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Advances in
MICROBIAL PHYSIOLOGY VOLUME 47
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Advances in
MICROBIAL PHYSIOLOGY Edited by
ROBERT K. POOLE West Riding Professor of Microbiology Department of Molecular Biology and Biotechnology The University of Sheffield Firth Court, Western Bank Sheffield S10 2TN, UK
Volume 47
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03 04 05 06 07 08 MP 9 8 7 6 5 4 3 2 1
Contents
CONTRIBUTORS
TO
VOLUME 47
ix
Physiological Diversity and Niche Adaptation in Marine Synechococcus David J. Scanlan
1. 2. 3. 4. 5. 6. 7. 8.
Abbreviations . . . . . . . . . Introduction . . . . . . . . . . Light-harvesting apparatus C metabolism. . . . . . . . . . Nutrient acquisition . . . . . Motility . . . . . . . . . . . . . . Cell cycle . . . . . . . . . . . . . Grazing/viruses . . . . . . . . Conclusions . . . . . . . . . . . Acknowledgements . . . . . . References . . . . . . . . . . . .
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Adoption of the Transiently Non-culturable State – a Bacterial Survival Strategy? Galina V. Mukamolova, Arseny S. Kaprelyants, Douglas B. Kell and Michael Young Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 2. Bacterial stress avoidance strategies . . . . . . . . . . . . . . . . . . . . . . . 69
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CONTENTS
3. 4. 5. 6.
. . . . . 92 . . . . . 94 . . . . . 96
Are there specific chemical inducers of ‘‘non-culturability’’? . Is ‘‘non-culturability’’ genetically controlled?. . . . . . . . . . . . Resuscitation of ‘‘non-culturable’’ cells. . . . . . . . . . . . . . . . Social behaviour of bacterial populations and ‘‘non-culturability’’. . . . . . . . . . . . . . . . . . . . . . . . . . . 7. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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103 106 109 109
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132 132 176 176
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188 188 192 219 236 239 239 240
The Biodiversity of Microbial Cytochromes P450 Steven L. Kelly, David C. Lamb, Colin J. Jackson, Andrew G.S. Warrilow and Diane E. Kelly Abbreviations . . . . 1. Introduction . . . . . Acknowledgements . References . . . . . . .
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The Tat Protein Translocation Pathway and its Role in Microbial Physiology Ben C. Berks, Tracy Palmer and Frank Sargent
1. 2. 3. 4. 5.
Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Journey to the translocase . . . . . . . . . . . . . . . . . . . . . . . . . . Transport across the membrane . . . . . . . . . . . . . . . . . . . . . . Biosynthesis of integral membrane proteins by the Tat system Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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Microbial Globins Guanghui Wu, Laura M. Wainwright and Robert K. Poole Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Globins – definition and the classical view . . . . . . . . . 2. The Vitreoscilla globin (Vgb) and other single domain myoglobin-like globins . . . . . . . . . . . . . . . . . . . . . . . 3. Truncated globins. . . . . . . . . . . . . . . . . . . . . . . . . . .
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CONTENTS
4. Flavohaemoglobins. . . . 5. Evolution of Globins . . 6. Summary. . . . . . . . . . . Acknowledgements . . . . References . . . . . . . . . .
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275 298 299 300 300
Author Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 311 Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 323
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Contributors to Volume 47
BEN C. BERKS, Department of Biochemistry, University of Oxford, South Parks Road, Oxford OX1 3QU, UK COLIN J. JACKSON, Wolfson Laboratory of P450 Biodiversity, Institute of Biological Sciences, Edward Llwyd Building, University of Wales, Aberystwyth, Ceredigion SY23 3DA, UK ARSENY S. KAPRELYANTS, Bakh Institute of Biochemistry, Russian Academy of Sciences, Leninsky pr. 33, 117071 Moscow, Russia DOUGLAS B. KELL, Institute of Biological Sciences, University of Wales, Aberystwyth, Ceredigion SY23 3DD, UK and (current address) Department of Chemistry, UMIST, Sackville St, PO Box 88, Manchester M60 1QD, UK STEVEN L. KELLY, Wolfson Laboratory of P450 Biodiversity, Institute of Biological Sciences, Edward Llwyd Building, University of Wales, Aberystwyth, Ceredigion SY23 3DA, UK DIANE E. KELLY, Wolfson Laboratory of P450 Biodiversity, Institute of Biological Sciences, Edward Llwyd Building, University of Wales, Aberystwyth, Ceredigion SY23 3DA, UK DAVID C. LAMB, Wolfson Laboratory of P450 Biodiversity, Institute of Biological Sciences, Edward Llwyd Building, University of Wales, Aberystwyth, Ceredigion SY23 3DA, UK
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CONTRIBUTORS TO VOLUME 47
GALINA V. MUKAMOLOVA, Institute of Biological Sciences, University of Wales, Aberystwyth, Ceredigion SY23 3DD, UK and Bakh Institute of Biochemistry, Russian Academy of Sciences, Leninsky pr. 33, 117071 Moscow, Russia TRACY PALMER, Department of Molecular Microbiology, John Innes Centre, Norwich NR4 7UH, UK ROBERT K. POOLE, Department of Molecular Biology and Biotechnology, The University of Sheffield, Firth Court, Western Bank, Sheffield S10 2TN, England, UK FRANK SARGENT, Centre for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich NR4 7TJ, United Kingdom DAVID J. SCANLAN, Department of Biological Sciences, University of Warwick, Gibbet Hill Road, Coventry CV4 7AL, UK LAURA M. WAINWRIGHT, Department of Molecular Biology and Biotechnology, The University of Sheffield, Firth Court, Western Bank, Sheffield S10 2TN, England, UK ANDREW G.S. WARRILOW, Wolfson Laboratory of P450 Biodiversity, Institute of Biological Sciences, Edward Llwyd Building, University of Wales, Aberystwyth, Ceredigion SY23 3DA, UK GUANGHUI WU, Department of Molecular Biology and Biotechnology, The University of Sheffield, Firth Court, Western Bank, Sheffield S10 2TN, England, UK MICHAEL YOUNG, Institute of Biological Sciences, University of Wales, Aberystwyth, Ceredigion SY23 3DD, UK
Physiological Diversity and Niche Adaptation in Marine Synechococcus David J. Scanlan Department of Biological Sciences, University of Warwick, Gibbet Hill Road, Coventry, CV4 7AL, UK
ABSTRACT During the twenty years or so since the discovery of tiny photosynthetic cells of the genus Synechococcus in marine oceanic systems, a tremendous expansion of interest has been seen in the literature pertaining to these organisms. The fact that they are ubiquitous and abundant in major oceanic regimes underlies their ecological importance as significant contributors to marine C fixation. Recent advances in the physiology and biochemistry of these organisms are presented here, focusing on strains of the MC-A and MC-B clusters; it is stressed that the data contained herein should be put into the context of the ecological niche occupied by particular genotypes in situ. This system is ripe for joining the often separate disciplines of molecular ecology and microbial physiology and provides a great opportunity to tease out the underlying processes that both mediate organism evolution and also the environmental factors that dictate this.
Abbreviations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 1.1. The phylogeny of marine Synechococcus . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 2. Light-harvesting apparatus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
ADVANCES IN MICROBIAL PHYSIOLOGY VOL. 47 Copyright ß 2003 Elsevier Science Ltd ISBN 0-12-027747-6 All rights of reproduction in any form reserved
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3. C metabolism. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 3.1. Photosynthetic physiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 3.2. C fixation: RuBisCO and carbon-concentrating mechanisms . . . . . . . . . . . . . 14 4. Nutrient acquisition. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 4.1. N acquisition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 4.2. P acquisition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 4.3. Micro-nutrient acquisition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36 5. Motility. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39 6. Cell cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39 7. Grazing/viruses. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 44 8. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48
ABBREVIATIONS PUB PEB PE CCM FCCP DCMU DBMIB
phycourobilin phycoerythrobilin phycoerythrin carbon-concentrating mechanism carbonylcyanide p-trifluoromethoxy-phenylhydrazone (3-(3,4-dichlorophenyl)-1,1-dimethyl urea) 2,5-dibromo-3-methyl-6-isopropyl-p-benzoquinone
1. INTRODUCTION Single-celled picoplankton assigned to the genus Synechococcus (Johnson and Sieburth, 1979; Waterbury et al., 1979) are the dominant phycobilisome-containing cyanobacteria in the world’s oceans. They have a ubiquitous distribution, though generally being more abundant in nutrient-rich rather than oligotrophic areas, and are important contributors to global marine carbon (C) fixation, e.g., contributing up to 20% of fixed C in some areas (Li, 1994). They also play a key role in pelagic food-web structure via energy transfer within the microbial loop, and particularly in oligotrophic regions through heterotrophic flagellate and ciliate grazing. Indeed it has been estimated that 35–100% of the Synechococcus standing stock can be grazed per day (Campbell and Carpenter, 1986a). Since the oceanic ecosystem encompasses a range of water bodies from eutrophic through mesotrophic to oligotrophic, and delineates a complicated light environment, the physiology of these organisms has
DIVERSITY AND ADAPTATION IN SYNECHOCOCCUS
3
largely focused on their ability to acquire nutrients at sub-micromolar concentrations, and to harvest light over a range of different intensities (down a water column) and qualities (e.g. coastal vs open-ocean waters). As well as diel periodicity and seasonal patterns in the solar light environment, there may be concomitant changes in water column structure e.g., stratified versus well-mixed waters, which also magnify or alter the light and nutrient gradients in situ. Such large-scale changes to an organism’s environment, typified by summer versus winter water column conditions in temperate waters, may well have selected for ‘plastic’ genotypes present throughout the year, capable of responding to a wide spectrum of environmental change. Conversely, such conditions may have provided strong selective pressure for specialisation to a specific niche, and given rise to genotypes which appear and disappear in tandem with the presence/ absence of that niche. Such a niche partitioning of genotypes is exemplified by the ‘surface’ (high light-adapted) versus ‘deep’ (low light-adapted) ecotypes well documented for the Prochlorococcus genus (Moore et al., 1998; West and Scanlan, 1999), with the underlying physiological basis to this adaptation beginning to be uncovered thanks to recent genomic information (see Hess et al., 2001; Scanlan and West, 2002; Ting et al., 2002). There is also evidence of freshwater Synechococcus strains with different light tolerance properties that may correspond to high and low light-adapted ecotypes (Postius et al., 1998). This partitioning of genotypes in situ is not restricted to cyanobacteria, however. Surface- and deep-water clades of marine bacteria capable of a novel type of proteorhodopsin-based phototrophy have also been recently recognised, with the idea that subsets of proteorhodopsin genetic variants in mixed populations have a selective advantage at different positions in the depthdependent light gradient (Be´ja` et al., 2000, 2001). Furthermore, the SAR 11 a-proteobacteria and SAR 202 green non-sulphur bacteria clusters have been shown to partition in the water column, though as yet to undefined environmental variables (Giovannoni et al., 1996; Field et al., 1997). Further examples of both niche-adapted strains and species can also be found in the eukaryote kingdom, including temporal variation of genetically different clones (Gallagher, 1980, 1982; Venrick, 1990, 1998). This genetic and physiological variation among eukaryotic algal species has already led some authors to re-address the species concept in phytoplankton ecology (Wood, 1988; Wood and Leatham, 1992). With the idea of niche adaptation in mind, recent advances in the physiology and molecular biology of the marine Synechococcus genus are specifically reviewed here (see Glover, 1985; Stockner and Antia, 1986; Waterbury et al., 1986; Fogg, 1987; Carr and Mann, 1994; Partensky et al.,
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1999a for earlier reviews on the genus). Where available, particular focus is put on the comparative physiology of strains of different phylogenetic origin, with the aim of presenting this genus as a group of niche-adapted ‘clades’. Ultimately, then, these physiological data would need to be put into the context of the specific niche occupied by individual ecotypes. At present, we are just beginning to obtain the molecular tools necessary to initiate such work through the development of molecular oligonucleotide probes for assessing the in situ distribution of individual genotypes (Fuller et al., 2003). Reference to this in the next section will lay the foundation to Sections 2 to 4 describing specific physiological adaptations of this genus to the light and nutrient gradients of the oceans. The recent completion of the entire genome sequence of Synechococcus sp. WH 8102 [see http:// bahama.jgipsf.org/prod/bin/microbes/syn/home.syn.cgi] will allow a more detailed characterisation of the physiology of these organisms, and in particular provide a basis for elucidating genomic differences amongst genotypes and identify niche-specific genes, as well as allowing complete transcriptomic and proteomic studies of the response of this specific genotype to environmental change.
1.1. The Phylogeny of Marine Synechococcus Within the cyanobacteria, the genus Synechococcus is attributed to unicellular rod-shaped to coccoid organisms less than 3 mm in diameter, dividing by binary fission into equal halves in one plane. They contain photosynthetic thylakoid membranes located peripherally and lack structured sheaths (Waterbury and Rippka, 1989; Herdman et al., 2001). The genus includes both marine and freshwater isolates and is clearly polyphyletic (Honda et al., 1999; Robertson et al., 2001). Hence, organisms currently classified as Synechococcus await re-classification into several different genera. The marine Synechococcus isolates have themselves been classified into three groups, designated marine cluster (MC) -A, -B and -C (MC-A, MC-B and MC-C), based on the composition of the major light harvesting pigments, an ability to perform a novel swimming motility (see Section 5), whether they have an elevated salt requirement for growth, and G þ C content (Waterbury and Rippka, 1989). The nomenclature of these groups has, however, been recently re-defined (Herdman et al., 2001) (see Table 1). For the purposes of this review (see Scanlan and West, 2002), we consider only the physiology of strains within the MC-A and MC-B clusters since together with Prochlorococcus (see Chisholm et al., 1988; Partensky et
DIVERSITY AND ADAPTATION IN SYNECHOCOCCUS Table 1
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Characteristics of the marine Synechococcus clusters MC-A, MC-B and MC-C1.
Marine cluster A* includes open-ocean and coastal isolates cells range in diameter from 0.66–1.7 mm some are capable of a swimming motility all contain phycoerythrin (PE) as their major light-harvesting pigment. Some are capable of chromatic adaptation all are obligate photoautotrophs incapable of using organic compounds as sole source of cell C all strains have elevated growth requirements for Naþ, Cl, Mg2þ, Ca2þ mol% GþC ranges from 55–62% Marine cluster B* coastal marine isolates cells range in diameter from 0.9–1.4 mm all are non-motile all contain phycocyanin as their major light-harvesting pigment, PE is absent all are obligate photoautotrophs incapable of using organic compounds as sole source of cell C contain mostly halotolerant strains e.g., WH 5701, but one, WH 8007, has an elevated salt requirement for growth mol% GþC ranges from 63–69% Marine cluster C** coastal marine or brackish water isolates cells range in diameter from 1.2–2.0 mm all are non-motile 4 of 5 strains contain phycocyanin as their major light-harvesting pigment. One strain (PCC 7335) produces C-phycoerythrin and is capable of chromatic adaptation. This strain can also synthesise nitrogenase under anaerobic conditions 4 of 5 strains are capable of photoheterotrophic growth 3 strains are halotolerant (e.g. PCC 7002), and two have an elevated salt requirement for growth (PCC 7335 and PCC 7003) mol% GþC ranges from 47–50% 1 Data from Waterbury and Rippka (1989), Herdman et al. (2001), Palenik (2001) *Marine clusters A and B (Waterbury and Rippka, 1989) have recently been combined as two sub-clusters into Synechococcus Cluster 5 (Herdman et al., 2001). Thus, MC-A becomes Synechococcus Cluster 5.1 and MC-B becomes Synechococcus Cluster 5.2. Strain PCC 7001, previously a member of MC-B has been transferred to the genus Cyanobium (Herdman et al., 2001). **Marine cluster C now corresponds to Synechococcus Cluster 3 (Herdman et al., 2001) but with strain PCC 7335 removed. This latter strain now comprises the only member of a new cluster, Synechococcus Cluster 4. The mean DNA base composition (47.4 mol% GþC) of PCC 7335 is slightly lower than other members of cluster 3 and the genome size is markedly greater (3.1 Gdal) (Herdman et al., 2001).
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al., 1999b), the sister taxon to Synechococcus, they constitute a single picophytoplankton clade. We also retain the MC-A and MC-B designations. Considerable genetic diversity is found amongst cultured isolates of these clusters as defined by sequencing of rpoC1 encoding a sub-unit of the DNA-dependent RNA polymerase (Toledo and Palenik, 1997; Toledo et al., 1999), the 16S rDNA (Urbach et al., 1998; Fuller et al., 2003) and the intergenic spacer (Rocap et al., 2002) as well as RFLP studies of other gene loci (Douglas and Carr, 1988; Wood and Townsend, 1990) [reviewed in Scanlan and West, 2002]. This work demonstrates a good phylogenetic congruence amongst the different gene loci and has identified at least ten distinct clades within MC-A (see Fig. 1), though one of these, (clade VIII), includes isolates, e.g., WH 8101, recognised as members of MC-B. This considerable genetic diversity observed within the MC-A Synechococcus group is manifest though in only small-scale variation in 16S rRNA gene sequence (Fuller et al., 2003). Such microdiversity, as it has become known (see Moore et al., 1998; Casamayor et al., 2002), can give rise to physiologically quite distinct ecotypes, typified by the high and low lightadapted Prochlorococcus clades (Moore et al., 1998) already mentioned. Since the extent of this microdiversity seems greater within the Synechococcus genus compared to Prochlorococcus, we might anticipate then a relatively large physiological diversity in these organisms. This might be expected given the ubiquitous distribution of Synechococcus in marine waters, probably reflecting the greater diversity of ‘niches’ occupied by strains of this genus. The type and shape of niche occupied by these phylogenetically defined clades largely remain to be identified. Even so, some insights with regard to the distribution of specific clades can be made based on molecular and immunofluorescence work. Thus, strains within clade I show a dominance, in rpoC1 clone libraries, in waters off the Californian coast following recent mixing events or during coastal water intrusion, and appear to be absent in clone libraries during highly stratified oligotrophic conditions (Ferris and Palenik, 1998). Similarly, antisera raised against Synechococcus sp. WH 8016, phylogenetically also within clade I, showed this serogroup to be abundant in coastal and estuarine stations off Long Island, whose appearance was correlated with water temperatures >15 C (Campbell and Carpenter, 1987). Further, polyclonal antibodies raised against strain Synechococcus sp. CC9605, a representative of clade II, hint at an oligotrophic surface distribution (Toledo and Palenik, in press), whilst other immunofluorescence assays have shown a differential distribution of WH 7803 (MC-A clade V) and WH 5701 (MC-B) serogroups (Pomar et al., 1998). A distribution of members of clade II biased towards
DIVERSITY AND ADAPTATION IN SYNECHOCOCCUS
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Figure 1 Consensus tree based on a neighbour joining tree using 1345 nucleotides with short sequences added by parsimony. Bootstrap values are from 100 data sets by neighbour joining analysis with Jukes–Cantor correction. Where full-length sequences were available 1345 nucleotides were used for analysis, the values represented as circles. Where only shorter sequences were available 681 nucleotides were used for analysis, the values represented as triangles. Closed objects represent bootstrap values >95%. Open objects represent bootstrap values 70–95%. Values 1 0.9
þ
1.3
þ
þ
0.8
þ
þ
1.6
þ
Fuller et al., 2003
þ
1.4
þ
Fuller et al., 2003
þ
1.8
þ
Fuller et al., 2003
RS9905 RS9915 RCC311 Max 42 Minos 12
þ
þ
þ
þ
þ
þ
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nitrate (þ) ammonium (þ) nitrate (þ) ammonium (þ)
Rocap et al., 2002 Fuller et al., 2003 Fuller et al., 2003 Fuller et al., 2003
23
(continued )
24 Table 3 Cont. Phycobiliproteins Phylogenetic clade & Strain No.
Isolation location
Mol % G þ C#
Phycoerythrin (PE)
Present (þ) Absent () MC-A Clade V WH 7803
RS9705 RS9708 MC-A Clade VI WH 7805
Sargasso Sea (33 44.9’N, 67 29.8’W)
61.3
Red Sea Red Sea
PUB:PEB ratio
þ
0.39****
þ þ
40% amino acid identity and the same subfamily if they share >55% amino acid identity (e.g. CYP52A1, CYP52A2, etc.). The rules can be relaxed if it is found that CYPs share the same function and the use of italics indicates that the gene is being described rather than the protein. Some CYP families have been found to overlap and no doubt there will be a revisiting of the nomenclature in the future. The current scale of biodiversity was not anticipated so that originally CYP1-49 were set aside for animal forms, only CYP51-69 were allocated for lower eukaryotes and the families CYP71-99 were set aside for plants. Bacteria were assigned family numbers ranging
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upwards from CYP 101. The lower eukaryotic CYP families are now expanding from CYP501 upwards and plant CYP families from CYP701 upwards.
1.1. Discovery of CYPs in Microbial Systems CYPs were first identified in 1958 in mammalian liver microsomal samples (Klingenberg, 1958). Reduction of the sample with sodium dithionite, followed by exposure to carbon monoxide, resulted in the appearance of a distinctive absorbance band at approximately 450 nm (Fig. 1). The name cytochrome P450 – i.e. pigment with an absorption peak at 450 nm – (Omura and Sato, 1962) has subsequently been retained despite debate as to whether the term ‘‘cytochrome’’ is strictly correct. The first microbial CYP identified was in S. cerevisiae (Lindenmayer and Smith, 1964) and subsequently experimental biology and genomic projects have confirmed the distribution of CYP in many microbes. The first bacterial CYP, from Bradyrhizobium japonicum, was reported by Appleby (1967). In general, bacterial CYPs are soluble enzymes being localised exclusively in the cytosolic fraction of the cell. Their eukaryotic counterparts are membranous, usually associated with the smooth endoplasmic reticulum (ER) through a single
Figure 1 The reduced carbon monoxide difference spectrum of cytochrome P450, in this case of purified CYP51 of Mycobacterium tuberculosis.
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Figure 2 The CYP catalytic cycle depicted for eukaryotic Class II forms. Substrate binds to ferric CYP and the first electron donation reduces CYP and, with oxygen binding, produces an oxyferrous state. With input of a second electron the oxygen is activated and an atom is incorporated into the substrate with the subsequent release of product and water. The primary electron donor is NADPH-cytochrome P450 reductase and the cytochrome b5/NADH-cytochrome b5 reductase can provide the second electron.
transmembrane spanning anchor located at the NH2-portion of the protein (Larson et al., 1991). For catalytic activity, CYPs must be associated with their electron donor partner proteins, either ferredoxin/ferredoxin reductase for prokaryotic CYPs (so called class I or type I CYP systems) or NADPH cytochrome P450 reductase (CPR) for ER-associated eukaryotic CYPs (class II), although there are exceptions to these general systems as will be discussed. One CPR supports many CYP reactions in eukaryotic cells in the ER and other locations although class I-type mitochondrial systems have been found in animals (but not in microbes). In mammals, it has been suggested that CPR must be present for CYP activity (Black and Coon, 1982); the extent of functional redundancy among electron donor systems within biodiverse cells, including among ferredoxins and ferredoxin reductases of bacteria, remains to be determined comprehensively. For completion of the CYP catalytic cycle, two electrons are required (Fig. 2). The cycle begins with substrate binding to ferric CYP and conversion to a ferrous CYP on reduction with the first electron transfer and subsequent formation of an oxygen-bound (and substrate-bound) CYP. Next, on further transfer of the second electron, a peroxy-ferrous CYP state is produced followed by addition of an atom of oxygen to the substrate and
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in to water. The general CYP reaction can be summarised below where R is the substrate: Hþ þRH þ NADPH þ O2 ! ROH þ NADPþ þH2 O
1.2. CYP51 (sterol 14a-demethylase), an ancestral CYP form CYP from S. cerevisiae was extensively characterised by Yoshida and colleagues (Yoshida et al., 1974; Aoyama et al., 1984; Yoshida and Aoyama, 1984). Through biochemical experiments lanosterol 14a-demethylase (P45014DM), part of the sterol biosynthetic apparatus, was investigated and subsequently designated CYP51 in the superfamily – the first microbial eukaryotic CYP to be cloned and sequenced (Kalb et al., 1987). As in other areas of eukaryotic biology, work on yeast provided insight into CYP biology generally as the CYP studied emerged as the first member of the only family found in animals, plants and fungi (for review, Kelly et al., 2001). Based on studies with different systems, CYP51 was found to catalyse three successive monooxygenation reactions resulting in the formation of 14-hydroxymethyl and 14-carboxaldehyde derivatives, followed by elimination of formic acid and the introduction of a 14,15 double bond (Fig. 3; Alexander et al., 1972; Akhtar et al., 1977, 1978; Aoyama et al., 1987, 1989). The initial hydroxylation appeared rate-limiting with the 3-OH and C8 double bond of the sterol substrate being essential for activity (for review, Yoshida, 1993, Fig. 2). At this time, another CYP was purified from yeast growing in high concentrations of glucose (20%w/v) by Wiseman, King and colleagues (King et al., 1984). This form was a benzo(a)pyrene hydroxylase and appears to have been the sterol 22-desaturase protein designated later as CYP61 (Kelly et al., 1997a,b). As sterol biosynthesis is generally an essential primary metabolic pathway of eukaryotes, the identification of similar enzymatic requirements in animals (Trzaskos et al., 1984) and plants (Kahn et al., 1996) were interesting with respect to an ancient ancestral form of CYP (Nelson et al., 1996). However, some had argued that convergent evolution might have given rise to different forms with the same activity. Also, some eukaryotic organisms such as nematodes and insects lack a sterol biosynthetic apparatus and obtain sterols from their diet. These organisms do not contain CYP51, but CYP51 has been found to be wide-spread in eukaryotes and, unlike other CYPs, is seen across the Kingdoms of Life. Figure 4 shows a
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Figure 3 The 14a-demethylation of sterol by CYP51 proceeding via three sequential monoxygenase reactions with the final release of formic acid.
phylogram of CYP51s from the different Kingdoms of Life. The absence of CYP51 is a novelty sometimes seen in microbial eukaryotes; for instance, G. lamblia lacks CYP51 (S.L. Kelly, unpublished). CYP51 has been identified in many microbes including bacteria, (Bellamine et al., 1999), but most sequences deposited are fungal forms, reflecting the importance of this CYP as the target of the azole antifungal compounds used in medicine and agriculture (Lamb et al., 1999a, 2002). It is accepted that bacteria constitute the domain of life where CYPs arose. One hypothesis is that CYPs arose to detoxify reactive oxygen intermediates within the early atmosphere on earth (Wickramashighe and Villee, 1975), although, if that CYP was a CYP51, squalene epoxidase that precedes CYP51 in the sterol pathway would already have developed a similar function. Most of the known prokaryotic CYPs participate in specialised aspects of secondary metabolism or carbon source utilisation and hence there is no direct evidence that a prokaryotic CYP has survived from the earliest era of evolution. However, analysis of CYPs from the completed genome of M. tuberculosis revealed a CYP with homology to eukaryotic CYP51s (33% sequence identity to human CYP51). Subsequently, sterol 14-demethylase activity was shown for the M. tuberculosis CYP51 gene product (Bellamine et al., 1999) and this protein has been crystallised (Podust et al., 2001). The presence of another, but less obvious,
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Figure 4 A phylogenetic tree of CYP51s was produced using clustal X 1.8 and TreeView 1.6.1. The sequences used were Candida albicans (P10613), Cunninghamella elegans (Q9UVC3), Mycobacterium tuberculosis (P77901), human (Q16850), Penicillium italicum (Q12664), pig (O46420), rat (Q64654), Schizosaccharomyces pombe (Q09736), Sorghum bicolor (P93846), Uncinula necator (O14442), S. cerevisiae (P10614), Penicillium digitatum (Q9P340), Aspergillus fumigatus (Q9P8R0), Phanerochaete chrysosporium, Cryptococcus neoformans (AAF35366), Venturia nashicola (CAC85409), Ustilago maydis (P49602), Botryotinia fuckeliana (AAF85983), Wheat (P93596), Candida glabrata (P50859), Candida tropicalis (P14263), Arabidopsis thaliana1 (AAK92797), Arabidopsis thaliana2, Issatchenkia orientalis (Q02315), Streptomyces coelicolor (NP_629370), Oryza sativa (BAA76438), Monilinia fructicola (AAL79180), Mouse (NP_064394), Tapesia acuformis (AAF18468), Mycosphaerella graminicola (AAF74756), Blumeria graminis (AAC97606), Tapesia yallundae (AAF18469), Dictyostelium discoideum, Trypanosoma brucei and Methylococcus capsulatus. Primary accession numbers have only been used for proteins deposited in the Swiss-Prot and TrEMBL databases.
