glucose that invoked the highest level, while in four out of nine Strep. mutans strains glucose and fructose were almost equally effective (Hardy et al., 1981). A similar variability in the effect of glucose and fructose was observed with different strains of Streptococcus sanguis and Lactobacillus sp. in batch culture (Wicken et al., 1982).

3. Phosphate Limitation Under conditions of phosphate limitation, various Gram-positive bacteria stop wall teichoic acid synthesis and, in place of it, produce phosphate-free teichuronic acid (for review see Ward, 1981). In contrast to teichoic acid, lipoteichoic acid was reported to continue to be synthesized by B. subtilis which suggested a vital role for lipoteichoic acid (Ellwood and Tempest, 1968). More recent studies with B. licheniformis (Button et al., 1975) and group-B streptococci (Nealon and Mattingly, 1984), however, showed a drop to 10% or less of the lipoteichoic acid content if compared to magnesium-

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

269

2-or G l u c o s e 5-01

z 3

0

I

Q

Sor bitot

lllllll 1.0 0

0.05 0.1 0.2 0.3 0.4 0.5

0.05 0.1 0.2 0.3 0.4 0.5 0.7

DILUTION R A T E ( h - ' ) FIG. 20. Effect of carbohydrate source and dilution rate on the amount of cellular lipoteichoic acid detected by rocket immunoelectrophoresis against lipoteichoic acid antiserum in a chemostat culture of Streptococcus mutans Ingbritt. Amount of lipoteichoic acid is given as rocket height (cm) standardized on 50 pg (dry weight) of cells. Extracts were obtained from cells grown at pH 6.0 in media containing limiting (O.SO/",w/v) glucose, fructose, or sorbitol. From Hardy et a f .(1981) where details may be found.

limited or non-restricted growth conditions. Noteworthy in this context is also the observation that, under conditions of phosphate limitation in B. subtilis, the proportion of phosphatidylglycerol, the glycerophosphate donor in lipoteichoic acid synthesis, decreased to 10% of the total lipid as compared to 54% found in magnesium-limited cells (Minnikin et al., 1972). Inversely, a stimulating effect of phosphate on synthesis of phosphatidylglycerol and elongation of lipoteichoic acid was reported for toluene-treated cells of Lb. casei(Brautigan et al., 1981; Taron et al., 1983). The effect was, however, not specific for phosphate and was also observed with other anions. Certainly, further studies are necessary to characterize the relationship between phosphate supply and lipoteichoic acid synthesis in more detail and in particular to define exactly the fraction of lipoteichoic acid that is dispensable. 4. Energy Deprivation

When ATP synthesis was poisoned in growing cells of Staph. aureus, lipoteichoic acid synthesis, although independent of energy supply, stopped

270

W. FISCHER

immediately (Koch et al., 1984). Turnover of phosphatidylglycerol, however, continued, but the label now appeared in cardiolipin whose synthesis is usually negligible during logarithmic growth (cf. Table 5, p. 259). The physiological significance of this switch-over in lipid amphiphile synthesis is not yet understood. In this context, it should be noted that in Staph. aureus the content of cardiolipin is increased at the cost of phosphatidylglycerol under various conditions, e.g. post-logarithmic growth (Short and White, 1971), growth in medium containing high concentrations of salt (Kanemasa et al., 1972), on protoplast formation during autolysis (Okabe et al., 1980) and in L-form cells (Ward and Perkins, 1968). It will be of interest to see whether, under these conditions, synthesis of lipoteichoic acid also becomes arrested.

5. Sporulation Metabolism of lipoteichoic acid during sporulation was studied in B. megaterium by labelling experiments, using 2-[3H]glycerol(Johnstone et al., 1982). At the end of logarithmic cell growth, synthesis of lipoteichoic acid rapidly decreased but increased again during forespore engulfment and remained at this elevated level throughout the sporulation process. During this period, lipoteichoic acid was apparently produced for spore membranes in which 40% of the total amphiphile glycerol is found to be present in lipoteichoic acid. The phosphate required for synthesis of phospholipids and lipoteichoic acid during sporulation was derived from endogenous sources, and it was proposed that degradation of lipoteichoic acid formed during exponential growth satisfied this requirement. G. CONDITIONS AFFECTING CHAIN SUBSTITUTION

The content of alanyl residues in teichoic acid (Heptinstall et al., 1970) and lipoteichoic acid (Fischer and Rosel, 1980) is in Staph. aureus greatly affected by the salt concentration in the growth medium. As shown in Table 7, the degree of alanyl substitution gradually decreases with increasing salt concentration, sodium and potassium chlorides being almost equally effective (Koch et al., 1985). Salt seems to act on the activity of rather than biosynthesis of the enzymes involved in alanine incorporation because toluene-treated cells of Staph. aureus that had previously been grown in the presence of low concentrations of salt, responded promptly to increasing concentrations of salt with a gradual decrease in D-alanine incorporation into lipoteichoic acid (Koch et al., 1985). An inverse correlation between salt concentration and incorporation of D-alanine was also observed with membrane fragments from Lb. casei (Reusch and Neuhaus, 1971). In view of the recently discovered D-alanyl transfer from lipoteichoic acid to teichoic acid (Haas et al., 1984;

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

27 1

TABLE 7. Effect of sodium chloride on the Dalanine-ester substitution of teichoic acid and lipoteichoic acid of Staphylococcus aureus

NaC' (g 1 - I ) 0 to 2 50 75 100

Teichoic acid" Lipoteichoic acidb Alanylglycerolc Alanine' Total glycerol Phosphate 0.6

0.73 0.03

0.2

0.32 k0.02 0.30k0.02

-

0.55

References: "Heptinstall et al. (1970); "Fischer and Rose1 (1980). 'Values are molar ratios.

Koch et al., 1985) the salt-induced decrease in the alanine-ester content of teichoic acid might be a direct consequence of decreased alanine substitution of lipoteichoic acid. Glucose-limited chemostat cultures of Staph. aureus displayed a dramatic decrease in the D-alanine content of teichoic acid and lipoteichoic acid when the pH value of the growth medium was increased (Archibald et al., 1973; McArthur and Archibald, 1984). A change in pH value from 6 to 8 decreased the ratio of D-alanine to phosphorus in teichoic acid from 0.65 to 0.02, and in lipoteichoic acid from 0.75 to 0.07 (McArthur and Archibald, 1984). This is in contrast to results obtained with toluene-treated cells of Staph. aureus which were able almost completely to compensate the considerable base-catalysed loss of alanine-ester residues from lipoteichoic acid at pH 7.5 by accelerated incorporation of new alanine residues (see Fig. 18). The reason for this difference between chemostat culture and toluene-treated cells of Staph. aureus is unknown. Growth temperature may also affect the alanyl ester content (Novitsky et al., 1974). Cells of Bacillus cougulans, grown at 55"C, had a lower ratio of D-alanine to phosphorus in their teichoic acid than cells grown at 37°C. Functional importance of the alanyl substitution at low pH values in the growth medium was derived from the observation that, at low pH values, the content of ester-linked D-alaninein wall teichoic acid of B. subtilis subsp. niger was not affected by nitrogen-limitation when presumably only those nitrogencontaining compounds are synthesized that are essential for growth (Ellwood and Tempest, 1972). In Enterococcus faecalis an increased substitution of lipoteichoic acid was observed with glycosyl residues (Kessler et al., 1983) during inhibition or protein synthesis by chloramphenicol or valine deprivation.

272

W . FISCHER

H. DEGRADATION AND EXCRETION OF LIPOTEICHOIC ACIDS

So far, only two enzymes have been described which attack the poly(g1ycerophosphate) chain of lipoteichoic acids. A phosphodiesterase, detected in Aspergillus niger (Schneider and Kennedy, 1978), releases the terminal glycerol residue distal to the lipid moiety, which leads to complete degradation of the chain if the resulting phosphomonoester is hydrolysed by phosphomonoesterase (Childs and Neuhaus, 1980; Fischer et al., 1980a, b). A glycerophosphodiesterase which releases glycerophosphate from unsubstituted poly(g1ycerophosphate) teichoic acid and deacylated lipoteichoic acid was detected in Bacillus pumilis (Kusser and Fiedler, 1984). Substitution of the poly(g1ycerophosphate) or an intact lipid moiety renders lipoteichoic acid unattackable. Teichoicases from a Gram-negative soil bacterium and from B. subtilis Marburg, which seem to hydrolyse poly(g1ycerophosphate) teichoic acid, have not yet been tested with lipoteichoic acid (Wise et al., 1972; Grant, 1979a,b). Since the phosphodiesterases of B. pumilis and B. subtilis are synthesized under conditions of phosphate limitation, they are thought to supply the cell with phosphate from endogenous sources. The metabolic fate of lipoteichoic acid has been studied by pulseechase experiments. When growing cells of B. subtilis (Koga et al., 1984) and Staph. aureus (Koch et al., 1984; Haas et al., 1984) were labelled with [2-3H]glycerol, there was after the chase no loss of radioactivity from cellular lipids and lipoteichoic acid. In contrast, from growing Ent. faecalis approximately 75% of the radiolabel of lipoteichoic acid, phosphatidylglycerol and cardiolipin disappeared within 60 min after the chase (Carson et al., 1981). A similar loss was observed in Bacillus stearothermophilus where 45 and 38% respectively of the label lost from the cellular lipid amphiphile pool were recovered in the culture fluid as lipoteichoic acid and lipid, respectively (Card and Finn, 1983). Lipoteichoic acid has also been detected in supernatant culture media from lactobacilli, oral streptococci (Markham et al., 1975), Strep. mutans, Ent. faecalis (Joseph and Shockman, 1975) and group-A streptococci (Alkan and Beachy, 1978). Several lines of evidence strongly suggest that this release of lipoteichoic acid into the medium is not dependent on cell lysis. The amount of extracellular lipoteichoic acid may rise during bacterial growth and drastically increase in the stationary phase of growth (Markham et al., 1975). In Strep. mutans, for example, 10-17% of the total lipoteichoic acid was found extracellularly during logarithmic growth, 55% at the onset of the stationary phase and 90% after incubation for another 24 h (Joseph and Shockman, 1975).When Strep. mutans BHT was grown in continuous culture at pH 6 and a low dilution rate, the amount of lipoteichoic acid in the culture fluid exceeded the amount of cell-associated lipoteichoic acid about eightfold. When the pH value was increased from 5.5 to 7, the amount of cellular and

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

213

extracellular lipoteichoicacid increased synchronously (Jacques et al., 1979b). In principle, similar but less marked results were obtained with Strep. mutans Ingbritt (Jacques et al., 1979a; Hardy et al., 1981) and Lactobacillus fermentum, whereas, in Lb. casei under nearly all conditions, the amount of extracellular lipoteichoic acid was negligible (Wicken et al., 1982). In addition to the inherent metabolic stability of the cellular lipoteichoic acid in Staph. aureus (see p. 272), a firm anchoring in the membrane of this organism is also suggested by the observation that, on sublethal heating, lipoteichoic acid was completely retained in cells whereas about 30% of the polar lipids were released (Hurst et al., 1975). Release of lipoteichoic acid into the culture medium from group-A streptococci (Alkan and Beachy, 1978; Kessler and van de Rijn, 1981), Strep. sanguis (Horne and Tomasz, 1977, 1979), Strep. mutuns BHT (Brisette et al., 1982)and Staph. aureus (Nealon et al., 1986)is greatly enhanced, without cells being lysed, following treatment with penicillin and other inhibitors of cellwall synthesis. In contrast to the spontaneous release from exponentially growing Strep. sanguis, penicillin-treated cells released the bulk of their cellular lipoteichoic acid and it was not replenished by synthesis during antibiotic treatment (Horne and Tomasz, 1979). In Strep. mutans, however, synthesis of lipoteichoic acid and lipids increased in the presence of penicillin and, because cell growth was arrested, overproduced lipid amphiphiles seemed to be excreted (Brisette et al., 1982). Extracellular lipoteichoic acid may consist of acylated and deacylated forms in proportions ranging from the almost entirely acylated form as found with Lb. fermentum (Markham et al., 1975) to the entirely deacylated form as found with growing cells of Ent. faecalis (Joseph and Shockman, 1975). Deacylated extracellular lipoteichoic acid is likely to be formed at the cytoplasmic membrane by action of endogenous lipases and, having lost the lipid anchor, may readily leave the cell. Accordingly, radiolabelling experiments with Ent. faecalis revealed a precursor-product relationship between acylated cellular and deacylated extracellular lipoteichoic acids (Kessler and Shockman, 1979a), and subsequently enzyme activities that catalyse the deacylation of lipoteichoic acid have been detected in the cytoplasmic membrane (Kessler and Shockman, 1979b). Membrane-associated enzymes capable of deacylating lipoteichoic acid were also observed in group-A streptococciwhich release acylated but predominantly deacylated lipoteichoic acid when live cells are suspended in buffer (Kessler et al., 1979). Whether lipoteichoic acid-deacylating lipases serve to regulate the content of cellular lipoteichoic acid, requires further studies. Of particular interest will be the substrate specificity of these lipases because a regulatory role would demand the operation of enzymes that are able to discriminate between lipoteichoic acid and membrane lipids.

214

W. FISCHER

Events at the cytoplasmicmembrane that result in excretion of native fully acylated lipoteichoicacid are not yet understood. In a number of experiments, secreted lipoteichoic acid was detected in the spent growth medium either by immunological techniques or following phenol/water extraction, so that it remains open whether lipoteichoicacid was excreted alone or along with other membrane components. Of interest in this context are observations with various Gram-positive bacteria which show that, under normal growth conditions and, greatly enhanced, on treatment with penicillin, in addition to lipoteichoic acid, lipids (Home and Tomasz 1977; Horne et al., 1977; Cabacungan and Pieringer, 1980; Brisette et al., 1982; Brisette and Pieringer, 1985) and proteins (Hakenbeck et al., 1983) are secreted. Secreted lipids were identical in composition and similar in proportions to membrane lipids (Horne et al., 1977; Brisette et al., 1982), while the proteins secreted on treatment with penicillin closely resembled membrane proteins, but even more closely resembled mesosomal proteins (Hakenbeck et al., 1983). These observations, together with the appearance of vesicles on the surface of penicillin-treated cells, led to the hypothesis that secretion of lipoteichoicacid, lipids and proteins may be manifestations of the same phenomenon, i.e. shedding of membrane material in the form of vesicles into the medium. Vesicles that contained lipoteichoic acids, lipids and protein were isolated from the culture fluid of penicillin-treated Lb. casei (Ntamere and Neuhaus, 1987). Most interestingly, small amounts of vesicles having a similar composition were also detectable in the spent medium of normally growing Lb. casei (F.C. Neuhaus, personal communication). Whether vesicle formation is generally involved in excretion of acylated lipoteichoic acid remains to be clarified. Secretion of acylated lipoteichoicacid alone, i.e. not in the form of vesicles, would be expected to occur through the monomolecular form which, outside the cell, may aggregate to form micelles if the critical micellar concentration of the respective lipoteichoic acid is exceeded:

-

Cellular LTA ,Extracellular LTA-Extracellular LTA membrane-associated' monomolecular micelles

Using dyestuffs, critical micellar concentrations have been determined for various lipoteichoicacids and found to be in the range of 5 PM (Courtney et al., 1986)and 0-28to 0.69 PM (Wicken et al., 1986), respectively. These values are three to four orders of magnitude higher than those for phospholipids (Smith and Tanford, 1972) and close to values for lysophospholipids containing a single fatty-acyl chain (Haberland and Reynolds, 1975). Since lipoteichoic acids like phospholipids usually contain two fatty-acyl residues per molecule and the fatty-acylresidues are similar in length, the comparatively high critical micellar concentration of lipoteichoic acids might be caused by the negative charge of the hydrophilic chain. Whether the relatively high critical micellar

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

215

concentration facilitates loss of monomeric lipoteichoic acids from the membrane requires further studies. Of particular interest in this respect might be the effective critical micellar concentration of lipoteichoic acid if, as in the membrane, it is associated with lipids. IV. Cellular Location

In lactobacilli (van Driel et al., 1973), lactococci (Forstn et al., 1985), Staphylococcus aureus (Aasjord and Grov, 1980), groups A, B, C and G streptococci (Miorner et al., 1983; Orefici et af., 1986) antigenic material has been found to be exposed on the surface of whole cells which reacts with antibodies specific to the poly(g1ycerophosphate) chain of lipoteichoic acid. About 10% of the glycerophosphate-containinglipoglycan of Bijidobacterium bifldum was also detectable with specific antibodies on whole cells of this organism (Op den Kamp et af.,1985b). Depending on the amount of antigen exposed, it is detectable by conventional agglutination of whole cells (van Driel et al., 1973), immuno-electron microscopy (van Driel et al., 1973; Aasjord and Grov, 1980),immuno-adsorbent techniques (Forstn et al., 1985; Op den Kamp et al., 1985b) or radio-immunological procedures (Miorner et al., 1983). In lactobacilli and Staph. aureus, immuno-electron microscopy shows poly(g1ycerophosphate) distributed over the entire surface of protoplasts, and in thin sections of whole bacteria the antibody-binding material fills the space between the cytoplasmic membrane and the outside of cell wall (van Driel et af., 1973; Dickson and Wicken, 1974; Aasjord and Grov, 1980). These observations, together with the amphipathic nature of lipoteichoicacid, locate the bulk of it in the outer leaflet of the cytoplasmic membrane and suggest that it may extend from the membrane through the wall and, in some cases, reach the surface of the cell. There is, however, some evidence that at least part of the lipoteichoic acid detectable as a surface component may, during excretion from the membrane, be a transient wall component (Wicken and Knox, 1975a), possibly fixed by divalent cations to anionic wall components. Part of the surface lipoteichoic acid may also be associated with protein because a considerable fraction of antibody-detectable material could be removed from whole cells of group-A streptococci by treatment with proteases (Miorner et af., 1983). In Staph. aureus, Strep. pneumoniae and various other Gram-positive bacteria, more than 80% of the lipoteichoic acid was found associated with mesosomal vesicles (Huff et af., 1974; Horne and Tomasz, 1985). This is difficult to reconcile with an extracellular location for lipoteichoic acids if these vesicles are identical with the intracellular membranes seen by electron microscopy (Reusch and Burger, 1973). A similar problem was provided by

216

W. FISCHER

the lipomannan of Micrococcus luteus, 80% of which was also found in association with mesosomal vesicles (Owen and Freer, 1972), whereas immuno-adsorbent techniques showed it to be the major antigen at the surface of intact protoplasts (Owen and Salton, 197%). Moreover, mesosomal vesicles of M . luteus were only capable of synthesizing lipomannan from Dmannosyl- 1-phosphorylundecaprenol but, unlike cytoplasmic membranes, not from the precursor GDP-D-mannose (Owen and Salton, 1975b). In addition to these observations, there are a number of reasons for considering mesosomal vesicles to be artifacts, possibly formed by extrusion from the outer layer of the cytoplasmic membrane and therefore enriched with lipid macroamphiphiles but poor in enzymes of the inner side of the membrane (Salton and Owen, 1976). Except for their association with the cytoplasmic membrane, little is known about the exact location of the enzymes involved in biosynthesis of bacterial lipid amphiphiles (Pieringer, 1983). By analogy with the situation in the endoplasmatic reticulum of mammalian cells (Bell et al., 1981), synthesis of glycerolipids may occur at the cytosolic side of the membrane where the enzymes have direct access to their nucleotide substrates. Using a doublelabelling technique together with chemical modification specific for the lipid in the outer layer of the membrane of intact cells, Rothman and Kennedy (1977) were able to show that, in B. megaterium, phosphatidylethanolamine is indeed synthesized on the cytoplasmic side of the membrane and appears a few minutes after synthesis in the outer layer. This transmembrane movement is four orders of magnitude faster than in artificial protein-free membranes and does not require metabolic energy (Langley and Kennedy, 1979). Intact protoplasts and partly autolysed, osmotically stabilized cells of B. subtilis W23 proved to be excellent systems for studying biosynthesis of extracellularly located polymers such as peptidoglycan and wall teichoic acid together with its linkage unit (Bertram et al., 1981; Harrington and Baddiley, 1983; Baddiley, 1985). Both cell-wall polymers were readily synthesized from externally supplied nucleotide precursors, and enzyme activity could be detected on the outer surface of the membrane. Since, however, neither transphosphorylation via carriers nor transfer of nucleotides across the membrane could be demonstrated, membrane-spanning enzyme complexes have been postulated that rotate or reorient themselves such that the active sites are first exposed on the inner surface of the membrane where polymer synthesis, presumably in linkage to polyprenolphosphate, occurs from nucleotide precursors. As loading of the lipid proceeds, the complex rearranges so that the growing polymer attached to its lipid emerges from the outer surface. By this rearrangement, the active sites are thought to become transiently accessible to externally added nucleotide. Synthesis of lipoteichoic acid differs from that of wall teichoic acid and

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

211

peptidoglycan in that nucleotide precursors and nucleotide by-products are not involved in chain synthesis. One might therefore hypothesize that lipoteichoic acid is synthesized directly on the outer layer of the membrane, its definitive location, if the lipid reactants are able to pass across the membrane as does newly synthesized phosphatidyl ethanolamine in B. megaterium (see above). Diacylglycerol formed during the synthesis would move from the outer to the inner layer, to be converted there via phosphatidic acid into phosphatidylglycerol, and phosphatidylglycerol as the “glycerophosphate carrier” could return to the site of synthesis. Some diacylglycerol would at the inner surface be converted into glycolipid which, after outward movement, could serve as the lipid anchor of the next lipoteichoic acid molecule to be synthesized. In contrast to the synthesis of the poly(g1ycerophosphate) chain, addition of glycosyl residues and alanyl-ester residues to the polymer is dependent on nucleotide precursors (see Section III.D), so that synthesis on the outer layer would necessitate transport of activated glycosyl residues and alanyl-ester residues through the membrane. Transport of activated D-alanine through the membrane has to be envisaged independently as a requirement in re-esterification of completed lipoteichoic acid (see Section 1II.E). V. Biological Activities

A.

LIPOTEICHOIC ACID CARRIER

Glycerophosphate polymerase and ribitol phosphate polymerase, the wall teichoic acid-synthesizing enzymes of B. subtilis and Staph. aureus, have been solubilized and purified from membranes (Mauck and Glaser, 1972; Fiedler and Glaser, 1974a). Both enzymes required an endogenous amphiphilic acceptor on which the polymer was synthesized. The acceptor was similar in composition to lipoteichoic acids and therefore designated “lipoteichoic acid carrier” (LTC). The LTC from both bacteria was interchangeable, and LTC, active with the ribitol phosphate polymerase from Staph. aureus, could be detected in various Gram-positive bacteria (Fiedler and Glaser, 1974b). Sodium dodecyl sulphate-polyacrylamide disc-gel electrophoresis provided evidence for a covalent linkage between ribitol phosphate teichoic acid and LTC, which by degradative studies was identified as a phosphodiester bond (Fiedler and Glaser, 1974b,c). Lipoteichoic acid carrier isolated from Staph. aureus was originally proposed to contain a poly(g1ycerophosphate) chain of 12-14 units linked to monoglucosyl monoacylglycerol (Fiedler and Glaser, 1974a,c). The lipoteichoic acid of Staph. aureus was then shown to contain 28-30 glycerophosphate units linked to gentiobiosyldiacylglycerol (Duckworth et al., 1975) but only a minor fraction, separated by anion-exchange chromatography, was reported to act as LTC, differing from the bulk of lipoteichoic acid by having a poly(g1ycerophosphate) chain of 22-24 units and

W. FISCHER

278

three instead of two fatty-ester residues (Lambert et al., 1977b). Later studies showed that total lipoteichoic acid from Staph. aureus, which had lost alanyl ester on purification, eluted on anion-exchange chromatography as a single peak and was in the reaction with purified ribitol phosphate polymerase on the whole active as LTC, each molecule being loaded with a chain of about 45 ribitol phosphate residues (Fischer et al., 1980a).Non-substituted lipoteichoic acids from a range of bacteria were equally effective, and structural differences in the lipid anchor concerning the carbohydrate moiety or the number of fatty-acyl residues did not affect LTC activity. Deacylated lipoteichoic acid was, however, inactive as LTC but acted as an inhibitor, which suggested a non-specific lipid moiety to be necessary presumably for appropriate orientation of LTC on the surface of the detergent-phospholipid micelles used in the assay system. It was further established that poly (ribitol phosphate) is assembled in linkage to the terminal glycerol residue of the carrier. Acceptor activity was lost when the terminal glycerol residue was converted into ethyleneglycol or removed, leaving a phosphomonoester. Hydrolysis of the latter with phosphomonoesterase generated a new glycerol terminus and restored LTC activity (Fischer et al., 1980a). The LTC-inactive derivatives

TABLE 8. Role of lipoteichoic acid chain length for LTC activity in the reaction of purified poly(ribito1 phosphate) polymerase from Staphylococcus aureus and Staphylococcus xylosus Ribitol phosphate (pmol) polymerized by the polymerase from Average chain length" (number of GroP residues)

Staphylococcus aureus'

Staphylococcus

20 15.4 12.1 8.3 4.6 3.4 1.2 blank

845 724 70 1 673 383 109 55 105

392 342 342 311 209 119

40"

472

20 1

xylosusd

~~

-

88

a Lipoteichoic acid from Leuconostoc mesenteroides (free from alanyl ester), untreated and systematicallyshortened with phosphodiesteraseand phosphomonoesterase from Aspergillus niger. Lipoteichoic acid from Lactobacillus casei (free from alanyl ester). References: 'Fischer et al. (l980a); dFiedler(1981).

"

Ala

I

ll-

GicNAc

I

-

TEICHOIC ACID

'NAC

NAC

LINKAGE U N I T

FIG. 21. Structure of the ribitol teichoic acid linkage unit sequence of Staphylococcus uureus (Coley et al., 1978; Kojima et al., 1983, 1985). @ is during biosynthesis polyisoprenylphosphate and, after linking of the polymer to the cell wall, (2-6 of muramyl residues in peptidoglycan. ManNAc(p 1-4)GlcNAc- 1-phosphate carrying mono-, di- or tri(g1ycerophosphate)residues has been found in a wide range of Grampositive bacteria as a common linkage unit for structurally different wall teichoic acids (Kojima et al., 1985; Yokoyama et al., 1986 and references therein).

P-GlcNAc-MonNAcSP-Grot

P-Rba---H

'[Gr",'",,,]

I

Lo

UDP-GLcNAc

--.GlcNAc-MurNAc-GlcNAc-MurNAc--pbptide p$pt\de

3 CDP-Gro

LTA A l a n LTA

Prenol-PP-GlcNAc-MonNAcSP-GroiSfP-RbofH

LO

LO

L o CMP LO C D P - R b o

FIG. 22. Pathway for biosynthesis of the linkage unit and of ribitol phosphate teichoic acid in Staphylococcus aureus (Harrington and Baddiley, 1985; Yokoyama et al., 1986; for reviews see text). For assembly of poly(ribito1 phosphate) in oitro two alternative pathways are shown: (1) direct polymerization from CDP-ribitol on the linkage unit lipid; (2) synthesis on LTC with subsequent transfer to the linkage unit lipid. The stage at which N-acetylglucosaminyl residues (Nathenson et al., 1966) and alanyl substituents (Haas et al., 1984; Koch et al., 1985) are added to poly(ribito1phosphate) has not been definitely established.

280

W. FISCHER

containing a modified terminus inhibited loading of non-modified LTC by the polymerase, which suggested that parts other than the terminus of the chain also play a role in recognition by the enzyme. This was confirmed by stepwise shortening of the hydrophilic chain which revealed a minimum of four glycerophosphate residues to be required on the glycolipid moiety for functioning as LTC (Table 8). Identical structural requirements of LTC have been established for the ribitol phosphate polymerase from Staph. xylosus (Fiedler, 1981). Soon after the discovery of LTC, it became evident by structural and metabolic studies that, in Staph. aureus, teichoic acid is attached to the cell wall through a linkage unit (Fig. 21). Structurally closely related linkage units for wall teichoic acids have been discovered in a wide range of Gram-positive bacteria. The linkage unit of Staph. aureus, as shown in Fig. 22, is synthesized in linkage to polyisoprenylphosphate by sequential addition of individual components from their nucleotide precursors UDP-GlcNAc, UDP-ManNAc and CDP-glycerol (for reviews see Bracha et al., 1978a; Ward, 1981; Hancock and Baddiley, 1985). N-Acetyl-D-mannosamine as a constituent of the linkage unit of Staph. aureus and other bacteria is a recent discovery (Kojima et al., 1983, 1985) because its requirement in biosynthesis has been obscured by the presence of UDP-GlcNAc-2-epimerase in membrane preparations (Harrington and Baddiley, 1985). When radiolabelled linkage unit, synthesized in membranes of Staph. aureus, was extracted with 70% ethanol and added to new membrane preparation in the presence of CDP-ribitol and Triton X-100, it was converted into polymeric material of higher molecular weight as determined by sodium dodecyl sulphate-polyacrylamide-gel electrophoresis (Bracha and Glaser, 1976). In another experiment in which membranes of Staph. aureus were incubated in the presence of UDP-GlcNAc, CDP-glycerol and CDP[3H]ribitol, two radiolabelled polymeric products separated on anionexchange chromatography and were considered to be a poly(ribito1 phosphate)-linkage unit lipid and a poly(ribito1 phosphate)-LTC (Hancock et al., 1976). Accordingly, only the latter compound was formed if the nucleotide substrates for linkage unit synthesis were omitted (Hancock et al., 1976) or if, as in Staph. aureus mutant strain 52A5, the first enzyme involved in synthesis of linkage unit was defective (Bracha et al., 1978b). These results suggested that ribitol phosphate might be polymerized on LTC and then transferred to linkage unit lipid. In support of this hypothesis, Baddiley and his coworkers demonstrated that membrane preparations from Staph. aureus and Micrococcus varians possess the capacity to transfer teichoic acid which had been preformed on LTC to a subsequently synthesized membrane-bound linkage unit (McArthur et al., 1981). As the purified ribitol phosphate polymerase from Staph. aureus appeared to be unable to accomplish transfer of polymer

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

28 1

from LTC to linkage unit lipid (Bracha et af.,1978a), a different enzyme seems to be responsible for this reaction. On the other hand, Glaser and his coworkers reported that isolated linkage unit lipid could be substituted with poly(ribito1 phosphate) by purified ribitol phosphate polymerase from Staph. aureus on incubation with CDP-ribitol in the absence of LTC (Bracha et af., 1978a). These authors considered LTC an in uitro analogue of linkage unit lipid. Using both LTC and linkage unit lipid as an acceptor, the polymerase must be unable to differentiate between sn-glycero- 1-phosphate and snglycero-3-phosphate residues which constitute the hydrophilic terminus of LTC and linkage unit lipid, respectively. Evidence that in a range of bacteria the total or at least the bulk of lipoteichoic acid is inactive as LTC came from studies on the influence of chain substituents on LTC activity. Following the observation that galactosyl substituents considerably lowered LTC activity in the reaction with purified ribitol phosphate polymerase from Staph. aureus (Fischer et al., 1980a), conditions were established for isolation of native lipoteichoic acid that retained its natural substitution with D-alanine-ester (Fischer et af., 1980b). Lipoteichoic acid isolated in this way from Staph. aureus and other bacteria proved to be inactive as LTC, but acquired activity if the alanyl substituents were removed by controlled hydrolysis. It was concluded that LTC activity TABLE 9. Influence of D-alanyl substituents on LTC activity in the reaction of poly(ribito1 phosphate) polymerase from Staphylococcus aureus. From Koch et al. (1982)

Alanine" Phosphorus Relative LTC activity 0 0.24 0.33" 0.50 0.68 0.72'

1 .oo

0.40 0.27 0.10 0.02 n.d.d

"Valuesare molar ratios. b,cNative lipoteichoic acid from Staphylococcus aureus grown in the presence of high (b)and low (c) concentrations of NaCI. For preparation of the other compounds see Koch et al. (1982). 'n.d.. not detectable.