CYP51-like protein was observed in S. coelicolor (Lamb et al., 2002b) and in the sterol-producing bacterium, Methylococcus capsulatus (Jackson et al., 2002). These forms are discussed later in this review. The possibility of horizontal gene transfer of CYP51 has been suggested, but is unlikely for organisms producing sterols by an unlinked set of chromosomal genes encoding the whole pathway. However, in those instances where the
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organism does not produce sterols, as seems to be the case in S. coelicolor, such a transfer can be envisaged.
1.3. Bacterial CYPs We discuss in this review some of the better-known CYPs in the bacterial and eukaryotic microbes, but many genome projects have also revealed new forms without any known function. The website of David Nelson (http:// drnelson.utmem.edu/nelsonhomepage.html) contains lists of the CYPs that have been given a formal name in the nomenclature, but many genes are not yet named under this system.
1.3.1. Cytochrome P450cam The most studied CYP to-date is P450cam (CYP101) from the bacterium Pseudomonas putida ATCC17453, begun through the pioneering work of Gunsalus and colleagues (Katagiri et al., 1968; Tyson et al., 1972; Rheinwald et al., 1973; Sligar et al., 1974). This work was important for the genetic interest in the role of CYP with the discovery of catabolic plasmids, as well as the biophysical studies that focused on CYP101 as the archetypal CYP. This enzyme catalyses the 5-exo hydroxylation of camphor, an initial step in the biodegradation of camphor to acetate and isobutyrate as a sole carbon source for energy (Gunsalus and Wagner, 1978). The CYP101 gene (camC) is located in an operon with genes for its electron donor proteins, namely putidaredoxin reductase (camA) and putidaredoxin (camB). The operon also contains camD, which encodes 5-exo hydroxy camphor dehydrogenase. These four genes, camDCAB, encode proteins essential for the early steps of camphor biodegradation and are controlled by camR, a repressor. The first CYP crystal structure was obtained in 1985 for CYP101 and at higher resolution in 1987 by Poulos and colleagues (Poulos et al., 1985, 1987). Subsequently, structures of CYP101 substrate- and inhibitor-bound complexes have been determined. The initial work with CYP101 revealed important features, which have subsequently been observed for other CYPs. Overall CYP topology is conserved with structures approximating a triangular prism. In the absence of substrate, water molecules occupy the active site with one molecule coordinated with the iron atom of the haem. The haem thiolate ligand via a cysteine residue was shown in this structure. The motif containing this cysteine is conserved and provides a reference for
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the identification of orphan CYPs present in sequenced genomes and for primary sequence alignments of CYPs. CYP101 also provided a unique opportunity to study the CYP catalytic cycle in extensive detail (Schlichting et al., 2000). Many other bacterial CYPs are implicated in the breakdown of foreign compounds to biomass and carbon dioxide (mineralisation) which require further enzymes downstream of the initial CYP metabolism. For example, P450TERP (CYP108A1, Peterson et al., 1992) metabolises terpineol and has also been crystallised and the structure determined (for review, Graham and Peterson, 1999). Mineralisation is valuable in bioremediation of pollutants such as that observed for a CYP from M. smegmatis involved in piperidine utilisation (Poupin et al., 1999). A Rhodococcus sp. has been observed to contain a CYP (CYP116, Nagy et al., 1995) that metabolises thiocarbamate herbicide and atrazine, while other CYP105s undertake the metabolism of pesticides without mineralisation (for example O’Keefe et al., 1994). Bacteria without a capability for complete breakdown of chemicals may still be useful within microbial populations that can break-down such chemicals co-operatively and where specific hydroxylated products are desired.
1.3.2. P450 BM-3 (CYP102A1), a Model for Eukaryotic CYPs During the early 1980s Fulco and colleagues identified a CYP in the bacterium Bacillus megaterium ATCC14581 (Ruettinger and Fulco, 1981; Narhi and Fulco, 1982, 1986) that was similar to eukaryotic CYPs, particularly of the fatty acid hydroxylase (CYP4A) family. This CYP, P450 BM-3 (CYP102A1), is a soluble enzyme, but, unlike other class I bacterial CYPs, it utilizes a eukaryotic type (class II) redox system: a FAD- and FMN-containing cytochrome P450 reductase. The BM-3 reductase domain is fused to the C-terminal of the CYP domain in a single polypeptide. Experiments probing the catalytic activity of this enzyme revealed that it hydroxylated a range of fatty acids with only substrate and NADPH required for activity (Narhi and Fulco, 1982, 1986). Furthermore, the catalytic turnover for this CYP is the highest for any CYP reported to date, which presumably is a reflection of efficient electron transfer for catalytic activity (Narhi and Fulco, 1987; Wen and Fulco, 1987). A rate of 1932 nmol/min/nmol P450 protein for myristate oxygenation was observed compared with many other CYPs that exhibit rates with their substrates closer to 1 nmol/min/nmol P450. However, the endogenous role of this enzyme within B. megaterium remains to be discovered. The resolved atomic structure of CYP102A1 revealed further insight into CYP structure/function
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(Ravichandran et al., 1993). In contrast to CYP101, CYP102, although again adopting the triangular prism shape, had a long access channel leading to the haem where fatty acid substrates were bound (Li and Poulos, 1997). Attempts to determine the structure of the full length CYP-reductase complex have so far failed although the crystal structure of the complex between haem- and FMN-binding domains have been determined (Sevrioukova et al., 1999). Detailed site-directed mutagenesis studies with CYP102A1, have revealed many key residues involved in CYP architecture and function. For instance, mutagenesis of Thr268, a residue conserved in the majority of CYPs, has established its function in oxygen activation and electron transport (Yeom et al., 1995) and Tyr97 has been found to play a role in association with the haem prosthetic group (Munro, 1994). Within the reductase domain, residues important for FMN binding have also been determined (Klein and Fulco, 1993). Recently another bacterial CYP has been discovered, P450CIN (CYP176A1) from Citrobacter brakii that undertakes cineol metabolism (Hawkes et al., 2002). This form of CYP was encoded by an operon with genes for a flavodoxin and flavodoxin reductase rather than the typical class I system of bacteria using ferredoxin/ferredoxin reductase. As such, it is interesting with regard to both CYP102A1 and eukaryotic forms requiring the FMN/FAD containing CPR, as these types of reductases may have resulted from the fusion of two proteins similar to those driving P450CIN.
1.3.3. Bacterial Genomes Reveal a Plethora of CYP Genes As mentioned earlier, it is possible that CYPs evolved to detoxify oxygen in the early atmosphere (Wickramashighe and Villee, 1975) or had roles as electron transfer proteins. Subsequently, CYPs may have been harnessed for the generation of metabolic energy, being involved in the initial degradative attack of various carbon sources. Exposure of the organisms to a wide variety of hydrocarbon xenobiotics presumably selected CYPs through evolution to provide a defensive mechanism for the initial metabolism and subsequent elimination of such compounds. Examination of the many prokaryotic genome sequences and databases dedicated to CYP identification has revealed that the distribution of CYP is extensive (Table 1). Many bacterial genomes such as E. coli contain no CYPs, although actinomycetes appear to be particularly rich in CYPs, many of which have been found to be associated with secondary metabolism. Even among actinomycetes, Corynebacterium diptheriae contains no CYPs and of course other types of
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STEVEN L. KELLY et al. Table 1 The numbers of CYPs in various completed bacterial genomes; many such as E. coli, contain no CYPs. Organism
CYP complement
Campylobacter jejuni Bacillus halodurans
1 1
Methanosarcinia barkeri Halobacterium species NRC1
1 1
Sulfolobus tokodaii
1
Sinorhizobium meliloti Agrobacterium tumafaciens
2 2
Pseudomonas aeruginosa Deinococcus radiodurans Bacillus subtilis
3 3 8
Streptomyces coelicolor Mycobacterium tuberculosis Streptomyces avermitilis
18 20 33
Mycobacterium smegmatis
40
bacteria also contain CYPs involved in secondary metabolism outside of the actinomycetes. For example, CYPs plays an important role in the synthesis of the anti-cancer drug epothilone by Sorangium cellulosum (Tang et al., 2000).
1.3.4. Actinomycetes: Biodiversity of CYPs in Streptomyces Coelicolor and Mycobacterium tuberculosis The actinomycete bacteria contain a diverse range of organisms with importance as pathogens, as producers of antibiotics, and in biocatalysis/ bioremediation as well as in other areas of biotechnology. The genome projects reveal an extensive pool of CYPs in these bacteria, beginning with M. tuberculosis which contains 20 CYPs (Cole et al., 1998). The closely related Mycobacterium bovis BCG contained 18 CYPs, but Mycobacterium leprae has lost many CYPs in the course of evolution with degenerating CYPs evident in its genome (Cole et al., 2001). Other actinomycetes appear to have no CYPs while in some mycobacteria such as M. smegmatis 1% of genes encode CYP (we have observed 40 CYPs in this genome sequenced at TIGR). Streptomycete bacteria are important producers of medicinal and veterinary antibiotics. The genome of the model species, S. coelicolor, has been completed recently (Bentley et al., 2002). We detected 18
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CYPs in this genome (Lamb et al., 2002a) and a number have also been reported from a partial sequence of the industrially important Streptomyces avermitilis that produces the anti-helminthic compound avermectin (Omura et al., 2001). The endogenous role of the majority of these CYPs remains to be elucidated, but clues to function are appearing. Many appear in regions and operons associated with antibiotic biosynthesis. These are found also in other bacteria such as the actinomycete Amycolatopsis mediterranei where the chromosomal region associated with rifamycin biosynthesis contains five CYPs (August et al., 1998).
1.4. CYP superfamily in Streptomycetes Streptomycetes produce a vast array of antibiotics applied in human and veterinary medicine and agriculture, as well as anti-parasitic agents, herbicides and pharmacologically active metabolites (e.g. immunosuppressants). Streptomycetes also catalyse numerous transformations of xenobiotics of industrial and environmental importance (Hopwood, 1999; Kieser et al., 2000). These oxidative transformations have been observed with alkaloids (Sariaslani et al., 1984), coumarins (Sariaslani and Rosazza, 1983a), retinoids (Sariaslani and Rosazza, 1983b) and other complex xenobiotics (Taylor et al., 1999). The most significant of these biocatalytic reactions include aromatic and aliphatic hydroxylations, O- and Ndealkylations, N-oxidation and C–C fission. Available experimental evidence has indicated that all of these oxidative reactions are catalysed by CYPs. Application has been exploited in the synthesis of pravastatin, utilising a streptomycete biotransformation step (Serizawa and Matsuoka, 1991), as well as in 16-hydroxylation of steroids (Berrie et al., 1999) and in the preparation of drug metabolites for toxicological evaluation (Cannell et al., 1995).
1.4.1. P450 EryF, (CYP107A1), a CYP Involved in Antibiotic Biosynthesis CYP107A1, isolated from Saccharopolyspora erythraea, catalyses the hydroxylation of 6-deoxyerythronolide B to generate erythronolide B (Fig. 5). This is the precursor of the erythromycin antibiotic that this bacterium produces (Andersen and Hutchinson, 1992). Although this organism was classified as distinct from the streptomycetes, this CYP from Saccharopolyspora is included here for convenience. This form, named CYP107A1, was the fourth CYP whose structure was determined (Cupp-Vickery and
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Figure 5 The biosynthetic step mediated by CYP107A1 (eryF) in erythromycin biosynthesis.
Poulos, 1995) and again shared common structural features of all CYPs having despite S (Favre et al., 1999), 467R>K (White, 1997; Lamb et al., 2000) and 471I>T (Kakeya et al., 2000) have been implicated in resistance. Many of the altered residues are present in other CYPs in these positions and so seem to contribute subtly to the CYP catalytic active site. The fluconazole resistance mutation Y132H is reported from many clinical sources (Sanglard et al., 1998) and has been studied at the level of heterologously-expressed microsomal protein from S. cerevisiae where the typical interaction reflecting binding of the fluconazole to low-spin CYP51 was not observed (Lamb et al., 1999c). However, using purified protein, normal spectra indicating co-ordination of fluconazole to the haem as a sixth ligand have been observed (Tyrrell J. and Kelly S.L., unpublished). Mutation at an equivalent residue has been associated with azole resistance in mildews (Delye et al., 1997). The C. albicans Y132H CYP51 protein exhibited an increased IC50 for fluconazole again proving the resistance phenotype of this mutation was through reduced affinity for the drug. Generally, the high affinity of drug for wild-type protein was reflected by an IC50 of half the concentration of CYP51 present in the assay (e.g. 0.5 nmol needed for 1 nmol CYP51), while resistant proteins require up to ten-fold more drug for this effect. The rationale for the effect of this mutation in the protein is by no means clear, but could be associated with access of the drug into the active site. Whether resistance can produce forms with greater resistance will only become clear when more is known about the proteins, all of which require characterisation. This will contribute to information regarding overcoming resistance, coupled with the pattern of crossresistance to other azoles.
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The overall picture developing for CYP51 residues implicated in azole resistance is one of a very diverse range of mutations, possibly restricted to the region around amino acids 125–150, 405 and 448–488 (around the conserved cysteine at 470). The effects are presumably often long range rather than in residues directly interacting with the drug. Many of the changes are difficult to rationalise via molecular modelling. The earliest modelling was undertaken in Professor W.G. Richard’s laboratory in Oxford requiring supercomputer capability (Boscott and Grant, 1994). The model explained the differential activity of stereoisomers of azole drugs and predicted fluconazole bound to CYP51 via an aromatic interaction between F233 or F235 and the drug (Lamb et al., 1997b). This region of the F-G loop has been recognised as important for substrate binding in CYPs (Cosme and Johnson, 2000). Subsequent models of C. albicans CYP51 have utilised the additional information provided by further bacterial CYP structures. One was made and rationalised some of the fluconazole resistance mutations observed (Ji et al., 2000). Besides rationalizing known information, this model interestingly predicted a kink in the major I-helix seen in all CYPs and, when the structure of soluble CYP51 from M. tuberculosis was resolved, this showed the I-helix was in two parts (Podust et al., 2001). The active site of this CYP51 was open to the protein surface; it is timely to model the C. albicans protein using the known M. tuberculosis CYP51. However, with less than 40% amino acid identity, the accuracy of even these models for the C. albicans CYP51 will remain questionable. Recently, the first microsomal eukaryote CYP structure was determined after production of a soluble version lacking a N-terminal membrane anchor and using E. coli as a host (Williams et al., 2000 a,b). A similar route may be successful in finally obtaining a structure of the fungal target of azole antifungals. We have already made a soluble derivative of the C. albicans CYP51 by introducing a protease site behind the N-terminal membrane anchor (Lamb et al., 1999d) and using yeast for expression. A C. albicans CYP51 structure will ultimately resolve the basis of the cellular phenotypes observed that are associated with clinical fluconazole-resistant CYP51 mutations.
1.10. Future Perspectives for Microbial CYP Research From the previous sections, it is clear that study of microbial CYPs is an area of diverse academic and applied interest. At the centre of strategies of biochemical deterrence and attraction, the roles of CYPs in nature provide
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two sides to the warfare that has driven the evolution of this biodiversity. They participated in making more diverse toxic metabolites as well as ways of detoxifying them. Many of the metabolites are current or future medicines and further molecular diversity can be achieved using CYPs as biocatalysts in biotransformations. These will be of value in industries concerned with pharmaceutical and agricultural production, fine chemicals, flavours, fragrances, pigments, biosensors and more. Of course, much can be achieved by using the current range of enzymes, but many constraints to activity exist, particularly the low turnover of substrate in many of the reactions. The power of directed evolution has been focused on CYP by a number of laboratories including altering the reductive method for CYPs using CYP101 (Joo et al., 1999), creating an indigo producing derivative of CYP102 (Li et al., 2000), as well as changing CYP102A1 fatty acid specificity (Lentz et al., 2001). Site-directed studies will also have value, for example, in altering CYP101 to metabolise unnatural substrates (Bell et al., 2001). However, it appears clear that the directed evolution approach is the stronger method for studies where improved phenotype is desired. Often, the improvements will be due to long-range effects on the CYP active site and give similar difficulties for rationalisation as with the azole resistance effects with CYP51. As well as biotransformation or bioremediation strategies for use with CYPs, the area of metabolic pathway engineering will also feature strongly in the future. By far the most impressive example of these techniques so far is the re-engineering of yeast ergosterol biosynthesis to steroid production (Duport et al., 1998). In this, CYPs played an important role as well as other reactions of sterol metabolism. For instance, the removal of CYP61 was required and the introduction of heterologous enzymes, including CYPs, for co-ordinated expression under conditions appropriate for industrial fermentations. It is clear that in future many pathways for existing and new drugs will involve manipulation of CYPs to influence the end-product and efficiency of metabolic pathways. Of course, microbes, mainly E. coli and yeast, still remain an important source of heterologously expressed CYP reagents for toxico- and pharmacological studies and are needed for information profiles used in drug safety registration. They are used in academia and in the drug industry to predict metabolic fates of chemicals and to harvest metabolites for toxicological evaluation. What is clear is that, with the extraordinary diversity of CYPs revealed in recent years by studies on metabolic pathways, and from genome sequence projects, experimental analysis of CYPs and CYPomes represents a considerable challenge in the coming decades of the twenty-first century.
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ACKNOWLEDGEMENTS Research in the Wolfson Laboratory of P450 Biodiversity is currently supported by the Biotechnology and Biological Research Council, the Natural Environment Research Council, The Wellcome Trust, The Wolfson Foundation and the University of Wales Aberystwyth.
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The Tat Protein Translocation Pathway and its Role in Microbial Physiology Ben C. Berks1, Tracy Palmer2 and Frank Sargent3 1
Department of Biochemistry, University of Oxford, South Parks Road, Oxford OX1 3QU, UK 2 Department of Molecular Microbiology, John Innes Centre, Norwich NR4 7UH, UK 3 Centre for Metalloprotein Spectroscopy and Biology, School of Biological Sciences, University of East Anglia, Norwich NR4 7TJ, UK
ABSTRACT The Tat (twin arginine translocation) protein transport system functions to export folded protein substrates across the bacterial cytoplasmic membrane and to insert certain integral membrane proteins into that membrane. It is entirely distinct from the Sec pathway. Here, we describe our current knowledge of the molecular features of the Tat transport system. In addition, we discuss the roles that the Tat pathway plays in the bacterial cell, paying particular attention to the involvement of the Tat pathway in the biogenesis of cofactor-containing proteins, in cell wall biosynthesis and in bacterial pathogenicity.
ADVANCES IN MICROBIAL PHYSIOLOGY VOL. 47 Copyright ß 2003 Elsevier Science Ltd ISBN 0-12-027747-6 All rights of reproduction in any form reserved
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Abbreviations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 188 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 188 1.1. Evidence for the translocation of folded proteins by the Tat pathway . . . . . . 191 2. Journey to the translocase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192 2.1. Mechanisms to avoid mis-targeting of Tat substrates . . . . . . . . . . . . . . . . . 192 2.2. Routing to the Tat pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195 2.3. The diversity of cofactor-containing Tat substrates and their preparation for transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 2.4. Exported proteins without cofactors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 215 3. Transport across the membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219 3.1. Components and organisation of the Tat transport apparatus. . . . . . . . . . . . 219 3.2. The Tat transport cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222 3.3. The TatA/B protein family. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 224 3.4. The TatC protein family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 230 3.5. Mechanism of the Tat transporter: some considerations. . . . . . . . . . . . . . . . 232 4. Biosynthesis of integral membrane proteins by the Tat system . . . . . . . . . . . . . . 236 5. Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 239 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 239 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 240
ABBREVIATIONS MPT MGD SQR GFOR TTQ
molybdopterin molybdopterin guanine dinucleotide sulphide-quinone oxidoreductase glucose-fructose oxidoreductase tryptophyl tryptophanquinone
1. INTRODUCTION A significant proportion of bacterial proteins function in extracytoplasmic compartments. Specific targeting and transport mechanisms are therefore required to move these proteins from their site of synthesis in the cytoplasm to their final subcellular destination. The initial step in the sorting pathway of extracytoplasmic proteins is transport across, or alternatively insertion into, the cytoplasmic membrane. For a long time, it was assumed that a single mechanism, the ‘Sec’ pathway, carried out these functions. However, it has recently become apparent that most bacteria possess a second, completely distinct, general export pathway that operates in parallel to the Sec system. It is this ‘Tat’ apparatus that is the subject of this review.
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In order to understand the unique role that the Tat system plays in bacterial protein transport it is first necessary to appreciate how its mode of operation differs from that of the Sec pathway. In the Sec mechanism (Pugsley, 1993; Manting and Driessen, 2000), the substrate protein is presented to the Sec translocase (transporter) in an extended conformation. The substrate protein is then threaded, amino terminus first, through the translocase and only folds when it reaches the aqueous compartment on the far side of the membrane (for simplicity we shall refer to this trans compartment as the periplasm in this review even though the Sec and Tat systems are also found in Gram-positive organisms). The core of the Sec translocase is probably composed of two copies of a SecYEG heterotrimer (Breyton et al., 2002) with the SecY and SecE proteins being homologous to the Sec61a and Sec61g components of the secretory apparatus found in the endoplasmic reticulum of eukaryotic cells. Water-soluble substrates of the Sec system are synthesised with amino-terminal signal peptides and are transported post-translationally. The signal peptide is recognised and bound by the ATPase SecA while various chaperones bind to the mature portion of the protein to maintain an unfolded configuration. Precursor-bound SecA is able to bind to the SecYEG translocase. Concomitant with an ATP hydrolysis cycle, SecA inserts the amino terminal region of the precursor into the translocase in a loop configuration. The first arm of the loop comprises the signal peptide. This is bound to a specific site in the translocase in such a way that the amino terminus of the signal peptide is at the cytoplasmic side of the membrane. The second arm of the loop is the amino terminus of the mature region of the protein. The junction between the signal peptide and the mature protein is positioned at the apex of the loop and is proteolytically processed by signal peptidases located at the periplasmic side of the membrane. Transport of the substrate protein is effected by threading the mature protein half of the loop through the translocase. In vitro this can be achieved by additional cycles of SecA insertion but in vivo is likely to be driven primarily, if not exclusively, by the transmembrane proton electrochemical gradient (p). Insertion of integral membrane proteins into the cytoplasmic membrane normally also involves the SecYEG translocase (de Gier and Luirink, 2001). In most cases, membrane proteins do not have a cleavable signal peptide but are instead recognised by the extended hydrophobic regions that will become the transmembrane helices in the mature protein. As these hydrophobic sequences emerge from the ribosome exit channel, they are bound by signal recognition particle (SRP) which, in concert with other components, directs the nascent chain-ribosome complex to the SecYEG translocase. Insertion of the substrate protein into the translocase is
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therefore co-translational and ensures that the highly hydrophobic transmembrane helices can partition into the bilayer before they aggregate in an aqueous phase. Recent research shows that the transmembrane helices move from the translocase into the membrane via an additional membrane protein termed YidC (Samuelson et al., 2000; Scotti et al., 2000). Indeed, for some integral membrane proteins, it has been found that the SRP pathway targets the protein directly to YidC without the protein passing through the SecYEG apparatus. In addition to its role in the biosynthesis of integral membrane proteins, it appears that SRP is involved in targeting at least some water-soluble proteins to the SecYEG translocase (e.g. Sijbrandi et al., 2002). Despite the complexity of inputs to the SecYEG translocase, all substrate proteins are ultimately transported in an unfolded state. In contrast, the Tat pathway catalyses the movement of folded protein substrates across the membrane. Other examples of protein transport systems that move folded protein substrates are known including those associated with peroxisomal import, type II secretion across the outer membrane of Gram-negative bacteria, as well as transport through the nuclear pores of eukaryotic cells. However, the Tat system is the only general protein transport pathway that moves folded proteins across a coupling membrane (one that sustains a proton- or ion-motive transmembrane electrochemical gradient). The Tat system thus faces the formidable challenge of transporting a range of folded proteins while maintaining the permeability barrier of the membrane to ions, especially protons. The physiological role of the bacterial Tat pathway is to allow the cell to complete the maturation of periplasmic proteins in the cytoplasm before they are exported. Now it turns out that the majority of Tat substrates are proteins containing cofactor molecules (indeed, it was the interest in the biosynthesis of such proteins that led to the discovery of the bacterial Tat system) and this association arises because stable insertion of cofactors into proteins normally requires the protein to fold around the cofactor. Thus, if a protein is to pick up a cofactor in the cytoplasm prior to transport, it must use the Tat rather than Sec pathway. Proteins are specifically targeted to the Tat system by cleavable aminoterminal signal peptides. The features of Tat signal peptides are considered in more detail below (Sections 2.1 and 2.2). However, it is pertinent to mention at this point that Tat signal peptides contain a pair of consecutive and essentially invariant arginine residues and that these provide the basis for the Tat designation; indeed, Tat stands for twin arginine translocation (Sargent et al., 1998a). An alternative Mtt (membrane transport and targeting) designation has also been employed (Weiner et al., 1998).