282

W. FISCHER

previously observed in in vitro experiments may have been an artifact due to rapid base-catalysed hydrolysis of alanine-ester in the buffer (pH 8) which had been used frequently for isolation of LTC and in assays of teichoic acid synthesis. Afterwards, variously alanylated lipoteichoic acids became available by growing Staph. aureus in medium containing increasing concentrations of NaCl and by fractionating alanyl lipoteichoic acids on DEAESephacel in the order of decreasing alanyl content (.Fischer and Rosel, 1980). These variously substituted lipoteichoic acids made it possible to show that LTC activity in the reaction with the ribitol phosphate polymerase from Staph. aureus drastically decreased in a non-linear fashion with increasing alanine-ester content, approaching zero at an alanine/phosphate ratio greater than 0.7 (Table 9). Alanyl lipoteichoic acids that were inactive as LTC were not inhibitory in the reaction of ribitol phosphate polymerase with alaninefree LTC. If, however, the alanine ester substituents were converted into uncharged lactyl- or N-acetylalanyl residues, previously LTC-inactive lipoteichoic acids either acquired LTC activity if the terminal glycerophosphate was not substituted or, if the terminus was substituted, became active as competitive inhibitors. It is obvious from these results that alanine esters through positive charge interfere with binding of LTC to the polymerase, and that for binding glycerophosphate residues other than the terminal one must not be substituted with alanine. A detailed analysis revealed that a terminal sequence is required that contains approximately five non-alanylated residues, i.e. four negatively charged phosphate groups (Koch et al., 1982). If one compares similar degrees of substitution, glycosyl substituents on lipoteichoic acids are less inhibitory on LTC activity than are alanine-ester residues (Tables 9 and 10). Native lipoteichoic acids, however, frequently contain D-alanine-ester residues in addition to glycosyl substituents and then also display negligible LTC activity (Table 11). A different situation, however, in which LTC might be active in uiuo (Fig. 22) occurs in M . varians because the lipoteichoic acid of this organism is in the native state unsubstituted (Fischer and Rosel, 1980). Moreover, with membrane preparations from this organism, it has been shown that poly(6GlcNAc-1-phosphate) teichoic acid can be assembled attached to lipoteichoic acid, and transfer of the completed polymer from lipoteichoic acid to linkage unit lipid has been demonstrated (McArthur et al., 1981). In spite of this, attempts in my laboratory to isolate lipoteichoic acid loaded with poly(6GlcNAc-1-phosphate) from M. varians have so far failed. Another bacterium in which teichoic acid assembly on LTC might be operative in vivo (Fig. 22) is Staph. aureus when grown in the presence of high concentrations of sodium chloride. Under these conditions, the lipoteichoic acid has an alanine/ phosphate ratio of 0.33 and shows 27% LTC activity as compared with the dealanylated derivative (see Table 9). Isolation of ribitol phosphate-containing

TABLE 10. Influence of glycosyl substituents on LTC activity in the reaction of poly(ribito1 phosphate) polymerase from Stuphylococcus uureus. From Koch et al. (1 982) Lipoteichoic acid" No.

I

I1 111

IV V

Substituentsb Gk-3 Gh.2 Glcz Gal Gal

Degree of substitutionc

Relative LTC activity

0.84

0.14

0.63

0.66 0.71 0.34 0.62

0.45 0.45

0.27

"Isolated from Enterococcusfaecalis (1-111) and Lactococlactis (IV, V) strains. D-Alanyl substituents had been removed. bGI~l.s:a-Glc( 1- , a-Glc( 1-Z)m-Glc(1- , a-Glc( I -2)a-Glc( I 2)a-Glc(l- ; Gal: a-Gal(l- . 'Determined after hydrolysis with 40% (w/w) aqueous hydrogen fluoride as the ratio (total glycerol - free glycerol)/ total glycerol. dLTC activity of de-alanylated Staphylococcus aureus lipoteichoic acid was set at 1.OO. cus

TABLE 11. Combined effect on LTC activity of glycosyl and D-alanyl substituents in the reaction of poly(ribito1 phosphate) polymerase from Staphylococcus aureus. From Koch et al. (1982) ~

Lipoteichoic acids" D-Alanineb No.

Phosphorus

111

0.38' 0.31'

IV

Total substitutionc

Relative LTC activityd

0.65

0.02 0.08

0.5s

ONative alanine-containing form of lipoteichoic acids I11 and IV described in Table 10. "Values are molar ratios. CDetermined after hydrolysis with 40% (w/w) aqueous hydrogen fluoride as the ratio (total glycerol-free glycerol)/ total glycerol. dLTC activity of de-alanylated Staphylococcus aureus lipoteichoic acid set at 1.00. 'In 111 and IV, 53 and 90%. respectively, of the alanine was linked to glycerol, the remainder to glycosyl substituents.

284

W. FISCHER

lipoteichoic acid from Staph. aureus has been reported (Arakawa et al., 1981), but chemical proof of a covalent linkage between the two polymers is lacking. The inhibitory effect of D-alanine-ester residue on LTC activity clearly indicates that the linkage unit lipid must be protected from alanine esterification if it were to serve as the primary acceptor of ribitol phosphate polymerase. How this is accomplished in the native membrane remains to be clarified because the enzymes which add alanine ester to lipoteichoic acid appear to lack chain length specificity (see Section III.D.2). B. INTERACTION WITH CELL-WALL LYTIC ENZYMES (AUTOLYSINS)

The first lipoteichoic acid shown to be inhibitory to a homologous autolysin was the Forssman antigen of Strep. pneumoniae (Holtje and Tomasz, 1975a,b). The autolysin of this organism is a N-acetylmuramyl-L-alanine amidase which physiologically seems to play a role in separation of daughter cells at the end of cell division (Tomasz er al., 1975).Activity of this enzyme is dependent on the presence of teichoic acid on the cell wall (for a structure, see Fig. 7). Teichoic acid is required for conversion of the enzyme into a form with higher catalytic activity (Tomasz and Westphal, 1971) as well as for cleavage of the cell-wall substrate itself (Gottschlich and Liu, 1967; Mosser and Tomasz, 1970). For both of these functions, choline residues in wall teichoic acid are essential and cannot be replaced by ethanolamine residues which are incorporated into teichoic acid if pneumococcal cells are grown in a medium containing ethanolamine in place of choline (Mosser and Tomasz, 1970; Holtje and Tomasz, 1975c; Tomasz et al., 1975). Choline residues seem to act in the hydrolytic process by physically adsorbing the enzyme to the insoluble wall substrate rather than by allosteric activation (Guidicelli and Tomasz, 1984).Accordingly, if soluble teichoic acid-containing muropeptides are used as substrate choline residues are dispensable (Garcia-Bustos and Tomasz, 1987). Pneumococcal lipoteichoic acid, although structurally related to teichoic acid (see Section II.E), was found to inhibit cleavage of the cell-wall substrate by the amidase and to prevent conversion of the enzyme into the fully active form (Holtje and Tomasz, 1975a, b). Choline residues were again of importance because the autolysin-inhibitory activity was drastically decreased if choline residues on lipoteichoic acid were biosynthetically replaced by ethanolamine (Horne and Tomasz, 1985). Gel-permeation chromatography and density-gradient centrifugation demonstrated binding of the purified amidase to micellar lipoteichoic acid (Horne and Tomasz, 1985). The enzyme was also adsorbed to lipoteichoic acid immobilized by covalent linkage to Sepharose, and was specificallyelutable from this material with choline (Briese and Hakenbeck, 1983, 1985). In contrast to lipoteichoic acid in solution, the immobilized form mediated conversion of the amidase

285

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

TABLE 12. Distribution of amidase activity in fractions of pneumococcal cells grown in media containing choline or ethanolamine, respectively" Choline-cells

Cell wall Membrane Cytoplasm Total amount (units)

Ethanolamine-cells

Enzyme activity (% of total)

Enzyme formh

Enzyme activity (% of total)

Enzyme form*

1.2 58.7 40.1

n.d." C

0.5 0.5

(El

99.0

n.d.' E E

18 000

I6 000

OFrom Briese and Hakenbeck (1983) where details may be found. "C and E possess high and low catalytic activity, respectively, the E-enzyme being interconvertible to the C-form by incubation at 4°C with cell walls carrying choline-containing teichoic acid. 'n.d.. not determined.

into the fully active form, and was no longer inhibitory to cell-wall hydrolysis by the enzyme. On the other hand, lipoteichoic acid in solution also lost inhibitory properties if detergent was added (Holtje and Tomasz, 1975a) or if its hydrophobic acyl residues were enzymically removed (Briese and Hakenbeck, 1983). These observations led to the suggestion that the inhibitory effect may reside in the micellar organization of lipoteichoic acid, which not being an inhibitor in the classical sense, might act by capturing the enzyme and thus prevent its adsorption to the cell-wall substrate. Since, in pneumococcal cells, most of the autolysin was found in association with the cytoplasmic membrane (Table I2), and could be released from it by choline, lipoteichoic acid was proposed to act in vivo as a topological barrier (Briese and Hakenbeck, 1985). The intracellular distribution of amidase in ethanolaminegrown cells (Table 12) suggested, further, that binding of amidase to cholinecontaining lipoteichoic acid may also be involved in translocation of the enzyme through the cytoplasmic membrane. Purified pneumococcal lipoteichoic acid in high concentration also acted on living cells. Added to growing cultures of pneumococci it caused chain formation, resistance to stationaryphase lysis and protection against lysis by penicillin (Holtje and Tomasz, 1975a,b). Since these phenomena were also observed with pneumococci whose autolytic systems were defective (Tomasz and Westphal, 1971 ;Tomasz, 1974), control of autolysin activity was proposed as a physiological role for lipoteichoic acid in vivo (Holtje and Tomasz, 1974, 1975a). Regulatory alterations in lipoteichoic acid content or in the content of its choline residues remain, however, to be demonstrated.

286

W. FISCHER

0

0.4

0.8

1.2

1.6

LI POTEl C H O l C ACID ( n m o l / m g wall 1 FIG. 23. Inhibition of lysis of cell walls from Enterococcus faecalis by purified lipoteichoic acids from various bacterial species. Lipoteichoic acids were from Enterococcus faecalis 39 ( 0 )and 8 191 (A), Lactococcus lactis 9936 (m), Lactobacillus casei 094 (0)and Lactobacillus fermenturn 6991 ( A ) . From Cleveland et al. (1975) where experimental details may be found.

Wall teichoic acids and lipoteichoic acids of other Gram-positive bacteria seem to have similar effects on the autolysins of these organisms. As shown with B. subtilis teichoic acids may generally act as specific “allosteric” ligands for homologous autolysins (Herbold and Glaser, 1975a,b). A variety of poly(g1ycerophosphate)lipoteichoicacids have been found to be inhibitory in uitro to autolysins from bacteria that contain this type of lipoteichoic acid (Cleveland et al., l975,1976a, b). Accordingly, in the autolytic systems tested, both homologous and heterologous poly(g1ycerophosphate) lipoteichoic acids were inhibitory (Fig. 23) whereas the succinylated lipomannan from M. luteus and the lipoteichoic acid from Strep. pneumoniae were completely ineffective (Cleveland et al., 1975). Of the two autolytic enzymes from B. subtilis 168, namely a N-acetylmuramyl-L-alanine-amidase (Herbold and Glaser, 1975a) and a B-N-acetylglucosaminidase,only the latter was strongly inhibited by lipoteichoic acid in uitro (Rogers et al., 1984). Since this enzyme

PHYSIOLOGY OF LlPOTEICHOlC ACIDS IN BACTERIA

287

seems to be located between cytoplasmic membrane and peptidoglycan its inhibition by lipoteichoic acid has been proposed possibly to prevent disastroushydrolysis of newly synthesized glycan strands. The amidase which appears to act at the outer surface of the wall and to liberate wall turnover products (Rogers et al., 1984) was not inhibited by lipoteichoic acid. Structural requirements in poly(g1ycerophosphate) lipoteichoic acid for anti-autolyticactivity were studied with cell lysis and extracellular autolysins of Sraph. aureus (Fischer et al., 1981). Comparing lipoteichoic acids from various sources, differences in lipid structure and variations in chain length between 18 and 40 glycerophosphate residues did not alter anti-autolytic activity. It was, however, drastically decreased if the glycolipid carried only a single glycerophosphate residue in place of the poly(g1ycerophosphate)chain or if, as in the structure depicted in Fig. 4, digalactosyl residues were TABLE 13. Modification of the inhibitory effect of lipoteichoic acids on extracellular autolytic activity (A) and cell lysis of Staphylococcus aureus (9) by chain substitution and chain structure Concentration ( p ~ ) effecting 50% inhibition Chain substitution

A

B

None

1.6

0.8

0.59 0.71

2.3 5.2 11.2 28.8 82.4 inactive

n.d. 6.0 n.d. 11.4 n.d. 32.5

GlycosylGro/Groc 0.33 0.47 0.82

1.7 2.0 n.d.

n.d. 0.8 1.4

42.0

12.0

AlaGro/Groh 0.23 0.33 0.47 0.55

- - -Gal+Gal-rGroPG a l t

--

"From Fischer et al. (1981) where details may be found. bMeasuredin the hydrolysatewith 40% (w/w) aqueous hydrogen fluoride as the ratio (glycerol after alkaline treatment - free glycerol)/total glycerol. CMeasuredin the hydrolysate with 40% (w/w) aqueous hydrogen fluoride as the ratio (total glycerol -free glycerol)/total glycerol.

288

W. FISCHER

intercalated between the glycerophosphate moieties (Table 13). Substitution of poly(g1ycerophosphate) lipoteichoic acids with D-alanine-ester residues also had a drastic effect. The anti-autolytic activity continuously decreased with increasing alanyl substitution, approaching zero if the ratio of alanyl glycerol to total glycerol was greater than 0.7 (Table 13). In contrast to alanine-ester residues, glycosyl substituents had virtually no effect, even if 82% of the glycerophosphate moieties were substituted with mono-, di- and triglucosyl residues. From these results it was concluded that alanine-ester residues may act through their positive charge rather than by steric hindrance, and that the anti-autolytic activity might therefore reside in the negative charges of the poly(g1ycerophosphate) chain. For inhibition of the 1-Nacetylglucosaminidase from B. subtilis, alanyl-ester substitution of lipoteichoic acid was reported as being unimportant (Rogers et al., 1984), but the

0

I

I

I

25 50 75 NMOLES LIPID/MG CELL DRY WEIGHT

0

.,

FIG. 24. Effect of phospholipids on rate of lysis of intact cells of Enterococcus faecalis. Control rate constants ( K ) were in the range 1.1-4.2 h-'. Symbols: 0 , bisphosphatidylglycerol, aminoacylphosphatidylglycerol,and A , phosphatidylglycerol from Enterococcus fuecufis; 0 , bovine bisphosphatidylglycerol; 0,deacylated bovine bisphosphatidylglycerol. Taken with permission from Cleveland el af. (1976b) where experimental details may be found.

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

289

alanine/phosphate molar ratio in the lipoteichoic acid tested was only 0.24, a degree of substitution that also had little effect in the system from Staph. uureus (Table 13). Autolysins, susceptible to inhibition by poly(g1ycerophosphate) lipoteichoic acids, were also strongly inhibited by anionic phospholipids (Cleveland et ul., 1976a,b; Fischer et al., 1981; Rogers et al., 1984). As an example, the effect on lysis of Ent. fuecalis cells is shown in Fig. 24. Diacylglycerol, glycolipids, phosphatidylcholine and phosphatidylethanolamine were usually not inhibitory (Cleveland et al., 1976a, b; Fischer et al., 1981; Rogers et al., 1984). When tested at higher concentrations, neutral lipids may have some effect in particular systems (Carson et al., 1981). The observation that lipoteichoic acids and phospholipids generally lose their anti-autolytic activity on deacylation (Cleveland et al., 1975, 1976a. b; Fischer et al., 1981; Rogers et al., 1984) suggests that inhibition of lytic enzymes is effected by micelles and liposomes, respectively (Fig. 25). Since

FIG. 25. Proposed interaction of autolytic enzymes with negatively charged micelles of lipoteichoic acid and liposomes of phospholipids (cross section).

290

W. FISCHER

addition of non-ionic detergent leading to formation of mixed micelles (Dennis, 1974; Robson and Dennis, 1978, 1983) had a similar effect as deacylation, one may hypothesize that a prerequisite for the anti-autolytic effect is the high density of negative charges on the surface of lipoteichoic acid micelles and phospholipid liposomes. Whether attachment of lytic enzymesto micelles and liposomes results in allosteric inhibition or an exclusion of the active enzyme from the cell-wall substrate remains to be established. It is also unclear whether studies on isolated lipoteichoic acid reflect the situation in vivo because it is not known whether lipoteichoic acid is aggregated in the membrane to clusters of high-charge density or homogeneously distributed over the whole surface. The natural occurrence of lipoteichoic acids in both acylated and deacylated forms was thought to be appropriate for a role in regulation of autolytic activity in vivo (Cleveland et al., 1975). Alanine-free lipoteichoic acid and alanyl lipoteichoic acid with an alanine/phosphate ratio greater than 0.7 constitute, at least for autolysins of Staph. aureus, another pair of inhibitory and non-inhibitory forms, and variations in the degree of substitution would be able to modify the anti-autolytic effect in a stepless manner (Table 13). However, so far there is no evidence that this potential is used for regulation in the living cell because in normally grown Staph. aureus the bulk of lipoteichoic acid has no inhibitory capacity owing to the high degree of alanyl substitution (Fischer er a/., 1981). Recently described mutant strains of Lactobacillus casei which had a greatly lowered alanine-ester content in their lipoteichoic acids showed defects in cell separation and, due to uneven deposition of wall polymers, bent or C-shape morphology (Ntamere et al., 1987). Although these mutants suggested that an increased net negative charge of lipoteichoic acids might act as a trap for autolytic enzymes, resulting in the observed aberrant morphology and defective cell separation, there were no detectable differences in the bulk rate of autolysis and cell-wall turnover between these mutants and the parent strain (Ntamere et al., 1987). Conclusive evidence has so far not been obtained in attempts to demonstrate a role for lipoteichoic acids in in vivo regulation of autolytic enzymes in Ent. faecalis ATCC 9790. In this particular strain, variations in the cellular content of lipoteichoic acids would be relevant for regulation because its lipoteichoic acids contain no or negligible amounts of alanine-ester residues (Fischer et al., 1981; Kessler et al., 1983). In autolysis-defective mutants a significant increase in contents of cellular lipoteichoicacids and phospholipids was observed, and was considered to be at least partially responsible for the cryptic state of autolytic activity which became demonstrable in the presence of Triton X-100 (Shungu et al., 1980). On the other hand, Ent. faecalis cells that had been deprived of more than one-half of their cellular lipoteichoicacid

PHYSIOLOGY OF LIPOTEICHOIC ACIDS IN BACTERIA

29 1

by exposure to 0.5 M sucrose solution failed to autolyse at a faster rate than cells having a normal lipoteichoic acid content (Wong et al., 1981).In the celldivision cycle a decrease in cellular lipoteichoic acid content was seen about the time when cellular autolytic activity should become active, but there was no corresponding burst in newly synthesized lipoteichoic acid when autolytic activity should be inhibited (Carson and Daneo-Moore, 1981). Decreases in cellular autolytic activity, observed with Ent.faecalis grown at slow rates, also could not be associated consistently and quantitatively with increases in cellular lipoteichoic acid content (Carson et at., 1981). Although in the organisms so far studied the bulk of lipoteichoicacid seems not to be involved in regulation of autolytic enzymes, the possibility remains that minor, perhaps topologically defined, fractions may be modified to serve this purpose. C. INTERACTION WITH DIVALENT CATIONS

Lipoteichoic acid, due to its location between the cell wall and cytoplasmic membrane, has been thought to function, in connection with wall teichoic acid or other anionic wall polymers for example teichuronic acids, as an integrated cation-exchange system and reservoir of bound divalent cations between the exterior of the cell and the cytoplasmic membrane (Heptinstall et al., 1970; Meers and Tempest, 1970; Archibald, 1974; Beveridge er al., 1982). At the membrane, high concentrations (10-30 mM) of Mg2+ ions are required for membrane stability (Rogers and Reavely, 1969; Owen and Freer, 1972), fixation of lipoteichoic acid on the membrane (Hughes et al., 1973), and for activity of various membrane-associated biosynthetic reactions (Hughes et al., 1973; Pieringer, 1983). Noteworthy in this context are experiments with magnesium-dependentenzymes in the cytoplasmic membrane of B. licheniformis, which showed that Mg2+ions bound to teichoic acid and lipoteichoicacid activated the enzymes and were preferentially used if both bound and free Mg2+ ions were present (Hughes et al., 1973). The hypothesis that teichoic acids may function in scavenging cations, and in particular Mg2+ions, from the surroundings has been based on a number of observations which have been reviewed by Ellwood and Tempest (1972), Archibald (1974) and Lambert et al. (1977a). Binding characteristics for Mg2+ions to teichoic acids, succinylated lipomannan and lipoteichoic acid are summarized in Table 14. Wall teichoic acids and lipomannan displayed similar apparent association constants for Mg2+ ions but differed in binding capacity. There is an unexpectedly large difference in the apparent association constants of Mg2+ ions for poly(g1ycerophosphate) wall teichoic acid and poly(g1ycerophosphate) lipoteichoic acid. Moreover, different binding capacities have been reported, that suggest divalent binding of Mg2+ ions to poly(g1ycerophosphate) wall teichoic acid but univalent binding to poly(g1ycerophosphate)

FIG. 26. Space-filling models of lipoteichoic acid chain structures. A-C, 1,3-linked poly(g1ycerophosphates): A, unsubstituted, B, substituted with D-alanine ester, C, substituted with tl( 1-2)-linked diglucosyl residues and alanineester residues; D, poly(digalactosyl,galactosylglycerophosphate) (see Fig. 4); E, lipoglycan with partially alanylated monoglycerophosphate side chains on the galactofuranosyl residues (glucopyranosyl residues unsubstituted; see Fig. 5 ) . Models are shown in extended conformation. The distribution of chain substituents is arbitrary.

294

W. FISCHER

TABLE 14. Binding characteristics for magnesium ions to teichoic acids, lipomannan and lipoteichoic acid Polymer PoIy(g1ycerophosphate) wall teichoic acid Poly(ribitolphosphate) wall teichoic acid Succinylated lipomannan Pol y(g1ycerophosphate) lipoteichoic acid

Binding capacityd

References

2.1.103

0.5'

Lambert et al. (1975a)

0.6-103h

1' 0.25-0.3

Lambert et al. (1975b) Powell et af. (1975)

KasSoF.(I (M-I)

1.5-

64'

1

Batley et a/. (1987)

'Karsoc..is the apparent association constant. 'Determined by equilibrium dialysis at constant ionic strength (10 mM NaC1) and constant pH values (pH 5 for teichoic acids; pH 7 for lipomannan). 'Determined by 31Pnuclear magnetic resonance spectroscopy. "Maximum amount of magnesium ions (mole) bound per one mole of negative charge. 'Values quoted are for alanine-freepolymers.

lipoteichoic acid. Whether these differences result, at least in part, from the monomolecular and micellar solution of teichoic acid and lipoteichoic acid, respectively, or whether methodological factors are involved, remains to be established. It has been suggested, from several lines of evidence, that alanyl substitution of teichoic acids and lipoteichoic acids may interfere with binding of Mg2+ ions, and it has therefore been thought that one of the functions of alanyl-ester substitution may be to control binding of cations in the wallmembrane complex (Heptinstall et ul., 1970;Archibald, 1974; Lambert et al., 1977a). There are, however, unclear points that make further studies desirable. From results obtained by equilibrium dialysis, it was concluded that alanine-ester substitution left the association constant of Mg2+ ions for poly(ribito1 phosphate) teichoic acid unaltered, but caused a decrease in the binding capacity in stoicheiometricproportion to the alanine/phosphate ratio (Lambert er ul., 1975b). This was taken as evidence that each alanyl-ester residue electrostatically interacts with one of its adjacent phosphate groups and interferes with binding of Mg2+ ions by direct competition. Since, however, electrostatic interaction between alanyl-ester residues and adjacent phosphate groups is freely reversible, one would expect that alanyl-ester residues, if they have an effect, would decrease the association constant rather than the binding capacity. From data obtained with 3'P nuclear magnetic resonance spectroscopy, Batley and his colleagues (1987) concluded that there is no detectable effect of alanine-ester substituents on the association constant

PHYSlOLOGY OF LlPOTElCHOlC ACIDS IN BACTERIA

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of Mg2+ ions to poly(g1ycerophosphate) lipoteichoic acid. These authors argue that mobile counterions can eliminate the intramolecular electrostatic interactions because, in extended conformation, lipoteichoic acid phosphate residues and alanyl amino groups are further apart than the Debye radius at Mg2+ion concentrations in the range of the association constant. VI. Concluding Remarks

About ten years ago, the widespread occurrence of lipoteichoic acids and their then assumed structural homogeneity, together with their singular location on the bacterial membrane, led to the hypothesis that they may play an essential part in cell physiology of Gram-positive bacteria (for a review see Lambert et al., 1977a). A carrier function in biosynthesis of wall teichoic acid, control of autolytic enzymesand binding of divalent cations were mentioned as potential roles for lipoteichoic acids. In the meantime, deviant lipoteichoic acid structures have been detected and, more importantly, glycosyl and alanyl substituents have been found to affect considerably biological activities of the classical poly(g1ycerophosphate)lipoteichoic acids. The space-filling models shown in Fig. 26 summarize the more recently realized structural diversities. As can be seen, the phosphate groups which are responsible for the biological activities so far known may be freely accessible in the absence of substituents, or shielded to different extents by positively charged alanine-ester residues and/or glycosyl substituents. Moreover, the distance of the phosphate groups from each other will vary if, for example, hexosyl residues are intercalated between the glycerophosphate residues or monoglycerophosphate residues are attached to lipoglycan. Whether lipoteichoic acids play a vital role in bacterial physiology, as previously suggested, is still an open question. Mutants defective in lipoteichoic acid biosynthesishave not yet been described. On the other hand, loss of rather substantial fractions of cellular lipoteichoicacid, caused for example by phosphate limitation (Nealon and Mattingly, 1984), osmotic shock (Wong et al., 1981) or treatment with penicillin (Home and Tomasz, 1977, 1979) can apparently be tolerated by certain bacteria without significantly affecting the ability of lipoteichoic acid-depleted cells to survive. This does not preclude that a minor fraction of lipoteichoicacid may be indispensableor, if the total is not essential, that its presence may provide selective advantages to the organism as it was concluded with a mutant strain of Staph. aureus, 52A5, that is totally deficient in wall teichoic acid and shows, as a major defect, incomplete cell separation (Park et al., 1974). Searches in the past for functions for lipoteichoic acids have focused on properties of the hydrophilic chain. Comparatively little attention has been

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paid to the fact that lipoteichoic acids are membrane components, structurally different from other lipid amphiphiles, and accordingly might affect properties of the lipid bilayer.

Vll. Acknowledgements I am grateful to colleagues for fruitful discussions and material from their works. Gratefully I acknowledge the contributions of my previous and present coworkers. Work in the author’s laboratory has been aided by grants from the Deutsche Forschungsgemeinschaft. REFERENCES

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Owen, P. and Salton, M.R.J. (1975a). Biochemical and Biophysical Research Communications63, 875. Owen, P. and Salton, M.R.J. (1975b). Biochimica et Biophysica Acta 406.235. Owen, P. and Salton, M.R.J. (1975~).Proceedings of the National Academy of Sciences of rhe United States of America 72, 371 1. Park, J.T., Shaw, D.R.D., Chatterjee, A.N., Mirelman, D. and Wu, T. (1974). Annals of the New York Academy of Sciences 236, 54. Pieringer, R.A. (1968). The Journal of Biological Chemistry 243,4894. Pieringer, R.A. (1 972). Biochemical and Biophysical Research Communications 49,502. Pieringer, R.A. (1983). In “The Enzymes” (P.D. Boyer, ed.), vol. XVI, pp. 255-306. Academic Press, New York, London. Pieringer, R.A., Ganfield, M.-C.W., Gustow, E. and Cabacungan, E. (1981). In “Chemistry and Biological Activities of Bacterial Surface Amphiphiles” (G.D. Shockman and A.J. Wicken, eds), pp. 167-179. Academic Press, New York. Pless, D.D.. Schmit. A S . and Lennarz, W.J. (1975). The Journal of Biological Chemistry 250, 1319. Powell, D.A., Duckworth, M. and Baddiley, J. (1974). FEBS Letters 41,259. Powell, D.A., Duckworth, M. and Baddiley, J. (1975). Biochemicul Journal 151,387. Rajbhandary, U.L. and Baddiley, J. (1963). Biochemical Journal 87,429. Reusch, V.M., Jr. and Burger, M.M. (1973). Biochimica er Biophysica Acta 300,79. Reusch, V.M., Jr. and Neuhaus, F.C. (1971). The Journal of Biological Chemistry 246,6136. Robson, R.J. and Dennis, E.A. (1978). Biochimica et Biophysica Acta 508, 513. Robson, R.J. and Dennis, E.A. (1983). Accounts of Chemical Research 16,251. Rogers, H.J. and Reaveley, D.A. (1969). Biochemical Journal 113,67. Rogers, H.J., Taylor, C., Rayter, S. and Ward, J.B. (1984). Journalof General Microbiology 130, 2395. Rothman, J.E. and Kennedy, E.P. (1977). Proceedings of the National Academy of Sciences of the United States of America 74, 1821. Ruhland, G.J. and Fiedler, F. (1987). Systematic and Applied Microbiology 9,40. Salton, M.R.J. and Owen, P. (1976). Annual Review of Microbiology 30,451. Scher, M. and Lennarz, W.J. (1969). The Journal of Biological Chemistry 244,2777. Scher, M.,Lennarz, W.J. and Sweeley, C.C. (1968). Proceedings of the National Academy of Sciences of the United States of America 59, 1313. Schleifer, K.H., Kraus, J., Dvorak, C., Kilpper-Balz, R., Collins, M.D. and Fischer, W. (1985). Systematic and Applied Microbiology 6, 183. Schneider, J.E. and Kennedy, E.P. (1978). The Journal of Biological Chemistry 253,7738. Shaw, N . (1974). Advances in Applied Microbiology 17,63. Shaw, N. and Baddiley, J. (1968). Narure, London 217, 142. Shockman, G.D. and Slade, H.D. (1964). Journal of General Microbiology 37,297. Short, S.A. and White, D.C. (1970). Journal of Bacteriology 104, 126. Short, S.A. and White, D.C. (1971). Journal of Bacteriology 108,219. Shungu, D.L., Cornett, J.B. and Shockman, G.D. (1980). Journal of Bacteriology 142, 741. Smith, R. and Tanford, C. (1972). Journal of Molecular Biology 67,75. Sud, I.J. and Schaechter, M. (1964). Journal of Bacteriology 88, 1612. Tanford, C. (1980). In “The Hydrophobic Effect: Formation of Micelles and Biological Membranes”. John Wiley, New York. Taron, D.J., Childs, W.C. 111 and Neuhaus, F.C. (1983). Journal of Bacteriology 154, I 1 10. Thomas, J. and Law, J.H. (1966). The Journal of Biological Chemistry 241, 5013. Tomasz, A. (1974). Annals of the New York Academy of Science US, 439. Tomasz, A. and Westphal, M.(1971). Proceedings of the National Academy of Sciences of the Unired States of America 68, 2627.

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Tomasz, A., Westphal, M., Briles, E.B. and Fletcher, P. (1975). Journal of Supramolecular Structure 3, 1. Toon, P., Brown, P.E. and Baddiley, J. (1972). Biochemical Journal 127,399. Uchikawa, K., Sekikawa, I. and Azuma, I. (1986). Journal of Bacteriology 168, 115. Van Driel, D., Wicken, A.J., Dickson, M.R., Knox, K.W. (1973). Journal of Ultrusfructure Research 43,483. Veerkamp, J.H. (1972). Biochimica et Biophysica Acta 273, 359. Veerkamp, J.H. (1974). Biochimica et Biophysica Acta 348,23. Ward, J.B. (1981). Microbiological Reuiews 4 5 2 1 1. Ward, J.B. and Perkins, H.R. (1968). Biochemical Journal 106,391. Ward, J.B. and Perkins, H.R. (1973). Biochemical Journal 135,721. Wicken, A.J. and Baddiley, J. (1963). Biochemical Journal 87,54. Wicken, A.J. and Knox, K.W. (1970). Journal of General Microbiology 60,293. Wicken, A.J. and Knox, K.W. (1975a). Science 187, 1161. Wicken, A.J. and Knox, K.W. (1975b). Infection and Immunity 11, 973. Wicken, A.J. and Knox, K.W. (1980). Biochimica et Biophysica Acta 604, I . Wicken, A.J., Gibbens, J.W. and Knox, K.W. (1973). Journal of Bacteriology 113, 365. Wicken, A.J., Broady, K.W., Evans, J.D. and Knox, K.W. (1978). Infection andlmmunity22,615. Wicken, A.J., Broady, K.W., Ayres, A. and Knox, K.W. (1982). Infection and Immunity36,864. Wicken, A.J., Evans, J,.D. and Knox, K.W. (1986). Journal of Bacteriology 166, 72. Wilkinson, S.G. and Bell, M.E. (1971). Biochimica et Biophysica Acta 249,293. Wise, E.M., Jr., Glickman, R.S.and Teimer, E.(1972). Proceedings of the National Academy of Sciences of rhe United States of America 69,233. Wong, W., Shockman, G.D. and Wicken, A.J. (1981). In “Chemistry and Biological Activities of Bacterial Surface Amphiphiles” (G.D. Shockman and A.J. Wicken, eds), pp. 247-257. Academic Press, New York. Yamamoto, T., Koga, T., Mizuno, J. and Hamada, S.(1965). Journal of General Microbiology 131,1981. Yokoyama, K., Miyashita, T.,Araki, Y. and Ito, E. (1986). European Journal qfBiochemistry 161,479.