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Tat transport is an active transport process with the driving force for transport deriving from the transmembrane p (Mould and Robinson, 1991; Musser and Theg, 2000a; Yahr and Wickner, 2001; Alami et al., 2002). The Tat system is not restricted to bacteria but is also found in certain eukaryotic cells. Plant chloroplasts possess a Tat apparatus (formerly styled the ‘pH-dependent pathway’) that is used for protein uptake into the thylakoids (reviewed in Mori and Cline, 2001). All substrates of the chloroplast Tat system are nuclear-encoded proteins that must first be transported from the cytoplasm into the chloroplast stroma before they can be imported into the thylakoids. The mitochondria of plants and algae probably also operate some sort of Tat-like pathway since their mitochondrial genomes encode homologues of the essential TatC component of the bacterial Tat system (Bogsch et al., 1998). No Tat homologues have, however, been found in animals or yeasts. Our purpose here is to review current knowledge of the bacterial Tat pathway, paying particular attention to the molecular features of the system and to the role of the Tat pathway in bacterial cellular processes. Although the plant chloroplast Tat system is not covered explicitly, important studies of direct relevance to bacterial Tat transport are described. Previous reviews that contain more detailed or complementary discussion of aspects of Tat transport include Berks (1996), Voordouw (2000), Berks et al. (2000a,b), Mori and Cline (2001), Sargent et al. (2002), Yen et al. (2002).
1.1. Evidence for the Translocation of Folded Proteins by the Tat Pathway The evidence that the Tat system translocates folded proteins has been considered in some detail elsewhere (Berks, 1996; Berks et al., 2000a). The major arguments can be summarised as follows. Firstly, it can be inferred that cofactors are inserted into Tat substrates in the cytoplasm. If cofactor insertion is blocked, the precursor protein is not transported (Santini et al., 1998) and, if Tat transport is blocked, then the substrate protein that accumulates in the cytoplasm contains cofactor (e.g. Sargent et al., 1998a; Hilton et al., 1999; Temple and Rajagopalan, 2000). Since stable cofactor insertion requires that the protein has folded around the cofactor, then the substrate protein must be folded prior to transport. It is also notable that cytoplasmic chaperones are required for cofactor insertion into some Tat substrates (Section 2.3). Secondly, it is quite common for a protein without a signal peptide to be transported via the Tat system as a complex with a specific signal peptide-bearing partner protein (e.g. Rodrigue et al., 1999).
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It is difficult to see how these proteins could be recognising each other if they are not folded. Finally, the Tat system has been shown to be capable of translocating heterologous substrates that are known to be tightly folded prior to transport. These include jellyfish green fluorescent protein GFP (Santini et al., 2001; Thomas et al., 2001) and c-type cytochromes (Sanders et al., 2001) in studies with bacteria and methotrexate-bound dihydrofolate reductase (Hynds et al., 1998) or disulphide bond-stabilised bovine pancreatic trypsin inhibitor (Clark and Theg, 1997) with the plant thylakoid Tat system.
2. JOURNEY TO THE TRANSLOCASE The Tat system is a post-translational pathway and the journey of a substrate protein from ribosome to translocase is both eventful and carefully controlled (Fig. 1).
2.1. Mechanisms to Avoid Mis-targeting of Tat Substrates The basic structure of a Tat signal peptide is very similar to that of the signal peptides employed by the post-translational SecA pathway and by some substrates of the co-translational SRP pathway. In all three cases, a basic amino terminal (n-) region is followed successively by a hydrophobic (h-) region and a second polar (c-) region (Fig. 2). How, then, does a newly synthesised Tat precursor protein avoid its signal peptide being recognised by the SecA or SRP pathways? Two possibilities have been suggested. The first proposal is that basic residues in the c-region of Tat signal peptides act as a ‘Sec-avoidance’ motif (Bogsch et al., 1997). Such positively charged residues are very rare in Sec signal peptides (von Heijne, 1986) and impede protein transport when experimentally introduced into a Sec signal peptide (e.g. Geller et al., 1993). Nevertheless, c-region basic residues cannot be the sole mechanism by which Tat signal peptides avoid mis-targeting simply because they are not present in many such signal peptides (Berks, 1996; Peltier et al., 2000). The second proposal is based on the observation that there are differences in the degree of hydrophobicity of the h-region of the signal peptides used by the three protein transport systems with Tat signal peptides having the least, and SRP targeting regions the highest, h-region hydrophobicity (Cristo´bal et al., 1999). This has led to the suggestion that Tat substrates avoid misrouting because their signal peptides are
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Figure 1 Model for the cytoplasmic events in the biogenesis of Tat pathway substrates. The Tat signal peptide is found at the amino terminus of the nascent polypeptide chain. Mis-targeting of the precursor to alternative protein transport pathways is avoided because the Tat signal peptide does not interact efficiently with either the signal recognition particle (SRP) or SecA, possibly because of the weak hydrophobicity of the Tat signal peptide h-region. The Tat precursor undergoes folding in the cytoplasm prior to transport. Transport of unfolded or poorly folded substrates is prevented by an unknown mechanism. In the figure this task is speculatively assigned to the translocase itself. Some Tat substrates bind cofactor molecules that are inserted into the protein in the cytoplasm concomitant with protein folding. Often cofactor insertion involves dedicated chaperone proteins. It is thought that in some cases these assembly chaperones prevent the protein being exported before cofactor insertion is complete. A possible mechanism by which this may occur is depicted in the figure. The chaperone is shown binding to both the partially folded mature region of the apo-precursor and the signal peptide. The signal peptide is thus sequestered and cannot direct export. Upon cofactor insertion, the chaperone is released from the precursor and the signal peptide becomes available for protein targeting. The Tat pathway recognises its substrates proteins via the twin-arginine consensus sequence of the signal peptide. It is certain that the Tat translocase itself contains a specific Tat signal peptide binding site(s). However, in contrast to the SRP and SecA pathways, there is currently no evidence for an additional, universal, cytoplasmic targeting factor that recognises Tat precursors.
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Figure 2 Comparison of consensus features of Tat and Sec signal peptides. This figure is based on the analysis reported in Berks (1996) and Cristo´bal et al. (1999). Since the consensus Sec signal peptide shown is based on processed amino terminal signal peptides, it is assumed that this represents the type of peptide that is initially bound by SecA rather than by SRP. Each peptide comprises three domains: a polar amino-terminal domain (n-region), a hydrophobic central domain (h-region) and a polar carboxyterminal domain (c-region). The lengths of the signal peptides and of the signal peptide domains are drawn to scale. Tat signal peptides are on average 37 residues in length with a n-region of 17 amino acids, a h-region of 12 amino acids and a c-region of 8 amino acids. The corresponding figures for Sec signal peptides are a total average length of 23 residues with a n-region of 6 amino acids, a h-region of 11 amino acids and a c-region of 6 amino acids. The n-region of Tat signal peptides contains the consensus S-R-R-x-F-L-K motif in which the arginine residues are close to invariant, x is a polar amino acid or glycine, and the other residues are found at frequencies exceeding 50%. The n-regions of both Tat and Sec signal peptides are basic (designated þ in the figure) with average charges of þ2.8 for Tat and þ1.7 for Sec signal peptides assuming an amino terminal formyl group. The c-region of Tat signal peptides is most often basic, sometimes is uncharged, but is almost never net acidic (designated þ/O in the figure) with an average charge of þ0.5. In contrast the c-region of Sec signal peptides is rarely charged (designated þ/O in the figure) with an average charge of þ0.03. The h-region of Sec signal peptides is significantly less polar than that of Tat signal peptides. The major reason for this difference is that the h-region of Tat signal peptide is significantly enriched in glycine and threonine residues, and deficient in leucine residues, relative to the h-region of a Sec signal peptide. An open arrow represents the site of signal peptide cleavage by the type I signal peptidase (Lep) (Yahr and Wickner, 2001) for which the primary recognition site is the presence of small side chain amino acids, especially alanine residues, at positions 3 and 1 relative to the site of processing (designated as A-x-A1 in the figure). Tat signal peptides have a high frequency of proline residues at position 6 (designated P6 in the figure).
insufficiently hydrophobic to be recognised by SecA or SRP. In support of this contention, Cristo´bal and co-workers have shown that it is possible to route a Tat substrate away from the Tat pathway by increasing the hydrophobicity of the signal peptide h-region (Cristo´bal et al., 1999). Comparable levels of h-region hydrophobicity have, however, also been
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found to reroute SecA substrates onto the SRP pathway (Valent et al., 1997; Lee and Bernstein, 2001) and it is therefore unclear from this experiment whether differences in signal peptide hydrophobicity are sufficient for the cell to distinguish Tat and SecA-targeted substrates. In conclusion, while both the c-region charge and h-region hydrophobicity are attractive hypotheses to explain discrimination against Tat signal peptides, further experimental testing is required to fully establish their contributions to the prevention of precursor mis-targeting. Nevertheless, it is clear that SecA is able to discriminate against Tat substrates since the ATPase activity of SecA is only weakly stimulated by Tat signal peptides (Kebir and Kendall, 2002). The recently determined structures of SecA and the signal-peptide binding portions of SRP (Keenan et al., 1998; Freymann et al., 1999; Cleverley and Gierasch, 2002; Hunt et al., 2002) hold out the promise that the structural basis of signal peptide discrimination by these proteins will soon be defined.
2.2. Routing to the Tat Pathway Having escaped targeting to alternative transport pathways, the Tat substrate needs to display the targeting information that will cause it to be recognised by the Tat system. This targeting information comprises a consensus amino acid sequence presented in an appropriate signal peptide context (Fig. 2). In Proteobacteria this consensus Tat box is best defined as Ser-Arg-Arg-Xaa-Phe-Leu-Lys where the arginine residues are almost invariant, the other amino acids are found at a frequency in excess of 50% and Xaa is a polar amino acid (Berks, 1996; Stanley et al., 2000). The corresponding sequence for plant chloroplast Tat substrates is Arg-ArgXaa-Hyd-Leu/Met where Hyd is a hydrophobic amino acid (Peltier et al., 2000). In both cases, this Tat or twin-arginine consensus motif is located at the amino-terminal side of the n-region/h-region boundary (Fig. 2). Several site directed mutagenesis studies have emphasised the importance of the arginine pair for bacterial Tat transport (Leu et al., 1992; Dreusch et al., 1997; Cristo´bal et al., 1999; Gross et al., 1999; Halbig et al., 1999; Stanley et al., 2000; Buchanan et al., 2001a; DeLisa et al., 2002; Ize et al., 2002; Rose et al., 2002). Substitution of either arginine normally blocks transport. However, the conservative substitution of a single arginine residue with lysine is sometimes tolerated though usually with a decrease in rate of protein transport (Halbig et al., 1999; Stanley et al., 2000; Buchanan et al., 2001a; DeLisa et al., 2002; Ize et al., 2002). A recent saturation mutagenesis study of the arginine pair has demonstrated that single glutamine or asparagine substitutions can also permit efficient Tat transport
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(DeLisa et al., 2002). It is important to note that while experimentally it has proved possible to change a single arginine residue while retaining some Tat targeting capability such substitutions are exceedingly rare in native Tat substrates (or at least that subset of such proteins that can be reliably predicted to be Tat substrates on the basis of bound cofactor). To our knowledge the only experimentally tested, and indeed only suspected, examples in bacteria are the lysine-arginine motif found in the TtrB subunit of the tetrathionate reductase of Enterobacteriaceae (Hinsley et al., 2001) and the very unusual signal peptide of Escherichia coli penicillin amidase where the substitution is probably a glutamine for an arginine (Ignatova et al., 2002). An additional example is found in plant chloroplasts where the Rieske protein of the cytochrome b6 f complex has a lysine-arginine pair (Hinsley et al., 2001; Molik et al., 2001). There appears, therefore, to be strong selective pressure to maintain an arginine pair even though certain other residues are mechanistically acceptable. No successful substitution of both arginine residues of the Tat motif with lysines has ever been reported. Since such conservative substitutions do not affect targeting of a SecA substrate (Sasaki et al., 1990; Cristo´bal et al., 1999), this particular amino acid change has been widely exploited as an experimental test to determine whether a precursor is a Tat substrate. The high sequence conservation across the entire Tat consensus sequence suggests that the whole Tat box, and not just the arginine residues, contributes to the Tat recognition signal. Nevertheless, the role of the nonarginine residues has been the subject of only one investigation. Site directed mutagenesis carried out on the Tat consensus motif of a single Tat substrate indicates that the consensus phenylalanine, and to a lesser extent the consensus leucine, are important for Tat targeting and that, for the phenylalanine position, a high hydrophobicity is probably the important chemical parameter (Stanley et al., 2000). The Tat signal peptide of the E. coli enzyme trimethylamine N-oxide (TMAO) reductase (TorA) has been used extensively to route heterologous substrates onto the E. coli Tat pathway (Sargent et al., 1998a; Cristo´bal et al., 1999; Santini et al., 2001; Thomas et al., 2001; Delisa et al., 2002; Ize et al., 2002), confirming that the signal peptide contains all the targeting information required for Tat export. Surprisingly, however, it has been found that the effects of site-directed mutagenesis within the TorA Tat consensus box differ when the mutations are performed on the chimeric precursors rather than on the native TorA precursor. For example, a variant ‘Arg-Lys’ signal peptide was unable to direct export of its native TorA passenger (Buchanan et al., 2001a) but still allowed export of both GFP (Delisa et al., 2002) and Colicin V (Ize et al., 2002) fusions through the Tat
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system. These results indicate that the passenger protein, while not essential for targeting, nevertheless appears to influence the transport events.
2.3. The Diversity of Cofactor-containing Tat Substrates and Their Preparation for Transport The major role of the Tat pathway is to transport cofactor-containing proteins. These proteins are normally involved in either respiratory or photosynthetic electron transfer chains and so the operation of the Tat pathway underlies metabolic energy generation in many bacteria. For each type of cofactor that is found in Tat substrates, the cell has evolved a specific biosynthetic pathway together with mechanisms for inserting the cofactor into the protein partner. These operations must be carried out before the transport step and this means that Tat targeting has a degree of complexity that is not found in the Sec system. In this Section we survey what is known about cofactor insertion into each class of Tat substrate and examine how these maturation events interact, and are integrated, with the Tat transport cycle.
2.3.1. Proteins with Molybdopterin Cofactors Molybdenum-containing enzymes are ubiquitous in bacteria where they catalyse a diverse range of redox reactions including those essential for the global carbon, nitrogen, and sulphur cycles. With the exception of the cytoplasmic enzyme dinitrogenase, molybdenum in enzymes is found co-ordinated to the dithioline group of an organic tricyclic pyranopterin cofactor which in its most simple form is termed molybdopterin (MPT). The biosynthesis of MPT occurs by an evolutionarily conserved multistep cytoplasmic pathway. In E. coli, MPT synthesis involves the moaABCDE operon, the moeAB operon, the modEABCD locus, the mobAB operon, and the mogA gene. The first step in the biosynthesis of the pyranopterin cofactor involves the rearrangement of GTP by the cytoplasmic MoaAC complex to form the sulphur-less pyranopterin ‘Precursor Z’. MoaA contains a redox-active [3Fe-4S] cluster (Menendez et al., 1996; Solomon et al., 1999) while its MoaC partner subunit displays a ferredoxin-like structure but contains no known cofactors (Wuebbens et al., 2000). The sulphur atoms of the dithiolene group of MPT originate from cytoplasmic cysteine molecules. In an elaborate process that has parallels in the ubiquitin cycle, the carboxy-terminal glycine-glycine
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dipeptide of MoaD is first adenylated by the MoeB ATPase (Lake et al., 2001). The adenylated carboxy-terminus of MoaD is then attacked by an uncharacterised protein-persulphide to give thiocarboxylated MoaD (Lake et al., 2001). Next, the thiocarboxylated MoaD interacts with the MoaE protein to form active MPT synthase which finally proceeds to transfer the carboxy-terminal thiol group from MoaD onto Precursor Z thus completing the tricyclic pyranopterin unit (Rudolph et al., 2001). Lastly, the cofactor must be combined with molybdenum. Molybdenum is present in the environment as the molybdate oxyanion and transport into the cell cytoplasm is accomplished by a highly conserved member of the ABC transporter superfamily (Walkenhorst et al., 1995). Following uptake, molybdate is probably first converted to thiomolybdate by the MoeA protein (Hasona et al., 1998) before the MPT-binding domain of MoeA and its homologue MogA (and/or MoaB in E. coli) together synthesise the basic Mo-containing molybdopterin cofactor, MPT (Liu et al., 2000; Xiang et al., 2001). Various cytoplasmic bacterial molybdoenzymes utilise MPT itself for activity, for example the xanthine dehydrogenase of Rhodobacter capsulatus (Truglio et al., 2002). However the majority of bacterial molybdoenzymes, including all known periplasmic proteins, utilize variant forms of MPT modified by addition of mononucleotides. The most common modification of MPT involves the attachment of GMP to form molybdopterin guanine dinucleotide (MGD). Two copies of MGD are then used to co-ordinate a single molybdenum atom to produce the final bis(MGD)Mo cofactor (often contracted to ‘MGD cofactor’). In E. coli, MGD formation is catalysed by the cytoplasmic protein MobA encoded by the mobAB operon (Lake et al., 2000; Stevenson et al., 2000). MobA physically interacts with the MoeB/ MogA MPT-binding proteins (Magalon et al., 2002) and thus MGD is synthesised following molybdenum incorporation into MPT and prior to acquisition of the cofactor by apomolybdoenzymes (Nichols and Rajagopalan, 2002). Although the cytoplasmic MobB protein has been shown to bind GTP with high affinity (Eaves et al., 1997), a definitive role for MobB-family proteins in any aspect of molybdoenzyme biosynthesis has never been uncovered (Buchanan et al., 2001b; Magalon et al., 2002). Some bacterial molybdoenzymes including the cytoplasmic carbon monoxide dehydrogenase from Oligotropha carboxidovorans (Dobbek et al., 1999) and its periplasmic E. coli homologue YagRST (Table 1) utilise molybdopterin cytosine dinucleotide (MCD) in which MPT has been modified by addition of CMP. The mechanism of MCD biosynthesis, while probably related to that of MGD assembly (Baitsch et al., 2001), has not yet been experimentally defined.
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Table 1 Representative examples of multisubunit protein complexes in which at least one subunit is transported by the Tat system. MGD is bis(molybdopterin guanine dinucleotide)Mo, Se-Cys is selenocysteine, TTQ is tryptophyl tryptophanquinone. Targeting pathway
Complex
Subunit
Cofactors
Signal peptide
Pathway
1. Tat only (heterooligomers)
Desulfovibrio gigas [Ni-Fe] hydrogenase HynA1B1
HynA
Tat
Tat
HynB
2x [4Fe-4S] [3Fe-4S] [NiFe]
none
Tat
Desulfovibrio desulfuricans [Fe] hydrogenase HydA1B1
HydA
H-cluster
none
Tat
HydB
None
Tat
Tat
E. coli YagTSR
YagT YagS YagR
2x [2Fe-2S] FAD MCD
Tat none none
Tat Tat Tat
E. coli formate dehydrogenase-N (FdnGHI)3
FdnG
MGD-SeCys [4Fe-4S] 4x[4Fe-4S]
Tat
Tat
none
Tat
2x haem b 2x [4Fe-4S] [3Fe-4S] [NiFe] 2x haem b
none Tat
SRP Tat
none none
Tat SRP
2x [4Fe-4S] [3Fe-4S] [NiFe] 4x [4Fe-4S] None
Tat
Tat
none Tat none
Tat Tat SRP
Tat
Tat
Tat none
Tat SRP
Tat
Tat
Sec
SecA
2. Tat and SRP
E. coli [NiFe] hydrogenase-1 (HyaABC)?
3. Tat and SecA
4. Tat (homooligomers)
FdnH FdnI HyaA HyaB HyaC
E. coli [NiFe] hydrogenase-2 (HybABCO)2
HybO
S. enterica tetrathionate reductase (TtrABC)?
TtrA
Paracoccus pantotrophus periplasmic nitrate reductase NapA1B1
NapA NapB
MGD [4Fe-4S] 2x c-haem
Paracoccus denitrificans methylamine dehydrogenase [MauAMauB]2
MauA MauB
TTQ none
Tat Sec
Tat SecA
P. denitrificans nitrous oxide reductase NosZ2 Alcaligenes faecalis copper nitrite reductase NirK3 Zymomonas mobilis Glucose fructose oxidoreductase Gfo4
NosZ
Cu2 [4Cu-S]
Tat
Tat
NirK
2x Cu
Tat
Tat
Gfo
NADPþ
Tat
Tat
HybC HybA HybB
TtrB TtrC
MGD [4Fe-4S] 4x [4Fe-4S] none
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A remarkable recent discovery is that biogenesis of at least one group of MGD-containing enzymes involves a specific signal peptide-binding protein. This protein was discovered during work on the assembly of E. coli dimethylsulphoxide (DMSO) reductase. DMSO reductase is a heterotrimeric enzyme in which the DmsAB subunits are transported by the Tat system using a signal peptide on the MGD-binding DmsA. Once in the periplasm, DmsAB associates with the integral membrane DmsC subunit (Stanley et al., 2002). A small cytoplasmic protein has now been identified that binds more-or-less specifically to the DmsA Tat signal peptide in affinity chromatography experiments (Oresnik et al., 2001). The protein, termed DmsD (formerly YnfI), is encoded by an operon ( ynfEFGHdmsD) encoding an apparent DmsABC homologue. DmsD was initially proposed to act as an ‘escort chaperone’ directing the DmsAB complex to the Tat translocase (Oresnik et al., 2001). However, while DmsAB are enzymatically active when their export is blocked by mutations in the Tat apparatus (e.g. Sargent et al., 1998a), a dmsD mutation abolishes all cellular DMSO reductase activity (Oresnik et al., 2001) suggesting that DmsD functions in enzyme maturation rather than protein transport per se. Indeed, DmsD is a member of the TorD family of molecular chaperones (Sargent et al., 2002; Tranier et al., 2002) that have previously been implicated in the insertion of MGD into the Tat substrate TorA (Pommier et al., 1998). An alternative model of DmsD function has therefore been proposed (Sargent et al., 2002) in which DmsD binds co-operatively to the cofactor binding site of the DmsA apoprotein and to the Tat signal peptide (Fig. 1). In this model, the signal peptide is masked and unavailable to direct protein export until DmsD has been released from the precursor protein by cofactor insertion (Santini et al., 1998; Figure 1). DmsD would therefore have the role of ‘proof reading’ the enzyme to ensure that the cofactor had been correctly inserted before allowing protein export to proceed. Several other MGD-dependent periplasmic enzymes have apparent accessory proteins encoded in their structural gene operons that are candidates to be DmsD/TorD-like chaperones. Examples include the NapD protein required for the activity of periplasmic nitrate reductase NapA (Berks et al., 1995a; Potter and Cole, 1999) and FdhE required for the biogenesis of E. coli formate dehydrogenase-N (Mandrand-Berthelot et al., 1988). It is noteworthy that in each case cytoplasmic homologues of these various enzymes (biotin sulphoxide reductase in the case of TorA; assimilatory nitrate reductase in the case of NapA; cytoplasmic formate dehydrogenase in the case of formate dehydrogenase-N) do not possess equivalent chaperones. However, there are two classes of MPT-dependent enzymes in which both periplasmic and cytoplasmic members appear to use
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specific assembly chaperones. For the cytoplasmic membrane-bound nitrate reductases, the NarJ protein has a well-documented role in MGD loading (Blasco et al., 1998). Homologues of NarJ are associated with a probable periplasmic nitrate reductase in Archaeoglobus fulgidus and the nitrate reductase-like enzymes dimethylsulphide dehydrogenase and selenate reductase (Krafft et al., 2000; McDevitt et al., 2002). A similar situation exists for the xanthine dehydrogenase family where the chaperone XdhC is required for biogenesis of cytoplasmic examples (Leimkuhler and Klipp, 1999) and is also found associated with exported members of the family (e.g. the YagQ gene product of the E. coli yagTSRQP operon). It is, therefore, unclear at this stage whether signal peptide-binding chaperones have evolved from cofactor insertion proteins or vice versa.
2.3.2. Other Proteins with Nucleotide-Based Cofactors Some periplasmic proteins with FAD cofactors appear to be exported by the Tat pathway. However, others are transported by the Sec system suggesting that bacterial cells have mechanisms to insert flavin cofactors into proteins in the periplasmic compartment. Intriguingly, there is a set of homologous flavoproteins, the fumarate reductase family of flavocytochromes c, in which some homologues use the Tat pathway and some the Sec pathway. In this protein family, the FAD and haem prosthetic groups are found in separate domains. In the methacrylate reductase of Wolinella succinogenes, each of the domains is found on a distinct polypeptide with the FAD-binding subunit having a Tat signal peptide and the cytochrome c subunit a Sec signal peptide (Simon et al., 1998; Gross et al., 2001). However in the highly homologous fumarate reductase of Shewanella frigidimarina, both the flavin and haem domains are part of a single Sec-exported polypeptide (Pealing et al., 1992). Thus FAD is probably inserted into these homologous enzymes in different subcellular compartments (Berks et al., 2000a). Another pair of homologous flavoproteins with distinct targeting mechanisms are the FADbinding subunit of the flavocytochrome c ‘sulphide dehydrogenase’ of thiosulphate-oxidizing bacteria and the FAD-binding peripheral membrane enzyme sulphide-quinone oxidoreductase (SQR). While the flavocytochrome subunit is translocated by the Tat pathway, SQR lacks any sort of aminoterminal targeting sequence and is apparently translocated by a currently uncharacterised mechanism (Dolata et al., 1993; Schutz et al., 1999). The periplasmic enzyme glucose-fructose oxidoreductase (GFOR) of Zymomonas mobilis contains a bound NADPþ as the active site cofactor (Kingston et al., 1996). This protein is exported by the Tat pathway. Indeed,
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analysis of the correlation between cofactor binding by GFOR and export has been important in forming our view of the folding state of Tat substrates (Section 2.3.9). As with other Tat substrates substitution of the twin arginine residues of the GFOR Tat consensus motif with lysines blocks export. The resultant mutant GFOR precursor is stable enough to be isolated and the crystal structure has been determined (Nurizzo et al., 2001). Unfortunately, the Tat (mutant) signal peptide is disordered in all of the four crystal forms that were analysed. This may indicate that Tat signal peptides do not have a defined structure except when bound to sites on their cognate receptor proteins. There is no evidence that Tat substrates binding FAD or NADPþ associate with specific chaperone proteins.
2.3.3. Proteins with Iron-Sulphur Clusters Proteins binding iron-sulphur clusters are the most common substrates of the Tat pathway. In most cases, however, these proteins also either contain a second type of cofactor or are transported as a tight oligomeric complex with partner subunits containing another type of cofactor (see below). A prominent exception to this generalisation is the Rieske component of the cytochrome bc1 and cytochrome b6 f electron transfer complexes which contains a single [2Fe-2S] cluster (Hinsley et al., 2001; Molik et al., 2001). The Tat signal peptide of the Rieske proteins is uncleaved and forms an amino terminal transmembrane region. As noted above (Section 2.2), the signal peptide of the chloroplast protein has a lysine-arginine rather than arginine-arginine pair in the Tat consensus motif. Intriguingly, it has been shown that the chloroplast Rieske precursor associates not only with the general stromal chaperones Hsp70 and Cpn60 (Madueno et al., 1993) but also with the precursor, but not mature, form of a Tat-targeted chloroplast FKBP immunophilin (Gupta et al., 2002). It is conceivable that the chloroplast Rieske protein is transported as a complex with the FKBP. Although the co-ordination of iron-sulphur cluster insertion and Tat transport is currently unstudied, it is probably fair to assume that ironsulphur clusters are assembled and inserted into Tat substrates by the same mechanisms as their cytoplasmic counterparts. Three pathways for the biosynthesis of iron-sulphur clusters have been identified in prokaryotes and these are termed Isc, Nif, and Suf (Frazzon et al., 2002; Takahashi and Tokumoto, 2002). Each pathway contains a homologous and functionally equivalent core set of proteins. For example, the IscS/NifS/SufS proteins liberate S0 from cysteine to provide the sulphur atoms of the iron-sulphur
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cluster (Zheng et al., 1994; Mihara et al., 1999). Two further components, IscU/NifU and IscA/NifIscA/SufA, appear to act as scaffolds on which ironsulphur clusters are transiently assembled before being inserted into the target apoproteins (Agar et al., 2000; Krebs et al., 2001). The requirements for cysteine and ferrous iron in the biosynthesis of iron-sulphur clusters would be hard to meet in the periplasmic compartment. This, together with the use of a protein scaffold for cluster assembly, makes it easy to understand why the cell assembles periplasmic iron-sulphur proteins on the cytoplasmic side of the cell membrane.
2.3.4. Hydrogenases A hydrogenase is, by definition, an enzyme capable of the reversible oxidation of molecular hydrogen. However, under physiological conditions, any given hydrogenase operates in a single direction only. As a general rule, enzymes operating in the direction of dihydrogen oxidation (‘uptake hydrogenases’) are found in the periplasm while those operating in the direction of dihydrogen production (‘evolution hydrogenases’) are found in the cytoplasm. Hydrogenases can also be split into two broad classes based on cofactor content: the [Fe] hydrogenases and the [NiFe] hydrogenases. Periplasmic hydrogenases of either type are exported via the Tat system. [Fe] hydrogenases contain a unique catalytic centre, the ‘H-cluster’, comprising a dinuclear iron centre co-ordinated by carbon monoxide and cyanide ligands as well as a nearby [4Fe-4S] cluster (Adams and Stiefel, 2000). Periplasmic [Fe] hydrogenases have a heterodimeric structure. The ‘Large’ subunit contains the H-cluster and undergoes a carboxy-terminal proteolytic processing event during biosynthesis (Hatchikian et al., 1999). It seems unlikely that this cleavage is required for H-cluster assembly since cytoplasmic [Fe] hydrogenases do not undergo a processing event. It is possible, therefore, that carboxy-terminal processing is linked in some way to protein export. The major periplasmic targeting motifs for the [Fe] hydrogenases can, however, be clearly located on the ‘Small’ subunits. The ‘Small’ subunits of periplasmic [Fe] hydrogenases contain no cofactors and apparently have no role in the enzymatic mechanism. However they, rather than the Large subunit, bear the Tat signal peptide. Structural studies of a periplasmic [Fe] hydrogenase hint that the periplasmic isoenzymes may have evolved via a clever modification of their cytoplasmic cousins (Nicolet et al., 1999; 2002). Sequence analysis suggests that the periplasmic [Fe] hydrogenase Small subunits are homologous to the extreme carboxyterminal domain of some cytoplasmic [Fe] hydrogenases and, therefore,
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this carboxy-terminal domain may have been detached and then modified by the addition of a signal peptide. This theory is supported by the available genetics which demonstrate that the genes encoding periplasmic [Fe] hydrogenase Small subunits are always distal to, and in an operon with, the genes encoding Large subunits (Vignais et al., 2001). It is noteworthy that despite the complex nature of the H-cluster cofactor and the peculiar proteolytic processing associated with the periplasmic [Fe] hydrogenase Large subunits, no system-specific accessory genes encoding factors required for assembly of [Fe] hydrogenases have so far been identified. The active site of [NiFe] hydrogenases is a unique dinuclear nickel-iron centre in which the iron atom is co-ordinated by one carbon monoxide and two cyanide ligands. The core of all [NiFe] hydrogenases is a heterodimer of a ‘Large’ subunit containing the [NiFe] centre and a ‘Small’ subunit containing [FeS] clusters (Vignais et al., 2001). The Large subunits of the periplasmic [NiFe] hydrogenases contain no identifiable targeting motifs. Instead, it is the Small subunits that are synthesised with Tat signal peptides. The mechanism of assembly of the [NiFe] centre is extremely complicated. Nevertheless, significant advances in our understanding of NiFe cofactor biosynthesis have recently taken place. An important finding is that nickel uptake into the cytoplasm is required for the biosynthesis of both periplasmic and cytoplasmic [NiFe] hydrogenases (Wu et al., 1991; Navarro et al., 1993; Fu et al., 1995; de Pina et al., 1999; Eitinger and MandrandBerthelot, 2000; Olson and Maier, 2000). This indicates that nickel insertion into hydrogenases is a cytoplasmic event even for periplasmic hydrogenases. Indeed, it is now clear that, for all [Ni-Fe] hydrogenases, active site assembly is conducted by a conserved set of six cytoplasmic ‘cofactor chaperones’. Those associated with biogenesis of the E. coli cytoplasmic hydrogenase-3 are termed HypA, HypB, HypC, HypD, HypE, and HypF. Most of this set of Hyp proteins are also involved in the biosynthesis of the periplasmic hydrogenases (below). Assembly of the NiFe active site has been most extensively investigated for the E. coli HycE protein which is the Large subunit of hydrogenase-3. These findings should, however, be applicable to all [NiFe] hydrogenases. The CO and CN non-protein iron ligands are first generated from carbamoyl phosphate by the crystallographically defined HypF protein (Paschos et al., 2002; Rosano et al., 2002). There is some indication that the otherwise uncharacterised HypE protein may also be involved in this process since a genetic screen of the Helicobacter pylori genome indicated that HypF and HypE interact strongly (Rain et al., 2001). Meanwhile, the HypC protein forms a complex with HypD (which contains an iron-sulphur cluster) in a reaction dependent upon inorganic Fe and a conserved
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cysteine side-chain at the extreme N-terminus of HypC (Blokesch and Bo¨ck, 2002). The HypCD-Fe complex then acts as a scaffold for the capture of the CO and CN ligands generated by Hyp(E)F (Blokesch and Bo¨ck, 2002). It is likely that the Fe associated with the HypCD complex is the same Fe atom that is incorporated into the hydrogenase active site (Blokesch and Bo¨ck, 2002). At this stage, the HypCD-Fe-[CO-(CN)2] complex is now ready to interact with the HycE apoenzyme. HycE is synthesised as a precursor with a long carboxy-terminal extension that is proteolytically removed following active site assembly. Unlike the [Fe] hydrogenase (above), this carboxy-terminal extension is definitely not involved in protein transport since HycE is not exported from the cytoplasm. Instead, the presence of the carboxy-tail is essential for docking of the active site biosynthetic machinery onto the apoenzyme (Binder et al., 1996). Attachment of the HypCD-Fe-[CO-(CN)2] complex onto HycE is followed by the transfer of the CO and CN ligands (or, most likely, the complete Fe-[CO-(CN)2] complex) into the apoenzyme. The energetic requirements for this transfer are not known. Finally, with at least the HypC protein still in place, the Ni(II) is delivered by the HypA and HypB proteins. The exact role of HypA is unknown although its role in nickel insertion is supported by the fact that a hypA mutant phenotype is partially suppressed by excess nickel (Hube et al., 2002). The HypB homodimer is a GTPase (Maier et al., 1993) and, under normal physiological conditions, nickel incorporation into hydrogenase is dependent upon this GTPase activity (Maier et al., 1995). However, hypB mutant phenotypes, including point mutations that prevent GTP binding or hydrolysis, can be completely suppressed by the addition of excess nickel to the growth medium (Waugh and Boxer, 1986; Maier et al., 1995). Although E. coli HypB apparently does not bind nickel directly, HypBfamily proteins from other bacteria often contain histidine-rich N-terminal domains that can chelate up to eighteen Ni(II) ions per monomer and thus double as intracellular nickel-storage proteins or ‘nickelins’ (Olson and Maier, 2000). Once the HypAB-dependent nickel incorporation into apoHycE is complete, the C-terminal extension used as a staging-post by the biosynthetic machinery is irreversibly removed by a specific protease (Theodoratou et al., 2000). In summary, the requirement for nickel, ferrous iron, volatile small molecules, nucleotide triphosphates, and (possibly redox regulated) protein–protein interactions means that the assembly of the [NiFe] centre is clearly restricted to the cell cytoplasm. It is notable that it is only after the extensive process of cofactor incorporation is completed that the HycE protein interacts with its Small subunit partner (Magalon and Bo¨ck, 2000).
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For periplasmic [NiFe] hydrogenases, maturation of the Large subunit cofactor is only the start of the assembly process. Our molecular understanding of these additional steps is still rudimentary but can be illustrated by reference to the organisation and biosynthesis of the two Tattargeted hydrogenases of E. coli, namely the membrane-bound uptake hydrogenases-1 and -2. The hydrogenase-1 isoenzyme is encoded by the hyaABCDEF operon (Menon et al., 1990). HyaA and HyaB are the hydrogenase Large and Small subunits respectively (Sawers and Boxer, 1986) while HyaC is a dihaem-containing protein that anchors HyaAB to the periplasmic face of the cell membrane (Berks et al., 1995b). The hydrogenase-2 isoenzyme is encoded by the hybOABCDEFG operon (Menon et al., 1994; Sargent et al., 1998b). HybC is the hydrogenase Large subunit and HybO is the Small subunit. No HyaC homologue is encoded by the hyb operon. Instead, HybOC associates with the Tatdependent periplasmic FeS protein HybA and the large, cofactorless, integral membrane protein HybB to form the final catalytic complex (Dubini et al., 2002). Cross-linking experiments demonstrated that the HybOC complex is formed prior to export from the cytoplasm (Rodrigue et al., 1999). Immunochemical analysis of hydrogenases-1 and -2 in various tat mutants showed that the core HybAB and HybOC enzymes were fully formed in terms of cofactor and subunit composition but completely mislocalised to the cell cytoplasm (Bogsch et al., 1998; Sargent et al., 1998a, 1999). Taken together, these data indicate that the core [NiFe] heterodimer is fully assembled in the cytoplasm before Tat transport takes place. The functions of the seven non-structural genes encoded in the hya and hyb operons are poorly characterised but would appear to be linked to further aspects of hydrogenase biosynthesis. The hyaD, hybD, hybF and hybG gene products are all shown, or predicted, to be ‘cofactor chaperones’. HyaD and HybD are the cytoplasmic proteases required for the carboxyterminal processing of the HyaB and HybC subunits, respectively, following [NiFe] centre assembly (above). The HybF protein displays 49% overall sequence identity with the HypA protein from E. coli and has been shown to perform an analogous role to that of HypA in nickel processing but specifically for the hydrogenase-1 and hydrogenase-2 isoenzymes (Hube et al., 2002). The HybG protein shares 55% overall sequence identity with the E. coli hypC gene product which is the key chaperone in the assembly of the [NiFe] hydrogenase-3 HycE subunit (above). Mutations in hypC have been reported to have little or no effect on the synthesis of hydrogenases-1 or -2 and HybG has recently been shown to perform the role of HypC as a cytoplasmic ‘cofactor chaperone’ for these isoenzymes (Blokesch et al., 2001). The remaining hya and hyb gene products HyaE, HyaF, and HybE
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proteins are of particular interest as there appear to be no equivalent gene products required for the biosynthesis of cytoplasmic [NiFe] hydrogenases. Homologues of these proteins are, however, found in other bacteria with periplasmic [NiFe] hydrogenases (Casalot and Rousset, 2001). In Ralstonia eutropha, the HyaE homologue HoxO and the HyaF homologue HoxQ have been shown to be required for both targeting and activity of the Tat-dependent hydrogenase. In contrast, the R. eutropha HybE homologue HoxT was not required for targeting or in vitro activity of the enzyme but, despite a lack of obvious cofactor, was apparently required for in vivo electron transfer from hydrogenase to the quinone pool (Bernhard et al., 1996). In the light of these observations the phenotypes of E. coli hyaE and hyaF mutant strains are surprising since assembly of both hydrogenase-1 and hydrogenase-2 proceeded apparently unhindered in each mutant background (Dubini et al., 2002). A strain simultaneously lacking both HyaE and HyaF, however, was completely devoid of hydrogenase-1 but retained some hydrogenase-2 activity (Menon et al., 1991). For the moment, therefore, the precise roles of HyaE-, HyaF-, and HybE-family proteins in Tat-dependent [NiFe] hydrogenase biosynthesis remain enigmatic.
2.3.5. Proteins with Copper Cofactors Biology uses copper as a cofactor almost exclusively in proteins found in extracellular compartments (Frau´sto da Silva and Williams, 2001). Copper sites in proteins are normally formed simply through direct co-ordination of copper ions by amino acid ligands. In general, it is assumed that the copper proteins found in the bacterial periplasm form their metal sites by pickingup copper ions from the surrounding medium and certainly the copper centres of many proteins can be easily reconstituted in vitro by mixing copper ions with an empty apoprotein. It is, then, perhaps surprising that targeting of copper proteins to the bacterial periplasm is complicated with systems dependent upon either Sec or Tat pathways having been identified. Most periplasmic copper proteins belong to the cupredoxin superfamily in which the copper ions are bound at one end of an antiparallel b-barrel structure. The simplest members of this family comprise a single b-barrel domain binding a single copper atom. Examples are azurins, pseudoazurins, plastocyanins and rusticyanins. These proteins all possess Sec signal peptides. More complex are the copper nitrite reductases which are homotrimeric enzymes with each subunit composed of two cupredoxin domains (Godden et al., 1991). Nitrite reductase has two mononuclear
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copper sites per subunit and, since one of these sites is at a subunit interface with ligands donated by two separate polypeptides, it is likely that copper is only inserted following oligomer formation. Some copper nitrite reductases are synthesised as precursors with Sec signal peptides. However, others have clear Tat signal peptides (Berks et al., 2000a). The most complex members of the cupredoxin family are the multicopper oxidases. In these proteins, a single polypeptide forms three cupredoxin domains and a mononuclear and a trinuclear copper centre are normally bound. All bacterial multicopper oxidases have Tat signal peptides. Indeed, the E. coli CueO protein (formerly YacK; Outten et al., 2000; Kim et al., 2001; Roberts et al., 2002) has been used in Tat targeting studies (Stanley et al., 2000). Thus, within the cupredoxin superfamily, targeting to both the Sec and Tat systems is observed. This suggests that the use of the Tat system by this protein superfamily is for reasons other than cofactor binding. Instead, it has been proposed that the Tat system is preferentially employed as the number of cupredoxin domains increases because the cell has increasing difficulty in either preventing or reversing rapid folding of multiple cupredoxin domains (Berks et al., 2000a). It is worthwhile noting that there is no evidence for the involvement of system-specific ‘cofactor chaperones’ in the biosynthesis of any of these members of the cupredoxin family. It is also interesting to note that many eukaryotes secrete multicopper oxidases (e.g. ceruloplasmin, ascorbate oxidase, laccase) and that this process must involve the Sec-like system of the endoplasmic reticulum. A further three families of copper protein are known to be periplasmically localised in bacteria. The periplasmic Cu, Zn superoxide dismutases, which contain both a copper atom and a zinc atom at their active site, are synthesised with Sec signal peptides (Imlay and Imlay, 1996; Pesce et al., 1997). In contrast, the enzymes tyrosinase and nitrous oxide reductase have been shown to be exported by the Tat pathway (Heikkila¨ et al., 2001; Schaerlaekens et al., 2001). Tyrosinases are single subunit enzymes with dinuclear copper active sites (Klabunde et al., 1998). The unusual biogenesis of the tyrosinase from Streptomyces species has been studied in some detail. The tyrosinase gene product itself lacks a signal peptide. Instead, the tyrosinase apoprotein is secreted in complex with a second protein, MelC1, that bears a Tat signal peptide (Lee et al., 1988; Chen et al., 1992; Leu et al., 1992). Following secretion, MelC1 is only released from the complex upon copper ion insertion into the tyrosinase (Chen et al., 1992; Tsai and Lee, 1998). The MelC1 protein is thus a bifunctional molecule that acts both as a targeting factor to control access to the Tat machinery and as a chaperone in the cofactor insertion process.
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Nitrous oxide reductase is a homodimeric enzyme in which each subunit possesses two types of copper-containing site (Brown et al., 2000a). The CuA centre is a dinuclear copper site located within a cupredoxin domain while the CuZ active site is a structurally unique tetranuclear copper cluster in which an inorganic sulphur atom bridges the four metal atoms (Brown et al., 2000b; Rasmussen et al., 2000). By analogy to the biosynthesis of other periplasmic proteins containing metal-sulphur clusters (above), it might be expected that the copper sulphide cluster in nitrous oxide reductase would be inserted in the cytoplasm. However, when Tat transport of nitrous oxide reductase in Pseudomonas stutzeri was blocked, no copper was found in the precursor protein that had accumulated in the cell cytoplasm (Dreusch et al., 1997; Heikkila¨ et al., 2001). This suggests that, as in other periplasmic copper proteins, the metal ions are inserted into nitrous oxide reductase in the periplasm rather than the cytoplasm. Tat targeting of the enzyme is not, therefore, linked to a cofactor binding-event. Such a conclusion raises the question of the biosynthetic origin of the sulphur atom found in the CuZ cluster. In all organisms so far examined, the structural gene for nitrous oxide reductase is linked to accessory genes, nosDFYLX, coding for proteins thought to be involved in cofactor provision. The periplasmically-located lipoprotein NosL may act as a copper donor during maturation of nitrous oxide reductase since it binds a single cuprous copper ion (McGuirl et al., 2001). However, there is some uncertainty about whether NosL is essential for nitrous oxide reductase biosynthesis (Dreusch et al., 1996; McGuirl et al., 2001). NosD is another predicted periplasmic protein while NosFY appear to form a transporter of the ATP-binding cassette (ABC) superfamily (Zumft et al., 1990). A specific role for these proteins in the biosynthesis and insertion of the CuZ cluster is suggested by the observation that nitrous oxide reductase is periplasmically located in a nosDFY strain but contains only the CuA and not the CuZ cluster (Riester et al., 1989; Zumft et al., 1990). Since only the CuZ centre contains an inorganic sulphur atom a plausible scenario is that NosFY function to transport the sulphur atom, from cytoplasm to periplasm with NosD acting as a chaperone for CuZ cluster assembly. NosX is a further predicted periplasmic protein and is essential for the production of an active nitrous oxide reductase (Saunders et al., 2000). Targeting of nitrous oxide reductase is unaffected in a nosX mutant but there is circumstantial evidence that insertion of at least the CuA centre is compromised in this strain (Saunders et al., 2000). One intriguing aspect of the biosynthesis of nitrous oxide reductase is that the enzyme possesses an exceptionally large Tat signal peptide. This is normally around 50 amino acid residues in length (Zumft and Kroneck,
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1996), with the additional residues over and above the ‘standard’ Tat signal peptide being evenly split between a longer c-region and that portion of the n-region that precedes the consensus twin-arginine motif. The reason for the extended signal peptide is not known. Another oddity is that the nitrous oxide reductase gene cluster of P. stutzeri, though not those of other organisms, encodes a TatA/E homologue (Glockner and Zumft, 1996). However, ablation of this P. stutzeri tat gene has only minor effects on the export of nitrous oxide reductase, presumably because of the presence of two other TatA/E homologues in this bacterium (Heikkila¨ et al., 2001).
2.3.6. The Tryptophyl Tryptophanquinone (TTQ) Cofactor of Methylamine Dehydrogenase Methylamine dehydrogenase is a periplasmic enzyme with an a2b2 quaternary structure (Chen et al., 1998). The active site tryptophyl tryptophanquinone (TTQ) cofactor is formed by the post-translational modification of two tryptophan side-chains in the b subunit. These tryptophan residues are linked by a covalent bond between their indole rings. In addition, two oxygen atoms are incorporated into the indole ring of one of the tryptophan residues to form a quinone moiety. While the a (MauB) subunit precursor has a standard Sec signal peptide, the TTQcontaining b (MauA) subunit possesses an apparent Tat signal peptide (Chistoserdov and Lidstrom, 1991; Chistoserdov et al., 1994). The b subunit signal peptide has two noteworthy features. Firstly, it shows unusually high sequence conservation between species. Secondly, like the signal peptide of nitrous oxide reductase, it is extraordinarily long. Indeed, at approximately 60 residues in length it is as much as half the size of the mature portion of the b subunit and is the largest known bacterial signal peptide. As with the nitrous oxide reductase signal peptide, the excess length over a standard Tat signal peptide is split between the c-region and the polypeptide sequence preceding the twin-arginine motif. While the sequence of the b subunit signal peptide suggests that it is a Tat targeting signal, several considerations indicate that at least some of the post-translational modification and folding of this subunit occurs in the periplasm after the transport step. Although the b subunit has only around 130 residues, it contains a remarkable six disulphide bonds (Chen et al., 1998). It is difficult to imagine that the b protein is stably folded until the disulphide bonds have been inserted, yet disulphide bond formation is a periplasmic process (Ritz and Beckwith, 2001). Indeed, as far as we are aware, the only other example of a disulphide bond in a Tat substrate is that
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found in the catalytic subunit of flavocytochrome c (Chen et al., 1994) and this is probably produced through the catalytic mechanism of the enzyme rather than by the general periplasmic disulphide bond forming machinery (Griesbeck et al., 2002). Biosynthesis of methylamine dehydrogenase involves five dedicated accessory genes, of which mauD, mauG and mauL appear to code for periplasmically located proteins, and the others code for membrane, and not cytoplasmic, proteins (Chistoserdov et al., 1994). This again suggests that cofactor maturation takes place predominantly following export. Sequence analysis indicates that the MauD protein contains the conserved Cys-Xaa-Xaa-Cys amino acid motif found in disulphide interchange proteins (van der Palen et al., 1997). MauD could, therefore, be involved in disulphide bond insertion into the b subunit. It has been shown that c-type cytochromes, a class of protein that occurs only in the periplasmic compartment, are required for maturation of the TTQ cofactor (Page and Ferguson, 1993). This phenotype could be linked to the fact that the MauG protein is a cytochrome c-containing peroxidase. However, it should be noted that the quinone group of TTQ has been reported to be present in the b subunit of a mauG mutant strain but not in cytochrome c-deficient mutants (Page and Ferguson, 1993; Chistoserdov et al., 1994). Inspection of the structure of the methylamine dehydrogenase b subunit shows that the quinone group of TTQ is relatively exposed at the surface of the protein (Chen et al., 1998). It therefore seems reasonable to suggest that formation of this portion of the cofactor could be catalysed by an accessory protein even if the b subunit has folded. In contrast, the tryptophan crosslink is buried and may be an important structural determinant in itself. It is possible the crosslink could be formed in the folded protein by a concerted reaction linked to quinone formation; however, it is also possible that formation of the crosslink precedes folding. If this occurs, and if it occurs in the cytoplasm, the protein is structured before transport and we have a rationale for the use of the Tat pathway. Nevertheless, given the extensive evidence for cofactor maturation and intramolecular crosslinking reactions following transport, there must be a question mark over whether the methylamine dehydrogenase b subunit really is a Tat substrate and direct experimental conformation that this protein is transported by the Tat pathway would be highly desirable.
2.3.7. Proteins with Cobalamin Cofactors The catalytic subunits of the reductive dehalogenases from strict anaerobes contain an active site cobalamin cofactor and two [4Fe-4S] clusters.
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Sequencing studies indicate that these proteins are synthesised with apparent Tat signal peptides and that these signal peptides are processed during enzyme assembly (Neumann et al., 1998; van de Pas et al., 1999; Suyama et al., 2002). Initially, these observations caused some confusion since the reductive dehalogenases had been assigned a cytoplasmic location on the basis of substrate accessibility experiments (Miller et al., 1997). However, a periplasmic localisation for reductive dehalogenases has recently been confirmed by subcellular fractionation (Suyama et al., 2002). Thus, cobalamin can be added to the list of cofactor types associated with Tat targeting.