Author Index Page numbers in bold refer to pages on which references are listed at the end of each chapter. A Aasjord, P.,274, 275, 295 Abdolrahimzadeh, H., 269,300 Abel, K.M., 135,157 Abraham, J.M., 75, 102 Abraham, S.N., 95, 109 Achtman, M., 60, 65, 69, 71, 72, 73, 88, 89, 91, 92, 93, 102, 103, 106, 107, 108, 112 Adams, M.H., 246,298 Adams, M.W.W., 2, 4, 16, 17, 18, 21, 47 Adhya, S.L., 73,102 Aguirre, R.,20, 21, 48 Aiba, S., 221, 228 Ainley, K., 58-59, 109 Ainsworth, S.K., 93, 110 Aitkin, D.M., 186, 206, 223 Aizawa, K., 127, 128, 142, 152, 157 Akagi, J.M., 271, 300 Akazawa,T., 122,132,138,139,145,146, 149, 154, 156,157, 160, 164 Albracht, S.P.J., 20, 47 Albrecht, S.L., 5, 46, 47 Aleem, M.I.H., 24, 27, 50 Alfaro, G., 88, 102 Alkan, M.L., 272, 273,295 Allen, L.H., 144, 164 Almon, H., 20,47 Aloni, H., 219, 228 Altekar, W., 179, 183,224,225 Amako, K., 96,112 Amarasingham, C.R., 212, 223 Andersen, K., 148, 149, 157 Anderson, A., 174, 224 Anderson, B.J., 82, 102, 108 Anderson, E.S.,54, 105

Anderson, J.S.,279, 300 Anderson, T.F., 54, 89, 93, 102, 110 Anderson, W.F., 70, 77, 103, 112 Andreesen, J.R., 177, 179, 223 Andres, I., 71, 112 Andrews, T.J., 132, 135, 136, 137, 138, 139, 140, 157, 161

Anemuller, S., 181, 223 Aoki, T. 58-59, 104 Appleby, C.A., 26, 28, 30, 31, 32, 34, 35, 36, 47, 48

Aragno, M., 121, 157, 164 Arai, T., 58-59, 104 Arakawa, H., 283,2% Araki, Y., 278,279,299,302 Arancia, G., 274, 300 Archibald, A.R., 233, 234, 235,237,241, 243, 263, 270, 271, 277, 279, 291, 294, 2%, 297,298,299 Argos, P.,218,228 Armstrong, G.D., 58, 59, 67, 69, 83, 85, 88,103, 106, 110,114 Armstrong, J., 87, 103 Armstrong, J.J., 233, 2% Arnold, W., 41,52 Arnon, D.I., 202, 225 Arnon, D.I., 189, 202, 225 Arp, D.J., 13, 14, 15, 16, 17, 18, 21, 22, 23,28, 48 Asami, S., 138, 157 Ashi, H.H., 270, 300 Atherly, A.G., 41, 48 Ausubel, F.M., 41,49,51 Avissar, Y.J., 40,48 Ayala, R.P., 121, 122, 160 Ayres, A., 267, 268, 273, 301 Azuma, I., 237, 241, 301

304

AUTHOR INDEX

B Baddiley, J., 233,234, 235, 236,237,239, 241, 245, 246, 251, 261, 262, 263, 265, 269, 270, 271, 273, 276, 277, 278, 279, 280,282, 291, 294, 296, 297, 298, 299, 300,301 Badger, M.R., 135, 136, 142, 143, 150, 157, 161 Baga, M., 55, 62, 63, 76, 77, 83, 94, 95, 103,108, 109, 113 Bai, J.N., 184, 223 Baker, T.S., 135, 159 Balch, W.E., 166, 167, 223, 225 Balestra, J.P.G., 267, 2% Balkwill, D.L., 121, 161 Ball, F., 116, 119, 120, 124, 125, 163 Ballantine, S.P., 20, 48 Ballas, L.M., 275, 296 Ballment, B., 138, 139, 140, 157 Banaszak, L.J., 220, 226, 230 Baratova, L.A., 170,226 Barclay, E.A., 77, 113 Barkowski, C., 181, 188, 189, 229 Barnaby, C., 198,228 Baron, L.S., 58-59, 85, 105 Baross, J.A., 222, 223 Barrell, B.G., 166, 229 Barrera, O., 79, 80, 112 Barth, P.T., 68, 69, 103 Bass, J.A., 63, 96, 114 Bassham, J.A., 143, 162 Bates, D.L., 200, 223 Batley, M., 242, 243, 291, 294, 296 Battersby, A.R., 217, 223 Bauer, W., 242, 243, 298 Baumberg, S., 58, 59, 105 Baxter, R.M., 185, 223 Bayer, M.E., 69, 92, 99, 103 Bayler, S.T., 218, 223 Beachey, E.H., 61,95,109,110,273,274, 297,300 Beachy, E M . , 272,273,295 Beattie, K.A., 129, 147, 163 Beaty, J.S., 41, 43, 49 Becker, R.R., 9,51, 134, 162 Beckwith, J., 73, 92, 105, 109 Bedbrook, J.R., 145, 158 Bedmar, E.J., 11, 12, 48 Beecroft, L.J., 210, 230

Beer, H., 72, 104 Bell, A.W., 210, 216, 228, 230 Bell, M.E., 235, 302 Bell, R.M., 275, 296 Bell, S.G., 129, 147, 163 Bender, R., 179,223 Benson, D.R., 15, 48 Bergersen, F.J., 26, 32, 35, 48 Bergmans, H., 62, 63, 74, 76, 105, 108, 111, 113 Bergquist, P.L., 69, 112 Bergstrom, S., 64, 79, 80, 81, 100, 101, 103, 112 Berhow, M.A., 122, 133,157 Bernhardt, G., 222, 223 Berry, J.A., 144, 157, 162 Berry, J.O., 41, 48 Berry, V.K., 63, 96, 114 Bertram, K.C., 276, 2% Beudeker, R.F., 116, 124, 125, 132, 150, 151, 153, 155,157, 160 Beutin, L., 69, 71, 72, 88, 103, 107, 108 Beveridge, T.J., 291, 296 Bhayana, V., 216,223,228 Biebricher, C.K., 72, 103 Biedermann, M., 124, 125, 126, 157 Bieler, S., 76, 107 Biesecker, G., 221, 223 Billyard, E., 64,79,80, 100, 101, 102,106, 109,111 Birch-Andersen, A., 61, 62, 63, 79, 107, 110 Biryuzova, V.I., 121, 129, 130, 153, 160, 162 Bisher, M.E., 68, 112 Bittman, R., 21, 50 Black, R., 57, 62, 108 Black, S.C., 187, 189, 213, 214, 215, 224 Blake, S.M., 100, 103 Blakemore, R., 166, 225 Blaschkowski, H.P., 202, 204, 223 Blom, J., 57, 103 Bloomer, A.C., 67, 103 Bloxham, D.P., 216,223,228 Blunden, E.A.G., 4, 25,49 Bock, A., 171,223 Bock, E., 118, 124, 125, 128, 129, 154, 157, 160, 163, 164 Bock, H.-G., 21, 50 Boedeker, E., 57, 62, 108

305

AUTHOR INDEX

Boersman-Finkelstein, M., 21 I , 228 Bogdanov, A.A., 170,226, Boger, P., 20,47 Bogorad, L., 145, 158 Bohnert, H.J., 129, 146, 157, 161 Bohrer, W., 250, 253, 297 Bonen, L., 166,225,230,231 Bongers, L., 8, 24, 27, 48 Bopp, L.H., 129, 160 Bothe, H., 2, 24,48, 49,204,223 Bothma, T., 58-59, 104 Bovre, K., 57, 103, 111 Bowen, C.C., 116, 121,160 Bowes, G., 144,164 Bowien, B., 27,28,48, 116, 133, 134, 135, 136, 148, 149,157, 158,160 Boxer, D.H., 20,48 Bracha, R., 278, 280, 296 Bradbury, J.H., 26, 48 Bradley, D.E., 54, 56, 57, 58-59, 60, 63, 67,69,70,85,96,97,103,104,10S, 146, 149, 156, 158, 159 Bradley, R., 73, 93, 110 Branden, C-I., 135, 137, 158, 162 Brasnett, A.H., 156, 162 Brautigan, V.M., 252, 262, 263, 269, 2% Brewin, N.J., 42, 44, 45, 46, 48, 49, 50 Bricogne, G., 67, 103 Bridger, W.A., 213,223 Briese, T., 284, 285, 2% Briles, E.B., 246, 247, 284, 296, 301 Brinton, C.C., 54, 57, 61, 62, 63, 65, 67, 69, 72, 74, 83, 86, 90, 91, 93, 104, 107, 109,112 Brisette, J.L., 273, 274, 296 Broady, K.W., 245,267,268,273,301 Brock, T.D., 183, 220,221,223 Brooks, G.C., 21 1, 229 Brose, E.C., 69, 112 Brown, A.D., 186,206,223 Brown, A.M.C., 69, 104 Brown, D.H., 116, 119, 120, 124, 125, 163,220,229 Brown, P.E., 234,237,239,251,261,301 Brown, R.S., 65, 87, 106 Brown, S.E., 41,49 Brownlee, G.G., 166,229 Brundish, D.E., 246, 2% Bryant, R.D., 119, 158 Bryceson, I., 156, 158

Buchanan, B.B., 189,225 Buchanan, J.G., 233,296 Buchanan, T.M., 54,55,63,64,100,104, 107, 110 Buckenham, A.H., 154,158 Budgen,N., 180, 181, 183, 196, 197,216, 224, 228 Buehner, M., 221, 228 Buikema, W.J., 41, 49 Bujard, H., 76, 106 Bundy, L., 73,93, 110 Bungard, S., 207,208,229 Bunick, G.J., 219, 231 Bunn, C.R., 21,48 Buonocore, V., 179, 183, 196,224,225 Burger, M.M., 249, 275, 296,300 Burger-Wiersma, T., 123, 132, 133, 158 Burke, J.M., 89, 93, 104 Burris, R.H., 3, 11, 13, 15, 16, 17, 18, 22, 23,24,28,48,49,50,51,52 Burrows, M.R., 63, 104 Button, D., 237, 238, 268, 2% C

Cabacungan, E., 241,247,249,261,274, 296,300 Calvin, M., 135, 162 Cammack, R.,19,20,21,48,51 Campbell, I., 54, 105 Campbell, L.K., 267,268,269, 273,298 Campbell, N.E.R., 6, 38, 50 Cangelosi, G.A., 8, 48 Cannon, F.C., 42,51 Cannon, G.C., 124, 125, 126, 127, 129, 150, 152,157, 158, 163, 164 Cannon, J.G., 101,112 Cantrell, M.A., 5, 41, 42, 43, 44, 46, 48, 49, 50 Canvin, D.T., 151, 152, 153, 155, 163 Card, G.L., 260, 272, 2% Carnahan, J., 54, 104 Caro, L., 58-59, 89,93, 104, 105 Carpenter, M., 64, 67, 96, 106, 110 Carrs, B., 233, 2% Carson, D.D., 247, 260, 267, 272, 288, 290,2% Carter, K.R., 4, 16, 46, 48 Cascio, D., 135, 158

306

AUTHOR INDEX

Caspar, D.L.D., 65, 108 Cate, R.L., 200, 224 Caton, J., 148, 149, 157 Cattlico, R.A., 146, 162 Cavalli, L.L., 88, 108 Cavanaugh, C.M., 156, 158 Cazzulo, J.J., 213, 214, 224, 226 Cedergren, B., 55, 62, 75, 107 Champness, J.N., 67, 103 Chan, M., 271,300 Chandra, T.S., 129, 158 Chang, M., 278,280,296 Chapman, M.S., 135, 158 Chas, S.F., 144, 162 Chase, D.G., 54, 56, 62, 105 Chastain, R.A., 221, 230 Chatt, J., 3, 48 Chatterjee, A.N., 295, 300 Cheath, K.-Y., 71, 104 Cheasty, T., 62, 106 Chen, J.S., 2, 4, 18, 47, 48 Chen, K.N., 166,225 Chen, R., 91, 104 Chen, R.F., 195, 196, 224 Chen, S.L., 260, 261, 299 Cheney, C., 57, 62, 108 Cheng, Y.S.E., 156, 158 Chen-Schmeisser, U., 91, 104 Child. J.J., 41, 48 Childs 111, W.C., 242, 247, 249, 252, 260, 261,262,263,269,272,296,301 Ching, T.M., 4, 24, 25, 47,49 Chiu, T.-H., 247, 261, 297, 299 Chollet, R., 139, 140, 160 Choudry, M.K., 268,296 Chow, C., 170,230 Christeller, T.J., 136, 146, 158, 161 Cifferi, O., 146, 149, 163 Clark, A.J., 60, 68, 70, 104 Clark, B.L., 64, 109 Clark, M.W., 170, 171,227 Claus, G., 123, 159 Clegg, S., 76, 104 Clements, J.R., 75, 102 Clements, M.L., 57, 62, 108 Cleveland, R.F., 285, 286, 288, 289, 290, 296,297 Clewell, D.B., 60, 104 Codd,G.A., 115-157,116,117,119,121, 122, 123, 124, 125, 126, 127, 128, 129,

131, 132, 133, 134, 135, 136, 137, 138, 140, 142, 146, 147, 148, 149, 150, 151, 152, 153, 154, 155, 157, 158, 159, 161, 163 Coen, D.M., 145,158 Coetzee, J.N., 58-59, 104, 105 Coetzee, W.F., 58-59, 104 Cohen, D.R., 58-59,104 Cohen, S.N., 69, 70, 112 Cohen-Bazire, G., 121, 143, 155, 163 Colbeau, A., 20, 48 Colby, J., 116, 158 Cole, R.M., 275, 298 Coleman, R.A., 275, 2% Coley, J., 234, 277, 279, 297, 299 Colletti, C., 142, 143, 163 Collins, J.H., 200, 224 Collins, M.D., 237, 239, 241, 243, 250, 30 1 Colman, R.F., 194, 195,224 Conroy, K., 208,224 Conti, S.F., 121, 159 Cook, C.M., 154,158 Cooke, M., 86,108 Cooper, R.A., 172, 174,224 Corbin, D., 41,48 Cornett, J.B., 247, 290, 301 Cornish-Bowden, A., 207,224 Corwin, D., 79, 80, 112 Cossar, J.D., 131, 158 Cossart, P., 73, 105 Costerton, J.W., 119, 158, 162 Counts, G.W., 9, 75 Courtney, H.S., 273, 274, 297, 300 Coward, J., 93, 110 Cowell, J.L., 68, 112 Cox, G., 122,158 Cox, J.C., 31,48 Cram, D., 71,105 Craviato, A., 62, 77, 78, 105, 106, 112 Crawford, E., 58-59, 86, 89, 105 Creeger, E., 71, 105 Criswell, J.G., 27, 48 Critchley, P., 233, 297 Cronan, J.E., Jr., 240, 297 Cryz, S.J., 97, 105 Cunningham, S.D., 46, 48 Cuozzo, M., 70, 105 Curtis, S.E., 146, 147, 149, 158, 161 Curtiss, R., 111, 93, 105

AUTHOR INDEX

Cutting, J.A., 40, 48 Czechowski, M.H.. 21,49 D

Daday, A., 20,48 Dalkziel, K., 196, 224 Dalton, D.A., 37, 50 Daneo-Moore, L., 247, 260, 267, 272, 285,286,288,289,290,296,297

Daniel, R.M., 28, 31, 48 Daniels, C.J., 171, 224 Daniels, L., 189, 190, 224 Danson, M.J., 165-223, 175, 180, 18 1, 183, 187, 189, 195, 196, 197, 198, 200, 202,206, 207,208, 209, 210, 211,212, 213,214,215, 216, 218,223,224,228, 229,230 Darling, A.J., 131, 158 Darlison, M.G., 207, 229 Das, N.D., 96, 111 Date, T., 54, 67, 83, 91, 105, 113 Datta, N., 58-59, 60, 70, 71, 104, 105, 108,109 David, V., 76, 108 Davidson, E., 134, 163 Davidson, M.S., 129, 158 Davidson, N., 69, 70, 112 Davidson, R., 280, 2% Davison, A.L., 233, 2% Dawes, E.A., 210, 229 Day, L.A., 65,105 Deal, C.D., 67, 105 De Biasi, M-G., 196, 225 DeBlock, M., 69, 107 Decker, G.L., 119, 163 de Graaf, F.K., 54, 57, 61, 62, 63, 78, 79, 94,95, 105, 106, 107, 109, 111 de Graaf, J., 75, 113 Deisenhofer, J., 216, 230 DeJong, T.M., 42, 44, 45, 46, 48 DeLange, R.J., 171, 226, 229 De Ley, J., 179, 227 Delius, H., 129, 146, 157 Deming, J.W., 222,223 Dempsey, W.B., 71, 105 Dempster, G., 54, 105 Den Dulk-Ras, H., 42, 49 Dennis, E.A., 289,297,301

307

Dennison, S., 58-59, 105 Depicker, A., 69, 107 De Ree, H., 74,76,108,113 De Ree, J.M., 76, 105 Deroo, P.W., 21, 50 De Rosa, M., 179, 183, 185, 196, 224, 225

DeRosier, D.J., 200, 224 DerVartanian, D.V., 20, 21, 49, 50 Deutsch, R., 248, 297 de Vries, W., 25, 26, 52 de Wit, R., 154, 164 Dhaese, P., 69, 107 Dhar, N.M., 183,224 Dibb, N.J., 45, 48 Dick, A.J., 93, 110 Dickson, M.R., 274,275,297,301 Diekert, G., 20, 52 Distler, E., 2, 49 Distler, J., 250, 299 Ditta, G., 41, 48 Dixon, R.O.D., 4, 10, 24, 25,429 Dogra, R.C., 39,51 Doig, P., 99 Diiker, R., 265,266,270,271,279,299 Donawa, A., 24,50 Donelli, G., 274, 300 Donnelly, M.I., 136, 159 Doolittle, W.F., 167, 170, 171, 205, 224, 226

Doudoroff, M., 172, 179,225,229 Dow, C.S., 133, 134, 159, 163, 195, 228 Doyle, R.J., 291, 2% Drake, D., 96, 110 Drevon, J.J., 25, 49 DSouza, S.E., 179, 183,225 Dubochet, J., 65, 106 Duckworth, H.W., 210, 211, 216, 223, 228,230

Duckworth, M., 237,245, 265, 273,277, 291,297,298,300

Dugan, P.R., 129, 161 Duguid, J.P., 54, 77, 105, 110 Duker, E.M., 72,103 Dunmore, P., 211, 230 Du Toit, L., 58-59, 105 Duvel, D., 118, 128, 157 Dvorak, C., 237,239,241, 143,250,301 Dwarte, D.M., 122, 158 Dyer, F.P., 65,109

308

AUTHOR INDEX

Dyer, T.A., 166, 225 Dykes, C.W., 94, 105

E Ebert, A,, 124, 133, 134, 159 Edbright, R.H., 73, 105 Eden, G., 202,225 Edenharder, R., 202, 227 Edie, S.A., 11, 48 Edmunds, P.N., 54, 105 Edwards, B.F.P., 83, 106,221, 228 Egawa, R., 71, 105 Egerton, J.R., 82, 102, 108 Egge, H., 243, 250 Ehfekhar, F., 81 Eisbrenner, G., 2, 15, 24, 27, 30, 33, 35, 36, 37,48,49 Eisenberg, D., 135, 158, 159 Eisenberg, H., 2 1 7, 2 19, 220, 225, 228, 229,230,231 Eisenstein, B.I., 74, 75, 102, 105, 106 Eisenthal, R., 206, 207, 224 Ekiel, I., 190, 225 Elkan, G.H., 21,48 Ellar, D.J., 267, 270, 299 Elleman, T.C., 64,67, 82, 105 Ellis, J.R., 132, 145, 159 Ellis, L.F., 156, 162 Ellwood, D.C., 234, 268,271, 291, 297 El Mokadem, M.T., 32, 34,49,50 Elo, J., 95, 113 Emdur, L.I., 247, 297 Emerich, D.W., 4, 16, 17, 24, 25, 47, 49 Emnova, E.E., 121, 129, 130, 153, 160, 162 Engel, A., 65, 106 Engel, R., 248, 297 Engelke, J.A., 4, 51 Englard, S., 195, 225 Engler, G., 69, 107 Entner, N., 172, 225 Eosemberg, H., 198, 218, 219, 228 Eoyang, L., 71, 111 Erbes, D.L., 18,49 Erdmann, V.A., 169,231 Ericsson, L.H., 216, 223 Eschbach, M.-L., 88, 111 Evans, D.G., 54, 56,62,78,105,111

Evans, D.J., 54, 56, 62, 78, 105, 111 Evans, H.J., 2, 3, 4, 5, 6, 7, 9, 13, 14, 15, 16, 17,20,21,24,25,26,27, 30,33,35, 36,37, 38, 39,41,42,43,44,46,47,48, 49,50, 51, 52, 134, 162 Evans, J.D., 245, 267,273, 274,298,301, 302 Evans, J.N.S., 182, 184, 225 Evans, M.C.W., 189,225 Everse, J., 198, 228 Every, D., 57, 63, 105 Eyzaguirre, J., 184, 225 F Fahrney, D., 184,229 Fairlamb, A., 208, 225 Falcone, D.L., 134, 146, 148, 149, 163 Falkow, S., 9,55-59,62,63,64,69,74,75, 76, 77,83,85,94,95, 103, 105, 106, 107, 108,109,110,112 Fay, P., 122, 156, 158,159 Fay, S.P., 170, 171, 227 Feary, T.W., 96, 105 Fee, B.E., 71, 105 Feeney, J., 144, 159 Feigel, M., 243, 298 Feilberg-Jorgansen, N.H., 60, 105 Ferencz, A., 61, 110 Fernandez, R., 94, 100, 101, 111 Ferranti, R.D., 96, 110 Fersht, A.R., 197, 225 Feruichi, T., 58-59, 106 Fewson, C.A., 166, 167, 170,225 Fiala, G., 187, 221, 225, 229 Fiedler, F., 231, 236, 237, 239, 241, 242, 263, 272, 277,278,280, 281, 282,296, 2w, 299,301 Field, A.M., 62, 78, 109 Figurski, D., 69, 109 Finch, J.T., 200, 226 Finck, A., 20,49 Finlay,B.B., 61,64,69,70,71,73,81,83, 84, 85, 88, 89, 90, 91, 97, 99, 106, 110, 111, 114 Finn, D.J., 260, 272, 296 Finnegan, D., 71, 106 Fischer, W., 147,231,233-295,235,236, 237, 238, 239, 240, 241, 242, 243, 247,

309

AUTHOR INDEX

248, 250, 251, 252, 253, 254, 255, 258, 259, 261, 263, 264, 265, 266, 267, 270, 271, 272, 277, 278, 279, 280, 28 1, 282, 286, 287, 288, 289, 290,297, 298, 299, 300,301 Fischetti, V.A., 82, 107 Fisher, C.R., 122, 159 Fisher, E., 96, 105 Fisher, T.N., 96, 105 Fitchen, J., 140, 163 Fitt, P.S., 218, 227 Fives-Taylor, P., 89, 93, 104, 110 Fleischer, B., 22, 49 Fleischer, S., 21., 22, 49, 50 Fleischmann-Sperber, T., 237 Fleming, J., 58-59, 103, 104, 105 Fletcher, P., 284, 301 Floener, L., 2, 49 Flourney, D.S., 203, 225 Fogg, G.E., 116, 122, 159 Folkhard, W., 65, 66, 67, 106, 108 Ford, G.C., 221,228 Forester, H., 267, 268, 269, 273, 298 Fornari, C.S., 146, 148, 159 Forsberg, C.W., 291, 296 ForsCn, R., 274, 275, 298 Fothergill-Gilmore, L.A., 174, 225 Fowler, T., 70, 71, 73, 106 Fox, G. E., 166, 167, 169,223,225,231 Fox, J.A., 21, 50 Freas, S., 144, 157, 162 Freer, J.H., 275, 291, 300 Freese, E., 197, 225 Freidrich, B., 8, 51 Freitag, C.S., 74, 75, 102, 106 Frey, P.A., 203,225 Friedman, A.M., 41,49 Friedrich, B., 20,41,42,49, 129, 148,157, 159 Friedrich, C.G., 9,20,41,42,49, 129,148, 157, 158 Froholm, L.O., 57,63, 103, 106,111 Frost, L.S., 53-102, 55, 57, 58-59,61,64, 67,69,70, 71, 73, 83, 84, 85,86,88, 89, 90,93,96, 103, 106, 110, 114 Frost, S.C., 209, 225 Fuchs, G., 181, 182, 184, 188, 189, 198, 202,215,225,226, 228,229,231 Fuerst, J.A., 54, 106 Fujimota, N., 205, 230

Fujitia, Y., 197, 225 Fujiwara, M., 246, 298 Fukuyama, K., 205,230 Fulco, A.J., 260, 261, 299 Furer, E., 97, 105 Fusco, P.C., 62, 107

G Gaastra, W., 54, 57, 61, 62, 63, 79, 94, 105, 106, 111 Gabriel, J.L., 194, 195, 228 Gaffney, D., 71,72, 106 Gait, M.J., 80, 106 Gambacorta, A., 179, 183, 185, 196, 224, 225 Ganfield, M.-C.W., 237, 247, 250, 25 1, 252, 261,298,300 Gantt, E., 121, 159 Garcia-Bustos, J.F., 284, 298 Garges, S., 73, 102 Garrett, R.A., 171, 227 Gasson, M.J., 72, 106 Gatenby, A.A., 146, 149, 156, 158, 159 Gazzotti, P., 21, 50 Gedorka-Cray, P.J., 127, 159 Gehring, U., 202,225 Gemski, P., 54, 104 Genetello, C., 69, 107 Gerbrandy, S.J., 174, 230 Germanier, R., 97, 105 Gerstenberg, C., 129, 159 Gest, H., 172, 191, 193, 225 Gesteland, R., 58-59, 86, 89, 105 Getzoff, E.D., 67, 105 Giardina, P., 179, 183, 196, 224, 225 Gibbens, J.W., 247,301 Gibbons, N.E., 185, 223 Gibbons, R.A., 62,63,104,112 Gibbs, S.P., 123, 132, 152, 161 Gibson, C.E., 116, 159 Gibson, J., 166, 225 Gibson, J.L., 134, 138, 142, 146, 148, 149, 159, 163 Gicquel-Sanzey, 73, 105 Giere, O., 156, 159 Giles, I.G., 216, 228 Gill, R.E., 76, 107 Gill, S.R., 127, 159

310

AUTHOR INDEX

Girard, A.E., 119, 161 Giroux, J., 198, 229 Glaser, L., 247, 249, 277, 278, 280, 285, 296,297,298,299 Glickman, R.S., 272, 302 Gliozzi, 185, 224 Goebel, W., 76, 106 Goebel, W.F., 246,298 Goguen, J.D., 95,109 Goldfine, H., 240, 298 Goldsmith, M.E., 87, 106 Goransson, M., 76, 77, 94, 95, 103, 106, 109 Gorbach, S.L., 54, 56, 62, 105 Gordon, G.L.R., 142,161 Gorisch, H., 187, 189, 198,214,215,221, 225,226 Gotschlich, E.C., 55, 61, 63, 79, 94, 100, 101,104,111,112 Gottschalk, E.M., 136, 157 Gottschalk, G., 177, 179, 223 Gottschlich, E.C.S., 284, 298 Gotz, F., 184, 225 Graeme-Cook, K.A., 209,228 Graf, E.-G., 20,49 Graham, L.A., 40,44,49 Graham, L.B., 118, 164 Grant, R.A., 87, 106 Grant, W.D., 272, 298 Gray, C.W., 65, 87, 106 Gray, J.C., 132, 145, 159 Gray, M.W., 167,205,226,229 Green, G.R., 171, 181, 226, 229 Greenawalt, J.W., 119, 163 Greenberg, G.R., 233,2% Greenwalt, J.W., 119, 160 Greenwood, D.M., 139, 157 Grinter, N.J., 68, 69, 103 Gross, R.J., 62, 77, 105, 106, 112 Grossebuter, W., 187, 189, 198,214,215, 221,225,226 Grosskopf, E., 46,47,51 Grov, A., 274,275,295 Guest, J.R., 199, 207, 229 Guidicelli, S., 284, 298 Gupta, R., 166, 170, 171,224,225,226 Gurevitz, M., 146, 159 Gusemann, M., 148, 149, 158 Gustow, E., 247, 300 Gutteridge, S., 133, 144, 146, 147, 159

Gyles, C.L., 63, 112

H Haaker, H., 27,50 Haas, B., 169,227 Haas, R., 64,79, 80, 100, 101, 102, 106, 109, 237, 247, 248, 253, 254, 258,259, 263, 264, 267,270,272,279,297,298, 299 Haberland, M.E., 274,298 Haddad, K.R., 44,50 Haddock, B.A., 31, 40,48,49, 172,202, 228 Hagberg, L., 75, 106 Hagblom, P., 64,79, 80, 100, 101, 102, 106,111 Hageman, R.H., 140, 161 Hageman, R.V., 3, 11, 49 Hahn, M., 43, 50 Hakenbeck, R., 274,284,285,2%, 298 Hale, G., 200, 223, 224 Hall, D.O., 19, 51, 205, 220, 226 Hall, S., 206, 207, 224 Hall, W.T., 123, 159 Halliday, I.J., 94, 105 Halvorson, H.O., 92, 111 Hamada, S., 245, 298, 302 Hamilton, P.B., 4,52 Hamlin, R., 135, 158 Hamlyn, P.H., 80, 106 Hamstra, H.J., 76, 113 Hancock, I.C., 234, 235, 276, 278, 280, 282,291,294,296,298,299 Hangland, R.A., 41, 43, 49 Hanley, J., 62, 106 Hannus, F.J., 5, 49 Hansen, B.S., 89, 107 Hansen, G.A., 57,103 Hansinge, R.P., 21, 50 Hansson, H.A., 77, 112 Hanstein, W.G., 217, 226 Hanus, F.J., 2,4, 5, 6, 7, 9, 13, 14, 16, 20, 21,37, 38, 39,41,43,44,46,47,48,49, 50

Hardy, J., 62, 63, 64,76, 77,94,103, 108 Hardy, L., 267,268,269,273,298 Hardy, R.W.F., 6,26,27,48,49 Harford, S.,94, 105, 210, 211, 224, 226, 230

AUTHOR INDEX

Harker, A.R., 5, 13, 14,21, 37,46,49,50 Hams, H., 117, 118, 124, 125,159, 160, 164 Harrington, C.R., 276, 280, 298 Hams, J.I., 221, 223 Hams, T.J.R., 156, 159 Harrison, D., 134, 159 Hartl, T., 187, 189, 198, 221, 225, 226 Hartman, F.C., 133, 136, 137, 147, 159, 162 Harwood, C.R., 72,107 Hase, T., 205, 220, 226 Haselkorn, R., 146, 147, 149, 158, 161 Hatch, M.D., 142, 159, 198, 226 Hatchikian, E.C., 20, 21, 48 Hatefi, Y., 217, 226 Haugland, R.A., 5, 43, 46, 48, 49 Hausinger, R.P., 205, 226 Havekes, L., 88,107 Havelka, U.D., 6, 26, 27, 48, 49 Hawthornthwaite, A.M., 122, 123, 127, 128, 131, 132, 146, 147, 148, 149, 152, 159,161,163 Hayashi, H., 270,299 Hayward, A.C., 54, 106 Heath, E.C., 174,226 Heckels, J.E., 100, 102, 108, 113 241, 298 Heckmann, M.O., 25,49 Hedges, R.W., 58-59, 60, 69, 104, 105, 112 Heine, E., 20, 49 Heine, H.-G., 209, 229 Heinhorst, S., 146, 147, 160 Heinrich, G., 154, 157 Helinski, D.R., 41, 48, 69, 109 Helmuth, R., 65,73,91,93, 102,106,107 Hemmings, N.L., 237, 238, 2% Henderson, C.E., 200,226, 229 Henderson, E., 170, 171,227 Henkel, J., 140, 164 Hennecke, H., 43,50 Henning, U., 87,88,91, 104, 109, 111 Hennings, N.L., 268,2% Henrichsen, J., 63, 107 Heptinstall, S., 270, 271, 291, 294, 296, 298 Herbold, D.R., 285, 298 Herdman, M., 123, 129, 160 Hermodson, M.A., 63,64,107