2.3.8. Biogenesis of Oligomeric Proteins Involving the Tat System Many Tat substrates are subunits of oligomeric complexes (Table 1). In some of these enzyme complexes, certain subunits are transported by the Tat pathway while others are translocated by the Sec pathway. Other complexes include Tat-dependent proteins that associate with partner subunits that are integral membrane proteins. Assembly of such complexes is presumably relatively straightforward as the Tat-directed proteins should encounter the binding sites on their cognate partner subunits only upon reaching the periplasm. Some complexes comprise multiple, non-identical, Tat-targeted subunits (Table 1). A striking feature of these complexes is that usually only one of the subunits bears a Tat signal peptide with export only occurring following formation of the hetero-oligomeric complex in the cytoplasm (Sections 1.1, 2.3.1 and 2.3.4). The use of such a piggy-back mechanism probably indicates these periplasmic proteins arose from the addition of a Tat targeting signal to a pre-existing cytoplasmic enzyme. Currently the only exception to the rule that only one subunit type in a complex possesses a Tat signal peptide is the Salmonella enzyme tetrathionate reductase in which the TtrA and TtrB subunits each have their own Tat signal peptide (Hensel et al., 1999; Hinsley et al., 2001; Table 1). It should be noted, however, that tight complex formation between TtrA and TtrB has only been inferred from genetic experiments. Homo-oligomeric Tat substrates also exist (Table 1). These complexes present the cell with a novel biosynthetic challenge because each protomer possesses its own Tat signal peptide. As these signal peptides have apparently all been correctly processed by a periplasmic signal peptidase in the final complexes, it is likely that the protomers are translocated separately and only come together in the periplasm (Berks et al., 2000a).
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Given the central dogma of Tat transport that the Tat-dependent protomers should already be folded in the cytoplasm prior to transport, the cell must have a mechanism to prevent their premature association. Thus, while hetero-oligomeric Tat substrates need to ensure that complex formation occurs prior to transport, homo-oligomeric substrates must prevent the oligomer assembling before translocation has taken place. Interestingly, E. coli formate dehydrogenase-N is an example of a protein that must undertake both types of oligomer control. Formate dehydrogenase-N (Fdn) comprises a basic trimeric unit, FdnGHI, in which the FdnGH (or ab) subunits form a Tat-targeted heterodimer with the twin-arginine signal peptide located on the FdnG protein while the FdnI (or g) subunit is an integral membrane protein. Recent structural work has revealed that the final mature form of the membrane-bound formate dehydrogenase-N adopts an (abg)3 ‘trimer-of-trimers’ architecture with all three different subunits providing extensive protein-protein contacts in the final assemblage (Jormakka et al., 2002). As a result, biosynthesis of this enzyme probably requires formation of FdnGH heterodimers prior to Tat transport but suppression of trimerisation of the FdnGH units. In the case of formate dehydrogenase-N, simple steric occlusion of the interfacial region by the Tat signal peptide may be enough to prevent trimer formation since the amino terminus of mature FdnG is at the trimer interface (Sargent et al., 2002).
2.3.9. General Proofreading Properties of Tat Transport It is important that the substrate proteins are not exported by the Tat system until all assembly processes have been completed. While the studies described above (Section 2.3.1) indicate that E. coli DmsD could act as a ‘proofreading’ factor for the Tat pathway, it is apparent that this is a very specific system that is restricted to a closely homologous set of molybdoenzymes. Putative chaperones have been suggested for some other periplasmic molybdoenzymes (for example TorD, NapD, and FdhE; Section 2.3.1) with the gene encoding the putative chaperone being linked to the structural genes of its cognate partner in most cases. However, based on this criterion, it is notable that many molybdoenzymes do not have associated assembly chaperones. Examples are the SoxC component of the thiosulphate-oxidizing multienzyme system of some purple bacteria (Wodara et al., 1997), and the Proteobacterial tetrathionate, thiosulphate and polysulphide reductases (Krafft et al., 1992; Heinzinger et al., 1995; Hensel et al., 1999). Furthermore, the [NiFe] hydrogenases appear to be the only other class of cofactorcontaining Tat substrates that might utilise a similar private chaperone-based
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system-specific proofreading protocol (Section 2.3.4). Thus, system-specific chaperones may be the exception rather than the rule. So is there a general proofreading mechanism that operates on other (or all) Tat substrates and, if so, why do certain enzymes require an additional dedicated proofreading mechanism? Several observations suggest that the bacterial Tat transport system has the general property of actively discriminating against improperly folded substrates. Fusions between reporter enzymes and Tat substrates have been observed to be export-incompetent when the fusion causes a partial truncation of the mature region (Stanley et al., 2002). However, if the entire mature protein region is removed and the reporter enzyme is fused directly after the Tat signal peptide, then export of the chimera is observed (e.g. Cristo´bal et al., 1999; Stanley et al., 2002). Since only the partial truncations are expected to present the Tat system with an improperly structured substrate, these experiments suggest that the Tat system only allows transport of folded substrates. Further evidence supporting the idea that proper folding of the protein is a prerequisite for Tat transport comes from the observation that point mutations in Z. mobilis GFOR that affect the binding affinity for the NADPþ cofactor slow Tat export (Halbig et al., 1999). Indeed, point mutations that affect Tat export, presumably as the result of structural alterations, have also been reported for P. stutzeri nitrous oxide reductase (Heikkila¨ et al., 2001). This report is particularly interesting because the export-incompatible mutations occur in the carboxy terminal CuA domain. This again suggests that the folded state of the entire substrate molecule is important and that correct folding of the domain adjacent to the signal peptide alone is insufficient to allow transport. A more direct piece of evidence that a folded substrate is required for transport comes from experiments in which cytochrome c was targeted to the E. coli Tat apparatus by fusion to a Tat signal peptide (Sanders et al., 2001). Covalent attachment of haem to c-type cytochromes normally occurs in the periplasm so when the fusion protein reaches the Tat apparatus it lacks a haem cofactor and is unstructured. Tat-dependent transport of the fusion was not observed. However, when the host E. coli strain was engineered to insert the haem cofactor into the cytochrome in the cytoplasm the fusion protein was exported. This experiment uses a heterologous substrate so there is no possibility that the observed rejection of the unfolded fusion protein is mediated by a substrate-specific chaperone. Instead, the results of this experiment strongly suggest that the Tat pathway possesses a general proofreading activity that prevents or rejects export of incorrectly structured proteins. There is currently no evidence for components additional to the translocase being required for the transport of all Tat substrates. It is,
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therefore, tempting to speculate that Tat translocase itself has intrinsic proofreading activity. Were this idea correct it might be expected that the homologous chloroplast Tat system would also exhibit proofreading activity. There is, however, conflicting evidence as to whether the thylakoid Tat system will accept unfolded proteins (Roffey and Theg, 1996; Hynds et al., 1998). The most obvious way that a general proofreading system could work would be by sensing the increased surface hydrophobicity of incorrectly folded substrates (Berks et al., 2000a). This might be monitored indirectly via the presence of general folding catalysts. Alternatively, the substrate could be forced to interact with highly hydrophilic portions of the translocase during a productive export event. Another physical attribute that might be sensed by the proofreading system is the rigidity of the protein. This could be feasible if the translocator mechanism has to do mechanical work on multiple regions of the substrate or if the substrate has to oppose the lateral pressure or the membrane bilayer when the transmembrane channel is open. If the Tat system has a general proofreading mechanism, why do some molybdoenzymes apparently utilise specific chaperones for the same purpose? One possibility is that these proteins fold sufficiently to overcome the general proofreading system before they receive their molybdenum cofactor. Certainly cofactor-free molybdoproteins can be isolated in a stable form (Temple and Rajagopalan, 2000; Leimku¨hler and Rajagopalan, 2001). An additional mechanism may, thus, be required to prevent export of the immature apoprotein. Pre-folding would allow partial pre-formation of the MPT binding site. Indeed, the crystal structure of an assembly intermediate of the nitrogenase MoFe protein shows how a complex cofactor can be inserted into its protein partner by limited refolding and rearrangement of preformed protein domains (Schmid et al., 2002). In the case of heterooligomeric Tat substrates the accessory factor may, alternatively, be required to prevent the signal peptide-bearing subunit being exported before it has bound its partner subunit(s) (Sections 2.3.1, 2.3.4 and 2.3.8). Finally, for those substrates containing transmembrane helices, an accessory factor may prevent the transmembrane region either aggregating or interacting with the membrane before the protein has reached the Tat transporter (Section 4).
2.4. Exported Proteins Without Cofactors While the bacterial Tat pathway was initially identified as a pathway dedicated to the export of proteins binding complex cofactors, it is now
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increasingly apparent that there also many Tat substrates that either lack cofactors entirely or contain isolated metal ions that can easily be picked up in the periplasmic compartment. Presumably, these proteins are targeted through the Tat system because they fold too rapidly or stably to be transported by the Sec apparatus. We estimate that E. coli K-12 strains contain twenty six proteins with putative Tat signal peptides of which nine apparently do not contain complex cofactors (Berks et al., 2000a; Stanley et al., 2000, 2001; Ize et al., 2003). Genome sequence analysis has suggested that the Gram-positive bacteria Bacillus subtilis and Streptomyces coelicolor as well as the Archaeon Halobacterium sp. NRC-1 possess many cofactorless extracellular proteins that are synthesised with plausible Tat signal peptides (Bentley et al., 2002; Jongbloed et al., 2002; Rose et al., 2002). It is, however, salutary to note that, in an experimental test of the targeting pathway of 14 such potential Tat substrates in B. subtilis, only one of the proteins was found to be transported by the Tat apparatus (Jongbloed et al., 2002). Thus, we still have a rather limited ability to correctly identify Tat substrates from the sequence of the signal peptide unless we also know that the mature protein binds a Tat-linked cofactor. Sequence analysis indicates that many of the predicted ‘cofactorless’ Tat substrates are metal-dependent (but redox-inactive) hydrolases. For some of these hydrolases Tat targeting has been experimentally demonstrated. Examples include the PlcH and PlcN secreted phospholipases C of Pseudomonas aeruginosa (Voulhoux et al., 2001), the PhoD phosphodiesterase of B. subtilis (Jongbloed et al., 2000), the E. coli R-plasmid-encoded penicillin amidase [which is initially synthesised as a catalytically inactive proenzyme (Ignatova et al., 2002)] and Thermus thermophilus alkaline phosphatase (Angelini et al., 2001). Phylogenetic analysis indicates that, while the Tat system has a wide distribution amongst both Eubacteria and Archaea, it is absent from those organisms (e.g. fermentative bacteria, some intracellular pathogens, some methanogens) that do not use extracellular redox proteins (Sargent et al., 1998a; Berks et al., 2000a; Yen et al., 2002). This observation strongly suggests that the primary function of the Tat system is the export of cofactor-containing proteins and that the pathway has subsequently been co-opted for use with difficult cofactor-free substrates. An extreme example of substrate conversion may occur in the plant chloroplast where the only cofactor-containing substrates of the thylakoid Tat pathway are the Rieske protein of the cytochrome b6 f complex (Molik et al., 2001), which binds a [2Fe-2S] cluster, and polyphenol oxidase, which is homologous to tyrosinase. As a consequence, it is thought that the main purpose of the
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thylakoid Tat pathway is to deal with nuclear-encoded proteins that fold irreversibly in the stroma during transit between the chloroplast inner membrane and the thylakoids. The realisation that many bacterial Tat substrates lack complex cofactors has prompted a reappraisal of the range of cellular process that might depend on the activity of the Tat pathway. In the remainder of this Section, we look at two areas of bacterial cell biology where the involvement of cofactorless Tat substrates has been generating a good deal of recent interest.
2.4.1. Biosynthesis of the Cell Wall and the Amidase Puzzle E. coli mutants pleiotropically defective in the transport of all twin-arginine signal peptide-bearing proteins displayed a distinctive chain-forming phenotype when visualised by microscopic techniques (Santini et al., 2001; Stanley et al., 2001; Thomas et al., 2001). Inspection of the long chains of cells suggested that bacterial cell division was blocked at a relatively late stage in the tat mutants since the murein septa were fully formed but apparently not yet separated (Stanley et al., 2001). This chain-forming phenotype was also linked with a defect in the integrity of the outer membrane, and was initially likened to that of an envA (lpxC) mutant (Stanley et al., 2001). However, it is now clear that this phenotype arises due to the failure of E. coli tat mutants to export two amidase proteins (Ize et al., 2003). E. coli expresses three sequence-related periplasmic N-acetylmuramyl-Lalanine amidases (AmiA, AmiB, and AmiC) (Heidrich et al., 2001). The physiological role of these enzymes is to cleave the murein septum at a late stage in cell division thus physically separating the two new daughter cells (Heidrich et al., 2001; Stanley et al., 2001). Both the AmiA and AmiC isoenzymes are synthesised as precursors with twin-arginine signal peptides and are translocated to the periplasm by the Tat pathway. A strain that carries deletions in the signal peptide-coding regions for AmiA and AmiC, thus synthesising mislocalised amidase A and C, has exactly the same chainforming phenotype and outer membrane defect as the tat mutant strains (Ize et al., 2003). Interestingly, the AmiB protein is apparently targeted to the periplasm on the Sec pathway, and overproduction of AmiB can compensate for the outer membrane and cell division defects of the tat mutants (Ize et al., 2003). It is not obvious why failure to cleave the division septum should lead to a loss in the integrity of the outer membrane, but these two phenotypes are
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clearly linked since strains deleted for all three amidase genes show a similar outer membrane defect as the tat mutant strain (Heidrich et al., 2002). Interestingly, the amidases of many other Gram-negative bacteria are also synthesised with putative twin arginine signal sequences. These include pathogens such as Salmonella species, Yersinia pestis, P. aeruginosa and Neisseria meningitidis. It is likely therefore that the pleiotropic outer membrane defect observed with E. coli tat mutants might be a common feature associated with tat mutants in other bacteria. Perhaps the most intriguing question to emerge from the study of periplasmic amidases is why homologous isoenzymes from the same host organism use such fundamentally different export pathways, particularly given the observation that AmiC can be successfully exported when the native Tat signal peptide is replaced with a Sec signal peptide (Ize et al., 2003).
2.4.2. Pathogenicity and the Tat Pathway Genome sequencing studies show that the Tat system is found in many bacterial pathogens of animals and plants including Bordetella pertusis, Mycobacterium tuberculosis, Y. pestis and H. pylori (Yen et al., 2002) and it is becoming increasing apparent that the Tat system is involved in the virulence of at least some of these organisms. The evidence to support this assertion is most compelling for the opportunistic pathogen P. aeruginosa where, in a mouse model, a tat mutant exhibited none of the lesions, abscesses, or histochemical responses associated with infection by a wildtype strain (Ochsner et al., 2002). The lack of virulence of the tat mutant could, in part, be ascribed to a failure to secrete extracellular phospholipases PlcH and PlcN, key virulence factors which degrade lipids in the host cell membrane (Ostroff et al., 1990; Darby et al., 1999; Rahme et al., 2000). These phospholipases are first transported into the periplasm by the Tat system before being delivered across the outer membrane by the Xcp ‘secreton’ (Voulhoux et al., 2001). It is likely that the Tat pathway is also required for the proper functioning of other virulence factors in P. aeruginosa, including proteins involved in iron acquisition and in biofilm formation (Ochsner et al., 2002). Additionally, if as predicted (Section 2.4.1), the N-acetylmuramyl-L-alanine amidases of P. aeruginosa are Tatdependent, then defects in the integrity of the cell envelope would also be expected to contribute to the avirulence of the P. aeruginosa tat mutant. Indeed, Ochsner and co-workers (2002) reported abnormal flagella and pili function in the tat mutant, a phenotype which could be the result of cell envelope defects.
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Attenuation of virulence in S. typhimurium and pathogenic E. coli strain K-1 has also been linked to ablation of the Tat pathway (Crooke et al., 2001). It is possible that, in these organisms, correct assembly of anaerobic electron transfer chains may contribute to virulence since Salmonella strains defective in anaerobic electron transport have been shown to be impaired in their ability to survive within human epithelial cells (Contreras et al., 1997). Indeed, it has recently been suggested that the respiratory nitrite reductase system of these bacteria (which includes the Tat-targeted component NrfC; Weiner et al., 1998) enables them to survive phagocytosis by white blood cells by detoxifying the nitric oxide produced during the phagocytic event (Poock et al., 2002). It is also pertinent to note that a mutant of the gastric bacterium H. pylori that does not produce the enzyme hydrogenase has a severely compromised ability to colonise a mouse model (Olson and Maier, 2002). Since biosynthesis of this hydrogenase uses the Tat system, it can be inferred that the Tat apparatus is required for H. pylori, and possibly other gut bacteria, to become established in their host organism. Given that the Tat system is not found in humans, and that it is apparently required for multiple virulence processes in bacteria, it is possible that the Tat pathway would be a good target for antibacterial drugs.
3. TRANSPORT ACROSS THE MEMBRANE 3.1. Components and Organisation of the Tat Transport Apparatus Genetic studies in E. coli suggest that the minimal components of the Tat translocation pathway are the integral membrane proteins TatA, TatB and TatC (Bogsch et al., 1998; Chanal et al., 1998; Sargent et al., 1998a, 1999; Weiner et al., 1998; de Leeuw et al., 2001). These proteins are assumed to come together in the cytoplasmic membrane to form the Tat translocase. TatA and TatB show weak (25%) sequence identity but each is essential for protein transport suggesting that they perform non-identical functions (Sargent et al., 2001). The structural genes encoding TatA, TatB and TatC are co-transcribed and form the major transcript of the tatABCD operon (Wexler et al., 2000). The remaining gene in the operon, tatD, encodes a DNase and is not involved in the Tat pathway (Wexler et al., 2000). An additional Tat-related protein is coded by the monocistronic tatE operon. TatE shares almost 60% amino acid sequence identity with TatA and the two proteins are functionally interchangeable (Sargent et al., 1998a, 1999).
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However, the level of expression of TatA is approximately one hundred times that of TatE (Jack et al., 2001) and as a consequence tatE is currently regarded as a cryptic gene duplication of tatA. An equivalent duplication of the tatA gene is also found in several bacteria that are closely related to E. coli including Salmonella typhimurium, Yersinia pestis and Vibrio cholerae. It is notable that the tatE gene products of these organisms show considerably lower sequence conservation between themselves than the tatA gene products do. This is consistent with the view that tatE is a cryptic gene duplication. The orthologs of E. coli TatA, TatB and TatC in plant thylakoids are Tha4 (Mori et al., 1999; Walker et al., 1999), Hcf106 (Settles et al., 1997) and cpTatC (Mori et al., 2001) respectively, where the equivalence of TatA with Tha4 and TatB with Hcf106 is based primarily on functional criteria (see below). The relative TatA:TatB:TatC stoichiometry in wild-type E. coli cells has been estimated to be 30:1:0.4 on the basis of chromosomal translational fusions (Jack et al., 2001) or 35:1:0.3 from measuring radiolabelled protein production from tatABC co-transcribed from a T7 promoter (our unpublished results using the experimental system described in Sargent et al., 1998a, 1999). Quantitative immunoblotting of native E. coli membranes found a TatA:TatB molar ratio of 19 4 (Sargent et al., 2001). Taken together, these measurements suggest that TatA is present at a considerable molar excess over the other Tat components. A somewhat different picture has emerged for the chloroplast Tat system where quantitative immunoblotting measurements suggests a relative Tha4:Hcf106:cpTatC stoichiometry of 8:5:1 in pea thylakoids (Mori et al., 2001). Chemical crosslinking studies indicate that E. coli TatA minimally forms homotetramers and TatB homodimers in their native membrane environment (de Leeuw et al., 2001). Since these crosslinks were observed even using the one carbon-containing reagent formaldehyde, it can be inferred that the constituent protomers are in intimate contact. The crosslinking patterns were unchanged in strains where either TatA or TatB was expressed without the other Tat components (de Leeuw et al., 2001), suggesting that the TatA and TatB homo-oligomeric complexes are fundamental structural units of the Tat pathway. There is circumstantial evidence that TatC may also function as an oligomer. Firstly, cross-complementation was observed between two otherwise inactive TatC point mutants (Buchanan et al., 2002). This indicates that there must be at least two TatC molecules in a functional Tat translocation pathway. Secondly, the genome of Halobacterium strain NRC-1, in addition to coding for a standard TatC-like protein, also specifies a molecule that is a tandem fusion of two TatC-like domains (Yen et al., 2002). This raises the possibility that, at least in this organism, TatC can function as a dimer.
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Several studies have used protein purification as a method to investigate the types of Tat complex found in the cytoplasmic membrane of E. coli (Bolhuis et al., 2001; Sargent et al., 2001; de Leeuw et al., 2002). In each case, the starting material was membranes from cells co-ordinately overproducing TatABC. Despite differences in the detergents used and the purification strategies employed, a reasonably consistent picture has emerged suggesting that in the resting Tat system the Tat proteins are found in two types of complex. The high apparent molecular masses of these complexes indicate that each contains multiple copies of the constituent Tat components as expected from the crosslinking studies described above. One complex has an apparent molecular mass around 700kDa and contains approximately equimolar amounts of the three Tat proteins, suggesting that each is present at around twelve copies per complex (Bolhuis et al., 2001; de Leeuw et al., 2002). We refer to this as the Tat(A)BC complex to indicate that it contains all the TatC and most of the TatB present in the membrane but only a minor proportion of the TatA protein. The major part of the TatA protein together with a small proportion of the TatB protein are found in a second type of complex which varies somewhat in apparent molecular mass between different purification protocols (350–650kDa) but is clearly always a very high order oligomer (Sargent et al., 2001; de Leeuw et al., 2002; and see discussion below). This we refer to as the TatA(B) complex to indicate that it is composed predominantly of TatA. Expression and purification of TatA by itself in the absence of TatB and TatC leads to a similar complex (de Leeuw et al., 2002; Porcelli et al., 2002). Sedimentation equilibrium measurements of this TatA complex purified in the detergent non-apolyoxyethylene dodecyl ether (C12E9) indicate that it has a molecular mass of 460kDa and therefore that it contains approximately 43 copies of TatA (Porcelli et al., 2002). The two types of Tat complex purified from E. coli correspond quite closely to those identified in pea thylakoids where blue native PAGE suggests the presence of separate Tha4 and Hcf106-cpTatC complexes (Cline and Mori, 2001). The apparent molecular mass of the Hcf106-cpTatC complex is 700kDa while that of the Tha4 complex decreases from a maximum value of 400kDa as the detergent to protein ratio is increased. The latter observation was interpreted as suggesting that detergent can disrupt the Tha4 oligomers (but see Section 3.5). The inference from purification studies that TatB and TatC form a complex is supported by several observations. Firstly, TatC is not stable in vivo in the absence of TatB (Sargent et al., 1999). Secondly, the tatB and tatC genes are more frequently genetically linked than either is to tatA. Thirdly, a chimera in which TatB is fused to the amino terminus of TatC, thus imposing both a physical association and an equimolar stoichiometry
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on the two Tat components, is functional (Bolhuis et al., 2001). It is also known that cpTatC protein can be crosslinked to Hcf106 in thylakoid membranes (Cline and Mori, 2001; a similar crosslink has not been observed between E. coli TatB and TatC). It is interesting to note that there is a discrepancy between the excess of TatB over TatC detected in the whole cell experiments described above and the TatA:TatB stoichiometry of 1:1 determined for the purified Tat(A)BC complex and inferred on the basis of the functional TatBC chimera (Bolhuis et al., 2001). Although this difference could be due to experimental errors in the whole cell experiments it is also possible, consistent with the purification of a TatA(B) complex, that only a portion of the TatB protein is permanently associated with TatC.
3.2. The Tat Transport Cycle Mechanistic analysis of the Tat translocase has until recently been dominated by in vitro studies on isolated plant thylakoids. However, the establishment of an in vitro transport assay for the E. coli Tat system (Yahr and Wickner, 2001; Alami et al., 2002) should now allow detailed biochemical analysis of the bacterial system to be undertaken. Tat precursor proteins are able to bind to E. coli inverted membrane vesicles or to plant thylakoids in the absence of p (Ma and Cline, 2000; Musser and Theg, 2000b; Alami et al., 2002). This binding depends both on a functional Tat consensus sequence on the precursor signal peptide and on the presence of Tat proteins in the membrane (Alami et al., 2002). If p is subsequently restored, the bound substrate is transported across the membrane indicating that the initial binding events are productive. These observations suggest that the initial step in Tat transport is the recognition of the Tat signal peptide by protein–protein interactions with the Tat translocase. Several lines of evidence indicate the thylakoid Hcf106-cpTatC complex, or the TatBC-containing equivalent in bacteria, is the site of initial precursor binding. Antibodies against Hcf106 and cpTatC, but not those against Tha4, inhibit precursor binding to thylakoid membranes (Cline and Mori, 2001). In addition, it has been shown that the bound precursor can be crosslinked to Hcf106 and cpTatC (Cline and Mori, 2001). More definitively, following detergent solubilisation of precursor-binding thylakoid membranes, the precursor protein co-migrates with the Hcf106-cpTatC complex during blue native PAGE (Cline and Mori, 2001). Intrinsic tryptophan fluorescence has been used to show that a Tat signal peptide elicits a specific conformational change in, and thus must bind to, the purified TatBC-containing complex from E. coli (de Leeuw et al., 2002).