311

Herrlich, P., 69, 88, 107 Herrmann, J., 235, 240,250, 251,297 Herskovitz, T.T., 226,217 Hespell, R.B., 166, 225 Hether, N.W., 237,298 Hewett, M.J., 272, 273, 299 Hewson, J.K., 211, 230 Hickock, R.E., 27, 30, 35, 36, 37,42,44, 48,49 Higa, A., 213, 226 Hill, D.F., 91, 107, 146, 158 Hill, E., 220, 226, 230 Himes, R.H., 271, 300 Hind, G., 132, 164 Hindennach, I., 87, 111 Hirai, Y., 270, 300 Hirata, A., 119, 120, 160 Hirota, Y., 71, 105 Hoare,D.S., 116, 117, 118, 119, 162,163 Hobden, A.N., 94,105 Hochstein, L.I., 176, 177, 179, 197, 226, 230 Hodges, R.S., 61,64,85,94,97, 113, 114 Hodler, LA., 97, 112 Hoekstra, W.,76, 88, 107, 113 Hofman, J.D., 170,226 Hofmann, T., 61,62,94, 106, 111 Hogenauer, G., 71, 108 Hogrefe, C., 42, 49, 148, 159 Hohn, T., 93, 108 Hollander, R., 181, 226 Hollingshead, S.K., 82, 107 Holmes, D.S., 129, 160 Holmes, K.K., 55, 104 Holsters, M., 69, 107 Holt, S.C., 119, 160 Hothuizen, Y., 119, 120, 124, 125, 126, 127, 129, 130, 152, 160 Holtje, J.-V., 283,284,285,286,289,290, 2%, 298 Hom, S.S.M.,40,44,49 Hombrecher, G., 42,45,48,49 Hones, J., 199, 226 Hontelez, J.G.G., 42, 50 Hooper, A.B., 127, 160 Hooper, E.A., 200,202,223,224 Hooykaas, P.J.J., 42,49 Hopner, T., 202, 227 Horecker, B.L., 174, 183, 226,228 Home, D., 273,274,284,295,298

312

AUTHOR INDEX

Houchins, J.P., 15, 49 Hough, D.W., 196, 207, 208, 214, 216, 228,229 Houwink, A.L., 54, 107 Howard, A., 135, 158 Howard, J.B., 205,226 Hoyne, P.A., 64,67,82,105 Hsu, P., 76, 107 Hsu, S.C., 247, 261, 299 Huber, G., 187, 229 Huber, R., 187, 216, 229, 230 Huff, E., 247,267,275,298 Hughes, A.H., 265, 273, 291,298 Hughes, V.M., 58-59, 104, 105 Hull, R.A., 55, 75, 76, 107, 109 Hull, S.I., 75, 76, 107 Hultberg, H., 55, 75, 95, 107, 108, 113 Hurst, A., 265, 273, 298 Hurwitz, J., 174, 226 Husemann, M., 148,160 Huynh, B.H., 20,21,49,50 I Iglewski, B.H., 97, 114 Iino, T., 72, 112 Iion, T., 92, 107 Imanaka, T., 221,228 Incharoensakdi, A., 138, 139, 154, 160 Inderson, I., 135, 158 Ingledew, W.J., 31, 48 Inuzuka, M., 54, 67, 83, 89, 91, 105, 112, 113 Inze, D., 69, 107 Ippen, K., 72, 107 Ippen-Ihler, K., 57, 60,68, 69, 74, 88,91, 92, 107, 108, 109, 112 Irvin, R.T., 96, 108 Isaacson, R.E., 63, 109 Isaacson, Y.A., 21,50 Ishaque, M., 24, 27, 50 Ishimoto, N., 279, 283, 295, 300 Ishizuka, I., 235, 240, 25 1, 297, 298 Issaacson, R.E., 54, 55, 62, 63, 77, 107 Isuzaki, N., 119, 160 Ito, E., 237, 238, 241, 278, 279, 283, 295, 298,299,302 Ivrin, R.T., 96,99 Iwasaki, H., 237,238,241,298 Iyer, V.N., 69, 108

J

Jackson, L.L., 237, 298 Jacobs, A.A.C., 95, 107 Jacobson, A., 90, 93, 107 Jacoby, G.A., 60, 107 Jacques, N.A., 267, 268, 269, 273, 298 Jaenicke, R., 21 7, 222, 223, 226 Jahneke, L.S., 127, 160 James, A.N., 100, 107 James, L.T., 100, 108 Jann, B., 254, 297 Jann, K., 254,297 Jannasch, H.W., 155, 160 Jansen, K., 184,225, 226 Jany, K-D., 199,226 Jarrell, K.F., 190, 225 Jenkins, T.M., 215 Jennings, H.J., 246, 299 Jennings, N.T., 4, 16, 46, 48 Jensen, T.E., 116, 121, 122, 160 Jephcott, A.E., 63, 79, 107 Jodal, U., 75, 106 Johanson, W.G., Jr., 63, 96, 97, 114 Johansson, G., 274, 275,300 Johnson, A.P., 100, 108 Johnson, H.S., 198, 226 Johnson, K., 97,98, 107 Johnson, P., 200, 224 Johnston, A.W.B., 42, 44,45, 46,48,49 Johnstone, K., 267,270,299 Jones, D., 210, 230 Jones, G.W., 54, 62, 107, 112 Jones, W.J., 182, 226 Jordan, D.B., 139, 140, 160 Jorgensen, B.J., 74, 108 Jorgensen, J., 75, 109 Joseph, R., 272, 273, 299 Jouanneau, Y., 134, 146, 148, 149, 160, 163 Jungkind, D.L., 240, 299 Junke, D.D., 247,261,299

K Kaback, H.R., 209,228 Kadam, S.K., 71,111 Kagan, S.A., 44,45,46,48,50 Kagrammanova, V.K., 170,226

AUTHOR INDEX

Kaine, B.P., 171, 226 Kaklij, G.S., 184, 226 Kalia, V.C., 25, 49 Kaling, M., 146, 163 Kalkman, M.L., 20, 47 Kallenius, G., 55, 61, 62, 75, 76, 95, 107, 111, 113 Kaltwasser, H., 20, 52 Kaluza, K., 43, 50 Kam, Z., 220,229 Kandler, O., 166, 170, 171, 187,223,226, 227 Kanemasa, Y., 270,299,300 Kanodia, S, 182, 184, 225, 227 Kaplan, N.O., 198, 228 Kaplan, S., 146, 148, 159 Kapulnik, Y.,46, 48 Katakura, Y., 221, 228 Katayama-Fujimara, Y.,119, 120, 160 Kates, M., 185, 227, 230 Kathir, P., 69, 92, 108 Kaufman, B., 250,299 Kawashima, I., 119, 160 Kawashima, N., 145, 160 Kay, C.M., 64,65,66,67,94,97,113 Keister, D.L., 32, 34, 49, 50 Kelemen, M.V., 233, 241, 296, 299 Kelly, D.J., 195, 228 Kelly, D.P., 180, 181, 183, 187, 189, 231 Kempen, H., 88, 107 Kenealy, W.R., 189, 198, 202, 227, 231 Kennedy, E.P., 242, 272, 276,299,301 Kennedy, L.T., 249, 299 Kennedy, N., 69,88,102,107 Kent, S.S., 141, 160 Kerby, R., 182, 231 Kerscher, L., 187, 189,202,203,204,205, 206,220,226,227,228,230 Kersters, K., 179, 227 Kessell, S.R., 206, 207, 224 Kessler, R.E., 239,267,271,273,290,299 Kettleborough, C.A., 133, 146, 147, 159 Keys, A.J., 133, 144, 146, 147, 159 Keyser, H.H., 10, 11, 50 Khan, A.A., 154, 160 Kies, L., 123, 160 Kikuchi, G., 28, 30, 51 Kilpper-Balz, R., 237,239,241,243,250, 301 King, M.D., 238, 299

313

Kinghorn, H.A., 210,213,230 Kirk, C.S., 63, 64,107 Kirsop, B.H., 182, 227 Kitao, T., 58-59, 104 Kitto, G.B., 198, 228 Kjems, J., 171, 227 Klassen, P., 63, 105 Klemm, P., 54, 57, 61, 62, 63, 74, 75, 76, 78, 79,94,95, 105, 106, 107, 108, 111 Klenk, H.-P., 169, 227 Kline, B.W., 119, 120, 163 Klink, F., 171, 227 Klintworth, R., 148, 149, 158, 160 Kloser, M., 87, 109 Klucas, R.V., 20,50 Klug, A., 67, 103 Knaff, D.B., 2,50 Knappe, J., 202,204,223,227 Knight, S., 135, 158 Knobloch, K., 24, 27,50 Knowles, J., 76, 111 Knox, J.M., 100,107 Knox, K.W., 234,235,236,239,242,245, 247, 267, 268, 269, 272, 273, 274, 275, 298,299,301,302 Knutton, S., 57, 62, 108 Kobayashi, H., 132, 138, 145, 146, 149, 156,157,160, 164 Koch, H.U., 231,236,237,239,241,242, 243, 247, 248, 251, 253, 254, 258, 259, 261, 263, 264,265, 266, 267, 270,271, 272, 277, 278, 279, 280, 281, 282, 286, 287,288,289,290,297,298,299 Koch, T.K., 177, 197,230 Koga, T., 245,302 Koga, Y., 248,258,260,272,299 Kojima, N., 21, 50, 278, 279, 299 Koller, B., 129, 146, 157 Komano, T., 58-59,106 Konarsak-Kozlowska, M., 69, 108 Konig, H., 166, 170, 187, 221, 222, 223, 226,229 Konigsberg, W.H., 87, 106 Konings, R.N.H., 129, 130, 160 Konings, W.L., 120, 124, 125, 126, 127, 152,160 Konings, W.N., 119, 120, 125, 129, 130, 160 Koomey, J.M., 63, 64,79, 80, 81, 100, 101, 103, 108, 112

314

AUTHOR INDEX

Koomey, M., 80 KOOPS,H.-P., 117, 118, 124, 125, 159, 160,164

Korhonen, T.K., 61, 62, 75, 76, 77, 95, 106, 108, 111,113

Kornberg, H.L., 175, 227 Koronakis, V., 71, 108 Korthals, H.J., 132, 133, 158 Kostrikina, N.A., 121, 129, 130, 153,160, 162

Kotani, S., 245, 298 Kow, Y.W., 13, 15, 16,50 Kraft, J., 129, 160 Krag, S.S., 250, 299 Krahn, P.M., 87, 93, 108, 110 Kramer, C., 88,91, 104, 109 Kraus, J., 55,104,237,239,241,243,250, 301

Kraus, S.J., 63, 79, 100, 112 Krebs, H.A., 210,227 Kremer, B.P., 123, 160 Krishnapillae, V., 69, 109 Kristo, C.L., 82, 102 Krnjulac, J., 77, 113 Krol, A.J.N., 42, 50 Kronvall, G., 274, 275, 300 Kriiger, H.-J., 20, 21, 49, 50 Krump, M.A., 144,162 Krzycki, J.A., 182, 231 Kuenen, J.G., 116, 119, 120, 124, 125, 126, 127, 129, 130, 132, 150, 151, 152, 153, 155, 157,158,160,161,162 Kumar, S., 73, 108, 216, 223 Kundig, F.D., 250, 299 Kuntz, I.D., 218, 219, 227 Kuraishi, H., 119, 120, 160 Kusecek, B., 88, 102 Kushner, D.J., 167, 217, 227 Kushwaha, S.C., 185,227 Kusser, W., 272, 299 Kwan, S., 69, 112

I Laane, C., 27,50 Lai, C.Y., 183, 226 Laine, R.A., 240,243,250,253,254,261, 297,299 Laine, S., 69, 92, 108

Laing, W.A., 136, 140, 146,158, 161 Laishley, E.J., 119, 162 Laishley, J.W., 119, 158 Lake, J.A., 170, 171,227 Lambden, P.R., 100,108 Lambert, D.M., 146, 149, 163 Lambert, G.R., 5, 37, 44, 46, 49, 50 Lambert, P.A., 235, 277, 291,284,299 Lambert, R., 118, 164 Lanaras,T., 124, 125, 126, 127, 128, 131, 150, 152,159,161

Lancaster, J.R., 20, 50 Landgraf, H.R., 235,240,250,251,297 Lane, M.D., 209,225 Lang, N.J., 121, 161 Langley, K.E., 276, 299 Langridge, R., 65, 109 Langworthy, T.A., 170, 185,227 Lanter, J.M., 170, 226 Lanyi, J.K., 217, 218, 227 Lark, D., 55, 62, 76, 77, 83,95, 108, 109, 110

Larsen, H., 176, 217, 227 Lascelles, J., 40, 50 Lau, R.H., 170,226 Laudelout, H., 118, 164 Lavin, G.I., 246, 298 Lavin, K., 93, 110 Law, J.H., 240,299,301 Lawford, H.G., 31,48 Lawlis, V.B., 142, 161 Lawn, A.M., 58-59,60,61,72,85,86,89, 108,109

Leach, J.M., 54, 62, 110 Leadbeater, L., 150, 154, 161 Lebherz, H.G., 184,227 Lecatsas, G., 58-59, 104, 105 Lederberg, E.M., 88, 108 Lederberg, J., 88, 108 Lee, E.H., 137, 147, 159 Lee, J.S., 61,70,84,85,86,89,90,93,106 Lee, S.W., 57,108 Leeven, R., 95, 107 Leffler, H., 62, 108 LeGall, J., 15,20,2 1,49,50,51,205,226 Leicht, W., 220, 230 Lennarz, W.J., 245, 250, 258, 299, 300, 301

Leonard, K.R., 65, 106 Lepidi, A.A., 42, 51

315

AUTHOR INDEX

Lepo, J.E., 39, 50 Levine, G.A., 143, 162 Levine, M.M., 57, 62, 108 Levine, R.P., 88, 89, 110 Lewin, R.A., 122, 161 Lewis, B.J., 166, 225, 230, 231 Lewis, H.M., 207,229 Lidin-Janson, G., 75, 106 Lienhard, G.E., 195,227 Lim, S.T., 7, 50,245, 299 Lin, F.J., 198, 228 Lin, T.C., 87, 106 Lindahl, P.A., 21, 50 Lindberg, F.P., 76, 77, 94, 95, 108, 109 Lindberg, U., 75, 106 Lindenmaier, W., 76, 106 Lindquist, Y., 135, 137, 158, 162 Lindsay, B., 247,298 Line R.A., 240, 252, 255, 297 Linggood, M.H., 62,63,112 Linzer, R., 262, 299, 300 Linstowsky, I., 195, 225 Liu, T.Y., 284, 298 Liu, Y.-P., 57, 74, 112 Ljones, T., 24, 50 Ljundahl, L.G., 220,227 Ljungdahl, P.O., 20, 21,50 LO, H.-S., 180, 228 Lobos, J.H., 129,160 Lodish, H., 92, 113 Loffelhardt, W., 129, 146,157,161 Loh, B.A., 96,99,110 Lombardi, F.J., 260, 261, 299 Londesborough, J.C., 181,227 Long, M.V., 133,162 Long, S.R., 41, 49 Lopez, J., 65, 87, 113 Lorimer, G.H., 132, 135, 136, 137, 139, 140, 142, 143, 144, 150, 157, 158,161, 162 Lory, S., 97, 98, 107 Lottspeich, F., 203, 205, 228 Lotz, W., 46, 47, 51 Louis, B.G., 218, 227 Low, D., 76, 108 Lowenstein, J.M., 210, 227 Lubben, M., 181,223,227,228 Ludermann, H.-D., 222,223 Ludvig-Festl, M., 202, 204, 223 Luehrsen, K.R., 166, 169,225

Lugowski, C., 246,299 Lugtenberg, B., 87, 88, 107, 108 Lujik, L.W., 2, 52 Lund, B., 76, 77, 94, 95, 108, 109 Lyman, C., 127, 160 M

Maathuis, F.J.M., 129, 130, 160 McArthur, A.E., 271, 299 McArthur, H.A.I., 280, 282, 299 McCallum, L.C., 14,48 McCarty, M., 234, 236, 239, 241, 273, 299,300

McConnell, M.M., 62, 77, 78, 109, 112, 113

McEachran, D.E., 96, 108 McFadden, B.A., 116,121,122,132,133, 139, 142, 151, 154,157, 161,162, 163

McGee, Z.A., 57, 100,108,112 McGroarty, K.M., 119, 158 McIntire, S.A., 71, 105 McIntosh, L., 140,146,147,159,161,163 McIntyre, J.O., 21, 50 McKeller, R.C., 198, 229 McKern, N.M., 64,67,82,105, 109 MacLaren, D.M., 75, 113 McMichael, J.C., 94, 109 Macnicol, P.K., 32, 35, 47 McQuattie, A., 206, 207, 208, 218, 224 Maeba, P., 211,231 Magrum, L.J., 166, 167,223,225 Mah, R.A., 170, 171,227 Maier, R.J., 2, 5, 6, 7, 8, 9, 10, 13, 14, 16, 17, 18, 19,20,21,22,26,27,28,29,30, 31, 32,33, 34,35,36,37,38,39,40,44, 45, 46, 47, 49, 50, 51, 52 Makela, P.H., 76, 77, 95, 111 Makowski, L., 65, 108 Malhotra, S.S.,154, 160 Malsey, S.,65, 106 Mancuso, D.J., 247, 261, 299 Mangeney, E., 123, 132, 152, 161 Maniloff, J., 166, 225 Mankin, A.S., 170, 226 Mann, M.B., 78, 111 Mannheim, W., 181,226 Manning, 88, 108 Manning, P.A., 71, 72, 88, 89, 93, 102, 103, 107

316

AUTHOR INDEX

Manoil, C., 87, 108 Markham, J.L., 272,273,299 Marley, G., 57, 62, 108 Marrs, B.L., 134, 163 Marrs, C.F., 63, 64, 108 Marsden, W.J.N., 116, 117, 119,124,127, 129, 134, 142, 147, 150, 151, 154, 155, 158,161, 163 Marsh, D., 238, 299 Marsh, S.S., 34, 50 Marston, F.A.O., 156, 161, 162 Martin, C., 274, 298 Martin, M.N., 140, 142, 162, 164 Martin, P.A.W., 129, 161 Martin, W.G., 198, 229 Martiny, H., 124, 125, 159 Marvin, D.A., 65,66,67,87,93,106, 108 Matheson, A.T., 170, 171, 228, 230 Mathews, S., 203, 205, 228 Matlib, M.A., 211, 228 Matsubara, H., 205, 220, 226, 228, 230 Matsumura, M., 221, 228 Matsushiro, A., 221, 231 Matthew, M., 60, 107 Matthews, B.W., 70, 77, 103, 112 Matthews, R.A., 89, 109 Mattick, J.S., 82, 102, 108 Mattingly, S.J., 268, 295, 300 Mauck, J., 277, 299 Maule, 5.,61,68,70,71,84,85,88,89,113 Maurer, L., 74, 95, 108 Maxwell, E.S., 146, 149, 163 Mayer,F., 54,65,108,121,133, 134, 135, 157,158,164 Mayhew, S.G., 18,52 Meek, J., 144, 157 Meers, J.L., 291,300 Mekalanos, J., 63 Menzel, E., 135, 157 Merberg, D., 6, 7, 8, 10, 30, 31, 50 Merrill, W.W., 96, 110 Mevarech, M., 198, 205, 218, 219, 220, 226,228,230 Meyer, M., 42,49 Meyer, R.J., 69, 109 Meyer, T.F., 64,79, 80, 100, 101, 102, 106,109,111 Meynell, E., 58-59, 60, 61, 70, 71, 72, 85, 86,89,104, 107, 108, 109 Meynell, G.G.,58-59,60,68,70,71,108, 109

Michaelis, S., 92, 109 Michalowski, C., 129, 146, 157 Midollini, S., 20, 51 Mikkelsen, L., 94, 108 Milanez, S., 137, 159 Mildvan, AS., 135, 161 Miller, A.G., 151, 152, 153, 155, 163 Milstein, C., 80, 106 Minami, Y., 205,228 Minchin, F.R., 26, 27, 50, 51 Minion, F.C., 95, 109 Minkley, E.G., 60, 68, 69, 87, 88, 89,91, 82,107,109,110,111

Minnikin, D.E., 269,300 Minshew, B.H., 9, 75, 76, 107 Miorner, H., 274, 275, 300 Mirelman, D., 280, 295, 296, 300 Mitchell, C.G., 211, 228 Mitsui, Y., 65, 109 Miyachi, S., 127, 128, 142, 152, 157, 164 Miyashita, T., 279, 302 Miziorko, H., 132, 135, 136, 137, 142, 150, 161

Mizuno, J., 245, 298, 302 Mlawer, N., 64, 79, 109 Mobach, H.W., 89, 112 Mockel, W., 202,227 Molinari, A., 274,300 Mollby, R., 55, 62, 75, 107 Moller, J.K., 60, 105 Mooi, F.R., 54, 78, 94, 95, 109 Moon, H.W., 55,62,63,107,109 Moore, D., 69, 83, 92, 108, 109 Moore, P.M., 89, 113 Moras, D., 221,228 Morelli, G., 93, 102, 274,298 Morgan, P.,195,228 Morii, H., 248,258,260,272,299 Moriyama, T., 21 1,228 Morona, R., 87,88,109 Morris, D.C., 221,228 Morris, D.W., 58-59, 109 Morris, J.A., 55, 109 Morris, J.G., 175, 228 Morris, R.J., 26, 48 Morse, D.E., 183, 228 Mortenson, L.E., 2, 4, 16, 17, 18, 21, 47, 48

Moshiri, F., 13, 14, 18, 19, 21, 22, 39,45, 51,52

Mosser, J.L., 284, 300

317

AUTHOR INDEX

Motokawa, Y.,28, 30,51 Mott, M.R., 82, 108 Mottl, M.J., 155, 160 Moura, I., 15, 20, 21, 50, 51, 205, 226 Moira, J.J.G., 15,20,21,50,51,205,226 Moura, L., 2 0 , s Mucke, H., 129, 146, 157,161 Mullany, P., 62, 78, 109 Mullineaux, P., 71, 109 Mur, L.R., 123, 132, 133, 158 Murphey, W.H., 198,228 Murphy, J.R., 119, 161 Murray, R.G.E., 118,161 Murray, S., 131, 158 Mutaftschiev, S., 16, 17, 18, 28, 51

Norris,P.R., 180,181, 183,187,189,231 Norton, I.L., 133, 162 Novak, P., 45,51 Novitsky, T.J., 271, 300 Novotny, C.P., 89, 93, 104, 110 Nowitzki, S., 187, 189, 202, 205, 227 Ntamere, AS., 248, 260, 262, 274, 290,

300 Numberger, E., 184, 225 Nuti, M.P., 42, 51

0

Oakes, M., 170, 171,227 Oasahara, K., 221,231 Oberlies, G., 18 1, 228 Obermeier, W., 240, 300 N O’Brian, M.R., 7, 16, 17, 18, 27, 28, 29, Nadkarni, G.B., 184,226 30, 31, 32, 33, 34, 35, 36, 37, 51, OCallaghan, R.J., 73, 87, 93, 108, 110 Nadler, K.D., 40, 48 Odom, J.M., 17,51 Nagle, D.P., 182, 226 O’Donnell, I.J., 64,109 Nagy, B., 55, 63, 107, 109 Nakano, M., 237,240,241,243,250,252, Oesterhelt, D., 187, 189, 202, 203, 204, 253,255,297,300 205,206,220,226,227,228,230 Nargang, F., 140, 146, 147,161, 163 Ofek, I., 61, 110 Ogata, R.T., 88, 89, 110 Nathenson, S.G., 279,300 O’Gorman, L., 71,105 Nealon, T.J., 268, 273, 295, 300 Ogren, W.L., 133,140,141,142,144,145, Nelson, L.M., 4, 24, 25, 46, 47, 51 160,161,162, 163 Ner, S.S.,216, 228 OHanley, P., 55,62,63,76,77,83,94,95, Neuer, G., 202, 204,223 103,109 Neuhaus, F.C., 233, 242, 247, 248, 249, 252, 260, 261, 262, 263, 269, 270, 272, OHanley, R.,55, 62, 110 O’Hara, E.B., 6, 7, 8, 20, 30, 3 I , 50, 52 274,290,296,299,300,301 Neumann, E., 198,218,219,228 Ohba, M., 171,228 Ohlendorf, D.H., 70, 77, 103, 112 Neurath, H., 216, 223 Newcombe, E.H., 122,157 Okabe, A., 270,300 Okabe, K., 122, 128, 158 Nicolaus, B., 179, 183, 185, 196, 224 Old, D.C., 77, 110 Niederman, M.S., 96, 110 Oliver, R.N., 200, 224 Niemitalo, S., 274, 275, 298 Nierzwicki-Bauer, S.A., 121, 146, 147, Olsen, G.J., 166, 167, 168, 169, 170, 231 Olsen, K.W., 221,228 149, 161 Olsen, R.N., 58-59, 68, 72, 110 Nishihara, M., 248, 258, 260, 272, 299 Olsson, O., 62, 63, 76, 77, 94, 103 Nisioka, T., 106 Oosterhof, A., 243, 258, 274, 275, 300 Niskasaari, K., 274,275,298 Op den Kamp, H.J.M., 243, 258, 274, Nolteersting, U., 204, 223 Norgren, B., 55, 76, 109 275,300 Norgren, M., 76, 77, 83, 94, 95, 109, 113 Opgenorth, A., 61, 70, 84, 89, 90, 106 Normark, S., 55,62,63,76,77,83,94,95, Orefici, G., 274, 300 Orlandini, A,, 20, 51 103, 108, 109, 113

318

AUTHOR INDEX

Orme-Johnson, W.H., 21, 50 Orndorff, P.E., 74, 75,95, 108, 110 Orosz, L., 89, 110 Orskov, F., 54, 55, 61, 62, 63, 108, 110 Orskov, I., 54,55,61,62,63,108,109,110 Orskv, I., 61, 110 Osborn, C.B., 122,157 Osen, E.G., 57, 108 Oshima, T., 171, 181, 196, 228, 230 Ottow, J.C.G., 54, 57, 61, 110 Ou, J.T., 87, 89, 93, 94, 109, 110 Oulevey, J., 21, 52 Owen, P., 209, 228, 245, 275, 291, 300, 301

Peat, A., 121, 162 Pechman, K.R., 166,225 Peck, H.D., Jr., 17, 20, 21, 49, 50,51 Pedersen, K.B., 57, 111 Peeters, P.A.M., 274, 275, 300 Peiringer, R.A., 247,250, 275, 291,300 Pelroy, R.A., 143, 162 Penaranda, M.E., 78, 111 Pentilla, M.E., 76, 111 Perham, R.N., 87,103,184,200,202,206, 223,224,226,228,229 Perkins, H.R., 249, 270, 301 Perlman, D., 92, 111 Perlman, R.L., 72,111 Perumal, N.B., 88, 111 Perutz, M.F., 220, 221, 228 Peters, K.R., 118, 124, 128, 157, 162 P Petersen, G.B., 91, 107 Packer, L., 2, 52 Pettit, F.H., 206, 228 Pfennig, N., 116, 162 Packman, L.C., 200,229 F'fleiderer, G., 199, 226 Pahwa, K., 39,51 Phares, E.F., 133, 162 Pai, R.M., 184, 223 Palmer, L.B., 96, 110 Phelps, A.S., 4, 51 Phillips, A.L., 133, 146, 147, 159 Panicker, M., 87, 110 Phillips, D.A., 11, 12, 42, 44,45, 46, 48 Paradisi, S., 274, 300 Paranchych, W., 53-102, 55, 57, 58-59, Pierce, J., 137, 144, 157, 161, 162 61,64,65,66,67,69, 70, 71, 72, 73, 81, Pierce, J.K., 76, 104 83,84,85,86,87,88,89,90,91,93,94, Pieringer, R.A., 237, 241, 247, 249, 250, 251, 252, 261, 267, 272, 273, 274, 288, 96,97,9a, 99,103,106,108, 110,113, 114 290,296,298 Parenchych, W., 64,67,68,94,97,98,99, Pieroni, P., 85, 114 Pinkwart, M., 16, 51 111 Parish, J.H., 58-59, 109 Pitt, T.L., 63, 97, 104 Park, J.T., 295, 300 Plaut, G.W.E., 194, 195, 196,224,228 Parker, J.M.R., 61, 85, 114 Pless, D.D., 245,300 Parker, M.L., 97, 107 Poerio, E., 179, 183, 196, 224 Parmelee, D.C., 216, 223 Polen, S., 69, 91, 109 Parry, M.A.J., 133, 144, 146, 147, 154, Pon, N.G., 135,162 158, 159, 162 Poole, R.K., 7, 51 Partridge, C.D.P., 20, 51 Pope, L.M., 118,162 Pasloske, B.L., 64,81, 97, 99, 106, 110, Porte, E., 24, 27, 28, 29, 31, 37, 51 111 Porter, R.D., 146, 149,163 di Pasquale, G., 146, 149, 163 Portis, A.R. Jr., 133, 144, 145, 162, 163 Pastan, I., 72, 111 Postgate, J.R., 4, 38, 50, 51 Patil, D.S., 20, 21, 48, 51 Postma, P., 75, 113 Patnaik, R., 57, 62, 108 Poulsen, F.M., 51, 103 Payton, M.A., 172,202,228 Powell, D.A., 245, 291, 300 Pearce, W.A., 54, 100, 104, 110 Prakash, R.K., 42, 51 Pearlstone, J.R., 64,67,68,94,96,97,98, Pratt, K.J., 209, 228 99,110,111 Priefer, U., 45, 52

AUTHOR INDEX

Priestley, I.M., 129, 147, 163 Promes, L., 76, 105 Pugh, E.L., 185,227 Piihler, A., 41,45,52 Pullen, A.M., 216, 228 Pun, T., 214 Pundak, S., 219, 228 Punsalang, A.P., Jr., 63, 100, 111 Purohit, K., 9,51, 116, 134, 151, 154,161, 162 Pyle, M., 96, 111

Q Quebedeaux, B., 6,26,27,48,49 Quivey, R.G. Jr., 132, 146, 148, 162, 163

R Rabin, B.R., 135, 162 Rabinowotz, J.C., 202,230 Rachlin, J.W., 121, 160 Racker, E., 172,228 Rahmsdorf, U., 69,88,107 Rainey, A.M., 134, 146, 148, 149, 163 Rajbhandary, U.L., 241,300 Raleigh, D.P., 182,225 Ramakrishnan, V., 136, 159 Ramaley, R., 197, 225 Ramphal, R., 96,111 Rao, J.K.M., 218,228 Rao, K.K., 205,220,226 Ray, A., 71,104,105 Rayter, S., 285, 286, 288, 289, 301 Read, M.J., 94, 105 Reaveley, D.A., 291, 301 Reddy, G.A., 137,162 Redmond, J.W., 242,243, 29 1, 294,2% Reed, L.J., 200, 206, 224, 228 Reeves, R.E., 180, 228 Rehemtullah, A., 71, 111 Reich, M.H., 200, 229 Reichert, R.W., 96, 111 Reistad, R., 218, 219, 229 Reith, M., 146, 162 Remington, S.J., 216, 229, 230 Remsen, C.C., 118,164

319

Reusch, V.M., Jr., 262, 270, 275, 300 Reyn, A., 63, 79, 107, Reynolds, D.M., 119,162 Reynolds, H.Y., 96, 110 Reynolds, J.A., 274, 298 Rhen, M., 61, 62, 76, 77, 111 Rhen, V., 76,111 Rice, D.W., 199,229 Rich, A., 145,158 Richards, H., 58-59,104 Richardson, C.C., 99-100,112 Richarme, G., 209,229 Richter, P., 62, 63, 107 Riede, I., 88, 111 Ristaino, P., 57, 62, 108 Rivera-Ortiz, J.M., 3, 51 Robbins, K., 64,79,80,81,100,101,103, 112 Roberts, M.F., 182, 184, 225, 227 Robertson, J.N., 100,113 Robertson, J.P., 64,111 Robertson, L.A., 116, 119, 161, 162 Robinson, M.D., 250, 299 Robinson, M.S., 21 1,229 Robison, P.D., 140, 162 Robson, R.J.,289, 301 Robson, R.L., 4,51 Roche, T.E., 200,224 Rogers, H.J., 285,286,288,289,291,301 Rogers, L.J., 134, 159 Rollin, F., 170, 230 Romanova,A.K., 121,129,130,153,160, 162 Romermann, D., 42,49 Roorda, I., 62, 63, 105 Roosendaal, B., 94, 111 Rose, LA., 195, 227 Rose, Z.B., 195,229 Rosel. P.. 231. 236. 237. 239. 241. 242. Roseman, S., 250, 299 Rosenbusch, J.P., 87, 108 Rosenthal, A.F., 21, 50 Rosenthal, S., 89, 112 Rossman, M.G., 221,228 Rostami-Rabet, A., 123, 160 Rothbard, J.B., 63, 64,94, 100, 101,111, 108 Rothfield, L., 71, 105

320

AUTHOR INDEX

Rothman, J.E., 276,301 Rowe, B., 62, 77, 78, 105, 106, 109, 112, 113 Rowell, P., 131, 158 Royer-Pokora, B., 76, 106 Ruhland, G.J., 237, 241, 301 Ruiz-Argueso, T., 4, 16, 17,24,25,44,47, 49,51 Russell, S.A.,4,5,6, 16, 17,20,37,38,41, 43,44,46,47,49,50,51 Rutherford, E.L., 58-59, 104 Rutter, J.M., 62, 112 Rutter, W.J., 184, 227 Ruvkun, G.B., 41,51 S