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Circumstantial arguments suggest that TatC is most likely to contain the Tat signal peptide binding site. Firstly, it has been argued that only TatC shows sufficient sequence conservation to provide the specific binding site that is undoubtedly required to recognise the Tat signal peptide consensus motif (Berks et al., 2000a). Secondly, thylakoid complexes containing only Hcf106 do not bind precursor protein (Cline and Mori, 2001) and bacterial complexes containing TatB but not TatC show no evidence of interaction with Tat signal peptides (de Leeuw et al., 2002). Once a Tat precursor has bound to the Hcf106-cpTatC receptor complex, subsequent transport of the substrate protein across the thylakoid membrane requires both p and Tha4 (Cline and Mori, 2001). Since Hcf106-cpTatC and Tha4 are separate complexes in the resting membrane, the implication is that the two complexes dynamically interact during the transport cycle. Strong support for this contention comes from a recent study in which it was shown that Tha4 could be crosslinked to Hcf106 and to cpTatC but only if both p and a precursor protein were present (Mori and Cline, 2002). Similar crosslinks were detected if a synthetic Tat signal peptide was substituted for the full-length precursor. Since the signal peptide lacks a transportable domain, this suggests that association of the Tat proteins precedes the transport step. If transport of the substrate was allowed to go to completion before crosslinking was initiated, the contacts with Tha4 were no longer formed. This suggests that the Tat complexes disassemble once substrate translocation is completed. These observations have led, by analogy, to the working model of the E. coli Tat transport cycle shown in Figure 3. Precursor binding is proposed to cause a conformational change in the Tat(A)BC complex (apparently supported by the signal peptide binding studies mentioned above) that exposes binding sites for TatA. In an energy-dependent step driven by p, TatA then assembles onto the receptor complex to form the functional translocation site. Next the substrate is transported to the far side of the membrane. It is an open question whether transport requires further energetic input from p or whether the assembled translocase is in a ‘tense’ state that is dissipated by passage of substrate through the translocase. With substrate gone, the receptor complex returns to its resting conformation and the translocation site disassembles. Given this sequence of events, it is likely that TatA forms all, or the bulk of, the translocation channel – an inference further supported by structural analysis of an E. coli TatA(B) complex (Section 3.5). If this is the case, the Tat(A)BC receptor complex can be considered to have the role of gating the translocation channel. While Tat(A)BC and TatA(B) complexes are the predominant species purified from overproducing E. coli strains, a minor TatABC complex containing a large molar excess of TatA
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Figure 3 A working model for the Tat translocation cycle in E. coli. This figure is based on the scheme developed for plant thylakoid Tat transport by Mori and Cline (2002). The periplasmic side of the membrane is at the top of each panel. In the resting state TatA (Tha4 in thylakoids) and TatBC (Hcf106cpTatC in thylakoids) form separate complexes (note however that in E. coli some TatA appears to be associated with the TatBC complex and some TatB with the TatA complex). In an initial step the Tat signal peptide of a substrate protein is bound by the TatBC complex and results in a conformational change in the TatBC complex. This allows TatA to assemble onto the TatBC complex in an energy-requiring step driven by the transmembrane proton electrochemical gradient (p). In this model TatA is depicted as a pre-existing complex that associates with the TatBC receptor complex. However, it is also possible that TatA is present as smaller subcomplexes that assemble around the substrate protein/TatBC complex. The TatA-TatBC complex dissociates following transport of the substrate molecule. It is not clear whether further energetic input from the proton electrochemical gradient is required for the protein translocation step or during the disassembly of the translocation site. This uncertainty is denoted p.
over the other two components has also been isolated (de Leeuw et al., 2002). The low yields of this complex have so far prevented detailed characterisation but it is possible that it corresponds to an assembled Tat translocation site.
3.3. The TatA/B Protein Family Proteins of the TatA and TatB families are predicted to have a structure in which an amino terminal transmembrane helix is followed by an amphipathic helix and then a predominantly unstructured tail (Settles et
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al., 1997; Sargent et al., 1998a; Berks et al., 2000a; Fig. 4). This predicted structure is consistent with the experimentally determined secondary structure content of the purified E. coli TatA complex (Porcelli et al., 2002). Application of the positive-inside rule (von Heijne, 1992) suggests that the amphipathic helices of these proteins are located at the cytoplasmic side of the membrane. Such an orientation is supported by a study in which it was shown that E. coli TatA is accessible to proteases from the cytoplasmic but not the periplasmic side of the membrane (Porcelli et al., 2002). It is also known that proteases and antibodies located at the stromal (equivalent of cytoplasmic) side of the thylakoid membrane can interact with chloroplast Tha4 and Hcf106 proteins (Settles et al., 1997; Ma and Cline, 2000; Cline and Mori, 2001). It is, however, important to note that these accessibility studies cannot prove that the entire carboxy-terminal region is cytoplasmic/ stromal since they only map the locations of the epitopes of the antibodies used to detect the Tat proteins. An independent argument for a cytoplasmic location for the carboxy terminus of TatB protein comes from the observation that a chimera in which the carboxy terminus of TatB is fused to the amino terminus of TatC can substitute for TatB and TatC in E. coli Tat transport (Bolhuis et al., 2001). Since the amino terminus of TatC is located in the cytoplasm (Section 3.4) this experiment demonstrates that the carboxy terminus of TatB must also be in that compartment. Let us now examine the structural features of TatA and TatB proteins in more detail. It is important to realise that it is not straightforward to use multiple sequence alignments to identify those sequence features that distinguish a TatA protein from a TatB protein (Sargent et al., 1999; Berks et al., 2000a; Yen et al., 2002). This is because it is often unclear whether a given prokaryotic TatA/B homologue is functionally a TatA or a TatB protein. Such assignments can, however, be made with confidence if the analysis is restricted to the Proteobacteria (which includes E. coli) and the results are presented in Figure 4. TatB proteins (89 to 246 amino acids) are usually considerably longer than TatA proteins (57 to 89 amino acids). The additional length in the TatB proteins arises mainly from a greatly enlarged tail region but the carboxy terminal end of the amphipathic helical region is also extended. For both TatA and TatB proteins, the transmembrane helix shows higher sequence conservation at the ends than in the middle. If one assumes that between ten and twelve amino acids in a helical conformation are required to traverse the hydrophobic core of the lipid bilayer, it becomes apparent that the amino terminus of the transmembrane helix contain several conserved polar residues that are likely to be located within the phospholipid head group region rather than protruding into the periplasmic space. Indeed, the protonatable group at position 8 (normally glutamate in
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Figure 4 Primary structure analysis of the homologous TatA and TatB proteins from E. coli. Predicted secondary structure elements are shown above each protein sequence. a-helical regions are represented as cylinders and b-strands by arrows. It should be noted that only helices a-1 and a-2 in each protein are predicted with a high level of confidence (>80%). Filled triangles under the sequence indicate positions at which single mutations that seriously inhibit or abolish Tat transport have been identified (Hicks et al., 2003). Starred vertical arrows indicate the longest carboxy-terminal deletions of each protein that still permit Tat transport while grey vertical arrows indicated the shortest carboxyterminal deletion that shows signs of impaired transport (Lee et al., 2002). Various analyses of position-specific amino acid conservation amongst Proteobacterial TatA/E proteins (25 sequences) and TatB proteins (21 sequences) are presented beneath each sequence. Only regions that show appreciable sequence similarity are detailed. Note that three TatB sequences (those of Legionella pneumophila, Helicobacter pylori and Campylobacter jejuni) could not be aligned to the other Proteobacterial TatB proteins past residue 48 and are therefore excluded from the analysis of the sequence positions following this point. Also note that the multiple sequence comparison is somewhat biased towards E. coli-like sequences since the genome sequences of bacteria that are closely related to E. coli are over-represented in the sequence databases. [1] Highly conserved features amongst the Proteobacterial TatA/E and TatB proteins. Invariant amino acids are shown in black. Those amino acids or classes of amino acids found in at least 85% of sequences are shown in grey either in single letter code for specific amino acids or as ‘þ’ for a basic residue, ‘’ for an acidic residue, ‘c’ for a charged side chain and ‘s’ for an amino acid with a small side chain (glycine or alanine). [2] Highly conserved features of
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TatB and glutamine, histidine or lysine in TatA) could even be within the hydrocarbon core of the membrane and is a prime site for possible mechanistic proton exchange. However, recent site-directed mutagenesis experiments have shown that, at least for TatB, the presence of an acidic residue at this position is not important for function (Hicks et al., 2003). The only amino acid residue that is invariant between TatA and TatB proteins is a glycine (position 21) located between the predicted transmembrane and amphipathic helices. It has been suggested that this residue could act as a bend or hinge between the two helices (Berks et al., 2000a). Consistent with this, substitution of the invariant Gly21 in either TatA or TatB for a more bulky side-chain drastically reduces Tat-dependent protein translocation (Hicks et al., 2003). The glycine kink may be reinforced in TatB proteins by two invariant prolines, one immediately following the invariant glycine residue and the second four residues later. Experimental evidence for the functional importance of the first of these prolines comes from the observation that substitution by leucine renders TatB transport-incompetent (Weiner et al., 1998; Hicks et al., 2003). Interestingly, introduction of a proline residue at an equivalent position (Thr22) of TatA almost abolished transport activity (Hicks et al., 2003). The amphipathic helix of TatA and the corresponding amino-terminal portion of the amphipathic helix of TatB have a net positive charge. If the amphipathic helix lies along the membrane surface, this basicity would allow favourable electrostatic interactions with the phospholipid head groups. In TatA proteins, the amphipathic helix exhibits higher sequence conservation than the transmembrane helix whereas the reverse is true of TatB proteins. A series of conserved small chain amino acids are found on the non-polar side of the amphipathic helix of TatA but not of TatB. This could indicate that the amphipathic helix of TatA is involved in packing interactions with other helices. A phenylalanine residue (Phe39 for E. coli TatA) is absolutely conserved amongst proteins of the TatA family and is critical for function. Substitution of this residue for
Proteobacterial TatA/E and TatB proteins that are also found in plant chloroplast Tha4/9 and Hcf106 proteins respectively. [2a] Highly conserved features of Proteobacterial TatB proteins that are also found in plant chloroplast Tha4/9 proteins. [3] The polarity of the sequence position. This is only given if at least 75% of the proteins have the same polarity at the position. Residues are designated either ‘p’ for polar or ‘h’ for hydrophobic where glycine is regarded as being compatible with either class. [4] A measure of the amino acid conservation. For each position the most commonly occurring amino acid was identified. The fraction of sequences containing that amino acid was then determined where complete occupancy was assigned a value of ten. For example, if serine is found at position 5 in TatA in greater than 90% of the sequences then that position has the value 9.
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alanine not only abolishes Tat transport activity, but also disrupts transport when introduced on a plasmid into the wild-type strain (i.e. it has a dominant negative phenotype; Hicks et al., 2003). A phenylalanine residue is never found at an equivalent position in TatB proteins. There are, however, a string of conserved glutamates in the amphipathic helix of TatB (Glu49, Glu53 and Glu58 in the E. coli protein). A triple substitution of these residues with alanine renders TatB all but incompetent for transport activity. It has been speculated this negatively charged patch of TatB may be involved in twin arginine recognition during signal peptide binding (Hicks et al., 2003; see Section 3.2). The carboxy terminal tail regions of both TatA and TatB proteins are highly variable in length and exhibit no sequence conservation even within each family. This lack of sequence conservation suggests that it is only the general physicochemical properties of the tail region that could be of importance for function. In this respect, it is notable that the tails are unusually highly charged (which may just be a reflection of their random coil configuration) with an overall net negative charge. Studies in which E. coli TatA and TatB have been systematically truncated from the carboxy terminus show that the carboxy-terminal tails of both proteins can be removed without seriously compromising Tat transport (Lee et al., 2002). Nevertheless, these regions might still have non-essential auxiliary functions, for example reducing proton leakage or involvement in proofreading by providing a hydrophilic region that a potential substrate must traverse to gain access to the transport pore (Section 2.3.9). Some additional experimental insights into the differences between E. coli TatA and TatB proteins are available from a study in which protein engineering was used to exchange the amino terminal transmembrane helices of the two proteins using the conserved ‘hinge’ glycine as the fusion boundary (Lee et al., 2002). It was found that a chimera comprising the transmembrane helix of TatA and the carboxy terminus of TatB could restore very low level Tat transport activity to either a tatAE or a tatB mutant but not to a tatABE strain. Thus the fusion could act as either a TatA or a TatB molecule, provided that all other Tat proteins were present. In contrast, a fusion between the transmembrane helix of TatB and the carboxy terminus of TatA was unable to complement either tatAE or tatB mutants. These data suggest that the functional features that are unique to TatA reside in the transmembrane helix while those functional features that are exclusive to TatB are found in other regions of the molecule. This is of course the opposite of the conclusion that would be inferred from sequence analysis based on higher sequence conservation in functionally important parts of the molecule. Clearly, we are some way off fully understanding the structural basis for the functional differences between TatA and TatB.
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The function of the predicted transmembrane helix in TatA/B proteins has been probed in experiments where this region of the molecule was genetically removed from E. coli TatA and TatB proteins. The truncated TatA protein was found to be completely water soluble in vivo suggesting that the amino terminal helix is responsible for the membrane association of native TatA (de Leeuw et al., 2001). In contrast, truncated TatB exhibits a weak and peripheral association with the cytoplasmic membrane under the same conditions (de Leeuw et al., 2001). The water-soluble TatA domain has been purified and further characterised (Porcelli et al., 2002). The protein was found to be monomeric and to lack defined secondary structure. It was inferred from these observations that the transmembrane helix is required both for the homo-oligomerisation and folding of TatA. The purified soluble domain could be shown to insert into phospholipid monolayers at physiological surface pressures and to form helical structure in the presence of phospholipid bilayers (Porcelli et al., 2002). Since in native TatA this domain will be localised to the membrane by the transmembrane helix, these observations suggest that, notwithstanding the water-soluble nature of the purified truncated protein, the carboxy-terminal portion of TatA interacts with the membrane. Many of the sequence features found in Proteobacterial TatA and TatB are also conserved in their chloroplast functional homologues Tha4 and Hcf106 (Fig. 4). However, with the exception of the conservation of Phe20, it is noticeable that the transmembrane and hinge region of Tha4 exhibits the features of Proteobacterial TatB and not TatA proteins. In other words, the amino terminal portion of both Tha4 and Hcf106 look like Proteobacterial TatB. This raises the question of whether chloroplast Tha4 and Proteobacterial TatA really are functionally equivalent proteins. If they are, the similarities between Tha4 and Hcf106 suggest that the transmembrane region is unimportant in determining the functional differences between the two classes of proteins and, therefore, that the differential interaction of the two proteins with cpTatC are likely to be mediated primarily through the amphipathic helices. It would also imply that not all the sequence differences between Proteobacterial TatA and TatB proteins are linked to the different functions of the two proteins but rather represent genetic drift. It is of course also possible that our inability to clearly correlate the sequences of chloroplast Tha4 and Proteobacterial TatA molecules is because TatA/B became functionally differentiated in multiple lineages rather than in a unique event or, alternatively, that the evolutionary picture has been complicated by gene conversion in the chloroplast lineage. It is interesting to note at this point that an analysis of whole microbial genomes suggests that not all Tat systems utilise distinct TatA and TatB
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proteins. For example, Rickettsia prowazekii (a-Proteobacterium) and Staphylococcus aureus only possess a single protein of the TatA/B family while B. subtilis has three extremely similar TatA/B homologues that are likely to have arisen from a recent gene triplication (Berks, 2000a; Yen et al., 2002). This raises the possibility that the ancestral Tat system contained a single type of TatA/B protein.
3.4. The TatC Protein Family TatC is the largest of the essential Tat components. Sequence analysis strongly suggests that TatC has six transmembrane helices arranged such that the amino- and carboxy-termini of the protein are found at the cytoplasmic side of the membrane (Sargent et al., 1998a; Yen et al., 2002; Fig. 5). Gene fusions between TatC and compartment-specific reporter enzymes have confirmed the overall predicted topological orientation of TatC as well as the positions of most of the transmembrane helices (Drew et al., 2002; Gouffi et al., 2002). However, these studies also indicate that the polypeptide loop between the fourth and fifth predicted transmembrane helices is located at the periplasmic, rather than the predicted cytoplasmic, side of the membrane (Gouffi et al., 2002) and suggests that TatC has only four transmembrane helices (Fig. 5). TatC is the most conserved component of the Tat pathway and, as discussed above (Section 3.2), this has led to the suggestion that TatC contains the binding site that recognises the Tat consensus motif of the substrate signal peptide. TatC proteins can be divided into three quite distinct sequence subgroups, the largest of which comprises prokaryotic and chloroplast TatC molecules involved in ‘standard’ Tat protein transport. We restrict our sequence analysis to this group. The other two sequence groups are mitochondrial TatC molecules and the duplicated TatC-like protein found in many Archaea. The function of the TatC homologues in these two latter sequence groups is unclear and may not be identical to that of the canonical E. coli TatC molecule. For example, while plant mitochondria possess a TatC homologue, no obvious mitochondrially-targeted TatA or TatB homologues are found in plants and so the mitochondrial TatC may interact with an alternative set of effector proteins. Two groups have probed the functional importance of the observed TatC sequence conservation by site-directed mutagenesis of conserved residues in the E. coli TatC protein (Allen et al., 2002; Buchanan et al., 2002). Where equivalent mutations were made by both groups, there is some discrepancy as to the severity of the mutant phenotypes reported, possibly due to the
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Figure 5 Topological model of E. coli TatC indicating functionally important residues. For the purposes of this schematic diagram the predicted transmembrane regions are depicted as helices while extramembranous regions are shown as regions of random coil. In reality it is very likely that parts of the extramembranous regions will in fact form defined elements of secondary structure. The amino acids making up predicted transmembrane helices four and five are depicted using dashed lines. This is to indicate that a recent study using reporter fusions has questioned the existence of these helices (Gouffi et al., 2002). The protein sequence begins at position 2 because the initiator methionine of E. coli TatC is post-translationally removed (Buchanan et al., 2002). Highly conserved or invariant amino acids are shown in single letter code together with their residue number. Each of these residues has been subject to mutation to an alanine by Buchanan and co-workers (2002). Residues for which this substitution abolished Tat transport activity are shown on a grey background in a black border. Those residues where Tat transport activity was seriously compromised but not abolished by the mutations are shown on a grey striped background with a grey border. Residues where substitution had no significant effect on Tat transport are shown on a grey background.
slightly different experimental systems employed. In our laboratory, we replaced the 21 most highly conserved residues individually by alanine (Buchanan et al., 2002; Fig. 5). We found that many of these substitutions impaired Tat transport but identified only three mutations that were critical for Tat function. None of these three mutations exhibited dominance. Two of these residues, Phe94 and Glu103, are found in the first cytoplasmic loop (between helices two and three; Fig. 5). Phe94 is invariant with the exception that it is substituted by tyrosine in one chloroplast TatC molecule. Indeed, it
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was found experimentally that Phe94 could be functionally substituted by tyrosine but not by leucine (Buchanan et al., 2002). Glu103, in contrast, is completely invariant. It could not be functionally substituted by another acidic residue, aspartate. However, trace Tat transport activity was detected in our study if this amino acid was replaced by glutamine (Buchanan et al., 2002). Remarkably, and contradictorily, Allen and co-workers have reported that the same Glu103Gln mutation had no effect on Tat function (Allen et al., 2002). The third amino acid that we identified as crucial to Tat function by site-directed alanine substitution is highly conserved Asp211 in the final periplasmic loop (Fig. 5). This mutation (but not the Phe94Ala or Glu103Ala substitutions) was found to destabilise the TatB-TatC interaction in detergent solution (Buchanan et al., 2002), suggesting that Asp211 has a crucial structural role. Co-expression studies demonstrate crosscomplementation of the inactive asp211ala allele with either of the inactive phe94ala or glu103ala alleles. This indicates that the two groups of mutations affect different functions of TatC. The ability to crosscomplement two inactive tatC alleles also shows that each functional Tat translocation unit must contain at least two TatC molecules (Section 3.1). The tatC mutagenesis study of Allen and co-workers identified fewer substitutions that were severely affected in Tat transport (Allen et al., 2002). Specifically, they found that mutation of highly conserved Arg17 in the amino terminal tail (Fig. 5) to alanine completely blocked Tat transport (though we found only a mild defect associated with the same mutation; Buchanan et al., 2002) and mutation of highly conserved Pro48 in the first periplasmic loop (Fig. 5) to alanine severely affected Tat transport. Notwithstanding the discrepancies between the two studies, it is clear that only some of the highly conserved or invariant amino acids in TatC are required for Tat transport. In addition, it is clear that the cytoplasmically located amino terminal tail and first cytoplasmic loop are particularly important for Tat function (Fig. 5).
3.5. Mechanism of the Tat Transporter: Some Considerations The mechanism by which the substrate protein is transported by the Tat apparatus is still almost completely obscure. Nevertheless, certain constraints on the operation of the Tat translocase are known (or can be deduced) and it is possible to make some comments on possible mechanisms by which Tat transport may be achieved. Transport is known to be energised by p. Given that most periplasmic proteins are acidic it is, in principle, possible that the substrate might be
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translocated by electrophoretic movement through a passive channel. However, this idea is inconsistent with the observation that Tat transport in thylakoids is independent of the electrical component of p (Brock et al., 1995). In addition, Tat transport in thylakoids is two-to-threefold slower in D2O than in H2O suggesting that translocation involves proton- rather than charge-transfer events (Musser and Theg, 2000a). It is more likely, therefore, that p drives Tat translocation by countertransport of protons and substrate. It is possible that the transporter exerts a unidirectional mechanical force on the substrate to achieve transport. However, it is also conceivable that transport occurs by biased diffusion in which random Brownian motion in the direction of transport is permitted but retrograde movements are blocked. In such a model, p would have to be assigned a role in converting the apparatus from the output to input conformation. Current estimates indicate that translocation of a substrate protein through the thylakoid Tat apparatus takes about a minute (Teter and Theg, 1998). The length of this process suggests that transport probably involves the repetition of small translocation steps. It is almost certain that the Tat apparatus provides a channel across the phospholipid bilayer through which the substrate proteins are transported (Berks et al., 2000a). The nature of this channel is constrained both by the properties of the substrate proteins and by the need to preserve the ionic integrity of the cytoplasmic membrane during the transport process. The largest E. coli Tat substrates have minimum diameters of around 60A˚ (Berks et al., 2000a; Sargent et al., 2002), suggesting that the Tat apparatus must be able to form a channel of this size. Low resolution images of the TatA(B) complex in detergent solution obtained by electron microscopy show, in some orientations, a ring of macromolecular density surrounding a cavity of about 65–70A˚ in diameter (Sargent et al., 2001). Since TatA is thought to form the bulk of the transport channel (Section 3.2), it has been speculated that the cavity seen in these images corresponds to the substrate transport channel. Clearly, such a large hole would abolish the permeability barrier of the membrane to ions and small molecules. It can, therefore, be inferred that the channel is gated and only opens in the presence of a suitable substrate. One aspect of this gating procedure has already been identified since it has been established that a substrate protein is not directly recognised by the TatA channel. Instead, the substrate initially binds to a TatBC receptor complex thereby activating the complex to recruit the TatA channel (Section 3.2 and Fig. 3). It is, indeed, conceivable that the TatA channel is constitutively competent for the transport of macromolecules but only encounters such molecules at a significant frequency if they are delivered by TatBC.
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The process of gating encompasses not just the signal for opening but the physical mechanism by which the Tat system prevents co-transport of ions and other small solutes with the substrate protein. The problem of sealing the channel is exacerbated by the fact that the substrate proteins vary up to two-fold in diameter (Berks et al., 2000a) as well as in shape and surface properties. This variation in substrate properties prevents the use of a lockand-key type approach to achieve tight packing between channel and substrate. In principle, two other possibilities exist to achieve the required seal. The Tat system could use either a static channel structure with gates at either end, or a dynamic channel that changes conformation to maintain tight packing around the substrate during the transport process. The observation that the TatA(B) complex appears to adopt a ring structure in detergent solution (Sargent et al., 2001) would be consistent with the static channel model. However, it does not exclude the dynamic channel model because there may be dynamic elements within a static structural framework. It should also be taken into account that a likely driving force for the closure of a dynamic channel is the lateral pressure of the phospholipid bilayer. The TatA(B) complexes in detergent solution do not experience this bilayer pressure and might, therefore, be expected to adopt a channel-open conformation even if a dynamic channel mechanism appertains. A static channel model requires that only one of the gates at either end of the channel can be open at any given time. This implies that the substrate molecule must be positioned entirely within the channel and the cytoplasmic gate closed before the gate at the periplasmic side of the membrane is opened. Thus, for a static channel, but not for a dynamic channel, the channel structure must be large enough to accommodate the entire substrate molecule. Since one of the E. coli Tat substrates with a 60A˚ diameter is also 90A˚ in length (Sargent et al., 2002), a static channel would also need to be at least 90A˚ long. This is almost double the width of a membrane bilayer (50A˚) and therefore a static channel would be expected to extend a considerable distance beyond the membrane surface into the aqueous compartments on either side of the bilayer. This may be feasible since images of the purified TatA(B) complex show a minimal dimension of 90A˚ regardless of the orientation of the particle (Sargent et al., 2001). In summary, current data are insufficient to distinguish between static and dynamic models for the Tat channel structure. One might, however, ask whether a dynamic channel could maintain a sufficient level of membrane impermeability during the transport process. To prevent leakage of ions in general, it is probably not necessary for the channel to pack particularly tightly around the substrate protein because the ions will need to retain at least their inner hydration shell (most ions will
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therefore have effective diameters of around 10A˚). However, the problem of preventing proton conduction through the Tat channel is considerably more challenging because, in addition to diffusing as hydronium or hydroxyl ions, protons can be transported orders of magnitude faster than the diffusion limit by the Gro¨tthuss mechanism. This rapid transport of protons requires a network of hydrogen-bonded water molecules and/or protonatable amino acid groups and it is, therefore, necessary to prevent formation of such ‘proton wires’ if proton movement is to be restricted to manageable rates. It is reasonable to assume that the inner walls of the translocation channel are predominantly hydrophilic in nature in order that the energetic barrier to the movement of the polar substrate protein is minimised (note that while the walls are expected to be polar they probably lack amino acids with charged side-chains in order that strong ion-pairing interactions with substrate side-chains are avoided). It seems unlikely that this polar sleeve could pack tightly enough around the substrate protein to exclude a layer of water molecules. Instead, the channel would have to contain a structural feature that would disrupt proton wire formation. In the aquaporin family of water channels, it has been suggested that formation of a proton wire is prevented by imposing opposite orientations on the ordered water molecules in each side of the channel (Tajkhorshid et al., 2002). A more likely option for the Tat system would be a hydrophobic ring on the channel wall to exclude water molecules. This would have to be sufficiently thin so that the energetic barrier to transport of the polar substrate could be overcome by the mechanical force exerted on the substrate protein by the Tat machinery. One structural possibility to maintain a close packing around the substrate would be for the TatA molecules to undergo a dynamic polymerisation around the substrate molecule, such that the number of protomers in the pore matches the current maximal diameter of the substrate within the pore (Berks et al., 2000b; Mori and Cline, 2002). Such a mechanism implies that TatA does not form a single type of distinct complex but exists as either monomers or small oligomers in the membrane. We believe that this is not consistent with current data since purification of TatA in the detergent C12E9 gave a single, distinct, highly oligomeric species of 460kDa during analytical ultracentrifugation (Porcelli et al., 2002). Although apparent heterogeneity has been observed for TatA complexes solubilised in the detergent 3-[(3-cholamidopropyl)dimethylammonio]-1propanesulphonate (Chaps) during gel filtration chromatography (de Leeuw et al., 2001, 2002) and the molecular mass of Tha4 during blue nativePAGE changes as the detergent to protein ratio is varied (Cline and Mori, 2001), these observations could indicate either differences in protein conformation or instability in detergent solution rather than different
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polymerisation states. Indeed, consistent with a detergent-induced conformational change, it is notable that the Tha4 complex is a single distinct species at different detergent-to-protein ratios even though the apparent molecular mass varies over a factor of three (Cline and Mori, 2001). In any case, the observation that the apparent molecular mass of the Tha4 complex is highest (400kDa) at low detergent-to-protein ratios strongly suggests that the native membrane-associated complex is a very high order oligomer. An iris-like mechanism, in which the diameter of the channel is altered by co-ordinately changing the tilt angle of the helices, would be an alternative mechanism for providing a tight seal around a transported protein. Precedents for such mechanisms exist, including wholescale structural rearrangement in the opening of mechanosensitive ion channels (Betanzos et al., 2002; Perozo et al., 2002) or gating of the TolC and potassium channels (Andersen et al., 2002; Jiang et al., 2002) where one end of each mobile helix is fixed. These examples, however, operate essentially a single step transition. In contrast, the Tat channel would need to operate a series of very small step sizes to maintain a good seal around the continuously varying diameter of the transported substrate protein. The very high number of protomers in the TatA complex (in excess of 40; Section 3.1) might allow the use of small incremental changes in channel diameter.