Sabesan, K.N., 221,228 Sacconi, L., 20, 51 Sadoff, H.L., 197,229 Sadoff, J.C., 96,111 Salemme, F.R., 68, 113 Salipigni, J.D., 96, 111 Salit, I.E., 57, 61, 62, 94, 106, 111 Salminen, SO., 4, 24, 25, 47, 51 Salnikow, J., 148, 160 Salsac, L., 25, 49 Salton, M.R.J., 245, 275, 300,301 Salton, R.J., 245, 299 Saluja, A., 121, 163 Salvucci, M.E., 133, 144, 145, 162 Sambucetti, L., 71, 111 Sanderson, K.E., 71, 111 San George, R.C., 217,226 Sanger, F., 166,229 Sani, A., 134, 162 Santos, M., 205,226 Sanwal, B.D., 211, 231 Sapshead, L.M., 28, 30,51 Sarles, L.S., 132, 163 Sarner, J., 185, 230 Sarvas, M., 76, 111 Sasaki, T., 28, 30, 51 Sastry, P.A., 58-59,64,67,68,94,97,98, 99,103,110,111, 113 Saunders, R.E., 116, 119, 120, 124, 125, 163 Saunders, V.A., 129,162 Sawyer, W.D., 63, 100, 111

Saxen, H., 75,108 Scardovi, V., 174,229 Schacht, J., 202,227 Schaechter, M., 267, 296, 301 Schafer, G., 181,223,227,228 Schafer, S., 181, 188, 189, 229 Schatz, G., 216, 229 Schauer, D., 74, 75, 110 Shaup, H.W., 166,230 Schavass, V., 169,227 Scheinman, A., 170, 171,227 Schell, J., 69, 107 Schenk, H.A.E., 123,162 Scher, M., 258,301 Schilperoort, R.A., 42,49, 51 Schink, B., 8, 14, 16, 51 Schlegel, H.G., 2,8, 14, 16, 18, 19,27,28, 42, 48, 49, 51, 116, 121, 129, 133, 135, 148,157, 158, 159,162, 164 Schleifer, K.H., 184, 225, 237, 239, 241, 243,250,301 Schlesier, M., 8, 51 Schloss, J.V., 133, 137, 161, 162 Schlosser, U., 118, 164 Schmidmayr, W., 91, 104 Schmidt, J.M., 55, 111 Schmidt, M.A., 254,297 Schmit, AS., 245, 300 Schmitt, R., 65, 108 Schmuck, L., 23 1,237,241,272,277,278, 280,297 Schnabel, R., 169,231 Schnare, M.N., 167, 229 Schneider, G., 135, 137, 162 Schneider, J.E., 242, 272,301 Schneider, K., 2, 16, 18, 19,20,41,49,51 Schnoss, M., 58-59, 89, 104 Schoemaker, J.M., 156, 162 Schoener, B.E., 156, 162 Schoener, R.G., 156, 162 Schonheit, P., 20, 52 Schoolnik, G., 55, 62, 63, 64, 76, 77, 83, 94,95,103, 108, 109, 110 Schoolnik, G.K., 64, 94, 100, 101, 111 Schopf, J.W., 172, 191,225 Schubert, K.R., 3, 4, 5, 24, 51 Schulman, H.M., 40,48 Schulte, T., 71, 105 Schulz, G.E., 199, 229 Schuster, D., 243, 250, 254, 297

AUTHOR INDEX

Schweizer, M., 87, 111 Schwillens, P., 76, 105 Schwuchow, S., 88,93,102 Scotland, S.M., 62, 77, 78, 105, 106, 109, 112 Scott, J.R., 82, 107 Scraba, D.G., 61, 85, 86, 90, 93, 106 Searcy,D.G., 171,180,181, 182,221,226, 228,229 Searle, J.W., 4, 25, 49 Seefeldt, L.C., 14,48 Seely, R.J., 184, 229 Seemann, J.R., 162, 144 Segal, E., 64,79, 80, 100, 101, 102, 106, 111 Segerer, A., 187, 229 Seifert, H.S., 79, 111 Sekikawa, I., 237, 241, 301 Sellwood, R., 62, 63, 104, 112 Senior, P.J., 210, 229 Servaites, J.C., 144, 159, 162 Shairer, H.U., 40, 49 Shanmugam, K.T., 7,50 Shapiro, J.A., 60, 112 Sharp, P.A, 69,70, 112 Shaw, D.R.D., 249,295,299,300 Shaw, K.M., 198,229 Shaw, N., 236,301 Shaykh, M.M., 151, 153, 162 Shedlovsky, T., 246, 298 Sheehy, J.E., 26,27,50,51 Sherod, D., 220,227 Sherwood, P.M.A., 294,296 Shimada, A., 237,238,241,283,295,298 Shinozaki, K., 146, 147, 149, 163 Shipley, P.L., 58-59, 63, 72, 110, 112 Shiraiwa, Y.,127, 164 Shively, J.M., 116, 117, 119, 120, 121, 124, 125, 126, 127, 129, 134, 142, 146, 147, 149, 150, 152, 154, 157, 158, 160, 163,164 Shockman, G.D., 234,239,247,267,271, 272, 273, 274, 285, 286, 288,289, 290, 295,296, 297,299,301,302 Shoham, M., 220,229 Short, S.A., 258,270,301 Shug, A.L., 4,52 Shungu, D.L., 247,290,301 Shuta, A., 89, 112 Siegelman, H.W., 132, 163

32 1

Sietsma, J.H., 41, 48 Siitonen, A., 95, 113 Silva, B., 69, 107 Silver, R.P., 54, 56, 62, 105 Silverman, P.M., 60, 68, 70, 71, 89, 92, 105,111,112 Sim, E., 15,52 Simion, F.A., 267, 270, 299 Simon, R.,41,45,52 Simpson, F.B., 3, 6, 9, 26, 38, 50,52 Simpson, W.A., 273,274,297,Singh, M., 211, 229 Singleton, R., Jr., 220, 229 Sirgel, F.A., 58-59, 104, 105 Skurray, R.S., 60,68, 69, 70, 71, 72, 83, 88,89,90, 92, 102, 104, 105, 106, 107, 113 Slack, C.R., 198, 226 Slade, H.D., 234, 245, 298, 301 Slater, E.C., 20,47 Sleeper, B.P.,127, 159 Sletten, K., 63, 106 Small, C.L., 146, 149, 156, 163 Smillie, L.B., 64,67,68,94,96,97,98,99, 110,111 Smith, A.J., 116, 117, 118, 134, 159, 162, 163 Smith, G.D., 20,48 Smith, H.R., 62, 77, 78, 109, 112, 113 Smith, H.W., 54, 62, 63, 110, 112 Smith, I.C.P., 190, 225 Smith, I.W., 54, 105 Smith, J., 200,224 Smith, L., 196, 207, 208, 214, 229 Smith, R., 274, 301 Smith, R.V., 116, 159 Smith, W.W., 135, 158, 159 Smyth, C.J., 54, 57, 62,78, 108, 112 Snead, R.M., 125, 142,163 So, M., 64,67, 79, 80, 81, 100, 101, 102, 105,106,109,111,112 Sodano, S., 185, 224 Sogin, M., 166,231 Sojka, W.J., 54, 55, 62, 109, 110 Somerville, C.R., 140, 144, 146, 147, 149, 156,159, 161,163 Somerville, S.C., 140, 146, 147, 149, 156, 163 Sone, N., 30,52 Sowa, B.A.D., 83,69,109

322

AUTHOR INDEX

Sparling, P.F., 101, 112 Spears, P.A., 74, 75, 110 Speert, D.P., 81, 96, 99, 110 Spencer, D.F., 167,229 Spierings, G., 76, 113 Spragg, P., 200,224 Sprott, G.D., 190, 198,225,229 Srere, P.A., 211, 228,229 Srivastava, S., 73, 108 Stackebrandt,E., 118,157,166,221,225, 229

Staden, R., 67, 103 Stahl, D.A., 166, 225, 231 Stallions, D.R., 93, 105 Stam, H., 25, 26,52 Stanfield, S., 41, 48 Stanier, R.Y., 121,123,143, 155,160,163 Stanley, C.J., 200, 229 Starnes, S.M., 146, 149, 163 Stein, D.B., 171, 181, 229 Steiner, I., 182, 225 Steinmuller, K., 146, 163 Stenderup, A., 60, 105 Stephens, D.S., 57, 112 Stephens, P.E., 207,229 Stetter, K.O., 167,169,177,187,221,222, 223,225,226,229,230,231

Steven, A.C., 68, 112 Stevens, S.E., Jr., 121, 146, 149, 161, 163 Stevenson, K.J., 206, 207, 208, 209, 218, 224,228

Stewart, D.J., 64, 67, 82, 105, 109 Stewart, W.D.P., 116, 117, 121, 122, 123, 124, 125, 129, 131, 133, 140, 142, 146, 147, 148, 149, 150, 154,158,159,163 Stezowski, J.J., 187, 189, 198, 221, 225, 226 Stieritz, D.D., 97, 112 Stocker, B., 71, 111 Storzbach, S., 64, 79, 80, 109, 111 Stouthamer,A.H.,25,26,31,52,174,230 Strauss, D.C., 63, 96, 114 Stribling, D., 184, 229 Stringer, C.D., 133, 136, 137, 147, 159, 162 Strominger, J.L., 279, 300 Stults, L.W., 13, 14, 18, 19,20,45,51,52 Stupperich, E., 182, 184, 189, 202, 215. 225,226 Sud, I.J., 267, 301

Suda, K., 216,229 Sugino, Y ., 22 1, 231 Suguira, M., 146, 147, 149, 163 Suh, S.W., 135, 158, 159 Suissa, M., 216, 229 Summers, A.O., 129, 158 Sundaram, T.K., 198,229 Sundermeyer-Klinger,H., 118, 157 Suryanarayana, M.P., 184, 223 Susskind, B., 180,228 Sussman, J.L., 217, 220, 229, 230 Svanborg-Eden, C., 62,75,76,77,83,95, 106,108,109, 112, 113

Svenson, S.B., 55, 61, 62, 75, 76, 95, 107, 108, 111,113

Swaney, L.M., 57,74,112 Swanson, J., 55, 63, 64, 79, 80, 81, 100, 101, 103, 104,112

Sweeley, C.C., 258, 301 Szymona, M., 179,229

T Tabillon, R., 20, 52 Tabita, F.R., 116,132, 133, 134,138, 140, 142, 143, 146, 148, 149, 156, 159, 160, 161, 162, 163,164 Tabor, S., 99-100, 112 Tai, J.Y., 94, 100, 101, 111 Tai, S., 245, 298 Tainer, J.A., 67, 105 Takabe, T., 132, 138, 139, 145, 146, 149, 154, 156,157, 160,164 Takahata, N., 146, 147, 163 Takeda, Y., 70,77,103, 112 Takeya, K., 96,112 Talamo, B., 245, 250, 258, 299 Talgoy, M.M., 210, 230 Tallgren, L.G., 95, 113 Tamura, Y., 97, 112 Tanaka, S., 97, 112 Taneja, A.K., 61, 85,114 Tanford, C., 238,274,301 Tanimoto, K., 72, 112 Tanner, R.S.,166, 225 Tarelli, E., 234, 279, 297 Taron, D.J., 249,252, 260,261,262,263, 269,2%, 300,301 Taylor, B.F., 119, 163

323

AUTHOR INDEX

Taylor, C., 285, 286, 288, 289, 301 Taylor, D.E., 58-59,69,104,112 Taylor, L., 70, 71, 73, 106, 112 Taylor, S.C., 133, 163 Taylor-Robinson, D., 100, 108 Teimer, E., 272, 302 Teixeira, M., 15, 20, 21, 50, 51 Tel-Or, E., 2, 52 Tempest, D.W., 234, 268, 271, 291, 297,

300 Teng, N.H., 64,101,111 Tennigkeit, J., 24, 48 Teo, B.K., 21,50 Terzaghi, B.E., 146, 158 Teti. G.. 274. 300 Thauer,’R.K., 20, 49, 52, 181, 189, 198,

Truelsen, T.A., 46, 52 Trus, B.L., 68,112 Tschape, H., 58-59,104, 113 Tsernoglou, D., 220, 226, 230 Tsolas, O., 183,226 TSO,M.-Y., 24, 52 Tsukihara, T., 205, 230 Tsuzaki, N., 119, 120,160 Tuner, G.L., 32,47 Tuovinen, O.H., 129,161 Turner, A.P.F., 116, 158 Turner, G.L., 26, 32, 35,48 Turpin, D.H., 151, 152, 153, 155, 163 Tweten, R.K., 127, 159

202,225,228,231

Theodore, T.S., 275, 298 Thiele, O.W., 21, 52 Thierry, J.C., 221, 223 Thomas, J., 240, 301 Thomm, M., 177,230 Thompson, R., 70, 71, 73, 106, 112 Thornber, J.P., 170, 171,227 Thorns, C.J., 55, 109 Tiboni, O., 146, 149, 163 Tichy, H.V., 46,47, 51 Tietze, E., 58-59, 113 Tilton, R.C., 119, 161 Timmis, K.N., 68,71, 112 Titani, K., 216, 223 Tolman, C.J., 182, 184, 225 Tomany, M.J., 141, 160 Tomasz, A., 246,247, 273, 274,283, 284, 285, 286, 289, 290,295, 2%, 298,300, 301 Tomlinson, G.A., 176, 177, 197, 230 Tommassen, J., 88, 107 Tomoeda, M., 54,67,83,89,91,105,112, 113 Tong, E.K., 211, 230 Toon, P., 234, 237, 239, 251, 261, 301 Torii, M., 245, 298 Torisky, R.S., 144, 162 Torres-Ruiz, J.A., 133, 139, 163 Toth, M.N., 129, 163 Trench, R.K., 122,159 Trent, J.D., 221, 230 Tropp, B.E., 248,297 Trovatelli, L.D., 174, 229

U Uchida, T., 166, 230 Uchikawa, K., 237, 241, 301 Uhlin, B.E., 76, 77, 83, 94, 95, 103, 106, 109,113

Uyeda, K., 202,230

V Vaisanen, V., 75, 95, 108, 113 Vakeria, D., 129, 146, 147, 148, 149,163 Valentine, R.C., 72, 89, 107, 112 Valle, E., 146, 160 Valois, F.W., 118, 164 van Alphen, L., 87,108 van Berkum, P., 10, 11,50 van Breemen, J.F.L., 1 19, 120, 124, 125, 126, 127, 152, 160

van Bruggen, E.F.J., 119, 120, 125, 160 van Brussel, A.A.N., 42, 49 van Den Bos, R.C., 42,50 van den Bosch, H., 76,113 van den Bosch, J.F., 75,76, 105,113 van den Handel, C., 76, 113 Van de Rijn, I., 239, 273, 299 van der Vies, S.M., 146,149,156,158,159 Van der Werf, A.N., 16,52 Van der Westen, H.M., 18,52 van de Wiel, C.C.M., 132, 133, 158 Van Die, I., 62, 63, 74, 76, 105, 108, 111, 113

Van Driel, D., 274,275,301

324

AUTHOR INDEX

van Eykelenburg, C., 121,164 van Gefen, B., 76, 113 van Gemerden, H., 116, 154,161,164 van Gool, A.P., 118,164 van Halbeek, H., 243,300 van Kamen, A., 42,50 van Megan, I., 76,113 Van Montagu, M., 69,107 van Pelt-Heerschap, H., 95, 107 van Slogteren, G.M.S., 42, 49 Van Verseveld, H.W., 25, 26, 31, 52 Van Vliet, F., 69, 107 Vavougios, J., 61, 62, 95, 111 Veeger, C., 18, 27,50,52 Veenhuis, M., 132, 133, 158 Veerkamp, J.H., 243,245,250, 258, 274,

Walsh, C.T., 21, 50 Walsh, K.A., 216, 223 Walther-Mauruschat, A., 121, 164 Wang, L., 64, 101,111 Warburg, O., 140, 164 Ward, J.B., 234, 235, 249, 262, 268,210,

275,300,301 Venema, J., 95, 107

Way, A.W., 57, 108 Wearne, S., 214 Weaver, K.E., 132, 134, 163, 164 Webb, L., 220,226 Weber, D.F., 10, 11,50 Weber, F., 20,52 Weber, G., 41,52 Weber, M.M., 205,220,226,230 Weber, P.C., 68, 113 Webster, L.T., Jr., 181, 227 Webster, R.E., 65,87,106,113 Wehrmann, H., 118,159,160 Weimer, P.J., 190, 214, 230 Weiss, R.L., 55, 63, 67, 113 Weitzman, P.D.J., 175, 193, 195, 198,

Venkitasubramania, T.A., 184, 223 Verboom-Sohmer, U., 75, 113 Vetter, H., Jr., 202, 227 Viale, A.M., 146, 149, 156, 160, 164 Vignais, P.M., 15, 20, 24, 27, 28, 29, 31, 37,48,51,52

Villarroel, R., 69, 107 Vilncent, P., 64,111 Virji, M., 102, 113 Virrankoski-Castrodeza, V., 58-59, 109 Visentin, L.P., 170, 230 Vogel, H.J., 69, 88, 103 Volpel, K., 81, 96, 99, 110 vu, C.V., 144,164

W Wachtel, E.J., 217, 219, 220, 225, 231 Wada, K., 205,228,230 Waddill, F.E., 132, 163 Wade, R.D., 216,223 Wagner, A.F.V., 216, 223 Wahlstrom, E., 61, 62, 76, 111 Wakabayashi, S., 205,220,226,228,230 Wakagi, T.W., 171, 181, 228,230 Walker, C.C., 24, 52 Walker, G.C., 68, 69, 72, 113, 114 Walker, J.E., 87, 103, 221, 223 Walker, J.M., 62, 106 Walker, N., 207 Walsby, A.E., 122, 159

278,285,286,288,289,301

Ward, M.E., 100, 113 Ward, M.W., 64,111 Warren, L.G., 180, 228 Wassef, M.K., 185, 227, 230 Waterbury, J.B., 118, 164 Watson, S.W., 118, 161, 164 Watt, P.J., 100, 108, 113 Watts, T.H., 64,65,66,67,94,97,99,106, 110, 113

200, 210, 211, 212, 213, 214, 215, 216, 224,226,228,229,230 Welch, G.G., 129, 160 Werber, M.M., 217, 230 Werneke, J.M., 145, 162 Westphal, K., 118, 124, 125, 126, 128, 129, 157, 163,164 Westphal, M., 284, 285, 301 Whatley, F.R., 180, 181, 229 Whatley, J.M., 122, 164 Wheelis, M.L., 8, 48 Whelan, J., 58-59, 104 White, D.C., 258, 270, 301 White, R.H., 222, 230 Whitman, W.B., 142, 164, 167, 182, 226, 230 Whittingham, C.P., 154, 158, 162 Whitton, B.A., 121, 161, 162 Wicken, A.J., 26,234,235,236,239,242,

325

AUTHOR INDEX

243, 245,247,267,268,269,271,272, 273, 274, 275, 285, 286, 288, 289, 290, 291, 294, 295,2%, 297,298,299,301, 302 Wickner, W., 92, 113 Wiegand, G., 216,229,230 Wildman, S.G., 145, 160 Wildner, G.F., 140 164 Wilke-Douglas, M., 148, 149, 157 Wilkins, B., 60, 68, 113 Wilkinson, A.E., 198,229 Wilkinson, S.G., 235, 302 Willetts, N.S., 57, 60, 61, 68, 69, 70, 71, 72,83,84,85,88,89,90,102,103, 104, 106,109,113 Williams, C.H., Jr., 200, 206, 230 Williams, D.L., 260, 207, 224 Williams. E.. 116. 158 Williams; P.A., 58-59,104 Williams, R.P., 100, 107 Willshaw, G.A., 77, 78, 112, 113 Wilson, P.W., 4, 51, 52 Wimpenny, J.W.T., 28, 30,51 Winans, S.C., 68, 69, 72, 113, 114 Winberg, J., 55, 62, 75, 107 Winblad, A., 62, 107 Windhovel, Y., 148, 149, 158 Winter, H., 182, 225 Winters, C., 88, 89, 110 Wise, E.M., Jr., 272, 302 Wiseman, G., 280, 298 Wiseman, R.L., 105 Wittenberg, B.A., 26, 48 Wittenberg, J.B., 26, 48 Wittig, W., 62, 110 Witty, J.F., 26, 27, 50, 51 Woese,C.R., 166, 167, 168, 169, 170, 171, 223,225,226,230,231 Wolf, G., 181, 226 Wolfe, R.S.,166, 167, 223, 225, 231 Wolk, C.P., 121, 122, 164 Wolters, J., 169, 231 Wonacott, A.J., 221, 231 Wong, M.-Y., 39 Wong, T.-Y., 27, 52 Wong, W., 290,295,302 Wood,A.P., 180, 181, 183, 187, 189,231 Wood, E.A., 45,48 Wood, P.A., 187, 189, 195, 196, 198,213, 214,215,224

Wood, R.C., 240,299 Woodland, D.L., 187, 189,213,214,215, 224 Woods, D.E., 63,96,97, 114 Wootton, J.C., 110 Worobec, E.A., 61,85,99, 113, 114 Wright, I.P., 198, 229 Wright, J.A., 21 1,231 Wright, P.E., 26,48 Wu, J.Y., 21 1, 231 Wu, T., 295, 300 Wullenweber, M., 117,118, 124,125,159, 164 Wyndaele, R.,46, 52

X Xavier, A.V., 15, 20, 2 1, 50,51, 205, 226 XU,L.-S., 5, 13, 14, 21, 46, 49 Xuong, N.-H., 135,158

Y Yagawa, Y., 127,164 Yaguci, M., 170, 230 Yamada, C., 146, 147,163 Yamakawa, T., 235,298 Yan, W., 69,112 Yanagita, Y., 30, 52 Yanamota, T., 245,302 Yang, J.T., 21 1,231 Yates, M.G., 16, 20, 24, 51, 52 Yayanos, A.A., 221,230 Yeaman, S.J., 206, 228 Yellowlees, D., 139, 157 Yokayama, K.,279,302 Yoshioka, T., 270,299 Young, C., 57,62, 108 Young, N.M., 246,299 Yura, T., 87, 110 Yutani, K., 221,231

Z

Zablen, L.B., 166, 225 Zablin, L., 166, 230

326

AUTHOR INDEX

Zaccai, G., 219, 220, 231 Zam, Z.S., 96, 111 Zeikus, J.G., 182, 189, 190, 198,202,214, 224,227,230,231 Zetsche, K., 146, 163 Zhang, J.M., 68,112

Zillig, W.,166, 167, 169, 177, 187, 227, 229,230,231 Zuber, H., 220, 221, 231 Zuber, M., 37,50 Zuidweg, E., 76, 113 Zykalova, K.A., 121, 129, 130, 153,162

Subject Index Abbreviations: 6PGLU, 6-Phosphogluconate; RuBisCO, Ribulose 1,5-bisphosphate carboxylase/oxygenase. A

Absorption spectra, hydrogenase from R. japonicum bacteroids, 14-15 R. japonicum bacteroid membranes, cytochrome reduction, 33 R. japonicum hydrogenasederepressed membranes, free-living, 29 Acetate production by Halobacterium saccharouorum, 177 in pulses-chase experiments of lipoteichoic acid synthesis, 252, 253 Acetazolamide, evidence for external carbonic anhydrase, 128 Acetone, extraction of lipid, 21 Acetylation, in propolin to pilin conversion, 69, 92 Acety I-CoA, affinity of citrate synthase for, ATP sensitivity, 210-21 1 conversion into acetate in T. acidophilum, 180 formation from pyruvate, 175 in archaebacteria, 186 evolution of reaction, 193 oxidation in citric acid cycle, 175, 176 synthesis in methanogenic archaebacteria, 184 Acetyl-CoA synthetase, ADP-forming, 180 AMP-utilizing, 181 N-Acetyl-D-mannosamhe in Staph. aureus linkage unit, teichoic acid synthesis, 278

Acetylene reduction, 12 8-N-Acetylglucosaminidaseinhibition, 285 alanyl-ester effect, 287-288 N-Acetylmuramyl-L-alanineamidase, 285 distribution in pneumococci, 284, 295 role in autolysin activity control, 285 teichoic acid requirement, 283-284 Achromobacter, modified EntnerDoudoroff pathway in, 179 Acinetobacter anitratum, 2 16 Actinomycetes, poly(g1ycerophosphate) lipoteichoic acids absent, 245 Active-site coupling, 200 Adenosine diphosphate (ADP), in conversion of acetyl-CoA into acetate, 180 specificity of succinate thiokinase, 215 Adenosine monophosphate (AMP), reactivation after NADH inhibition of citrate synthase, 210 Adenosine monophosphate (AMP)utilizing acetyl-CoA synthetase, 180-181 Adenosine triphosphatase (ATPase), absence from Thermoplasma acidophilum, 18 1 Adenosine triphosphate (ATP) citrate synthase inhibition, in archaebacteria, 214 in eubacteria and eukaryotes, 21021 1 concentration in nitrogen fixation reaction, 3 formation, in Embden-Meyerhof pathway, 172, 191

328

SUBJECT INDEX

Adenosine triphosphate (ATP) (contd.) in Entner-Doudoroff pathway, 172 reoxidation of NADH and FADHZ, 175 hydrogen oxidation coupled to, hup probes, 47 increase, in host control of hydrogenase, 13 requirements of hydrogen evolution by nitrogenase, 24 synthesis, by uptake hydrogenase action, 4 electron transport coupled to, 35 hydrogen oxidation-dependent, 2425,47 Adhesin, 83 K88 pili, 95 N. gonorrhoea pili, 100, 101 Pap pili, 55, 61, 94-95 genes for, 76, 77 pilE locus expressing, 74 Type I pili, 95 x, 95 Aerobic micro-organisms, 2, see also Eubacteria Alanine, codons, 218 in halophilic enzymes, 2 18, 2 19 Alanine/phosphate ratio, lipoteichoic acid, 290, 294 Alanyl residues in lipoteichoic acids, addition of, 262-263,276 anti-autolytic activity, effect on, 287, 290 base-catalysed hydrolysis, 263, 264 distribution of, 242, 243 effect on lipoteichoic acid carrier activity, 280-28 1, 283 glucose effect on, 271 magnesium ion binding, effect on, 294 model of structure, 292 pH and temperature effect, 271 re-esterification after loss, 265, 266,276 salt effect on, 270, 271 site of incorporation, 263 species with, 240, 241 transport to teichoic acid, 263-265, 276 Alaska, pea cultivar, 1 1, 12

Alcaligenes, modified EntnerDoudoroff pathway in, 179 Alcaligenes eutrophus, hydrogenase, absorption spectrum, 14 composition and antibody crossreactions, 14 electron acceptor reactivity, 16 femcyanide stability of, 19 K , value, 16 NADH-linked hydrogen oxidation, 28 nickel in, 20 oxygen stabilization of, 18 RuBP correlation absence and, 9 hydrogen as energy source with low oxygen, 8 hydrogen oxidation-dependent ATP synthesis, 24 hydrogen oxidation in, electron transport, 27 mixotrophic growth, 8 oxygen sensitivity negative (Ose-) mutants, 8 RuBisCO, activation, 136 gene cloning, 149 gene number, 148 structure in, 134-135 Alcaligenes eutrophus ATCC, 148, 149 Alcaligenes eutrophus H 16, hydrogenase, genes on megaplasmid, 148 phosphoribulokinase and RuBisCO gene location, 148 plasmids in, 42 Alcaligenes eutrophus TF93, 42 Alcaligenes eutrophus TF93 1, Hoxmutants, 42 Alcaligenes latus hydrogenase, composition and antibody crossreactions, 14 electron acceptor reactivity, 16 K , value, 16 Alcaligenes vinelandii, hydrogenase, composition and antibody crossreactions, 14 electron acceptor reactivity, 16

329

SUBJECT INDEX

Alcaligenes vinelandii (contd.) molecular weight, 13 hydrogen oxidation in, electron transport, 27 Aldolase, class I and 11, 183-184 Amino acids, polar, in halophilic enzymes, 218 (p-Aminophenyl) dichloroarsine, 207 Ammonia, dinitrogen reduction to, by nitrogenase, 2 , 4 Ammonia-oxidizing bacteria, carboxysome distribution and structure, 117-1 18 Amphiphiles, 234, 236 Anabaena, carboxysome association with microtubules, 121-122 Anabaena 7 120, RuBisCO genes, 146 Anabaena cylindrica, 20 hydrogen oxidation, reducing quivalent donation, 24 RuBisCO in carboxysomes, evidence, 131 Anabaena PCC7120, plasmid in, 129 Anabaena variabilis, carbonic anhydrase in, 127 K,,, (COS values of RuBisCO, 142 nickel in, 20 Anacystis nidulans, see also Synechococcus RuBisCO, gene cloning and location, 146, 149 L subunit probe, hybridization, 147 nucleotide sequence of gene, 146147 Anacystis nidulans R2, RuBisCO in carboxysomes, evidence, 131 Anaerobes, glycolysis in, 172 hydrogen evolution and oxidation, 2 obligate, methanogens as, 167 pyruvate metabolism, 175 Anaerobic conditions, 2-0x0 acid oxidoreductases in, 202, 203,204 2-0x0 dehydrogenase (NAD+) unsuitable in, 202 Antibodies to poly(g1ycerophosphate) chain of lipoteichoic acids, 274

Anti-F pilus antiserum, 86 Antigenic determinants, adhesive pili, 62, 94 conjugative pili, 85-86 NMePhe pili 97, 99 N . gonorrhoeae, 63,94, 101-102 Antigenic variation, adhesive pili, 62, 94 gonococcal pilin genes, 80-8 1, 100, 102 Antimycin A, 28 Anti-PAK antiserum, 8 1 Anti-PA0 antiserum, 8 1 Anti-pED208 pilus antiserum, 85, 86 Anti-pilus antibodies, 72, 93 Aphanothece halophytica, RuBisCO heterologous subunit reconstruction, 138-1 39 RuBisCO S subunit function, 138 Arabidopsis thaliana, mutant, RuBisCO not activated in light, 144 RuBisCO activase, 144, 145 Arabinose in repression of hydrogenase activity, 6 oxygen-insensitive mutants, 7 Archaebacteria, 166-172, see also individual species biochemistry, 170-1 71 cell-envelope in, 170 central metabolism, 176-193 citric acid cycle, 186-190 evolutionary origins, 191 gluconeogenesis, 183-1 85, 191 glycerol synthesis, 185-1 86 hexose catabolism, 176-182 patterns of, 190-193 concept of phylogenetically distinct group, 167, 168 dihydrolipoamide dehydrogenase in, 206-209 elongation factor, (EF-2) in, 171 enzyme diversity, 194-2 17, see also Dehydrogenases with dual cofactor specificity citrate synthase and succinate thiokinase, 213-216 2-0x0 acid: ferredoxin oxidoreductases, 193, 199-205 %

330

SUBJECT INDEX

Archaebacteria (conrd.) enzymes, adaptation of for extreme conditions, 217 halophilic, 2 17-220 salt bridges in, 221 structure, 217-222 thermophilic, 220-222 enzymology in, 171 eubacterial features, 171 eukaryotic characteristics, 171 evolution, 167, 172, 205 halophilic, 167, 169, 170, 217 2-0x0 acid oxidoreductases in, 202203,204-205,209 alkaliphilic, 206, 2 13 amino acid and protein utilization, 176, 183, 186 carbohydrate-metabolizing strains, 176, 186 central metabolic pathways summary, 192 citrate synthase in, 213-214, 215 citric acid cycle in, 186-1 87 class I aldolase in, 183, 184 classical, 206, 213 dihydrolipoamide dehydrogenase activity in, 206-207 ferredoxins [2Fe-2S] in, 205 gluconeogenesis in, 183 glycerol synthesis, 185 hexose catabolism in, 176-179 M6 strain, see Halobacterium saccharovorum succinate thiokinase in, 213, 215216 hexose catabolism, 176-182 histone-like proteins in, 171 lipids in, 170, 185 methanogenic, 167, 168, 169 central metabolic pathways summary, 192 citrate synthase in, 214, 215 citric acid cycle in, 189-190 dihydrolipoamide dehydrogenase in, 207 gluconeogenesis in, 182, 183, 191 glycerol synthesis in, 186 hexose catabolism, 182 malate dehydrogenase in, 198

reverse Embden-Meyerhof pathway, 182, 183, 184, 191 succinate thiokinase in, 215 methanogens as, Lake’s terminology, 170 mRNA in, eukaryotic features, 171 phenotypes, 167 phylogenetic tree, based on 16S/18S rRNA sequences, 167- 168 based on 16s rRNA sequences, 168, 169 based on hybridization homologies, 169 phylogeny, 167- 170 main divisions, 167, 169 protein synthesis initiation in, 171 ribosome and rRNA in, eubacterial features, 170 sulphur-dependent, 167, 170 thermoacidophilic, 217 central metabolic pathways summary, 192 citrate synthase in, 213-214, 215 citric acid cycle in, 187-189 dihydrolipoamide dehydrogenase in, 207 dual cofactor specificity of enzymes, 1 9 4198 ferredoxins [4Fe-4S] in, 205 gluconeogenesis in, 183 glucose dehydrogenase in, 177, 196- 197 glycerol synthesis, 185 heterotrophic growth on yeast, 183, 187 hexose catabolism in, 178, 179-181 isocitrate dehydrogenase in, 194196 malate dehydrogenase in, 198 phenotyw, 169 iange df(iptima1 temperatures), 221-222 similarity in enzymology of species, 216 succinate thiokinase in, 214, 215 transcription in, eukaryotic features, 171 tRNA in, 170, 171 Arsenate, piliated cell sensitivity, 73

SUBJECT INDEX

Ascorbate, 30, 32 Aspartate, codons, 218 in halophilic enzymes, 2 18 Aspergillus niger, 249, 272 Atebrin, 33, 34 Autolysins, lipoteichoic acid interaction, 283-290 chain substitution effect, 286-287, 290 inhibition by Forssman antigen, 283284,285 inhibition due to, 283-286 micelles and liposomes role, 289 phospholipid inhibition, 288-289 Autotrophs, 2, 115, 132, see also Ribulose 1,5-bisphosphate carboxylase/oxygenase carboxysomes in, 115, 155 as ecological markers for, 116, 155-156 evolution, 141-I42 methanogenic archaebacteria, 182 RuBisCO enzyme function, 116, 136, 155 Azotobacter, hydrogenase activity in, 2, 4 Azotobacter chroococcum, electron acceptor reactivity of hydrogenase, 16 hydrogen oxidation, reducing equivalent donation, 24 nickel in, 20

B Bacillus cereus, dual-specificity glucose dehydrogenase, 197 Bacillus coagulans, lipoteichoic acid, diacylglycerol as lipid anchor, 238 effect of temperature on alanine content, 271 Bacillus licheniformis, alanylated short-chain homologues of lipoteichoic acid, 263 glycerophosphoglycolipids, glycolipids and lipoteichoic acids in, 235, 236 lipoteichoic acid, diacylglycerol as lipid anchor, 238

33 1

lipoteichoic acid synthesis, phosphate limitation, 268 magnesium-dependent enzymes, 293 Bacillus megaterium, lipoteichoic acid, diacylglycerol as lipid anchor, 238 estimates of content, 247 lipoteichoic acid synthesis, effect of growth state, 267 sporulation effect, 270 phosphatidylethanolamine synthesis, site of, 276 phosphatidylglycerol pools, 26 1 Bacillus pumilis, glycerophosphodiesterase from, 272 Bacillus stearothermophilus, metabolic fate of lipoteichoic acids, 272 salt bridges in thermophilic enzyme, 22 1 Bacillus subtilis, 8-N-acetylglucosaminidaseinhibition, 285,287-288 dual-specificity glucose dehydrogenase, 197 glycerophosphoglycolipids, glycolipids and lipoteichoic acids in, 235,236 lipoteichoic acid synthesis, phosphate limitation, 268, 269 in phosphatidylglycerol biosynthesis inhibition, 248 metabolic fate of lipoteichoic acids, 272 poly(g1ycerophosphate) lipoteichoic acids in, chain composition, 242 teichoic acids, as ligands for autolysins, 285 teichoic acid-synthesizing enzymes, 277 teichoicases from, 272 Bacillus subtilis Marburg, lipoteichoic acids in, 234 Bacillus subtilis subsp. niger, 271 Bacillus subtilis W23, biosynthesis of teichoic acid, location, 276 Bacteriophage, fl, 87 attachment to F pili, 90 fd, 65

332

SUBJECT INDEX

Bacteriophage (contd.) filmentous DNA, attachment to F-like pili, 89 evidence for pilus retraction, 93 F pili interaction, 87 interactions with F-like pili, 86-87 lambda, 41 Pf, 65 Ps. aeruginosa pili, attachment, 96 QB, 86 attachment to F-like pili, 89, 91 R 17, 54, 56 attachment to F-like pili, proteins in, 89, 91 interaction with F-like pili, 86-87 sensitivity of conjugative pili, 58, 6061 Bacteroides nodosus, genetic organization of pili, 8 1, 82 NMePhe pili in, 63, 64,81 non-conjugative pili, 57, 63 Bacteroids, 6, see also Rhizobium japonicum, bacteroids Bayer’s junctions, 69 Benzyl viologen, 16, 17 Bicarbonate, RuBisCO stimulation, 142 B$dobacterium bijidum, glycerophosphate-containing lipoglycan, fatty-acyl composition, 239 structure, 243-245 surface location, 274 synthesis, 258 lipoteichoic acid, 243 Binary association coefficient, in oligonucleotide catalogue, 166 Bisphosphatidylglycerol, 250 Bisphosphoglycerate mutase, 174 “Black smokers”, 222 BIasia pusilla (liverwort), 122 Bligh-Dyer phase partition, 252 Blood group, P system, 55, 61,94 Blue-green prokaryotes, 123 Bordetella, non-conjugative pili, 57 Bordetella pertussis, pili, structure, 68 Bradley system, pili classification, 57 C

Calvin cycle, 116, 155

absence, in Sulfolobus, 187 carboxysomes and, 1 15, 1 16, 155 enzymes, absent from carboxysomes except RuBisCO, 152 in chemolitho-autotrophic bacteria from dark environments, 156 intermediates, in RuBisCO regulation, 142 Carbamate, in RuBisCO activation, 136, 137 Carbohydrate, effect on lipoteichoic acid content and synthesis, 267-268 metabolism, by halophiles, 176, 186 synthesis, by methanogenic archaebacteria, 182, 184, 191 Carbon, in derepression of hydrogenase activity, 6, 9 oxygen hypersensitivity mutants, 7 oxygen-insensitivemutants, 7 hydrogenase repression insensitivity by Alcaligenes eutrophus Osemutants, 8 limited, carboxysome numbers, 152, 154 organic, production by Calvin cycle prokaryotes, 116 in regulation of cytochrome pattern of Hupc mutant, 31 in regulation of hydrogen metabolism of Rhizobium, 6-9 Carbon dioxide, assimilating enzyme, see RuBisCO concentrating mechanisms, 142 carboxysomes as, 152 enhancement of hydrogenase activity, 6,9 limitation, RuBisCO activity increase, 150-1 5 1 microbes utilizing, carboxysomes in, 115 in RuBisCO activation, 135, 136, 145 S subunit role, 138 Carbon dioxide fixation, 9-10, see also Ribulose, 1,s-bisphosphate carboxylase carboxysomes as sites in uiuo, 150152

SUBJECT INDEX

Carbon dioxide fixation (contd.) in hydrogen-derepressed cells, 9 hydrogen as reductant for, 2 by methanogenic archaebacteria, 182, 184, 191 mutants (Cfx-), Hup+ and Hup- (R. japonicum), 9-10 oxidoreductase reaction, 202 reductive citric acid cycle in thermophiles, 187, 188, 189 RuBP-dependent, 124, 140 inhibition, 140 Carbon dioxide/oxygen specificity of RuBisCO, 140-142 Carbonic anhydrase, 152 inhibitors, 128 as possible carboxysomal protein, 127-128 Carbon monoxide, competitive inhibitor of hydrogen in hydrogen evolution, 23 spectra, 6-type cytochrome in hydrogen oxidation, 29-30, 32 2-Carboxyarabinitol 1,5-bisphosphate (CABP), 139, 150 2'-Carboxyarabinitol- 1-phosphate (2CAIP), 144 2'-Carboxy-3-keto-~-arabinitol 1,5bisphosphate (CKABP), 137 Carboxylation reaction by RuBisCO, see RuBisCO Carboxysomes, 115-164, see also Cyanobacteria; individual organisms abundance and photosynthetic characteristics, 15I, 152 assembly, DNA role in?, 130 Calvin cycle enzymes absent except RuBisCO, 152 as carbon dioxide concentrating mechanism, 152 in chemolitho-autotrophic prokaryotes, 117-121, 153 in colourless sulphur-oxidizing bacteria, 119-120, 153 composition, 124-132 DNA in, 128-130 proteins in, 124-128 in cyanelles, 123, 153 in cyanobacteria, 121-122

333

distribution and structure, 117-123, 153 ecological marker for autotrophy, 116, 155-156 function, 116, 149-155 as carbon dioxide concentrating mechanism, 127 different in different autotrophs, 153 protection of RuBisCO from inhibition, 152, 153-154 as site of carbon dioxide fixation in vivo, 150-152 as storage bodies, 154-1 55 in hydrogen-oxidizing bacteria, 120121,153 immuno-electronmicroscopy, 130- 132 isolation and studies in vitro, 124130. man-made containing RuBisCO, 156157 membrane, permeability, 150, 152 in nitrite- and ammonia-oxidizing bacteria, 1 17-1 18, 153 number/cell, in carbon limitation, 152, 154 in Oscillatoria (Trichodesmium) erythraea, 156 in photo-autotropic prokaryotes, 121-123 in Prochlorophyta, 122 RuBisCO in, see also Ribulose 1,5bisphosphate carboxylase designation dependent upon, 1 17 evidence for, 124- 125 stability in vitro, 124 Cardiolipin, 22 content in Staph. aureus, in energy deprivation, 270 Cations, lipoteichoic acid interaction, 29 1-294 CDP-glycerol, 233-234 no rule in lipoteichoic acid metabolism, 247 structure, 235 CDP-ribitol, 234 Cell-enveloDe. in Archaebactetia. 170 Cell-wall I y k ' enzymes, interaction with lipoteichoic acid, see Autolysins

334

SUBJECT INDEX

Central metabolism, Archaebacteria, see Archaebacteria in eubacteria and eukaryotes, see Eubacteria; Eukaryotes CFA/I, CFA/II, see Pili Cfx- mutants, 9-10 Chemoautotrophs, R . japonicum oxygen-insensitive mutants, 7 Chemoheterotrophic growth, RuBisCO production in, 154 Chemolitho-autotrophic prokaryotes, 116 aerobic, RuBisCO substrate specificity, 142 carboxysomes in, distribution and structure, 117-121, 153 in dark deep-sea environments, 155 Chemolithotrophic growth, cytochrome spectra, 36 nickel in, 20 oxygen-dependent hydrogen oxidation rate, 36 Chlamydomonas reinhardtii, RuBisCO activase polypeptides, 145 Chlorella, inhibition of carbon dioxide assimilation, 140 Chlorobium thiosulfatophilum, 189 Chloroform, permeabilization of carboxysome membrane, 150 Chlorogloeopsisfritschii, carbonic anhydrase in, external enzyme, 127-128, 152 carboxysomes in, phosphoribulokinase activity, 127 polypeptide composition, 126 role in carbon dioxide fixation, 151-152 stability in vitro, 124 extrachromosomal DNA absent, 129, 147 immuno-electronmicroscopic localization of RuBisCO, 130, 13 1 phosphoribulokinase in, localization, 127, 131 RuBisCO number of genes, 147 pool localization, 13 1 Chlorophyll a and b, 122 Chloroplast, C . paradoxa cyanelles genome

similarity, 123 eubacterial origin, 167 protein, RuBisCO activity increase, 144-145 RuBisCO L and S subunit genes in, 145, 146 Choline in teichoic acid, in action of Nacetylmuramyl-L-alanine, amidase, 284 Chromatium vinosum, 138, 139 nickel in hydrogenase, 20 RuBisCO 8L molecules, catalytically competent, 139 gene cloning, 146 S subunit function, 138, 139-140 Chromosomally encoded control elements, pilus expression, 71-72 Chromosome, RuBisCO L and S subunit genes in, 145, 146 Citrate synthase, 209-217 in archaebacteria, 21 3-2 15 ATP, NADH and 2-oxoglutarate effect, 214 ATP inhibition in Gram-positive bacteria, 2 10-2 1 I control, in eubacteria and eukaryotes, 2 11212 in methanogenic archaebacteria, 214 in eubacteria and eukaryotes, 210213 properties (summary), 212 in halophilic archaebacteria, 186, 213-214,215 large and small, 21 1 in methanogenic archaebacteria, 214, 215 NADH inhibition in Gram-negative bacteria, 210 reaction catalysed by, 210 in S . acidocaldarius, 189 sensitivity to ATP and acetyl-CoA affinity, 210-21 1 structure, sequencing, 21 5-216 in thermophilic archaebacteria, 187, 214,215 Citric acid cycle, in archaebacteria, 186-190

SUBJECT INDEX

Citric acid cycle (contd.) enzyme diversity in, 175, 209 in eubacteria and eukaryotes, 175, 176 evolution, 193 in halophilic archaebacteria, 192 incomplete, in methanogenic archaebacteria, 189-190 in Thermoproteus neutrophilus, 188, 189 in methanogenic archaebacteria, 189190, 192 oxidative, in halophilic archaebacteria, 186187, 191 incomplete, 190 in Sulfolobus, little evidence for, 189 in thermophilic archaebacteria, 187, 191 reductive, incomplete in M. thermoautotrophicum, 189, 191 origin, 193 in Sulfolobus spp., 187, 189, 191 in Thermoproteus neutrophilus, 188, 189 in thermophilic archaebacteria, 187, 191, 192 Clams, bacterial, symbionts, carboxysomes in, 156 Cloning vector, 41 Clostridium, modified EntnerDoudoroff pathway in, 179 Clostridium pasteurianum, hydrogenase, nickel absence, 21 hydrogenase I, oxygen sensitivity, 18 hydrogenase 11, electron acceptor reactivity, 17 K, value, 16 Colicin B2 (ColB2) pilin, 83, 84, 85 Colicin El (ColEI) replicon, 41 Coli surface antigens (SCI, CS2, CS3), see Pili, CFA/II “Colonization factor antigen” (CFA/I, CFA/II), 54, 62, see also Pili Component 559-H2,35 evidence against, 35-38

335

low redox potential, evidence against, 37 quantification in presence of cyanide, 36 Conjugation, 57, 68, 87, 89 deficient cells (Con-), 87 Cosmid, complementing Hup- mutants, 4 3 4 pHUI, 43,44 pHU52,44 pLAFRI, 41,43,44 pSH22,44 Cowpea (V.unguiculata), hydrogenase activity control, 10 cp x AB gene product, 71 Critical micellar concentration of lipoteichoic acids, 274 Cyanelles, see also Cyanophora paradoxa; Glaucocystis polyhedral bodies in, 123, 153 RuBisCO gene location and number, 146, 147 gold immunoelectronmicrosopy, 132 Cyanide, F-pili disappearance, 90, 93 inhibition of cytochrome c oxidase, 34 in quantification of component 559-H2,36 in R. japonicwn bacteroid hydrogen oxidation, 34 spectra, b-type cytochrome in hydrogen oxidation, 29-30, 32 Cyanidium caldarium, 146 Cyanobacteria, see also Carboxysomes; individual species aerobic nitrogen-fixing filamentous, RuBisCO absent from heterocysts, 122, 131 carboxysomes in, 121-122 chromosomal DNA association, 129 function, 153 phosphoribulokinase in, 127 role in carbon dioxide fixation, 151 endosymbiotic, cyanelles derived from, 123 extrachromosomal DNA in, 129, 147

336

SUBJECT INDEX

Cyanobacteria (contd.) filamentous, carboxysome size and shape in, 121-122 hydrogenase activity in, 2 2-0x0 acid dehydrogenase and 2-0x0 acid oxidoreductase in, 204 protein reserves, carboxysome storage function, 155 R uBisC0 gene cloning and location, 146 localization of, 131 regulation, by effectors, 143 Cyanophora paradoxa, DNA attachment to, 129 RuBisCO in, gold immunoelectronmicroscopy, 132 L and S subunit genes, 146 polyhedral bodies and, 123 subunit gene, number of, 147 Cyanophycin, 155 Cycas revoluta, 122 Cyclic AMP, cya (synthesis) and crp (receptor protein) mutants, 72 hydrogenase expression in R. japonicum, 7 pilus expression, 72-73 Cyclic AMP-CRP complex, regulation of pap B transcription, 77 transcriptional control in pilus expression 73 Cyclic AMP receptor protein (CRP), 72,73 binding site, 73 Cyclopropanization, 240 2,3-Cyclopyrophosphoglycerate,184 Cytochrome, component 559-H2, see Component 559-H2 Cytochrome aa3, in hydrogen oxidation in R. japonicum, 28, 29, 30 in P. denitrificans, 28, 29 proposed electron-transport pathway in R. japonicum (free-living), 32 repression, at low oxygen tensions, 28,30 as terminal oxidase, 28, 30 Cytochrome b-type, 14, 30-3 1, see also

Cytochrome o in electron-transport pathway in R. japonicum (free-living), 32 in hydrogen oxidation R. japonicum, 29-30 in R. japonicum bacteroids, 32, 33,34 in Thermoplasma acidophilum, 181 unique in hydrogen oxidation?, 3538, see also Component 559-H2 Cytochrome c, CN- and Co- reactive, 34 electron acceptor reactivities, 16, 17, 28 reduction by R. japonicum (freeliving), 32 in R. japonicum bacteroids, 32, 33 Cytochrome c3, electron acceptor in Desulfovibrio, 17 Cytochrome c-552, 34 Cytochrome c oxidase, inhibition by atebrin and cyanide, 33, 34 Cytochrome d, 27 Cytochrome 0,7-8, 27, 29, 30 arguments that cytochrome 559-H2 are like, 35-36 in election-transport of free-lining R. japonicum, 32 in electron-transport of R. japonicum bacteroid, 34 in hydrogenase-constitutive strains, 37 in low oxygen concentrations and oxygen affinity, 30 P. denitrificans expression, 37 reducibile by succinate and NADH, 37 reduction rate, 35-36 Cytochrome, P-450, 35 Cytochrome spectra, 33, 36, 38-39

D Dark, chemolitho-autotrophic bacteria in, 155 Deazaflavin derivative, 202, 204 Deep-sea, microbial growth, chemolitho-autotrophic bacteria, 155

337

SUBJECT INDEX

Dehydrogenases with dual cofactor specificity, 194- 199 glucose dehydrogenase, 177, 196-197 isocitrate dehydrogenase, 194-196 malate dehydrogenase, 197-198 Desulfovibrio desulfuricans, nickel in hydrogenase, 20 Desulfovibrio gigas hydrogenase, nickel in, 20 nickel and iron-sulphur clusters, 21 Desulfovibrio vulgaris hydrogenase, aerobically stable, 18-19 composition and antibody crossreactions, 14 nickel absent, 21 Desulfurococcus mobilis, introns in tRNA genes, 171 2-0x0acid oxidoreductases, 202 Desulfurococcus mucosa, dihydrolipoamide dehydrogenase in, 207 Deuterium, hydrogenase oxidation, 23 Deuterium-water exchange reaction, 24 Diacylglycerol, conversion into glycolipid, 250, 259, 276 as lipid anchor in Bacillus spp. lipoteichoic acid, 238 in M . luteus, lipomannan, 245 recycling in Staph. aureus, 248, 259260 enzyme in, 260-261 recycling to phosphatidylglycerol, 258-259,276 synthesis, 250 location of, 276 in Staph. aureus, 259-260 Diacylglycerol kinase, 260 Diacylglycerol phosphatidylglycerol, as acceptor substrate in lipoteichoic acid synthesis, 250, 254 Diarrhoea, CFA/I, CFA/II pili in E. coli, 62 K99 in E. coli, 63 Dichlorophenol-indophenol (DCPIP), 16, 17 Digalactose receptor, 77 Digalactosyl residues in lipoteichoic acids, 243

Diglycerophosphoglycolopid, galactosylated, 261 Dihydrolipoamide, 200, 206 Dihydrolipoamide dehydrogenase, 200, 204,206-209 in halophiles, 206-207 lack of homology with E. coli enzyme, 207 membrane association, 208, 209 metabolic function, 208-209 in other archaebacteria, 207-208 purification and structure, 206 reaction catalysed by, 200, 201, 206 Dihydrolipoyl acyltransferase, 200 Dihydroxyacetone phosphate, 172, 185 Dimannosyldiacylglycerol, 258 Diplosoma virens, 122 Dithionite, 18-19, 29, 33 DNA, attachment to exterior of carboxysomes, 129-1 30 in carboxysomes, 128- 130 extrachromosomal, see also Plasmid absent from Chlorogloeopsis fritschi, 129, 147 in autotrophic prokaryotes, 129 in cyanobacteria, 129, 147 RuBisCO genes on, evidence, 148 metabolism in conjugation, genes involved, 69-70 recombinant technology, microbial RuBisCO production, 149, 156-157 transfer, cloning vectors for Rhizobium, 4147 plasmid-specific genes (orlT) in conjugative pili, 68 DNA-dependent RNA polymerase, 169 properties, 13-15

E Ecophysiological marker, carboxysomes as, 116, 155-156 EDTA, inhibition of hydrogenase derepression, nickel role, 20 Electron, acceptors, in 2-0x0 acid dehydrogenases in eubacteria, 200

338

SUBJECT INDEX

Electron (conr0.l in Zoxo acid oxidoreductase reactions in archaebacteria, 202, 205 allocation by nitrogenase, host control of, 11 carrier, in model for hydrogenase mechanism, 23, 24 flux in nitrogen fixation, reaction, 3 host control, 11 transport, in free-living R. japonicum, 28-32 in hydrogen oxidation, 27-38 in R. japonicum bacteroids, 32-35 unique cytochrome b in, 35-38 Elongation factor (EF-2), in Archaebacteria, 171 Embden-Meyerhof pathway, 172, 173 absent from halophilic archaebacteria, 177, 179 ATP yields, 172, 191 enzymes, in eubacteria and eukaryotes, 1 7 4 175 not detected in H. halobium, 179, 183 not detected in T. acidophilum, 181 evolutionary origin, random association of enzymes, 174 simple energy-conversion process, 191 in methanogenic archaebacteria, 182 reversal, in methanogenic archaebacteria, 182, 183, 184, 191 in thermophilic archaebacteria (possible presence), 181 Endometrial carcinoma, 100, 101 Endosymbionts, 123 Energy, deprivation and lipoteichoic acid synthesis, 269-270 Entamoeba histolytica, 180 Enterococcus faecalis, lipoteichoic acid, autolysin inhibition, 286 biosynthetic sequence, 250, 251 chain composition, 242 chain elongation, 249 chain structure, 240, 241 estimates of content, 247

extrace>iular, heacylateh, 273 fatty-acyl composition, 239 glycosylation of, 243, 261 metabolic fate, 272 metabolism, 247 role in autolysin regulation in vivo, 290 substitution, protein synthesis effect, 271 synthesis, effect of growth stage, 267 synthesis, membrane lipid metabolism, 259-260 phosphatidyldiglucosyldiacylglycerol incorporation, 250, 252 phospholipid inhibition of autolysins, 288-289 Enterococcus faecalis ATCC 9790, 290 Enterococcus faecalis NCIB 8 191, NCIB, 39 243,242 Enterotoxin, heat-stable (ST enterotoxin), 77, 78 Entner-Doudoroff pathway, 175 ATP yields, 172, 191 classical, 173, 178 in eubacteria and eukaryotes, 172, 173 evolutionary origins, 191 modified in halophiles, 176-1 79, 177, 178, 191 enzymes in, 179 modified in non-archaebacterial species, 179 non-phosphorylated pathway, reversal in thermoacidophiles, 183 in thermoacidophiles, 177, 178, 179-181, 191 Enzymes, see also Archaebacteria: individual enzymes archaebacteria, 171, 194-2 17 diversity, 216 in glucose catabolism in eubacteria and eukaryotes, 172-174 halophilic, 2 17-220 thermophilic, 220-222 Eocytes, 170 Escherichia coli, adhesive pili, see Pili, adhesive citrate synthase sequence, 216 class I aldolase in, 184

339

SUBJECT INDEX

Escherichia coli (contd.) conjugative pili, see Pili, conjugative cya crp mutants, 72 enterotoxigenic, CFA/I, CFA/II, 56, 62 K99 positive, 63 eukaryotic polypeptides expressed, in inclusion bodies, I56 incompatibility groups of plasmids, 60 in uiuo mutagenesis of citrate synthase, 21 1 nickel in hydrogenase, 20 NMePhe pilin gene expression in, 82 2-0x0 acid dehydrogenase and 2-0x0 acid oxidoreductase in, 204 pili, X-ray diffraction studies, 65 plasmids, 41 pRK290 infecting, 41 pyruvate dhydrogenase complex, 203 RuBisCO expression in, 149, 156 sphaeroplasts, pili assembly, 92 Type I pili genes (JimA-0,pilA-E), 74 uropathogenic, 55, 61, 75, 94 Escherichia coli 987P, 57, 62, 63 Escherichia coli CA8000, 73 Escherichia coli H 10407, adhesive pili, 56 Escherichia coli HBl1, conjugative pili, 56 Ethanolamine, choline in teichoic acid replaced by, autolysin inhibition and, 284 Ethyl methane sulphonate (EMS), 39 Eubacteria, 167, 168 aerobic, 2-0x0 acid dehydrogenase in, 200 6-phosphofructokinase absence, 172 pyruvate metabolism, 175 anaerobic, see Anaerobes central metabolism in, 172-176 enzymes in glucose catabolism, 174- 175 metabolic fate of pyruvate, 175176 sugar catabolism, 172-174 citrate synthase, 210-212 ferredoxin in, 205

isocitrate dehydrogenase, 195 malate dehydrogenase (NAD+specific) in, 197 2-0x0 acid dehydrogenase in, 200202,203 rRNA features in archaebacteria, 170 succinate thiokinase in, 212-21 3 Euglena gracilis, 140 Eukaryotes, 166, 167, 168 aerobic, puruvate metabolism, 175 central metabolism in, 172-176 enzymes in glucose catabolism, 174-175 metabolic fate of pyruvate, 175176 sugar catabolism, 172-174 citrate synthase in, 210-21 1 ferredoxin in, 205 isocitrate dehydrogenase, 194-195 2-0x0 acid dehydrogenase in, 200202,203 RuBisCO L and S subunit genes, 145 succinate thiokinase in, 212-213 Evolution, 2-0x0 acid: ferredoxin oxidoreductase, considerations, 204-205 archaebacteria, see Archaebacteria central metabolic pathways, archaebacterial considerations, 191-193 ferredoxins, 205 significance of dual specificity of isocitrate dehydrogenase, 196 Extended X-ray Absorption Fine Structure (EXAFS), 21 F FAD, dihydrolipoamide dehydrogenase containing, 200, 203 FADH2, formation in citric acid cycle, 175 Fatty-acyl residues in lipoteichoic acids, 237.238 cyclopropanization, 240 in diacylglycerol recycling, 260 percentage composition of, 239 pneumococcal, 246

340

SUBJECT INDEX

Feltham First, 11 Ferredoxin, 202, 204, see also 2-0x0 acid : ferredoxin oxidoreductase amino-acid sequences, 205,220, 221 “bacterial-type” [4Fe-4S], 205 “chloroplast-type” [2 Fe-2S1, 205 electron acceptors of 2-0x0 acid oxidoreductases, 202, 205 in eubacteria and eukaryotes, 205 evolutionary considerations, 205 Thermoplasma acidophilum, 22 1 Ferredoxin/thioredoxin system, 145 Ferricyanide, 19 reduction, 16, 17 Fertility inhibition (fin+),70, 71, 72 fimA, C, D genes, 74 fimB gene, in phase variation, 75 Fimbriae, 54, see also Pili antigen serology in nomenclature, 55 P, 55 Fimbrillin, 77 j m E gene, 75 f i n 0 gene product, 71 finOP genes, 69, 70-71 FinOP repressor system, molecular basis, 70-71, 72 finP gene product, 71 Flavin mononucleotide (FMN), 16, 17 Flavoprotein, hydrogen uptake and, 28, 33-34 F-like plasmids, see Pili; Plasmid Forssman antigen, pneumococcal, 246, 275 autolysin inhibition, 283, 284, 285 structure, 246 F pili, see Pili, F F pilin, see Pilin Fructose, effect on lipoteichoic acid content of cells, 268, 269 Fructose 1,6-bisphosphatase (FBPase), 145, 181, 183 Fructose 1,6-bisphosphate, 143 in methanogenic archaebacteria, 184 Fructose 1,6-bisphosphate aldolase, 181, 183 class I and 11, 183-184 in M . thermoautotrophicum, 182 F-transfer operon, 70-71 genes in, 69, see also individual tra genes

Fumarate reductase, in methanogens, 189

G Galactosyl substitution of lipoteichoic acids, 243 effect on lipoteichoic acid carrier activity, 280, 282 Gal-gal pili, 55, 61, 94 GC pili, see Neisseria gonococcus; Pili GDP-a-D-mannose, 258 Gene duplication, glycolytic enzymes, 174 Gene rearrangements, gonococcal pilin genes, 80, 100-102 Gentiobiosyldiacylglyercol, 28 1 Glaucocystis, RuBisCO in, gold immunoelectronmicroscopy, 132 Glaucocystis nostochinearum, 123 Glaucosphaera oacuofuta, RuBisCO in, but no polyhedral bodies, 123 Globoseries glycolipids, 61, 94 Gloeobacter violaceus, no extrachromosomal DNA in, 129 Gloeochaete wittrockiana, 123 Gluconate, derepression of hydrogenase activity, effect on, 6 glucose oxidation to, 177 Gluconeogenesis in archaebacteria, 183-185, 191 halophilic, 192 methanogenic, 192 thermophilic, 192 Glucose, in F pili, 83, 85, 87 in bacteriophage attachment, 91 limitation, lipoteichoic acid alanine content, effect on, 271 in lipoteichic acid, see Glycosyl residues lipoteichoic acid content of cells, effect on, 268, 269 pilus production (cyclic AMP suppressed), 73 Type I and K99 pili expression reduced, 77

SUBJECT INDEX

Glucose catabolism, see also EmbdenMeyerhof; Entner-Doudoroff pathways in archaebacteria, 176182 enzymes, see individual enzymes and pathways in eubacteria and eukaryotes, 172174 in halophilic archaebacteria, 176-1 79 summary, 192 in methanogenic archaebacteria, 182 summary, 192 non-phosphorylated pathway, 177 in thermoacidophilic archaebacteria, 177, 178, 179-181 summary, 192 Glucose dehydrogenase, dual specificity, 177, 196-197 NAD+-dependent, 177, 196, 197 in Sulfolobus solfataricus, 177, 196197 in thermoacidophilic archaebacteria, 177, 196-197 in Thermoplasma acidophylum, 197 Glucose 6-phosphate, oxidation, absent from H. saccharovorum, 177 Glutamate, codons, 2 18 in halophilic enzymes, 2 18 Glyceraldehyde, fate in S. solfataricus, 179, 186 fate in T. acidophilum, 180, 186 formation from 2-keto-3-deoxygluconate in S . solfataricus, 179, 191 reduction to glycerol by glycerol : NADP+ oxidoreductase, 186 Glyceraldehyde 3-phosphate, see also Entner-Doudoroff pathway enzymes metabolizing, in H . saccharovorum, 177 metabolism to pyruvate in eubacteriajeukaryotes, 172, 173, 174 Glyceraldehyde 3-phosphate dehydrogenase, amino-acid sequences from thermophilic organisms, 221 not detected in halophiles, 179, 183

34 1

not detected in T. acidophilum, 181 Glycerol, hydrogen inhibition of heterotrophic growth of Osemutants, 8 synthesis, in archaebacteria, 185-186 Glycerol dehydrogenase, in H . cutirubrum, 195 Glycerolipids, archaebacterial, 185 Glycerol: NADP+ oxidoreductase, 186 Glycerol phosphate, in teichoic acids, 234 Glycerol phosphate dehydrogenase, 185 sn-Glycero-1-phosphate, carrier, 250, 276 linkage to glycolipid in lipoteichoic acid synthesis, 253 linkage of units in lipoteichoic acid synthesis, 253 in lipoglycan, synthesis from phosphatidylglycerol, 258 in lipoteichoic acid, see Lipoteichoic acid structure, configuration, 235, 240,243 Glycerophosphate-containing lipoglycan, see Lipoglycan Glycerophosphate polymerase, 277 Glycerophosphodiesterase,lipoteichoic acid degradation, 272 Glycerophosphoglycolipid, glycolipids and lipoteichoic acid relationship, 235, 236 in Lactococcus garvieae, 254, 257 in Staph. aureus, lipoteichoic acid synthesis, 252-253, 254 structural studies, 240 Glycolipid, as acceptor substrate, in lipoteichoic acid synthesis, 250 diacylglycerol conversion into, 250, 259,276 glycerophosphoglycolipid and lipoteichoic acid relationship, 235, 236, 251 in lipoteichoic acids, structures and occurrence, 236,237 membrane, lipoteichoic acids attached to, 234, 235 synthesis, 259

342

SUBJECT INDEX

Glycolipid (contd.) in Gram-positive bacteria, 250 Glycolysis, 172, 173, see also EmbdenMeyerhof pathway Glycosylation of lipoteichoic acids, see Glycosyl residues; Lipoteichoic acid Glycosyl residues in lipoteichoic acid, 240,241,261-262,276 anti-autolytic activity, effect, 287 effect on anti-autolytic activity, 282, 283 effect on carrier activity, 282, 283 incorporation of, 240, 261-262, 276 model of structure, 293 Gold immunoelectronmicroscopy, 130, 131 Gram-negative bacteria, citrate synthase (large), NADPH inhibition of, 210, 211 conjugative pili, 60 succinate thiokinase in, 21 3 Gram-positive bacteria, citrate synthase, 210-21 1 conjugative pili not identified in, 57, 60 glycerophosphoglycolipids in, 235 lipoteichoic acid, 234 structure, 235 lipoteichoic acid, lipid and protein secretion, 274 membrane lipid turnover, 258 succinate thiokinase in, 213 teichoic acid in, 234 Growth stage and rate, effect on lipoteichoic acid synthesis, 267 extracellular lipoteichoic acid, 272 lipid amphiphile composition, 258, 259 Growth yield studies, 25-26