4. BIOSYNTHESIS OF INTEGRAL MEMBRANE PROTEINS BY THE TAT SYSTEM It is now apparent that the Tat system does not just transport water-soluble proteins but is also involved in the biogenesis of certain integral membrane proteins comprising a single transmembrane helix and a large periplasmic domain (Sargent et al., 2002). The most compelling evidence for the existence of such Tat-dependent membrane proteins comes from the recently solved structure of E. coli formate dehydrogenase-N (Jormakka et al., 2002). Minimally, formate dehydrogenase-N comprises the subunits FdnG, FdnH, and FdnI (Section 2.3.8). The FdnG catalytic subunit binds the molybdopterin cofactor and is synthesised with a Tat signal peptide (Berg et al., 1991). FdnG forms a tight complex with the iron-sulphur clustercontaining FdnH protein at the periplasmic face of the cytoplasmic membrane. The structure shows that membrane attachment of the FdnGH dimer is mediated by a transmembrane helix located within the last 40 amino acids of FdnH and by protein–protein interactions (via both
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the transmembrane helix and the water-soluble domain of FdnH) with the integral membrane FdnI subunit (Jormakka et al., 2002). Sequence analysis suggests that 5 of the 26 probable E. coli Tat substrates are integral membrane proteins. In each case, as found for FdnH, the proteins possess a single carboxy-terminal transmembrane helix (Sargent et al., 2002). Perhaps the most notable of this set of ‘tail-anchored’ Tatdependent membrane proteins are the well characterised membrane-bound [NiFe] hydrogenases (Sargent et al., 2002). In general, such hydrogenases comprise a trimeric structure not unlike that of formate dehydrogenase-N in which the Large/Small subunit pair (Section 2.3.4) sits on an integral membrane protein. In the hydrogenases, the Small subunit possesses both the Tat signal peptide and the transmembrane helix. Early sequence analysis had predicted the existence of the tail-anchors on the peripheral subunits of membrane-bound formate dehydrogenase and hydrogenases (Berg et al., 1991; Wu and Mandrand, 1993; Berks et al., 1995b). However, attempts to obtain experimental evidence for their function in membrane association yielded conflicting results. For example, the peripheral subunits of the hydrogenases of W. succinogenes, R. capsulatus, and E. coli are still membrane-associated in strains lacking the fully integral membrane subunits (Gross et al., 1998; Magnani et al., 2000; Dubini et al., 2002). However, analogous experiments with the R. eutropha membrane-bound hydrogenase (Bernhard et al., 1997) and E. coli formate dehydrogenase-N (Stanley et al., 2002) led to the accumulation of soluble periplasmic proteins with no strong membrane affinity. While non-specific proteolytic degradation of the carboxy-tail anchors could not be discounted in these experiments, it should perhaps be considered that periplasmic intermediates play a role in the assembly of these tail-anchor complexes (Sargent et al., 2002). In such a ‘periplasmic re-entry’ model, the entire enzyme, including the carboxy-terminal helix, would be exported to the periplasm. The carboxy-terminal tail-anchor would then associate with the inner membrane from the periplasmic side. An alternative mechanism for the assembly of membrane proteins by the bacterial Tat translocase would envisage the transmembrane helices acting as ‘stop-transfer’ domains in a manner analogous to membrane integration by the SRP/Sec pathway. The hydrophobicity of the carboxy-terminal helix would cause a stalling of the transport process as it passed through the Tat pore, with the extreme carboxy-terminus of the helix still at the cytoplasmic side of the membrane. The now channel-located hydrophobic helix would then move laterally into the lipid bilayer. Such a stop-transfer model could require a channel-clearing mechanism similar to that employed by the Sec-system during transmembrane helix integration
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(Sargent et al., 2002). The recently discovered YidC protein is thought to play this role for the Sec apparatus (Luirink et al., 2001; Section 1). However, a role for YidC in the integration of Tat-dependent membrane proteins has not yet been tested. Tat-dependent integral membrane proteins are not restricted to prokaryotes. The thylakoid membrane-bound Pftf zinc-dependent proteases of plant chloroplasts are targeted and integrated in a Tat-dependent manner (Summer et al., 2000). The Pftf proteins have a topology analogous to that of the E. coli tail-anchored Tat substrates, with a large hydrophilic domain at the trans side of the membrane followed by a single hydrophobic transmembrane anchor. Intriguingly, site-directed mutagenesis, and even complete deletion, of the Pftf Tat signal peptide did not severely impair Tattargeting and membrane integration of the Pftf protein in vitro (Summer et al., 2000). This suggests that the Tat-dependent transmembrane segment of Pftf constitutes a novel Tat-targeting element. The Tat-targeting activity of the carboxy-tails of bacterial Tat substrates has not been tested directly. However, it should be noted that membrane transport and integration of the W. succinogenes hydrogenase was completely blocked when the conserved arginine pair in the Tat signal peptide was replaced by glutamines (Gross et al., 1999), indicating an obligatory role for the signal peptide in Tattargeting of this enzyme. In addition to carboxy-tail proteins, the Tat system of chloroplasts, and probably also bacterias, assemble a second type of integral membrane protein. This is the Rieske iron-sulphur protein of the cytochrome b6f and bc1 complexes. As discussed (Section 2.3.3) this has an uncleaved Tat signal peptide that functions as an amino-terminal transmembrane helix to anchor the periplasmic iron-sulphur cluster domain to the membrane. It has been observed that if a Tat signal peptide is fused to an SRPtargeted integral membrane protein membrane targeting remains Tatindependent, suggesting that the SRP-targeting overrides Tat-targeting (Cristo´bal et al., 1999). This is quite logical since the transmembrane regions of the mature protein require the co-translational mode of membrane integration provided by SRP to avoid aggregation in the aqueous cytoplasmic environment. Nevertheless, this observation raises the question of how the transmembrane helices of Tat proteins avoid being targeted to the SRP pathway. In the case of Rieske-type signal-anchors, the reduced hydrophobicity of the Tat signal anchor when compared to standard transmembrane helices probably prevents SRP recognition (Cristo´bal et al., 1999). However, we can find no significant differences in the overall hydrophobicity index between other Tat-dependent and Sec-dependent transmembrane segments.
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A key difference between membrane protein integration by the Tat and Sec pathways is that the Tat mechanism is apparently post-translational (Summer et al., 2000). As a result, it will be necessary to prevent the hydrophobic transmembrane regions of the Tat precursor proteins from engaging in self-aggregation or premature interactions with the cytoplasmic face of the cell membrane prior to membrane integration. Perhaps masking of exposed transmembrane helices is another role for enzyme-specific accessory proteins such as, in E. coli, FdhE (formate dehydrogenases; Section 2.3.1) and HyaE/F/HyB (hydrogenases; Section 2.3.4).
5. CONCLUDING REMARKS It is just over five years since the Tat system was first experimentally demonstrated in bacteria (Settles et al., 1997; Santini et al., 1998; Sargent et al., 1998a; Weiner et al., 1998) and the length of this review testifies to the massive progress that has been made in the meantime. It is, however, safe to say that what we do not know about the Tat pathway still considerably exceeds that which we do know. We realise that Tat is unique in moving folded proteins across an ionically tight membrane but we do not yet know the structural or operating principles underlying the Tat mechanism. It is becoming apparent that the transport cycle of at least some Tat substrates involves cytoplasmic factors, including ‘cofactor chaperones’, and also that the Tat system is involved in the biogenesis of integral membrane proteins. Molecular details of these processes are eagerly awaited. There is also bound to be a good deal of interest in whether the involvement of the Tat pathway in bacterial pathogenesis is a general phenomenon, as well as in determining the molecular basis for the dependence of virulence on Tat in each pathogen. Finally, it is likely that the unique operating features of the Tat pathway will attract biotechnological applications in the near future.
ACKNOWLEDGEMENTS We thank members of our laboratories past and present for their contribution to our research on protein transport and our many colleagues with whom we have discussed the Tat system. Research in the authors’ laboratories has been or is currently supported by the Biotechnology and
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Biological Sciences Research Council, the Commission of the European Community, the Leverhulme Trust, the Wellcome Trust, the John Innes Centre, the University of East Anglia, the University of Oxford and The Royal Society. Ben Berks is RJP Williams Senior Research Fellow at Wadham College, Oxford. Tracy Palmer and Frank Sargent are Royal Society University Research Fellows.
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Microbial Globins Guanghui Wu, Laura M. Wainwright and Robert K. Poole Department of Molecular Biology and Biotechnology, The University of Sheffield, Firth Court, Western Bank, Sheffield S10 2TN, England, UK
ABSTRACT Globins are an ancient and diverse superfamily of proteins. The globins of microorganisms were relatively ignored for many decades after their discovery by Warburg in the 1930s and rediscovery by Keilin in the 1950s. The relatively recent focus on them has been fuelled by recognition of their structural diversity and fine-tuning to fulfil (probably) discrete functions but particularly by the finding that a major role of certain globins is in protection from the stresses caused by exposure to nitric oxide (NO) – itself a molecule that has attracted intense curiosity recently. At least three classes of microbial globin are recognised, all having features of the classical globin protein fold. The first class is typified by the myoglobin-like haemprotein Vgb from the bacterium Vitreoscilla, which has attracted considerable attention because of its ability to improve growth and metabolism for biotechnological gain in a variety of host cells, even though its physiological function is not fully understood. The truncated globins are widely distributed in bacteria, microbial eukaryotes as well as plants and are characterised by being 20–40 residues shorter than Vgb. The polypeptide is folded into a two-over-two helical structure while retaining the essential features of the globin superfamily. Roles in oxygen and NO metabolism have been proposed. The third and best understood class comprises the flavohaemoglobins, which were first
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discovered and partly characterised in yeast. These are distinguished by the presence of an additional domain with binding sites for FAD and NAD(P)H. Widely distributed in bacteria, these proteins undoubtedly confer protection from NO and nitrosative stresses, probably by direct consumption of NO. However, a bewildering array of enzymatic capabilities and the presence of an active site in the haem pocket reminiscent of peroxidases hint at other functions. A full understanding of microbial globins promises advances in controlling the interactions of pathogenic bacteria with their animal and plant hosts, and manipulations of microbial oxygen transfer with biotechnological applications.
Abbreviations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 257 1. Globins – definition and the classical view . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 257 1.1. Myoglobin and haemoglobin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 257 1.2. Enzymes, or not? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 258 1.3. Distribution and classification of microbial globins . . . . . . . . . . . . . . . . . . . . 258 2. The Vitreoscilla globin, Vgb, and other single domain myoglobin-like globins . . . . 260 2.1. Molecular characteristics of single domain globins. . . . . . . . . . . . . . . . . . . . 260 2.2. Vitreoscilla haemoglobin (Vgb) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 262 3. Truncated globins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268 3.1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 268 3.2. The two-over-two a-helical fold . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 3.3. Conserved residues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 3.4. Coordination of the haem. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 3.5. Ligand binding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 3.6. Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 273 4. Flavohaemoglobins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 4.1. Discovery of the flavohaemoglobins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 4.2. Composition of Hmp and other flavohaemoglobins . . . . . . . . . . . . . . . . . . . 279 4.3. Structure of the flavohaemoglobins Fhp and Hmp . . . . . . . . . . . . . . . . . . . . 280 4.4. Enzymic properties of Hmp . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 282 4.5. Functions of Hmp . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 285 4.6. Regulation of hmp transcription . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287 4.7. Phenotypes of hmp mutants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 291 4.8. NO-Detoxifying activities of Hmp and other flavohaemoglobins . . . . . . . . . . 291 4.9. Flavohaemoglobins of yeasts and fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . 296 5. Evolution of globins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298 6. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 299 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 300 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 300
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ABBREVIATIONS DTT GSNO Hcy NOC-5 NOS SIN-1 SNP TrHb
dithiothreitol S-nitrosoglutathione homocysteine 3-[2-hydroxy-1-(1-methylethyl)-2-nitrosohydrazino]1-propanamine Nitric oxide synthase 3-morpholinosydnonime hydrochloride sodium nitroprusside truncated haemoglobin
1. GLOBINS – DEFINITION AND THE CLASSICAL VIEW 1.1. Myoglobin and Haemoglobin Any standard textbook of biochemistry devotes several pages to haemoglobin and myoglobin (note the use of the singular in each case) – arguably the best studied of all proteins. In contrast, standard textbooks of microbiology rarely mention these proteins. This is despite the recognition by Keilin over half a century ago of the presence of globin-like proteins in yeast (Keilin, 1953), other fungi (Keilin and Tissieres, 1953) and protozoa (Keilin and Ryley, 1953). In fact, Keilin (1953) pays due credit to what is probably the first description of haemoglobin in yeast, that made by Warburg and Haas twenty years earlier. Keilin examined several strains of baker’s yeast but found in only two ‘‘the weak absorption band between 580 and 583 nm’’ described by Warburg. Keilin observed that this band (which was presumably the a-band of the oxygenated globin) ‘‘is visible only during very strong aeration, and that it disappears when aeration is interrupted or when the yeast suspension is treated with carbon monoxide’’. The band ‘‘can be observed even in a moderately aerated suspension provided it is kept at about 0 C’’. Keilin argued that this band did not arise from Warburg’s ‘respiratory ferment’ (i.e. the terminal oxidase) and that the ‘‘band at 583 nm belongs to a ferrous oxygenated haemoprotein such as oxyhaemoglobin’’. In this important paper, Keilin not only drew attention to the apparently haphazard distribution of haemoglobin in invertebrates and microorganisms and coined the term ‘non-circulating’ globin, but raised a point that, many decades later, still warrants consideration, as follows. ‘‘The mode of life of micro-organisms, their small size, their high respiratory activity and the low concentration of haemoglobin they contain preclude the
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possibility that their haemoglobin may also serve as a temporary store of oxygen.’’ This prophetic comment has been followed by the realisation that some microbial globins, which are much more widespread than perhaps Keilin anticipated, do indeed have alternative functions, particularly in enzymic activities related to protection from the potential damage caused by nitric oxide (NO) (Poole and Hughes, 2000; Ascenzi et al., 2001) and its congeners and perhaps in facilitating oxygen delivery.
1.2. Enzymes, or Not? Although myoglobin is generally considered a physiologically important oxygen store and oxygen carrier, its function has recently been re-evaluated. It is suggested that myoglobin can also act as an intracellular scavenger of NO (Brunori, 2001). Since this gas inhibits mitochondrial respiration, myoglobin may serve to protect respiration in tissues where myoglobin is found, such as in heart and skeletal tissue. Recently, the Ascaris haemoglobin, famed for its high oxygen affinity (binding oxygen nearly 25,000 times more tightly than does human haemoglobin), has been demonstrated enzymatically to consume oxygen in a reaction driven by NO. It is proposed that this reaction, involving the haem and a thiol, serves to maintain the perienteric fluid of the nematode hypoxic (Minning et al., 1999). Thus this globin ‘detoxifies’ oxygen. A further example of an enzymic function is that described for the globin, DHP, of the marine worm Amphirite ornate (Lebioda et al., 1999; Lebioda, 2000), which catalyses the rapid oxidative dehalogenation of polyhalogenated phenols in the presence of H2O2.
1.3. Distribution and Classification of Microbial Globins Despite detailed biochemical investigations of the haemoglobin (YHB1) of yeast in the 1970’s by Chance and others, our knowledge of microbial globins has been slow to develop and information on their probable physiological functions is only now emerging. Many globin genes from microbes, especially from bacteria, have now been sequenced and in some cases structures have been determined, revealing at least three quite distinct major classes of bacterial globins (Fig. 1). The first class is typified by the Vitreoscilla globin (Vgb or VtHb or Vhb) Although once described as a ‘‘soluble cytochrome o’’, on the basis of its O2reducing ability and the presence of haem B, sequencing of the vgb gene and the protein product showed it to be a true globin homologue of 153 amino
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Figure 1 The domain organisation of microbial globins. The three major classes are described in the text. Class 1B in this classification separates the C. jejuni globin (Cgb) because of its monomeric structure (G. Wu and R. K. Poole, unpublished). Note that this globin is distinct from the truncated globin in this bacterium. In class 1, the possible participation of soluble globin reductase(s) is indicated. Species names underlined indicate those from which the globin structure has been solved. FNR is plant ferredoxin-NADPþ reductase and is not to be confused with the the Fnr protein of E. coli.
acids, having the persistent ‘globin fold’ (three-on-three a-helical sandwich) of the polypeptide and the conservation of key amino acid residues in and around the haem pocket. Considerable interest has been directed at this protein even though its physiological function is not fully understood. The second are the truncated globins in which the textbook picture of the globin fold is dramatically altered by a simpler two-over-two a-helical structure while retaining key residues required for haem binding and interactions with small ligands. The functions of these globins, which are widely distributed in eukaryotic and prokaryotic microbes, as well as plants, are poorly understood. Members of the third class, the flavohaemoglobins, are distinguished by the presence of an additional reductase domain that contains flavin (FAD) and provides a binding site for NAD(P)H. Much has been learned in the past decade of these fascinating proteins, which are now widely acknowledged to be involved in resisting nitric oxide and related stresses.
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An unusual protein has been reported in yeast in which a putative haembinding domain of about 140 amino acids matches fairly well a sequence motif for globins proposed by Moens et al. (1996). The flanking C- and N-terminal portions do not show any significant similarities with known proteins, making this protein apparently unique (Sartori et al., 1999). Deletion of the gene elicits no phenotype. Although transcription of the gene (YNL234w) is weak under normal aerobic growth conditions, it is increased on O2 or nitrogen source limitation and various stresses. Globins are widely distributed in vertebrates, invertebrates, ciliated protozoa, eukaryotic algae, yeasts and other fungi, and higher plants (leguminous and non-leguminous). Some of these are outside the scope of this chapter but are described and reviewed elsewhere (Takagi, 1993; Andersson et al., 1996; Watts et al., 2001; Weber and Vinogradov, 2001). The best studied of these is leghaemoglobin (Appleby, 1984) or ‘symbiotic haemoglobin’ found in nitrogen-fixing nodules on legumes as the result of symbiotic association of rhizobia with the plant. The protein delivers oxygen to the bacteroids at very low poised O2 concentrations compatible with nitrogen fixation and respiration catalysed by the bacteroid oxidases, which have very high O2 affinities (Poole and Cook, 2000). We do not cover these plant globins since, although in the past it has been thought that the haem is synthesised by the symbiont bacteria, this is now doubted (Santana et al., 1998). The non-symbiotic haemoglobins of plants occur in the roots, stems or germinating seeds of many plants at low concentrations and their possible functions are under intense scrutiny (Andersson et al., 1996; Nie and Hill, 1997; Watts et al., 2001). To the best of our knowledge, this is the first comprehensive review of microbial globins. Additional references, particularly to the older literature are found in previous reviews on bacterial respiration and oxygen metabolism (Webster, 1987; Poole, 1994; Poole et al., 1994a; Poole and Cook, 2000) and in Wei et al. (2003).
2. THE VITREOSCILLA GLOBIN (VGB) AND OTHER SINGLE DOMAIN MYOGLOBIN-LIKE GLOBINS 2.1. Molecular Characteristics of Single Domain Globins Vitreoscilla is an obligately aerobic bacterium that grows in low oxygen environments such as stagnant ponds and rotten vegetables. Vitreoscilla haemoglobin (Vgb) was the first bacterial globin to be isolated. The protein
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is normally a dimer consisting of two identical subunits (but see Section 2.2.1) and two protohaems IX per molecule (Wakabayashi et al., 1986; Webster, 1987). Its physiological role has long been thought to be similar to that of myoglobin in that it could transport oxygen to terminal oxidases. In agreement with this proposal, Vgb is effective in promoting microaerobic growth and product formation in heterologous hosts (see Section 2.2.6 for details). Interestingly, a chimeric protein comprising Vgb and the flavoreductase domain from the Ralstonia eutropha flavohaemoglobin Fhp was found to be able to relieve nitrosative stress in E. coli. It was proposed that when Vgb is in the dimeric form, it participates in oxygen transport; when it is in a monomeric form, it could associate with a reductase and relieve nitrosative stress (Kaur et al., 2002). Eight other single domain bacterial globins were revealed in a BLAST search using the Vgb sequence against 98 finished and unfinished microbial genomic sequences at NCBI (National Center for Biotechnology Information) (Fig. 2). Interestingly, the one most similar to Vgb is from an obligate anaerobe, Clostridium perfringens. The next closest relative is Campylobacter jejuni Cgb (49% identity) from another obligate microaerobe. There are two similar globin sequences in Rhodopseudomonas palustris. Presumably, C. perfringens does not require an oxygen transport protein and haemoglobin must play other roles, e.g. resistance to nitrosative stress. These proteins should possess the typical globin fold, and they share sequence identities with Vgb ranging from 22% (from Thermobifida fusca) to 66% (from C. perfringens). All contain the conserved His-F8, Phe-CD1 and Tyr-B10 residues. However, the residue at position E7 is less conserved. Gln-E7 is located out of the haem pocket in Vgb (Tarricone et al., 1997b) and is not involved in ligand binding (Dikshit et al., 1998). In addition, most of those residues identified in R. eutropha flavohaemoglobin (Fhp) that are involved in interaction with lipids (Ollesch et al., 1999) are conserved in these globins (Fig. 2) This may suggest a possible interaction of these globins with membrane phospholipids.
2.2. Vitreoscilla Haemoglobin (Vgb) 2.2.1. Is Vitreoscilla Haemoglobin Equivalent to the Soluble ‘‘Cytochrome o’’? Vgb is undoubtedly the best-studied single domain bacterial haemoglobin. Vitreoscilla (named V. stercoraria in later papers) contains, in addition to a haemoglobin, a membrane-bound cytochrome o that functions as a
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Figure 2 ClustalW alignment (MacVector version 7.0, Oxford Molecular) of single-domain bacterial haemoglobins identified by a BLAST search using Vgb against the microbial genomic protein database at NCBI. Identical or conserved replacements are in shadowed boxes. The residues underneath the alignment are those that interact with lipid in R. eutropha Fhp (Ollesch et al., 1999). The last 14 Cterminal residues of the T. fusca globin are not shown. Tyr-810, Phe-CD1, E7 and His-F8 are at positions 50, 64, 73 and 106, respectively.
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terminal respiratory oxidase (for more details, see reviews by Poole (1983,1988). We consider it well established that the so-called soluble ‘‘cytochrome o’’ from Vitreoscilla was in fact the globin Vgb (Wakabayashi et al., 1986; Webster, 1987). However recently, Giangiacomo et al. (2001) have pointed out that the soluble ‘‘cytochrome o’’ studied by Webster and others before the protein was sequenced appears different from the cloned and heterologously expressed haemoglobin in terms of amino acid composition and kinetic data. Therefore, Giangiacomo et al. (2001) called the cloned Vgb the recombinant Vitreoscilla haemoglobin to distinguish it from the protein isolated directly from Vitreoscilla. Nevertheless, the vgb gene sequenced by Khosla and Bailey (1988a) ‘‘is in complete agreement with the known amino acid sequence of the protein’’ and so the source of any difference between the earlier ‘native’ and later recombinant forms of Vgb cannot be readily explained. The calculated molecular mass based on amino acid sequence is 15,773 Da, whereas the result obtained from SDS gel analyses is approx. 13,000 Da for each identical subunit (Webster, 1987), slightly smaller than the calculated value. Previously, the soluble ‘‘cytochrome o’’ was purified as a dimer (Tyree and Webster, 1978). However, according to the sedimentation equilibrium analyses carried out by Giangiacomo et al. (2001) Vgb is mainly present in the monomeric state (equilibrium dimerisation constants, 6 102 and 1 102 M1 for deoxygenated and carbonylated derivatives respectively at pH 7.0 and 10 C) in the presence of DTT, which prevents the formation of the disulfide-linked dimer. Ligand binding causes only a slight shift of the equilibrium and is unlikely to have any functional relevance in vivo (Giangiacomo et al., 2001). On the other hand, according to gel filtration analysis, Kaur et al. show that Vgb is dimeric at high concentration (10 mg ml1, 6.3 104 M, according to the monomer molecular mass) and partially monomeric at lower concentration (0.05 mg ml1, 3.1 106 M according to the monomer molecular mass) (Kaur et al., 2002). There are some obvious discrepancies between the reported amino acid composition of the purified protein (Tyree and Webster, 1978) and the amino acid sequence (Wakabayashi et al., 1986); for example, there is one cysteine in the amino acid sequence, but no cysteine was found in the amino acid analysis. However, the cysteine analysis may be problematic and any impurity in the protein sample will affect the amino acid composition analysis. Perhaps the most striking difference between the preparations is the oxygen dissociation rate constant (koff), but the reported values have been measured in two ways. The value calculated according to the effect of oxygen concentration on the rate of binding oxygen (the ‘on’-rate) by the NADH-reduced enzyme is 5,600 s1
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(Orii and Webster, 1986). However, the kinetics obtained by mixing oxygenated Vgb with dithionite are biphasic between 10 and 40 C and the values are 4.2 s1 and 0.15 s1 respectively at pH 7.0 and 20 C (Giangiacomo et al., 2001; see 2.2.2). Moreover, the recombinant Vgb displays anti-cooperative binding to cyanide, azide, thiocyanate, and imidazole (Bolognesi et al., 1999), whereas the soluble ‘‘cytochrome o’’ binds cyanide non-cooperatively (Tyree and Webster, 1978). The second order rate constants of O2 combination are not so different between two groups: 2 108 M1s1 (Giangiacomo et al., 2001) and 7.8 107 M1s1 (Orii and Webster, 1986).