H Haemagglutination, pili capable of, 74, 75,76 Haemoglobin, amino-acid sequences from thermophilic organisms, 221 Halobacterium, citrate synthase and succinate thiokinase in, 213

Halobacterium cutirubrum, glycerol synthesis, 185 Halobacterium halobium, ferredoxin, amino-acid sequences, 220 glycolytic enzymes in, 183 malate dehydrogenase from, 198 oxidative citric acid cycle in, 186-187 2-0x0 acid oxidoreductases, 202 6-phosphofructokinase absence, 179 Halobacterium marismortui, ferredoxins, sequences of, 220 malate dehydrogenase, structure and characteristics, 219 Halobacterium saccharovorum, acetate production, pyruvate: ferredoxin oxidoreductase in, 177 ATP not required in pyruvate production, 177 dual specificity glucose dehydrogenase, 197 glucose catabolism in, 177 glyceraldehydxe 3-phosphate metabolic enzymes in, 177 modified Entner-Doudoroff pathway, 176-179, 191 NAD+ reduction, 177 Halobacterium volcanii, introns in tRNA genes, 171 Halophiles, see Archaebacteria Hatch-Slack (C4-dicarboxylicacid pathway), 141, 142 Heme biosynthesis, 40 SR143 mutant deficient in, 40 Heterocysts, 122 absence of RuBisCO and carboxysomes from, 122, 13 1 Hexose catabolism, see also Glucose in Archaebacteria, 176-182 Hexose-monophosphate pathway, 175 in eubacteria, 172, 173, 174 Hexosyl-1 -phosphorylundecaprenol, 261-262 Hfr strains, E. coli, 72, 73 High Frequency of Transfer (HFT), 70 Hind111 restriction endonuclease, 81 Histone-like proteins, in archaebacteria, 171 Host control, of hydrogenase, 10- 13

SUBJECT INDEX

Host control (contd.) of piliation, 72 HQNO, 31-32 HTa protein, Thermoplasma acidophilum, I7 1 Hup gene, see Hydrogenase; Rhizobium species HV sequences in gonococcal pilin genes, 79, 80 Hybridization, 44,47 homologies of archaebacteria, 169 RuBisCO subunit probe, 147 Hydrogen, absence, derepression of hydrogenase in oxygeninsensitive, mutants, 7 cytochrome o reduced, 30, 37 cytochromes b and c reduced, 28, 3233, 36 in bacteroid membranes, 32-33, 37-38 free-living R. japonicum, 28, 29 P . denitrificans, 28, 31, 37 succinate comparison, 36, 37-38 effect, 8, 9 evolution, 2 host control of, 11-13 in leguminous and non-leguminous nodules, 4 low by R . japonicum strains, 4 in nitrogen fixation reaction, 2-3 rate dependence on electron flux, 3 relative efficiency of, 5 increase in derepression of hydrogenase activity, 6 inhibition of, A . eutrophus heterotrophic growth, 8 hydrogen evolution, 4, 23 nitrogenase, 4 metabolism in Rhizobium, 1-52 regulation, 6-13 nitrogenase inhibition, 4, 25 oxidation, 2 by Alcaligenes eutrophus in mixotrophic context, 8 ATP production, comparison with oxygen, 25 ATP synthesis coupling, 24-25, 47

343

cytochrome 6-type reduction, succinate and NADH comparison, 37 cytochrome c- and b- reduction rate, 36 efficiency, 16, 17 electron transport, 27-38, see also Electron transport: individual cytochromes energy conservation, 24 genes on plasmids, 129 by legume root nodules, 4-5 mutants failing to, 39 mutants (Hox-), Alcaligenes eutrophus, 42 nitrogen fixation not increased in R. leguminosarum, 5,46 oxygen-dependent, rate, 36 scavenger, 16 Hydrogenase, 2 aerobic purification, 18 amino acid composition, 14 antibody cross-reactivities, 14 beneficial effects (nitrogen fixation increase), 4, 5 , 9 derepression by low oxygen, 6, 7, 9 electron acceptor reactivity, 16-1 8 energetics, 24-38 electron transport, 27-38 physiological considerations, 24-27 enzymology, 13-24 genes, Rhizobium leguminosarum, 4547 genetics, 38-47 Hup+ strains, 2, 6 carbon dioxide fixation (Cfx-) mutants, 10 Hup genes on indigenous plasmids, 42-43 in R . japonicum, 43-45 in R. leguminosarum, 45-47 site-directed mutagenesis method, 41-42 Hup- mutants, carbon dioxide fixation (Cfx-) mutants, 10 component 559-H2 reduction in, invalidity of, 36-37 mixing of, reconstitution of activity, 39

344

SUBJECT INDEX

Hydrogenase (contd.) as mutants of Hup+, 43 selection method, 38-39 Hup probes, 47 hydrogen in derepression of, 6, 7 hydrogen oxidation, 2, see also Hydrogen efficiency, 16, 17 iron, content, 21 kinetic mechanism, 22-24 K,,, value, 15-16, 17 lipid requirement, 21-22 membrane-bound, absorption spectrum, 14 electron transport, 2, 27 Ose trait action, 8 midpoint potential, 16-17 molecular genetics, 40-47 mutants, 3 8 4 0 nickel in, 21 as structural component, 21 oxygen consumption, protective mechanism, 4, 25, 34, 35 oxygen-hypersensitive mutants, 6-7, 40 carbon repression hypersensitivity, 7 oxygen-insensitive mutants, 7,40 carbon repression insensitivity, 7 oxygen lability, 18-19 proteolysis, 14 purification and properties, 13-1 5 reducing agents in air, effect, 19 regulation, 6-13 by oxygen and carbon, 6-9 carbon dioxide fixation, 9-10 host control, 10-13 relative efficiency of nitrogen fixation, effect on, 5 in Rhizobium spp., 4 RuBP carboxylase correlation, 9-10, 25 soluble, carbon dioxide fixation, 2 subunit stoicheiometry, 13-14 symbiotic advantage, 4, 5, 9 Hydrogenase-constitutive mutants (HuPc), 7-8 bacteroids, hydrogenase and RuBP activity, 10 cytochrome b-type in, 35, 37

cytochrome o in, 7, 30, 35, 37 cytochrome o not component, 559-H2, 37 gene mutated in, 10 nature of mutations, 7-8 regulation by oxygen, 30-3 1 RuBP carboxylase expression, 10 Hydrogen-oxidizing bacteria, see also Hydrogen, oxidation; Hydrogenase; Rhizobium spp. aerobic, hydrogen metabolism, 2 carboxysomes absent from, 120-121, 153 Hydrophobic bonds, in halophilic enzymes, 2 18-2 19 Hydrophobicity, enzyme stability and, 22 1 hyp gene, 74, 75 I Immuno-electron microscopy, of carboxysomes, 130-132 location of lipoteichoic acids, 275 IncF plasmid, see Plasmid Inclusion bodies, see also Carboxysomes E. coli expressed eukayotic polypeptides in, 156 man-made RuBisCO in, 156-1 57 Incompatibility (Inc) in pili classification, 60, 68, see also Pili; Plasmid E. coli, 58-59 Intestinal epithelium, 987P positive E. coli, 63 K99 positive E. coli, 63 pili specific for, 61, 62, 95 Iodoacetate, 25 Ion-exchange fast protein liquid chromatography (FPLC), 133 Iron, in R. japonicum bacteroid hydrogenase, 14-1 $ 2 1 Iron-sulphur clusters, in hydrogenase, 21 Iron-sulphur proteins, 15, 18 IS3 element, 70 Isocitrate dehydrogenase, dual specificity, 195-196

345

SUBJECT INDEX

Isocitrate dehydrogenase (contd.) evolutionary significance, 196 NAD+-linked, 194 NADP -linked, 194-1 95 pig heart, 196 in S. acidocaldarius, 189,195-196, +

198 K , values, 195,196,198 in thermophilic archaebacteria, 187

K Kanamycin nucleotidyl transferase, 22I 2-Keto-3-deoxygluconate,177 cleavage to pyruvate in S. sovataricus, 179,191 K,,, values, hydrogenases, 15-16 bacteroid, 17

L Lactate dehydrogenase, hydrophobicity causing stability, 221 Lactobacillus casei, class I aldolase in, 184 glycolipids and glycerophosphoglycolipids, structure, 253,255 Lactobacillus casei lipoteichoic acid, alanylation of, 262 salt effect on, 270 chain elongation, 249 content growth stage effect on, 267 pH effect, 267 extracellular, negligible, 273 glycolipid and fatty acids in, 238 low alanine, autolysin activity, 290 metabolism, 247 mutant, lacking D-alanyl ester, 262 short-chain homologue, synthesis,

252 structure, 255 synthesis, 248 membrane lipid metabolism, 259-

260

phosphate limitation effect, 269 vesicles containing, 248,274 Lactobacillus fermentum lipoteichoic acid, content, growth stage effect on, 267 pH effect, 267 extracellular, 273 acylated, 273 poly(glycerophosphate), alanyl residues, 243 Lactococcus aarvieae. glycerophosphoglycolipids, structure,

256-257 lipoteichoic acid, biosynthesis, glycerophosphoglycolipids in,

254,256-257 digalactosyl residues in, 243 fatty-acyl composition, 239 structure, 244 synthesis, 254 Lactococcus lactis, galactosylated diglycerophosphoglycolipid in, 261 lipoteichoic acid, chain composition, 242 estimates of content, 247 fatty-acyl composition, 239 structure, 242 Lambda (A) bacteriophage, 41 Leghaemoglobin, 26-27,39 heme synthesis for, 40 Legume root nodules, hydrogen oxidation, 4-5 Light, RuBisCO activation, 144,145 Lipid, absence from carboxysomes, 126 anchor, lipoteichoic acids, 236-240,

243,249 in archaebacteria, 170, 185 membrane, 274 turnover and lipoteichoic acid synthesis, 258-261 requirement of hydrogenase, 21 -22 secretion by Gram-positive bacteria, penicillin effect, 274 Lipocarbohydrate, 246 Lipoglucogalactofuranan, 243 synthesis, 258

346

SUBJECT INDEX

Lipoglycan, glycerophosphatecontaining, 234, 243-245 fatty-acyl composition, 239 lipomannin relationship, 245 structure, 243-245 model of, 293 surface location, 274 synthesis, 258 Lipoic acid, in eubacteria and eukaryotes, 200 search for in archaebacteria, 209 Lipomannan, biosynthesis, 258 glycerophosphate-containing lipoglycan, relationship, 245 magnesium ion binding, 291 mesosomal vesicle association, 275 succinylated, 245-246 autolysin activity and, 285 Lipopolysaccharide (LPS), association with F-like pili receptor protein, 87,88 Liposomes, inhibition of autolysins, 289 Lipoteichoic acid, 233-302, see also individual bacterial species acylated, 238, 239 autolysin control, 289-290 extracellular, 273, 274 alanine/phosphate ratio, 290, 294 alanylation concomitant with poly(g1ycerophosphate)synthesis, 263 D-alanyl residues, see Alanyl residues in lipoteichoic acids autolysin inhibition, 283, 284, 285, 286 micellar organization role, 284, 290 biological activities, functions, 277293,294-295 biosynthesis, 234, see also Lipoteichoic acid, cellular content conditions affecting, 265-270 energy deprivation effect, 269-270 growth rate and stage, effect on, 267 location of enzymes, 275-276 membrane lipid metabolism and, proposed relationships, 260 membrane lipid turnover, 258-261 novel system for studying, 248

phosphate limitation effect, 268269,295 site, 276 sporulation effect, 270 carrier (LTC), 277-283 inhibitor, 277, 281, 283 as in vitro analogue of linkage unit, 280 in ribitol phosphate polymerization, 280 structural requirements, 277-278 cellular content, 247 effect of growth stage and rate, 267 osmotic shock effect, 295 pH and carbohydrate source effects on, 267-268 cellular location, 274-276, 29 1 cell-wall lytic enzymes interaction, 283-290 chain substitution, 240, 241 addition of residues, 261-263, 276 conditions affecting, 270-27 1 protein synthesis effect on, 271 choline residues, effect on autolysin inhibition, 284, 285 critical micellar concentration, 274 deacylated, 239 autolysin control, 289-290 extracellular, 273 inactive as carrier (LTC) but inhibitor, 277, 281, 283 loss of anti-autolytic activity, 289 definition, 234 degradation and excretion, 272-274 divalent cation interactions, 291-294 extracellular, 272, 273 acylated, secretion of, 274 acylated or deacylated forms, 273 deacylated, formation, 273 glycerophosphate-containing lipoglycan, see Lipoglycan glycerophosphoglycolipids and glycolipid relationship, 235, 236, 257 glycolipids in, 236, 237 glycosylation, 240, 261-262 glycosyl residues, see Glycosyl residues in lipoteichoic acid lipid anchor, 236-240, 243 chain synthesis, 249

347

SUBJECT INDEX

Lipoteichoic acid (contd.) magnesium ion binding, 291, 294 metabolic fate, 272-274 metabolism, 247-274 conditions affecting, 265-270 related macroamphiphile biosynthesis, 256-258 negative charge and autolysin inhibition, 289 occurrence, 235-247 phenol-water extraction, 239, 274 pneumococcal, see Forssman antigen poly(digalactosy1, galactosylglycerophosphate), structure, 243 poly(glycerophosphate), 234 acceptor substrates, 250, 276 alanyl residue distribution, 242, 243 anti-autolytic, structural requirements, 286-287, 290 antibodies to, 274 autolysin inhibition, 285 chain elongation, mode, 248-249 chain structure, 240-243 chain structure model, 292-293 diacylglycerol, see Diacylglycerol fatty acids in, 237, 238, 239 groups based on chain substitution, 240,241 length and substitution of chain, 240,241 lipid anchors, 236-240, 243, 249 location of, 274,275 metabolism, 247-254, 276 non-galactosylated species, 242 occurrence and structure, 235-243, 292,295 synthesis, linkages in, 253-254 synthesis, alanylation concomitant with, 263 synthesis of short-chain homologues, 252 unsubstituted, 242 poly(glycosylglycerophosphate), synthesis, 254 poly(hexosylglycerophosphate), 234 quantitative aspects, 247 release, penicillin effect, 273-274, 295 reviews on, 235 short-chain homologues, 252

alanylated, 263 space-filling models, 238,292-293, 295 Streptococcus pneumoniae, 246-247 structure, 235-247, 292-293,295 succinylated lipomannan in lieu of, 245-246 surface, proteins associated with, 275 teichoic acids relationship, 234 unsubstituted, 234, 241, 242, 282 action as carrier, 281, 282 model, 292 in vesicles, due to penicillin, 248, 274 Lipoteichoic acid-deacylating lipases, 273 Litho-autotrophs, megaplasmid role, 148 LTC, see Lipoteichoic acid, carrier Lysine, residue in halophilic proteins, 218 Lysophospholipids, 274

M M6 halophilic archaebacterium, see Halobacterium saccharovorum Magnesium, binding characteristics of teichoic and lipoteichoic acids, 291 binding to lipoteichoic acids, alanyl substitution effect, 294 limitation, phosphatidylglycerol content of cells, 269 RuBisCO activation, 135, 136 S subunit role, 138 teichoic and lipoteichic acid, role in enzyme activation, 293 role in scavenging, 291 Magnesium-dependent enzymes, 293 Malate dehydrogenase, 197-198 dual specificity, 197-198 H. marismortui, characteristics, 219 in methanogenic archaebacteria, 189 citrate synthase control, 214 in S. acidocaldarius, 189 in thermophilic archaebacteria, 187, 189,221 Thermoplasma acidophilum, 22 1 Mammalian cells, pili promoting adherence to, 54, 61, 83, 95 NMePhe pili, 96, 102

348

SUBJECT INDEX

Mannose-resistant haemagglutination (MRHA) activity, 75, 94 gene clusters, 76 Mannose-resistant (MR) pili, 61 animal-specific, 62 Mannose-sensitive (MS) pili, 61, 75, 94 Mating type, conjugative pili and, 58, 60, 87 Megaplasmid, 148 Mesophilic organisms, 220 Mesosomal vesicles, lipoteichoic acid associated, 275 Methane, formation, 182 Methanobacterium bryantii, nickel in hydrogenase, 20 Methanobacterium thermoautotrophicum, 181 class I aldolase in, 184 glucose catabolism in, 182 incomplete reductive citric acid cycle, 189, 191 malate dehydrogenase, 198 nickel in hydrogenase, 20 nickel and iron-sulphur clusters, 21 2-0x0 acid oxidoreductases, 202 succinate thiokinase, 21 5 Methanococcus voltae, 190 Methanogens, see Archaebacteria Methanosarcina barkeri, citrate synthase in, 214 dihydrolipoamide dehydrogenase in, 207 ferredoxin sequences, 205 incomplete oxidative citric acid cycle, 190 Methanospirillum hungatei, 190 dual specificity malate dehydrogenase, 198 Methylene blue, binding site, 23, 24 -dependent hydrogen oxidation, oxygen as inhibitor, 18, 24 oxidized, inhibition of hydrogen oxidation, 23 reduction, 16, 17, 18 Methylglyoxal pathway, 174 N-Methylphenylalanine (NMePhe) pili, see Pili Methyl viologen, 16, 17, 23 light activation of RuBisCO, 145

Micelle, lipoteichoic acid, autolysin inhibition, 284, 289 Microccus agilis, lipomannan in, 245 Microccusjavus, succinylated lipomannan, 245 Micrococcus luteus lipomannan, 245, 246 biosynthesis, 258 effect on autolysins, 285 location in mesosomal vesicles, 275 Micrococcus sodonensis, succinylated lipomannan, 245 Micrococcus varians, fatty-acyl composition of lipoteichoic acids, 239 transfer of preformed teichoic acid, 280 unsubstituted lipoteichoic acid, action as carrier, 282 Microcystis, cryptic plasmids, RuBisCO genes absent, 147 DNA, Anacystis nidulans L subunit probe hybridization, 147 Microtubules, carboxysome association with, 121-122 Mitochondria, eubacterial origin, 167 NAD+-linked isocitrate dehydrogenase in, 194 Mixotrophic growth of A . eutrophus, 8 mob site, 41, 45 Molecular genetics, techniques to study Rhizobium, 40-42 Molybdenum, in active site of nitrogenase, 3 nitrogen reaction within fixation reaction, 3 4 Monoclonal antibody, F pili, 86 gonococcal pili, 102 JEL92,90 Monophosphoglycerate mutase, 174 Moraxella, non-conjugative pili, 57, 63 Murein, 170 N NAD , dehydrogenases specific for, 194 +

SUBJECT INDEX

NAD+ (contd.) dihydrolipoamide dehydrogenase specific, in halophiles, 206 glucose dehydrogenase utilizing, 196197 isocitrate dehydrogenase utilizing, 194-197 malate dehydrogenase utilizing, 197198 in 2-0x0 acid dehydrogenase reactions, 200, 201 reduced, by H. saccharovorum, 177 NADH, b-type cytochrome reduction, 37 citrate synthase sensitivity, in archaebacteria, 214 cytochrome o reduction, 37 formation in citric acid cycle, 175 in hexose-monophosphate pathway, 172 hydrogen oxidation linked, 28 inhibition of citrate synthase in eubacteria, 2 10 NADH dehydrogenase, 28 NADP+ dehydrogenases specific for, 194 glucose dehydrogenase utilizing, 196197 isocitrate dehydrogenase utilizing, 194-197 NADPH, RuBisCO, effect on, 143 Neisseria, non-conjugative pili, 57, 63 Neisseria gonorrhoeae MSI 1 strain, 79, 80, 101 Neisseria gonorrhoeae pili, 63, 64, 79, see also Pili, NMePhe gene rearrangements, 80, 101 genetic organization, 79-8 1, 101 phase switching, 79, 102 pilin, see Pilin protein structure-function relationship, 100-102 receptor binding domain, 95, 102 silent ( p i l S ) and expressed (pilE) regions, 79, 102 as virulence factor, 100 X-ray diffraction studies, 67 Neisseria gonorrhoeae R10 strain, 101 Neisseria gonorrhoeae recA GC strain, 80

349

Neutron scattering studies, halophilic enzymes, 219, 220 Nickel, content of hydrogenase from bacteroids, 21 in EDTA inhibition of hydrogenase derepression, 20 in hydrogen metabolism, 19-21 role, 21 S subunit (RuBisCO) requirement, 138 Nickel hybride, 20 nifgene, 41,42,45 insertion sequences, 43 Nitrite-oxidizing bacteria, carboxysomes in, distribution and structure, 117-1 18, 153 numbers per cell, 1 18, 154 organisms having, 1 18 size and shape, 118 Nitrobacter agilis, carboxysomes in, appearance in, 1 18 lipid absence from, 125- 126 polypeptide composition, 126 stability in vitro, 124 DNAase-sensitive filament in, 128- 129 RuBisCO structure in, I34 Nitrobacter hamburgensis, X14,I29 RuBisCO structure, 134 Nitrobacter winogradsky, carboxysome numbers in stationary phase cultures, 154 carboxysomes appearance in, 118 DNAase-sensitive filament in, 128129 Nitrococcus mobilis, carboxysomes in, 117 Nitrogen, fixation, 2-4 in aerobic nitrogen-fixing filamentous bacteria, 122 efficiencyincrease by hydrogenase, 4, 5, 9 Hup- mutants lacking ability (Hup- Nif-), 3 9 4 0 Hup phenotype effect in R. leguminosarum, 46 in Oscillatoria (Trichodesmium) erythraea, I56

350

SUBJECT INDEX

Nitrogen (contd.) oxygen as limiting factor, 26-27 ratio of nitrogen to hydrogen produced, 3 reaction with molybdenum, 3-4 relative efficiency, 4-5 limitation, carboxysome numbers, 151,155 reduction, host control, 11 Nitrogenase, 2 4 , see also Nitrogen fixation dinitrogen reduction to ammonia, 2 electron allocation by, host control of, 11 host control, 11 hydrogen evolution in nitrogen fixation reaction, 2 4 energy cost, 24 hydrogen inhibition of, 4, 25 hydrogen sole source in R. japonicum, 16 molybdenum in active site, 3 protection from oxygen, by hydrogen oxidation, 4,25, 34, 35 reducing equivalents donated, 24 site-directed mutagenesis, 41 SR139 mutant lacking, 40 Nitrosomonas, carboxysomes in, 1 17, 118 Nitrosomonas europea, carbonic anhydrase in, 127 Nitrospina gracilis, carboxysomes absent, 117 Nitrospira marina, carboxysomes absent, 117 MNePhe, see Pili, NMePhe nod gene, 42,45 Nodulation, genes, 42, 45 Nostoc, chemoheterotrophic, RuBisCO levels, 154 Nostoc cyanobionts, carboxysomes in, 122 Nucleotide precursors, in chain substitution of lipoteichoic acids, 276

0 Oligonucleotide cataloguing, 166, 167, 168

Olisthodiscus luteus, 146 ompA protein, 87 TraTp protein contact, 88 oriT sequence, 68 Oscillatoria ( Trichodesmium) erythraea, 156 Ose- phenotype, 8 Osmotic shock, 295 Oxaloacetate, conversion into succinate, 193 Oxidase, terminal, 34, 35 flavoprotein as, 34 2-0x0 acid, 199 2-0x0 acid: ferredoxin oxidoreductase, 193, 199-205, see also under 2Oxoglutarate; Pyruvate in anaerobic conditions, advantage, 202,204 in archaebacteria, 202-203 in eubacteria and eukaryotes, 202 evolutionary considerations, 204-205 proposed catalytic mechanism, 201, 203 reaction mechanism, 201 size, 202 unique catalytic mechanism, 203-204 2-0x0 acid decarboxylase, 200 2-0x0 acid dehydrogenase, absent from anaerobic conditions, 202,212 in aerobic conditions, 201 dihydrolipoamide dehydrogenase role, 200, 201, 208 in eubacteria and eukaryotes, 203 evolutionary aspects, 204 reaction mechanisms, 200, 201 similarities with 2-0x0 acid oxidoreductases, 203-204 2-Oxoglutarate, 189, 193 effect on citrate synthase in archaebacteria, 214 inhibition of citrate synthase in eubacteria, 21 1 synthesis, 194 2-Oxoglutarate dehydrogenase, activesite coupling, 200 2-Oxoglutarate: ferredoxin oxidoreductase, in H . halobium, 202 in S . acidocaldarius. 189

351

SUBJECT INDEX

2-Oxoglutarate: ferredoxin oxidoreductase (contd.) in thermophiles, 187 2-Oxoglutarate synthase, evolution of, 193 in methanogens, 189 Oxygen, carbon dioxide specificity of RuBisCO, 140-142 consumption, by hydrogenase, detrimental effects (speculation), 25 hydrogen oxidation as protective mechanism by, 4, 25, 34, 35 dependent hydrogen oxidation, rate, 36 diffusion resistance, in nodule, 27 Hupc mutant regulation, 30-3 1 hydrogenase repression, mixotrophy and, 8 hypersensitivity mutants (Rhizobium), 6-7,40 inhibitor of, methylene blue-dependent hydrogen oxidation, 18, 24 RuBisCO carboxylation reaction, 137, 140, 153 inhibitory effect on photosynthesis, 141, 142 insensitivity mutants (Rhizobium),7, 40 lability of hydrogenases, 18-19, 27 as limiting factor in nitrogen fixation, 26-27 low, derepression of hydrogenase activity, 6, 7 electron transport coupled to ATP synthesis, 35 molar growth yield, hydrogen effect, 25-26 radical formation, 19 in regulation of hydrogen metabolism of Rhizobium, 6-9 repression insensitivity by Alcaligenes eutrophus mutants, 8 uncompetitive inhibitory of hydrogen in hydrogen oxidation, 23 Oxygenase reaction, RuBisCO, see Ribulose 1,5-bisphosphate carboxylase

P papA gene, 76, 77 papB gene, 77 papCD, papEFC and pap1 genes, 77 Pap (pyelonephritis associated pili), 55, see also Pili, Pap Paracoccus denitrijcans, aerobic, cytochromes in, 31 autotrophic, cytochrome o expression, 37 cytochromes in, 3 1 cytochrome aa3 in, 28 electron transport in hydrogen oxidation, 27 hydrogen oxidation-dependent ATP synthesis, 24 K,,, value of hydrogenase, 15- 16 Paulinella chromatophora, 123 P blood group, 5 5 , 61, 94 Pea cultivars, Alaska, 1 1, 12 Feltham First, 1 1 in host control of hydrogenase, 1 1 - 13 JI1205, 11-13 Peltigera canina, 122 Penicillin, effect on lipoteichoic acid, lipid and protein release, 273-274, 295 vesicle formation due to, 248, 274 Pentose-phosphate pathway, see Hexose-monophosphate pathway Peptidoglycan, biosynthesis, location of, 276 chain extension of, 249 Peptococcus aerogenes, 184 PH, effect of lipoteichoic acid content and synthesis, 267-268 effect of re-esterification of lipoteichoic acids, 265 thermoacidophilic archaebacteria, 167,217 Phase switching, in gonococcal pili, 79, 80, 102 Phase variation, 74, 77 Phenazine methosulphonate (PMS) reduction, 16, 17 Hup- mutant selection, 38, 39 Phenylarsine oxide, 209

352

SUBJECT INDEX

Phosphate, in F pili, 83, 85, 87 in bacteriophage attachment, 91 limitation, effect on lipoteichoic acid synthesis, 268-269,295 phosphodiesterase synthesis during, 272 Phosphatidic acid, synthesis, 260 Phosphatidic acid phosphatase, 250 Phosphatidylcholine, hydrogenase activity, 21-22 Phosphatidylethanolamine, in F pili, 83 synthesis location in B. megaterium, 276 Phosphatidylglucosyldiacylglycerol,250, 252 as acceptor substrate in lipoteichoic acid synthesis, 250, 252 Phosphatidylglycerol, biosynthesis inhibition, lipoteichoic acid synthesis block, 248 decrease, in phosphate limitation in B. subtilis, 269 formation from diacylglycerol, 259260 site of, 276 as glycerophosphate carrier, 250, 252, 276 glycerophosphate residues in lipoteichoic acids from, 234, 247, 250 in lipoteichoic acid synthesis, 234, 247,254 pools in B. megaterium, 261 structure, 235 synthesis of sn-glycero-1-phosphate of lipoglycan, 258 turnover, 258 for synthesis of lipoteichoic acids in bacterial doubling, 258 Phosphatidylglycolipid, membrane, lipoteichoic acids attached to, 234, 235 Phosphodiesterase, lipoteichoic acid degradation, 242, 249, 272 Phosphodiester bond, in poly(g1ycerophosphate) lipoteichoic acids, 234, 235, 240

Phosphoenol pyruvate (PEP), synthesis from pyruvate in methanogenic, archaebacteria, 184 Phosphoenol pyruvate (PEP) carboxykinase, 175 Phosphoenol pyruvate (PEP) carboxylase, 175, 189 6-Phosphofructokinase, not detected, in aerobic eubacteria, 172 in H . halobium, 179 in M . thermoautotrophicum, 182 in S. solfataricus, 179 in T. acidophilum, 181 6-Phosphogluconate (6PGLU), 142, 143 inhibitor of RuBisCO, 143, 144 oxidation, absent from H . saccharovorum, 177 2-Phosphoglycerate, formation in T. acidophilum, 180 Phosphoglycolipids, glycolipid and lipoteichoic acid relationship, 25 1 2-Phosphoglycollate, 143 Phospholipid, inhibition of autolysins, 288-289 Phosphomonoesterase, 242, 277-278 Phosphori bulokinase, absent from cyanelle inclusions, 132 in Chlorogloeopsis fritschii, 131 in cyanobacterial carboxysomes, 127 genes, Alcaligenes eutrophus, H 16, 148 Phosphorylated effectors, RuBisCO, 142-1 43 Photo-autotrophic prokaryotes, carboxysomes in, 121-123 Photorespiration, 140, 152 Photosynthate, as limiting factor in nitrogen fixation, 26 Photosynthesis, anaerobic, 193 carboxysome abundance and, 151, 152 improved nitrogen fixation, 13 inhibitory effect of oxygen, 141, 142 photorespiration interaction, 140 Photosynthetic bacteria, low C 0 2 / 0 2 specificity of RuBisCO, 141 Photosynthetic purple non-sulphur bacteria, RuBisCO in, 133 Phycobilisomes, 155

353

SUBJECT INDEX

Phylogenetic relationships, see also Archaebacteria rRNA sequence comparisons, 166, 167, 168 pilA gene, 74 in phase variation, 75 Pi1A’-lacZ fusion, 75 pilR, C, D genes, 74 pilE gene, 74 pilE region, pilin genes in gonococcus, 79, 80 Pili, 53-54, see also Pilin; individual bacteria 987P, animal-specific, 62 morphology and molecular weight, 57, 63 adhesive, 54, 55, 56, 61-63 antigenic determinants, 62, 94 of Escherichia coli, 61-63, 94-95 mannose-resistant (MR), 61, 62, 94,95 mannose-sensitive (MS), 61, 94, 95 protein structure-function relationships, 62, 94-95 receptor-binding domains, 94-95 antigenic determinants, see Antigenic determinants in Bacteroides nodosus, see Racteroides nodosus CFA/I, 56 morphology, 57 organization and expression of genes, 77-78 pilin subunit primary sequence, 62 CFA/II, CS2 N-terminal sequence, 62 organization and expression of genes, 77-78 CFA/II (CSl, CS2), morphology, 57 CFA/II (CSl, CS2, CS3) components, 54, 62 genes, 78 host serotype effect, 78 CFA/II (CS3), 57 classification, 54-55, 55-64 criteria for, 55 electron-microscope appearance, 57 function and biochemical properties, 57-64

incompatibility (Inc) in, 60 morphology, 55-57 conjugative, see also Plasmid-encoded elements; individual pili antigenic determinants, 85-86 bacteriophage sensitivity, 58 chemical composition, 83-85 derepressed mutants, 70 expression rate, HFT, 70 function and biochemical properties, 57-61 gene encoding pilus retraction, 68 morphology, 55, 56, 57, 58 nomenclature, 54, 55 organization and expression of genes, 68-73 plasmid-encoded elements, 68-71 role in identifying recipient and inducing contact, 60, 68, 87 summary of properties, 58 surface obligatory or surface preferred, 58, 60 tip of, functions, 87, 88, 89, 91 universal mating type, 58, 60 F, 54, 83 antigenic determinants, 85, 86 assembly and retraction, 87, 92-93 cleavage of RI 7-A protein, 87 cyclic AMP effect on, 72 functions, bacteriophage attachment, 89 interactions with recipient bacteria,

87-88 model based on high-resolution studies, 65, 66-67 monoclonal antibodies to, 86 pilin, see Pilin, F receptor on recipient cells (OmpAp), 87,88 sheared, fl bacteriophage attachment, 90 surface exclusion system, 88 F41, 57 animal-specific, 62 pilin subunit size, 63 flexible, mating type, 60 F-like, 85 antigenic determinants 85-86 C-terminus, 85, 91 glucose and phosphate in, 83, 85,87

354

SUBJECT INDEX

Pili (conrd.) phage attachment, 91 leader sequence, 92 N-terminus, 85, 89-90 phage interactions, 86-87 phage plating efficiency and cyanide effect, 90 protein associated with tip, 91 surface exclusion systems, 88-89 surface features, 89-9 1 unique configuration pilin subunits at tips, 89-90 functions, 54, 82, 87 Gal-gal, 55, 61, 94 gonococcal (GC), see Neisseria gonorrheae pili high-resolution studies, 6 4 6 8 IncF, see Plasmid, IncF K88 54 animal-specific, 62 antigenic variants (K88ab, K88ac, K88ad), 62-63,95 F41 relationship, 63 morphology, 57 organization and expression of genes, 78-79 pilin, see Pilin K99 54 animal-specific, 62 diarrhoea in animals, 63 expression, glucose reduction of, 77 molecular weight, sequence of pilin, 63 morphology, 57 organization and expression of genes, 78-79 mammalian cell adhesion, 54, 61, 83, 95 NMePhe pili, 96, 102 mannose-resistant, see Mannoseresistant NMePhe, 55, 56, see also N. gonorrhoea; Ps. aeruginosa antigenic determinants, 63, 94 function and biochemical properties, 63-64 genetic organization, 79-82 leader sequence, 99 N-terminal sequence, 64, 67, 99 nucleotide-sequence, 64

organisms expressing, 96 protein structure-function relationship, 96-102 X-ray diffraction studies, 64-68 nomenclature, 54-55 non-conjugative, morphology, 57 nomenclature, 54-55 PAK, antigenic determinants, 97, 99 model of, 66,67 pilin amino-acid sequence, 98, 99 pilin gene, 81 pili serotypes, 97 X-ray diffraction studies, 67 PAO, molecular weight of pilin, 82 pilin amino-acid sequence, 98, 99 pilin gene, 81 pili as virulence factor, 97 X-ray diffraction studies, 67 Pap, 55,75 adhesing, 55, 95 binding to P blood group, 5 5 , 61, 94 gene, 76-77 cluster, 75-76 similarity with K88 and K99, 78 homology with Type I, 62 morphology, 57 organization and expression of genes, 75-77 synthesis without adhesion function, 76 pED208, antigenic determinants, 85 surface exclusion system, 88 protein structure and function, 82102 Ps. aeruginosa (NMePhe), see Pseudomonas aeruginosa R100-1,84, 85 bacteriophage attachment, 90 receptor on recipient cells, 88 TraTp protein, 88 RI-19, 84, 85 antigenic determinants, 86 receptor on recipient cells, 88 R538-1, 84,85 receptors, 87-88, 88-89

SUBJECT INDEX

Pili (contd.) retraction, 96 evidence for, 93 genes encoding, 68 rigid, mating type, 60 synthesis, chromosomally, encoded control elements, 71-72 gene products (plasmid-encoded) involved in, 69-71 plasmid encoded control elements, 72 small effector molecules, 72-73 Type 1, adhesin, 95 CS 1, CS2 similarity, 62 genes, similarity with K88 and K99, 78 high-resolution studies, 64-65 homology with Pap pili, 62 mannose-sensitive, 61 MS haemagglutination, 94 organization and expression of pilin genes, 74-75 phase variation, 74 Pilin, ColB2, composition, 83, 84, 85 F, AP7 and Ap7*, 92 chemical composition, 83-85 molecular weight, 65 F-like, 61 central region, 91 K88, structure, 63, 94 subunit, 62, 63, 95 size, 63 K99, structure, 63, 94 MS and MR, 95 N. gonorrhoeae, as adhesin, 100, 101 antigenic determinants, 63, 94, 101-102 antigenic variation, 64, 100 hypervariable region, 101 nucleotide sequence, 79, 80, 100101 structure-function, 100-102 Pap, structure, 62, 94 pED208, composition, 83, 84, 85

355

processing from propilin, 69, 92 Ps. aeruginosa PAK and PAO, 98 transport across membranes, 82 Type 1, molecular weight, 64 structure, 62, 94 Pilin genes, 92 adhesive pili, sequence, 62, 63, 95 conjugative pili, 68-73 F pilin, sequencing of, 61, 68 NMePhe pili, nucleotide sequences, 64 organization and expression, 68-82 CFA/I and CFA/II pili, 77-78 chromosomally encoded elements, 7 1-72 K88 and K99 pili, 78-79 NMePhe pili, 79-82, see also Pili Pap pili, 75-77 plasmid encoded control elements, 72 in Ps. aeruginosa and B. nodosus, 81-82 small effector molecules, 72-73 Type I pili, 74-75 PAK, PAO, 81 sequence in Ps. aeruginosa, 99 Type I pili, 7 4 7 5 pilS region, pilin genes in gonococcus, 79, 80, 102 Pisum sativum, 10 Plasmid, see also DNA, extrachromsomal in autotrophic prokayotes (species with), 129 CFA/I pili, 77-78 DFA/II pili, 78 ColB2, insensitive to cyanide, no pilus retraction, 93 complementation in mutants, fertility inhibition, 70 control elements for organization and expression of pilin, 72 cryptic, 129, 147 DNA transfer to Rhizobium, 41 F, 68 F-like, 72 genes for hydrogen oxidation, 129 IncF, 60-61,68-71 map, 68

356

SUBJECT INDEX

Plasmid (contd.) N-terminus acetylation, 61 surface exclusion, 68 transfer region, genetic analysis, 69 transfer regions in one segment, 69 TraTp protein encoded, 88 IncI, 60, 72 receptor on recipient cells, 88 IncN, 60 pKM101,69, 72 R46,69 transfer regions, 69 incompatibility, 58-59, 60, 68 IncP, 60 IncW, 60 indigenous, Hup genes on in Rhizobium spp., 42-43 Nod-containing, R16J1, 42 pACYC177,41 pACYC184,41 Pap gene cluster cloned, 76 pBR325,4l, 43 pED208,70 pIJ1008,45 pRK2O 134,4I pRK290,41 pRL6J1, 45, 46 pVWJ31, pVW51,45 R27,69 R538-1, R124rd, R1-19, R136-1, 72 R91-5,69 RK2,41,69 RPI (IncP-I), 72 RP4, 4 1, 42,45 transfer region mapping, 69 self-transmissible encoding conjugative pili, 57, 68 in some cyanobacteria spp., 129 Ti, 69 Plasmid-encoded elements, pilin genes of conjugative pili, 68-71 surface exclusion, 68 transfer regions, 68, 69 Plastoquinone, 33 Poly(glycerophosphate), see Lipoteichoic acid Polyhedral bodies, see Carboxysomes Poly(hexosy1 glycerophosphate) lipoteichoic acids, 234, 243 Porphyridium cruentum, 146

Prophyrin, 193 ppGpp, in piliation control, 72, 73 Primer extension technique, 80 Prochloron, I22 RuBisCO heterologous hybridization of subunits, 139 RuBisCO present but phosphoribulokinase absent from, 132 Prochlorophyta, carboxysomes in, 122 free-living planktonic in Dutch freshwater lakes, 123 Prokaryotes, 166, see also individual species Calvin cycle, organisms, 1 16 chemolitho-autotrophic, see Chemolitho-autotrophic prokaryotes photo-autotrophic, 116, 121-123 RuBisCO L and S subunit genes in, 145, 146 Promotor, traJ, 73 tra Yz, 70-7 1, 73 Propilin, gene encoding, 69 NMePhe pili assembly, 64 pED208, 85 in pilus assembly, 69, 92 Protein, carboxysomes as storage bodies, 155 halophilic, 218 secretion by Gram-positive bacteria, penicillin effect, 274 synthesis, effect on lipoteichoic acid substitution, 271 initiation, in archaebacteria, 171 Proteolysis, 14 Proton release, from hydrogenase, 24 Protoplasts, biosynthesis of teichoic acids in, 275, 276 Pseudomonas aeruginosa CD4 and PA103, pilin amino-acid sequence, 98,99 Pseudomonas aeruginosa PAK/2Pf mutant, 96 Pseudomonas aeruginosa pili, bacteriophage receptors, 96

357

SUBJECT INDEX

Pseudomonas aeruginosa pili (contd.) conjugative, summary of, 59 genetic organization, 81-82 incompatibility groups, 60 NMePhe, 56, 63 non-conjugative, 57, 63 PAK, PAO, see also Pili, PAK; Pili, PA0 X-ray diffraction studies, 66, 67, 68 pilin, antigenic determinants, 63,94 gene nucleotide sequence, 99 structure-function relationship, 96100 twitching motility, 63, 96 as virulence factor, 96-97 Pseudomonas echinoides, nonstarforming (sta-) mutant, pili in, 65 Pseudomonas oxafaticus, I42 Pseudomonas saccharophilia, hydrogen oxidation-dependent ATP synthesis, 24 Pseudornonas thermophila carboxysomes, 129, 153 containing RuBisCO in, 121 DNA attachment to, 129 PstI restriction endonuclease, 81 Pulse-chase experiments, diacylglycerol recycling, 248, 259-260 lipoteichoic acid, metabolic fate, 272 metabolism, 252, 253, 260 synthesis, 247, 248 Pyrococcus, 221 Pyrodictium, 22 1 Pyruvate, formation, in archaebacteria, 177, 178-180 from glucose, ATP not required in H . saccharovorum, 117 from glyceraldehyde 3-phosphate in halophiles, 183 from 2-keto-3-deoxygluconate in S . solfataricus, 179, 191 glucose metabolism to, in eubacteria and eukaryotes, 172-1 74 metabolic fate, acetyl-CoA, 175

in archaebacteria (acetyl-CoA formation), 186 in eubacteria and eukaryotes, 175176 six types of reactions, I75 Pyruvate decarboxylase, 175 Pyruvate dehydrogenase, 175 acetyl-TPP formation in E. coli, 203 active-site coupling in, 200 in eubacteria and eukaryotes, diversity, 209 Pyruvate: ferredoxin oxidoreductase, see also 2-0x0 acid: ferredoxin oxidoreductase in archaebacteria, 177, 180, 186,202 in H. halobium, 202 in H . saccharovorum, I77 in T. acidophilum, 180 Pyruvate formate lyase, 202

Q Quinone, 181

R “Reactive Red” affinity columns, 13, 18 Redox potential, 17 component, 35,37 cytochrome c3, 17 cytochrome c, 33 flavoprotein, 33 Reducing agents, 19 Reducing equivalent, 24 Relative efficiency, 4-5 Rhizobium, “cowpea”, uptake hydrogenase in, 4 DNA transfer, cloning vehicles, 41 Hup+ strains, hydrogenase activity in, 2 hydrogen metabolism in, 1-52 molecular genetics techniques for, 4042 site-directed mutagenesis, 41-42 Tn5 mutants, 41-42 Rhizobium japonicum, autotrophic, 2, 6, 9

358

SUBJECT INDEX

Rhizobium japonicum (contd.) cytochrome pattern, 30-31 bacteroid, absorption spectrum and iron content, 14-15 component 559-H2, absence, evidence for, 37-38 cytochromes b- and c-reduction, 32,33 electron-transport system, 32-35 Hup' mutants, RuBp carboxylase activity absence, 10 hydrogenase, effect of nickel, 20 kinetic mechanism, 23-24 K , value, 17 nickel and iron content, 21 properties, 13 hydrogen oxidation, ATP increase, 24-25 hydrogen sole source due to nitrogenase, 16 hydrogen uptake (Hup) activity, 6, 7 oxygen-insensitive mutants, 7 free-living hydrogenase expression, 38 hydrogen oxidation electron transport, 28-32 oxygen and carbon regulation of hydrogenase, 6 gene bank, 43 Hup+ strains, 2, 6 beneficial effects, 5, 9 symbiotic advantage, 5 , 46 Hup' mutants, see Hydrogenaseconstitutive mutants Hup genes, 43-45 on indigenous plasmids in, 42-43 site-directed mutagenesis, 4 1 4 2 Hup- mutants, hup DNA excised from chromosome to create, 43 plasmids in, 42-43 Hup-specific DNA, homology with R. Ieguminosarum, 47 hydrogenase, 4, see also Hydrogenase anaerobic purification, half-life, 18 carbon dioxide fixation and, 9-10 carbon regulation, 6-9

detrimental action in oxygen consumption, 25 electron acceptor reactivity, 16-17 expression, cyclic AMP in, 7 high affinity for hydrogen, 16 host control, 10-1 1 increased efficiency of nitrogen fixation, 4, 9 iron-sulphur clusters, 15 K , value, 16 lipid requirement, 21-22 nickel in, 21 oxygen lability, 18-19, 27 oxygen regulation of, 6-9 purification and properties, 13-1 5,

18 hydrogen oxidation, cytochrome a03 and o in, 28, 29-32 efficiency, 16 maximal C2H2 reduction, 25 proposed electron-transport pathway, 31-32 ubiquinone in, 3 1 without nitrogen fixation, 2 hydrogen oxidizing, electron transport system, 27, 28-38 membrane particles, 27-28 nifgene, 42,43 oxygen-hypersensitive mutants, 6-7, 40 oxygen-insensitive mutants, 7, 40 RuBisCO structure in, 134 RuBP carboxylase in, 6, 9, 25 sphaeroplasts, 41 Rhizobium japonicum PJ17, PJ18, PJ20, 43 Rhizobium japonicum PJI 7nal, PJ18na1, 43 Rhizobium japonicum SR106, SR166, 39 Rhizobium japonicum SR 1 18, SR 146, 39 Rhizobium japonicum SRI 39 mutant (Hup- Nif-), 40,44 Rhizobium japonicum SR143 mutant (Hup- Nif-), 39-40 Rhizobium japonicum SRl, SR2, SR3,43 Rhizobium japonicum SU30647,45 Rhizobium japonicum USDA 122, 10-1 1, 39 Rhizobium japonicum USDA61, USDA74, 10-1 1

SUBJECT INDEX

Rhizobium leguminosarum, Hup+ strains, 4 Hup genes, 4 5 4 7 on plasmids, 42 Hup phenotype, effect on nitrogen fixation, 46 hydrogenase, host control of, 10, 1 1 hydrogen oxidation, ATP synthesis coupling, 25 nitrogenase protection from oxygen, 25 nitrogen fixation not increased with hydrogen oxidation, 5, 46 possible benefits of hydrogen oxidation, 5, 46, 47 Rhizobium leguminosarum strain 12, 300, 11 Rhizobium leguminosarum strain 128C53,44,45,47 host control of hydrogenase, 11 Rhizobium leguminosarum strain 16015 , Nod and Hup genes cotransferred, 42 Rhizobium leguminosarum strain 3960, 45 Rhizobium leguminosarum strain CNA 311, 10 Rhizobium leguminosarum strain ONA 311, 10 Rhizobium meliloti, site-directed mutagenesis, 41 Rhizobium meliloti 102F51, Hup genes, 44 Rhizobium ORS, 571, 25-26 Rhizobium strain 32H 1, 4 Rhodomicrobium vannielii, isocitrate dehydrogenase in, 195 RuBisCO structure, 133 Rhodopseudomonas (Rhodobacter) , RuBisCO structure, 133 Rhodopseudomonas blastica RuBisCO, S subunit function, 138 structure, 134 Rhodopseudomonas capsulata, nickel in hydrogenase, 20 RuBisCO S subunit function, 138 structure, 134 Rhodopseudomonas sphaeroides, cytochrome aa3 in, 28

359

Form I, 134, 138 RuBisCO genes on plasmid, 148 substrate specificity, 141 Form 11, 134, 138 RuBisCO gene on chromosome, 148 modified Entner-Doudoroff pathway in, 179 RuBisCO, gene cloning, 146 S subunit function, 138 structure, 134 Rhodospirillum rubrum, carbonic anhydrase in, 127 RuBisCO, as per cent of total protein, 132 activation site, 136 catalytic site, 137 gene cloning, 146 gene probe, 148 L subunit, amino acid sequence, 147 L subunit, gene number, 147 nucleotide sequence of gene, 146147 regulation, 140 structure, 133, 135 structure (model), 135 stimulation of inactive RuBisCO by 6PGLU, 142 Ribitol phosphate, 234, see also Teichoic acid polymerization, 280 Ribitol phosphate polymerase, 277, 278 Ribosome, eubacterial features, in archaebacteria, 170, 171 morphology, in phylogenetic analysis, 169 rRNA in, archaebacteria, 170 Ribulose 1,5-bisphosphate (RuBP), 135, 136 carboxysome membrane permeability, 152 Ribulose 1,5-bisphosphate carboxylase/ oxygenase (RuBisCO), 6, 39, 132149, see also Carboxysomes 8L8S, 133-134 in vitro construction, 139 occurrence, 133, 134, 135

360

SUBJECT INDEX

Ribulose 1,s-bisphosphate carboxylase/ oxygenase (RuBisCO) (contd.) removal of S subunits (catalytic core), 138 absence from, heterocysts, 122, 131 T. neutrophilus, 189 activase, 144-145 activation, 135-136, 144, 147 in oivo, 150 in light, 144, 145 site (lysine-201 of L subunit), 136, 137,147 activity, under carbon dioxide limitation, 150-1 51 antiserum, 125, 131 carbon dioxide fixation activity, 9-10 carbon dioxide/oxygen specificity, 140-142 carboxylation reaction, 136, 137 oxygen inhibition, 137, 140, 153 proposed scheme, 137-138 in carboxysomes, evidence for, 124-1 25 catalysis, 136-138, 147 catalytic site, 137, 147 metabolic effector interaction, 143 in chemoheterotrophic growth, 154 coregulation with hydrogenase in R . japonicum, 9, 10 in cyanelles, 123 diurnal changes, 144 function, 136, 137, 138, 155 gene, copy number, 147, 148 expression, 146, 149 on extrachromosomal DNA, evidence, 148 location and cloning, 145-146 nucleotide sequence, 146-147 probe, 147, 148 gene regulating, 10 genetics, 145-149 growth yield on oxygen, 25-26 heterologous subunit reconstruction, 138-1 39 importance, 116, 140, 155-157 inhibition, by 6PGLU, 143, 144 by sodium chloride, 154 inhibitors, 153, 154

endogenous, 144 kinetics, 140 large (L) subunit, 125, 133 function, 136, 137, 138 lysine-201, 136, 137, 147 man-made bodies containing, 156157 M , values, 133, 134 mutants lacking, 39 oxygenase reaction, 136, 137 advantages of abolition of, 140 oxygenation/carboxylation,enzymic partitioning, 139 phosphorylated effectors in regulation, 142-143 evidence and significance of, 143 in Prochlorophyta, 122 protection by carboxysomes, 152, 153-154 in Pseudomonas thermophila, 121, 129, 153 purification, 132-1 33 purification from R. japonicum, 9 regulation, carbon dioxide/oxygen, 140-142 endogenous inhibitors, 144 phosphorylated effectors, 142- 143 RuBisCO activase, 144-145 site-directed mutagenesis, 137, 138, 142, 149 glutamic acid change, 147 small (S) subunit, 125, 133 function, 138-140 to renature L subunits, 140 specificity and regulation, 140-145 spinach, activation site, 136 catalytic site, 137 hybridization with Synechococcus RuBisCO, 139 L subunit amino acid sequence, 147 stability in vitro, 132 storage in carboxysomes?, 154 structure, 133-135 L subunit heterogeneity, 134 microbial versus plant, 135 model (Alcaligenes eutrophus), 134135 multiple forms, occurrence, 134

SUBJECT INDEX

Ribulose 1,s-bisphosphate carboxylase/ oxygenase (RuBisCO) (contd.) subcellular distribution, 130, 131 symbiotic repression of, 10 in T . neapolitanus, see Thiobacillus neapolitanus tobacco, inhibitors, 144 structure, 135 toxic sulphur compound effect, I54 RNA, messenger (mRNA), eukaryotic features in archaebacteria, 171 ribosomal (rRNA), 16S/18S, measurement of phylogenetic relationships, 166, 167, 168 16s sequences, phylogenetic tree for archaebacteria, 168, 169 5S, in phylogenetic relationship analysis, 169 eubacterial features in archaebacteria, 170 hybridization homologies of archaebacteria, 169 Shine-Dalgarno sequence in halophiles, 170 transfer (tRNA), in archaebacteria, 170, 171 introns in genes in archaebacteria, 171 Root genotype, in host control of hydrogenase, 11, 12 Rotenone, 28 RuBisCO, see Ribulose 1,s-bisphosphate carboxylae/oxygenase

S Sacchaomyces ceruisae, 2 16 sfrA gene product, 71 sfrB gene product, 7 I Shine-Dalgarno sequence, 170 Shoot factor, in host control of hydrogenase activity, 1 I , 12 Sodium chloride, bridges, in thermophilic enzymes, 221 effect on alanyl residues in teichoic acid, 270, 271 halophile requirements, 167, 217

36 1

RuBisCO inhibition, 154 Sodium dodecyl sulphate (SDS), 83, 93 carboxysome dissociation, 125 Sodium dodecyl sulphate (SDS)polyacrylamide-gel electrophoresis (SDS-PAGE), 125,214,219 Sodium pyrophosphate, 67 Sorbitol, effect on lipoteichoic acid content of cells, 268,269 Soybean (Glycine max), hydrogenase activity control, 10 nickel effect on urease and hydrogenase, 20 oxygen as limiting factor in, 26 R. japonicum symbiosis, Hup+ trait effect, 5 Spinach, RuBisCO, see Ribulose 1,s-bisphosphate carboxylase oxygenase Spirulina platensis, 146, 220 Sporulation, lipoteichoic acid synthesis and, 270 Staphylococcus aureus, alanine ester turnover and transfer to teichoic acids, 263-264 composition of lipid amphiphiles in log growth, 258, 259 diacylglyerol recycling to phosphatidylglycerol, 248, 259-260 extracellular lipoteichoic acid, penicillin effect, 273 glycerophosphoglycolipids, glycolipids and lipoteichoic acids in, 235, 236 lipoteichoic acid, 234, 235, 236 acting as carrier, 277 alanyl content, glucose effect on, 271 alanyl content, salt effect on, 270, 271 alanyl residues in, 242 anchoring, sublethal heating effect, 273 content, effect of growth stage, 267 estimates of content, 247 location, 274, 275 mesosomal vesicles associated, 275 metabolic fate, 272 metabolism, 247, 248 modifications and anti-autolytic activity, 287, 290

362

SUBJECT INDEX

Staphylococcus aureus (contd.) poly(g1ycerophosphate)chain, 277 re-esterification, 265, 266 substituted, inactive as carriers, 280-28 1,282 synthesis, energy deprivation effect, 269-270 synthesis, membrane lipid metabolism, 259-260 synthesis (pulse-chase experiments), 252, 253 unsubstituted, 242 ribitol teichoic acid linkage unit, 278, 279 teichoic acid, 234 assembly on LTC in viuo, 283 preformed, transfer of, 280 teichoic acid-synthesizing enzymes, 277 toluene-treated, re-alanylation of teichoic acid, 265 Staphylococcus aureus 52A5 strain, teichoic acid deficiency, 295 Staphylococcus pneumoniae, see also Forssman antigen Staphylococcus xylosus, ribitol phosphate polymerase requirements, 278 ST genes, 77, 78 Streptococcus lactis Kiel 42 17 1, see Lactococcus garvieae Streptococcus mutans, lipoteichoic acid, content, carbohydrate source effect on, 268 growth stage effect on, 267 pH effect, 267 extracellular, growth stage and, 272 metabolic fate, 272 Streptococcus mutans BHT, 267, 268 extracellular lipoteichoic acid, 272 penicillin effect, 273 Streptococcus mutans Ingbritt, 267, 268, 269 extracellular lipoteichoic acid, 273 Streptococcus pneumoniae, Forssman antigen inhibitory to autolysin, 283, 284, 285 lipoteichoic acid, mesomal vesicles associated, 275 “lipoteichoic acid” from, 246-247

Streptococcus pyogenes M protein, 82 Streptococcus sanguis ATCC 10556 strain, 261 Streptococcus sanguis biotype B, poly(g1ycerophosphate) lipoteichoic acids absent, 245 Streptococcus sanguis lipoteichoic acid, glycosylation, 261 metabolism, 247 release, penicillin effect, 273 Succinate, cytochrome o reduction, 37 cytochrome reduction, evidence against component 559-H2, 36 cytochrome reduction in bacteriod membranes, 37-38 molar growth yield, RuBP carboxylase induction, 26 oxaloacetate conversion into, in evolution of citric acid cycle, 193 in porphyrin and amino acid synthesis, 193 in repression of hydrogenase activity, oxygen-insensitive mutants, 7 Succinate thiokinase, in archaebacteria, 213, 215-16 in eubacteria and eukaryotes, 210213 properties (summary), 212 in halophilic archaebacteria, 186, 215 in methanogenic archaebacteria, 189, 215 reaction catalysed by, 212 in thermacidophilic archaebacteria, 187,215 Succinyl-CoA, 189 Sugar catabolism, see Glucose Sulfolobales, 169 Sulfolobus, autotrophic, 187 heterotrophic growth on yeast, 183, 187 non-phosphorylated pathway of glucose catabolism, 177 oxidative citric acid cycle evidence lacking, 189 strain LM, non-phosphorylated modified Entner-Doudoroff pathway in, 180

363

SUBJECT INDEX

Sulfolobus acidocaldarius, citrate synthase and succinate thiokinase in, 214 citric acid cycle enzymes in, 189 glycerol, in ether lipids of, 185 glycerol synthesis in, 185-186 isocitrate dehydrogenase of, 189, 195-196, 198 malate dehydrogenase from, 198 2-0x0 acid oxidoreductases, 202 respiration-coupled phosphorylation, 181 triose phosphate isomerase in, 183 Sulfolobus brierleyi, non-phosphorylated modified Entner-Doudoroff pathway in, 180 reductive citric acid cycle in, 187, 189, 191 Sulfolobus solfataricus, glucose dehydrogenase (dual specificity), 196-197 non-phosphorylated modified EntnerDudoroff pathway, 178, 179 tRNA in, 171 Sulphur, toxic, effects on RuBisCO, 154 Sulphur-dependent Archaebacteria, see Archaebacteria Sulphur-oxidizing bacteria, colourless, carboxysome distribution and structure, 119-120, 153 dark environments, carboxysome absent, 155-156 RuBisCO in, 1 16 Surface exclusion, 68, 88-89, 89 F pili (genes and proteins in), 69, 88 SV sequences in gonococcal pilin genes, 79, 80 Symbiosis, hydrogenase synthesis and RuBP carboxylase repression, 10 Synechococcus, carboxylation mechanism of RuBisCO, 137 removal of RuBisCO S subunits, activity loss, 138 RuBisCO heterologous subunit reconstruction, 138-1 39 RuBisCO structure, 135 Synechococcus ieopoliensis, in carbon limitation, carboxysome numbers, 152, 154

carboxysome abundance versus photosynthetic characteristics, 15 1, 152, 154 nitrogen limitations, growth effect, 151, 155 oxygen protection mechanism for RuBisCO, 153-1 54

T Tamm-Horsfall protein, 61 Teichoic acid, 233 in action of N-acetylmuramyl-Lalanine amidase, 284, 285 D-alanyl residue transfer to, 263-265 salt effect on, 270-271 biosynthesis, location, 276 CDP-glycerol role in, 234 deficiency in Staph. aureus mutant, 295 definition, 234 degradation, 272 glycerophosphate residues in, 234 intracellular (membrane), 234 lipoteichoic acids relationship, 234 magnesium ion binding, 291 mode of chain growth, 249 poly(hexosy1 glycerophosphate), 234, 243 re-alanylation in Staph. aureus, 265 ri bi to1 phosphate, influence of substitution on LTC activity, 282 linkage to lipoteichoic acid carrier, 277-278 polymerization and transfer to linkage unit, 280 synthesis (pathway for), 277-278, 279 synthesis, 234 enzymes in, 277 linkage unit (structure and synthesis), 278, 279 transfer of preformed, 280 Teichoic acid lipid complexes, 234 Teichoicases, 272 Teichuronic acid, 268 Temperature, effect on alanine content of lipoteichoic acids, 271

364

SUBIECT INDEX

Temperature (contd.) pilus retraction and, 93 thermophile growth, 220,221-222 Terminal oxidase, 27, see also individual cytochromes cytochromes aa3 and o as, 29, 30 in R. japonicum bacteroids, 32 Tetra ethers, 170 Thermoacidophiles, 167 Thermocaccales, 169 Thermococcus celer, dihydrolipoamide dehydrogenase in, 207 Thermophilic marine bacteria, 222 Thermophily, 221 Thermoplasma, 221 non-phosphorylated pathway of glucose catabolism, 177 Thermoplasma acidophilum, 167 acetyl-CoA generation and conversion into acetate, 180 acetyl-CoA synthetase (ADPforming) in, 180 citrate synthase in, 213-214 dihydrolipoamide dehydrogenase in, 207 ferredoxin, 22 1 glucose dehydrogenase (dual specificity) in, 197 glycolytic enzymes not detected, 181 HTa protein in, 171 isocitrate dehydrogenase (dual specificity) in 196 malate dehydrogenase, 198, 221 non-phosphorylated modified EntnerDoudoroff pathway, 180, 181 oxidative citric acid cycle, 187, 191 2-0x0 acid oxidoreductases, 202 respiratory chain in, 181 succinate thiokinase in, 214 triose phosphate isomerase in, 183 Thermoproteales, 169 Thermoproteus neutrophilus, 181 carbon dioxide fixation pathways, 188, 189, 191 Thermostability, salt bridges, 221 Thermus aquaticus, 2 13 Thiamine pyrophosphate (TPP), 200, 202,204 Thiobacillus, carboxysomes in, I 19, 120

size and structure, 119 as taxonomic tool, 119 cryptic plasmids in, species having, 129 in dark deep-sea environments, 155 Thiobacillus albertis, carboxysomes in, 119 Thiobacillus intermedius, 151 Thiobacillus kakobis, carboxysomes in, 119 Thiobacillus neapolitanus carboxysomes, 119, 120 carbonic anhydrase absence, 127, 152 DNA associated, 129- 130 glycoproteins in, 125 large subunit heterogeneity in, 125 lipid absence from 126 polypeptides in, 126 role as storage body, 155 RuBisCO, 116, 120 as per cent of total protein, 132 activity, in carbon dioxide limitation, 150-1 51 carbon dioxide fixation rates, 150 distribution and levels, in oxygen changes, 153 K,,, (C02) values, 142 levels in nitrogen limitation, 155 shape, 1 19- 120 stability in vitro, 124 Thiobacillus thiooxidans carboxysomes, 119 Thiobacillus versutus, plasmids in, 129 Thiomicrospira, 155 Thiosphaera pantotropha, carboxysomes absent from, 119 plasmids in, 129 Thylakoids, 131 Tobacco mosaic virus, 67 Toluene-treated cells, 252 alanylation mechanism of lipoteichoic acid, 262 cyanobacteria, RuBisCO in, 143 lipoteichoic acid metabolism, 247, 249 re-esterification of lipoteichoic acid after alanine ester loss, 265, 266 traA gene, 69 mutants and bacteriophage - attachment, 89

365

SUBJECT INDEX

traG gene, 69, 92 traJ gene, 69, 70 TraJp protein, 70-71 sfrA and cpxAB affecting, 71 traM, 69, 72 traMYI gene, 69 Transduction, 41 Transcription, control of gonococcal pilin gene expression, 80 eukaryotic features of, in archaebacteria, 171 F transfer operon, 71, 72 in Hfr strain, 73 pap genes, 77 regulation of pilus expression by pilA and hyp genes, 75 RuBisCO subunit genes, 146, 149 Transferase, in glycolipid and glycerophosphate linkage in lipoteichoic acids, 253 Transposon, Tn5,41, 43,44,45 hyp gene inactivation in E. coli, 75 traQ gene, 69 in pilin processing, 69, 92 traS gene, 88 traST gene, 69 traT gene, 88 TraTp protein, 88 traYZ, 69, 70, 72 Triose phosphate isomerase, 183 Triphenyltetrazolium chloride, 38-39 Triton X-100,280,290 Trypanosoma brucei, 208 Tryptophan synthetase, hydrophobicity causing stability, 221

“Twitching motility”, 63, 96

U Ubiquinone in hydrogen oxidation, 31 UDP-glucose, 261 Urease, nickel effects on, 20 Urinary tract infections, adhesive pili of E. coli in, 55, 61 V

Vesicles, lipoteichoic acids, lipids and proteins in, 248, 274 mesosomal, lipoteichoic acid associated, 275 Vibrio cholera, NMePhe pili, 63 Vicia bengalensis, 10 Viciafaba, 10 Vicia unguiculata (cowpea), 10

X X-adhesins, 95 Xanthobacter autotrophicus, hydrogenase, cytochrome associated, 14 X-ray diffraction, pili structure, 64-65, 67 X-ray scattering, halophilic enzymes, 219,220

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