2.2.2. Biochemical Characterisation of Vgb and Clues to its Function All the early spectroscopic work in Webster’s laboratory was done with Vgb directly isolated from Vitreoscilla rather than with the recombinant protein. Much of the early work has been comprehensively reviewed (Webster, 1987), some of which focussed on the ligand-binding properties of the protein and the information that such data provided on cooperativity between the subunits in the presumed dimer. While cyanide binding to the oxidised protein suggested that the haems behaved independently, CO binding to the Fe(II) protein suggested cooperativity. The haems were reported to have very different midpoint potentials (þ118, 122 mV) (Webster, 1987). A high molecular mass form (50 kDa) was observed when the anaerobic Fe(II) protein was titrated with O2 at 0 C. This species (‘‘Compound D’’) has 1 O2 per 4 haems. Further addition of O2 gave a dimeric ‘‘oxygenated’’ compound but with oxidation of 50% of the haem. Hydrogen peroxide is the final product during the oxidation of NADH by ‘‘cytochrome o’’ (Webster, 1975), as in Hmp (Mills et al., 2001). Note that recent studies on the monomer-dimer equilibrium and O2-binding properties of the recombinant protein suggest that, when expressed in Escherichia coli, the protein is predominantly monomeric (Giangiacomo et al., 2001). Later work described ligand-binding properties of the Fe(II) recombinant Vgb with CO (Dikshit et al., 1998) and the Fe(III) recombinant Vgb with cyanide, azide and other ligands (Bolognesi et al., 1999). The kinetics of O2 recombination in the monomer measured by laser photolysis give a simple, second-order rate constant of 2 108 M1 s1 (Giangiacomo et al., 2001). The kinetics of O2 release, however, are biphasic between 10 and 40 C. Two different conformers may account for this biphasicity, possibly differing in the stabilisation of the haem-bound ligand by Tyr-B10. The existence of two conformers in thermal equilibrium in
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Vgb is mirrored in Hmp, for which both kinetic data (Gardner et al., 2000b) and resonance Raman data (Mukai et al., 2001) support such heterogeneity (see Section 4.3). The O2 affinity of Vgb, as estimated from the kinetic properties, is in the nM range. As noted by Giangiacomo et al. (2001), this indicates that Vgb is not likely to be involved in O2 delivery or storage. Nevertheless it has often been suggested that the growth- and metabolism-stimulating effects of Vgb expression are attributable to the more effective delivery of O2 within the host cells (see Section 2.2.6). An alternative proposal, namely that Vgb has functions akin to the flavohaemoglobins (Section 4), albeit requiring association with a separate reductase, gains support from the finding that a chimeric construct (Vgb fused to the reductase domain of Fhp) counters nitrosative stress (Kaur et al., 2002). Immunogold labelling reveals Vgb to be bound to the cytoplasmic side of the membrane (Ramandeep et al., 2001). Recently, the location has been suggested to be specifically one of the terminal oxidases (Park et al., 2002). This suggests that the previous finding that Vgb has both cytoplasmic and periplasmic locations (Khosla and Bailey, 1989a) may be an artefact arising from over-expression of the protein. Vgb stimulates the ubiquinol-1 oxidase activity of everted Vitreoscilla membranes by 68%, consistent with a membrane association. Inclusion of cytochrome bo in proteoliposomes increases the Vgb binding affinity 2.4- to 6-fold. The results suggest a direct, perhaps temporary (Wittenberg and Wittenberg, 1990), interaction of Vgb with the respiratory chain, most likely the cytochrome bo branch (Ramandeep et al., 2001). Consistent with this idea is that Vgb was reported to complement a mutant of E. coli defective in both terminal oxidases (Dikshit et al., 1992). There are at least two possibilities: one is that Vgb presents O2 to the oxidase for reduction (but see the above concerns over the very high O2 affinity). A second is that Vgb in association with the membrane, binds and transfers electrons from membrane-integral components to O2 (Wittenberg and Wittenberg, 1990).
2.2.3. Crystal Structures The ferric homodimeric Vgb was the first bacterial haemoglobin to be crystallised (Tarricone et al., 1997a,b). The three-dimensional structure conforms to the well-defined globin fold. However, the C and E helices are disordered, residues E7-E10 do not adopt the usual a-helical conformation, and Gln-E7 is located out of the haem pocket. Instead, residues Phe-CD1, Pro-E8 and Leu-E11 fill the distal site of the haem pocket. Binding of azide
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causes conformational rearrangement at the haem pocket distal site i.e. residue Pro-E8 moves out of the haem pocket and there are shifts of Gln-E7 to Ala-E10 by 2–4 A˚. The three dimensional structures of the thiocyanate and imidazole derivatives of recombinant ferric Vgb have also been determined (Bolognesi et al., 1999). In agreement with the crystallographic studies, site-directed mutagenesis shows that Gln-E7 does not appear to stabilise the haem iron-bound dioxygen through hydrogen bonding (Dikshit et al., 1998). The quaternary structure of Fe(III) Vgb and of its azide, thiocyanate and imidazole derivatives is unique in that the protein is dimeric in the crystal lattice. The association of the two subunits appears to be weak: they interact through van der Waals contacts at a very small molecular interface area of only 434 A˚2. It has been suggested that the disordered CE region is a potential site of interaction with the FAD/NADH reductase partner (Tarricone et al., 1997b) (see below).
2.2.4. A Putative Reductase for Vgb Vgb is reducible by NADH via a separate reductase isolated from Vitreoscilla (Gonzales-Prevatt and Webster, 1980; Kroneck et al., 1991; Jakob et al., 1992). It has one FAD per mol (Mr 61,000) and no iron. However, this enzyme reduced only partially purified ‘‘cytochrome o’’ not the pure ‘‘cytochrome o’’ (i.e. Vgb) and additional protein factor(s) appear to be needed (Gonzales-Prevatt and Webster, 1980). This reductase ability, if active in vivo, could confer on Vgb the oxidase, oxygenase and NOdetoxifying abilities seen in the flavohaemoglobins (Section 4) which possess an ‘‘on-board’’ reductase domain.
2.2.5. Regulation of vgb Gene Expression Expression of the vgb gene (Dikshit and Webster, 1988; Khosla and Bailey, 1988a) has been studied in Vitreoscilla sp., strain C1 (Dikshit et al., 1989), and in the heterologous hosts E. coli, Pseudomonas aeruginosa, Pseudomonas putida and Azotobacter vinelandii (Khosla and Bailey, 1988b; Dikshit et al., 1990; Joshi and Dikshit, 1994). In all these organisms, the vgb promoter is preferentially activated in response to O2 limitation according to the results obtained by analysing mRNA levels and using gene fusion techniques. However, expression of vgb is switched off under anaerobic conditions (Khosla and Bailey, 1989b). The anaerobic gene regulator Fnr was suggested to be required for vgb expression since Vgb
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was not expressed in an fnr mutant background (Joshi and Dikshit, 1994; Tsai et al., 1995). However, the proposed Fnr-binding site 50 TTTTAAGAGGCCAAT 30 is not similar to the full Fnr consensus sequence 50 TTGAT. . ..ATCAA 30 , where, the G and C (underlined) are considered to be crucial in interacting with the Fnr protein (Spiro and Guest, 1991). Catabolite repressor protein CRP may also be needed for full expression either directly or indirectly (Khosla and Bailey, 1989b, Joshi and Dikshit, 1994). Since Vgb is now proposed to participate in protecting cells from nitrosative stress (Kaur et al., 2002), the regulation of vgb expression by nitrosative stress is certainly worth pursuing.
2.2.6. Heterologous Expression of Vgb and Biotechnological Implications Vgb was originally reported to improve the microaerobic growth (Khosla and Bailey, 1988b) and protein synthesis (Khosla et al., 1990) of E. coli. Subsequently the globin has been expressed in a variety of organisms relevant to biotechnology including bacteria, yeast, as well as cultured plant and animal cells, to increase growth and product yields. Some of the biotechnological applications of Vgb published before 1996 have been summarised by Bailey et al. (1996) and are expanded and updated by Wu et al. (2003). Recently, error-prone PCR was used to improve the efficiency of Vgb function (Andersson et al., 2000). Cells expressing two copies of the vgb gene, joined by a short linker of six base pairs, had higher levels of ribosomal and tRNA and reached a cell density 19% higher than cells expressing the native Vgb (Roos et al., 2002). Interestingly, wild-type E. coli cells expressing either R. eutropha flavohaemoprotein (Fhp) or a chimeric protein comprising Vgb fused to the flavin reductase domain of Fhp reached a higher cell density under hypoxic conditions than control cells containing the native Vgb (Frey et al., 2000). This work prompted the screening of other microbial haemoglobins for the ability to promote microaerobic growth and substrate utilisation in E. coli; however, none of those haemoglobins out-performed Vgb (Bollinger et al., 2001), nor did yeast flavohaemoglobin or horse heart myoglobin (Kallio et al., 1996). Generally speaking, heterologous expression of Vgb increases the host’s growth and productivity under hypoxic environments, but the effects of Vgb also depend on medium composition and growth conditions (Wei et al., 1998). Future commercial use of this engineering will require
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a better understanding of the underlying mechanisms and a case-by-case assessment of the possible enhancements of the growth and metabolic advantages. A significant recent finding is that Vitreoscilla Vgb binds, in a yeast two-hybrid assay, specifically to the O2-reactive subunit I of the cytochrome bo-type terminal oxidase from Vitreoscilla. Binding sites are also present on the cytochromes bo from E. coli and P. aeruginosa (Park et al., 2002). These data strongly support the view, advanced many years ago by Webster and colleagues, that Vgb facilitates O2 delivery to the respiratory system.
3. TRUNCATED GLOBINS 3.1. Introduction The truncated globin family comprises the most recently discovered group within the haemoglobin protein superfamily and has been reviewed very recently (Wittenberg et al., 2002). These compact haemoglobins are found in eubacteria, cyanobacteria, protozoa and plants. It is of note that truncated haemoglobins (trHbs) are found in many pathogenic bacteria. It has been hypothesised that trHbs, proteins capable of binding O2, supply pathogens with the ability to survive within the low O2 environment of their host. Remarkably the globin fold of trHbs is formed of only four a-helices arranged in a two-over-two a-helical sandwich. This results from considerable editing of the traditional globin fold to engender a functional haemoglobin protein. As the name suggests, the trHbs are shorter by 20–40 residues than the single domain non-vertebrate globins. So far more than 40 presumed trHb genes have been discovered. Wittenberg et al. carried out a phylogenetic analysis and distinguished three groups within the trHb family (I, II, and III). TrHbs from Mycobacterium tuberculosis (trHbN), the cyanobacteria Synechocystis and Nostoc commune, the unicellular alga Chlamydomonas eugametos, and the protozoan Paramecium are found in group I. Group II is larger: among its members are numbered the other M. tuberculosis trHb (trHbO) and the globins of Staphylococcus aureus and Bordetella pertussis. The least is known concerning the members of group III. Though there is seemingly little amino acid identity between members of separate groups (for instance, there is only 18% between the two trHbs of M. tuberculosis) the extent of identity becomes much higher when comparing amino acid sequences of trHbs within the same group, for
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instance 79% between trHbO and the trHb from Mycobacterium avium (Wittenberg et al., 2002).
3.2. The Two-Over-Two a-Helical Fold The crystal structures of trHbs from Chlamydomonas, Paramecium, and of M. tuberculosis trHbN have been solved. The antiparallel helix pairs in the 2-on-2 sandwich are comprised of B/E and G/H, connected by an extended polypeptide loop. There are many unique features within the trHb globin fold, particularly the almost total absence of the A helix. The instability implied by extensive deletion of the A helix is prevented by the presence of a hydrophobic amino acid cluster in the AB region. This allows efficient sealing of the proximal side of the haem pocket. Indeed, the single conserved turn of the A helix allows the N-terminus of the trHb to become tethered to the protein core. There is a strongly conserved GlyGly motif in the AB hinge that is believed to further stabilise the trHb fold (Pesce et al., 2000). Two other Gly-Gly motifs, also thought to provide stabilisation, are located at the EF hinge and the pre-F loop (Milani et al., 2001a). Only group I and II trHbs possess the Gly-Gly motifs; little is known concerning the putative motifs that replace them for stabilisation within group III. The F-helix is deleted, excepting a single turn, replaced by a polypeptide segment in an extended conformation known as the pre-F loop (Pesce et al., 2000). The group II trHbs have a 15- to 16-residue EF loop region that carries a highly charged and polar sequence motif. In contrast to this, the group I trHbs have a shorter pre-EF loop and do not show any sequence conservation within the F-loop region. Because both the EF loop and the F helix influence the orientation of the proximal His-F8 imidazole and hence the O2-binding properties of the trHb, differences in these regions in the group I and II trHbs may lead to functional distinctions (Pathania et al., 2002a). Some members of group I utilise a strong hydrogen-bonded salt bridge as a means of support of the pre-F loop (Pesce et al., 2000; Milani et al., 2001b). The C helix is all but deleted, the CD-D region being reduced to approximately three residues. This is arguably the smallest polypeptide span that could join the C and E helices (Wittenberg et al, 2002). However, the NMR structure of Synechocystis trHb shows that the two-over-two globin fold is not inviolate. In this trHb the fold is slightly distorted; the F helix extends for more than two turns, the B and E helices are at a pronounced angle and the axes of the two H helix segments are not collinear (Falzone et al., 2002). Despite this, the trHb globin fold
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remains an example of how such editing of the amino acid sequence is distributed over the whole protein in order to retain a functional globin fold.
3.3. Conserved Residues There are a minimal number of strictly conserved amino acids within the known trHb sequences. The proximal haem ligand His-F8 is the only totally invariant residue. At the B9-B10 sites there is a strongly conserved Phe-Tyr couplet. Tyr-B10 is involved in haem ligand stabilisation. Position CD1 is conserved as Phe in group I and III trHbs; those residues in group II inhabiting the position can be Phe, Tyr or His. The E7 position is more variable. Group I trHbs mostly have Gln-E7, Group III have His-E7, and in Group II Ala, Ser or Thr can occupy the E7 position. The E14 position is almost always Phe and this residue is believed to shield the haem from solvent in a role comparable to that of Phe-CD1 in other haemoglobins (Wittenberg et al., 2002).
3.4. Coordination of the Haem In all trHbs, as in all haemoglobins, the proximal ligand to the haem is the His-F8 residue. The stretching frequency of the Fe-His-F8 bond has a relatively high value (220–232 cm1) as compared to mammalian haemoglobins (Couture et al., 1999a; Das et al., 2000; Mukai et al., 2002). This means that the proximal His residue is unstrained, indicative of high ligand affinity. Indeed, the stretching frequency of Fe-His-F8 within mammalian Hb in the R state is 222 cm1 (Das et al., 2000). The two trHbs, HbN and HbO, of M. tuberculosis (Couture et al., 1999b; Mukai et al., 2002) and that of Paramecium (Das et al., 2000) exist as a 5-coordinate ferrous high-spin species at all pH values. In some trHbs, the haem is hexa-coordinated by a second endogenous ligand to form a low-spin complex. In the cyanobacterium Synechocystis sp. PCC6803 trHb, residue His-E10 is proposed as the distal ligand. The 6-coordinate low-spin form is in equilibrium with the 5-coordinate high-spin form that predominates at pH 12. Under extreme alkaline conditions, both endogenous ligands dissociate and hydroxide becomes the fifth ligand. Exogenous ligand is only capable of binding to the 5-coordinate complex. At high ligand concentration, the rate of conversion of the 6-coordinate form to the 5-coordinate form is the rate-limiting factor in terms of ligand
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binding. In Synechocystis, the 6-coordinate form is in the majority as the rate of conversion is low at 30 s1 (Couture et al., 2000). It is thought that while His-E10 is the distal ligand, Gln-E7 is also required for rapid formation of the 6-coordinate complex. Hvitved et al. postulated that the role of Gln-E7 is to increase both the rate of association and dissociation of the hexa-coordinating side chain. This ensures rapid equilibration of the 6-coordinate trHb with the exogenous ligand (Hvitved et al., 2001). TrHbs in the cyanobacterium Synechococcus sp. PCC7002 and in Chlamydomonas too are hexa-coordinated by endogenous residues, i.e. His-E10 and Tyr-B10, respectively (Scott et al., 2002). It is thought that in Chlamydomonas trHb, the residue is in the tyrosinate form and that LysE10 helps to stabilise the interaction through the sharing of a proton (Das et al., 1999). The situation is a little different to that in Synechocystis as the 6-coordinate low-spin form predominates only at alkaline pH; at neutral pH the trHb is primarily a 5-coordinate high-spin complex. Ligand binding is not dependent to the same extent on the rate of conversion of the 6-coordinate to 5-coordinate form (Couture et al., 1999a). It may be that the N. commune trHb is also endogenously hexa-coordinated. NMR spectroscopy has identified a pH-directed structural transition that ostensibly involves alternate orientations of the His-E10 side chain (Yeh et al., 2000b). This endogenous hexa-coordination is rare in globins, most likely because it hinders the binding of O2. However, there are exceptions other than those among the trHbs. The ‘‘non-symbiotic’’ Hb found in rice uses a distal His-E7 to ligate the haem iron in the ferrous state (Lecomte et al., 2001), and there are also two hexa-coordinate (hx) Hbs to be found in humans and mice. Neuroglobin, which is localised to the brain (Burmester et al., 2000), is hypothesised to supply O2 to the retina (the visual process is highly ATPdemanding) (Schmidt et al., 2002). Less is known concerning the second hxHb, histoglobin (Trent and Hargrove, 2002). In the context of this review the hxHbs are interesting because they present similar geometry around the haem if the alignment is based on 3D structure rather than sequence. This may suggest related function within the group and further demonstrates the adaptability of the globin fold (Falzone et al., 2002).
3.5. Ligand Binding The O2-bound forms of trHbs have Fe-O2 stretching modes lower than those reported for any other haemoprotein with an axial His ligand. The frequencies range between 554 and 563 cm1 (Couture et al., 1999b; Das
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et al., 2000, 2001; Mukai et al., 2002). By comparison, the Fe-O2 stretching mode of mammalian Hb is 568 cm1. In the Chlamydomonas and Synechocystis trHbs, replacement of Tyr-B10 or Gln-E7, respectively, with Leu or Glu caused the frequencies to increase to values approaching those of mammalian haemoglobins. The smaller values for trHb were attributed to a strong hydrogen bonding network stabilising the bound O2 on the distal side of the haem (Das et al., 2001). Correspondingly, such mutation of the TyrB10 or Gln-E7 residues in the Chlamydomonas and Synechocystis trHbs and M. tuberculosis trHbN also caused the O2 dissociation rates of the proteins to increase by up to 100-fold (Couture et al., 1999a; Yeh et al., 2000a; Hvitved et al., 2001). Crystal structures of aquo-met Paramecium and cyano-met Chlamydomonas trHbs have shown that Tyr-B10 is buried in the inner haem pocket. It is oriented through hydrogen bonds to Gln-E7 and Thr or Gln-E11 in the protozoan and algal trHbs respectively. The Gln-E7 side chain points into the distal cavity providing an additional hydrogen bond to the haem-bound ligand. Paramecium trHb has a lower O2 affinity than Chlamydomonas and this is reflected in the looser contacts and hydrogen bonds of the algal trHb distal site (Pesce et al., 2000). The distal site of M. tuberculosis trHbN has been studied in some detail. Bound O2 is hydrogen-bonded to Tyr-B10 through the sp2 orbital of the proximal O2 atom (that bound to the haem iron). This causes constraint and weakening of the Fe-O2 bond resulting in a low stretching mode (Yeh et al., 2000a). TrHbN possesses a Leu residue at the E7 site; as Leu cannot form a hydrogen bond, the E7 position does not have a stabilising influence as it does in most other trHbs where Gln occupies E7. However, the crystal structure of HbN indicates that the Gln-E11 NE2 atom forms a hydrogen bond to the phenolic OH group of Tyr-B10 (Milani et al., 2001b). The hydrogen bonding pattern involving Tyr-B10 and Gln-E7 resembles that of oxyhaemoglobin from Ascaris suum, a parasitic nematode. This Hb has the high affinity for O2 that characterises many of the trHbs. The phenolic oxygen of the Tyr-B10 forms a strong hydrogen bond with the distal oxygen atom of the ligand and Gln-E7 makes a weaker connection with the proximal atom (Yang et al., 1995). The other M. tuberculosis trHb, trHbO, has a lower O2 affinity than trHbN. This is reflected in the residues occupying its distal site. It possesses Ala-E7 and Leu-E11. Both are likely to be less effective at generating a network of hydrogen bonding to stabilise the bound O2 (Pathania et al., 2002a). The hydrogen bonding network that exists in trHbO is slightly different to those outlined above. There is a hydrogen bond between the
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proximal oxygen atom of the bound dioxygen and Tyr-CD1, in direct contrast to the situation in Chlamydomonas and Paramecium trHbs and trHbN where Tyr-B10 has the primary stabilising influence on the bound ligand. However, Tyr-B10 is still involved in the hydrogen bonding network of trHbO. Mukai et al. postulated that it forms a hydrogen bond to the terminal oxygen atom of the bound dioxygen. Indeed, mutation of Tyr-B10 causes a smaller increase in the Fe-O2 mode than does mutation of Tyr-CD1 (Mukai et al., 2002). The mode of entry of ligands into the distal haem pocket of trHbs is also distinct from that of mammalian globins. The crystal structures of Paramecium, Chlamydomonas, and M. tuberculosis (trHbN) trHbs reveal the presence of a protein tunnel or cavity that appears suited to the diffusion of ligands to the haem. Phe-E14, a more-or-less invariant residue among the trHbs, is almost orthogonal to the porphyrin ring. It provides a rigid closure to the lower part of the haem pocket. This and the hydrogen bond connections between part of the E helix and the pre-F region afford total inhibition of access to the haem distal site via the E7-gate (Pesce et al., 2000; Milani et al., 2001b). In trHbN, the tunnel connects the haem distal pocket to the protein surface at two sites. It has a diameter of 5–7 A˚ and the separate branches of the tunnel are approximately 20 and 8 A˚ in length, respectively originating from between the GH and AB hinge areas and a gap flanked by residues of the G and H helices (Milani et al., 2001b). Residues located within the cavity and at the access sites are hydrophobic in nature and reasonably conserved over the trHb family. This is suggestive of a significant role for the tunnel. Its size and apolar nature renders it well suited to the storage or diffusion of small uncharged ligands such as O2 and NO (Wittenberg et al., 2002).
3.6. Function The functional roles of the trHbs as a group remain mysterious but their low concentrations in vivo hint at catalytic function(s) rather than O2 storage. Recent work concerning the mycobacterial trHbs has provedelucidatory. TrHbN has the higher oxygen affinity (P50 ¼ 0.013 mm Hg at 20 C) of the two, indeed to such an extent that a role in oxygen transport seemed highly unlikely. Another possibility concerns the detoxification of NO (Couture et al., 1999b). The conversion of NO to nitrate is an established detoxification reaction carried out by Hmp, the E. coli flavohaemoglobin (Poole and Hughes, 2000) (Section 4.8). There is evidence that reactive nitrogen intermediates produced by host macrophages play
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an important role in the induction of dormancy in the bacilli and that host NO synthase protects against tuberculosis (MacMicking et al., 1997). However, tuberculosis infection is a dynamic balance between the host immune system and growth of the bacilli, indicating the presence of an NO resistance mechanism within M. tuberculosis. TrHbN in Mycobacterium bovis has been shown to be necessary for the consumption of NO in vivo, with Tyr-B10 as a crucial residue. The presence of trHbN within M. bovis was able to protect aerobic respiration from the effects of NO and in vitro trHbN stoichiometrically oxidises NO to nitrate (Ouellet et al., 2002). M. smegmatis also expresses a trHbN homologue. The NO uptake of NO-exposed M. smegmatis cells is nearly 7.5 times higher than that of cells which have not previously been exposed to NO (Pathania et al., 2002b). TrHbO has a very different distal pocket, and unlike trHbN it is expressed throughout the growth phase, suggesting a different physiological function (Mukai et al., 2002). When expressed in E. coli (which possesses no trHbs) it increased the O2 uptake of membrane extracts approximately two-fold. However, in E. coli deficient in the terminal oxidase cytochrome bo’ there was no such improvement. This is indicative of a role in the facilitation of oxygen transport to the terminal oxidases, particularly cytochrome bo. Such a function appears rational for a globin within a pathogen that must survive in the low O2 environment of the granuloma (Pathania et al., 2002a). Nostoc commune is a photoautotrophic heterocyst-forming cyanobacterium that is capable of aerobic nitrogen fixation. It synthesises a monomeric 12.5 kDa globin called cyanoglobin or GlbN, which is synthesised under conditions of low O2 and only in cells grown in the absence of combined nitrogen (Potts et al., 1992). The glbN gene is located adjacent to nif genes on the chromosome, and is upregulated by NtcA, a positive regulator responsible for the control of genes involved in nitrogen fixation and there is a correlation between the accretion of GlbN and NifH (Hill et al., 1996). The purified protein binds O2 reversibly with high affinity and non-cooperativity (Thorsteinsson et al., 1996, 1999). In view of these properties, and the fact that the globin is located along the cytosolic face of the cell membrane, it has been suggested that GlbN may be part of, or present O2 to, a microaerobically functioning electron transfer chain. Similar functions have been attributed to leghaemoglobins in the case of symbiont rhizobia (Appelby, 1984), whereas in another N2-fixer, Azotobacter vinelandii, it is the rapid turnover of an integral membrane terminal oxidase (cytochrome bd ) that is believed to allow aerotolerant N2 fixation (Poole, 1988; Poole et al., 1994a; Poole and Cook, 2000).
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The trHb of Chlamydomonas is expressed in response to light and photosynthesis is required for its full expression. It is present predominantly in the chloroplast thylakoid membrane at low concentration (Gagne´ and Guertin, 1992; Couture et al., 1994). This trHb has been hypothesised to function in the protection of the photosynthetic machinery from small O2 leaks. Feasibly it could convert the O2 to H2O2 which would be degraded by a peroxidase (Das et al., 1999). Less is known concerning the other trHbs. A protein believed to be a trHb has recently been identified in another nitrogen fixing bacterium, the actinomycete Frankia. It has an O2 dissociation rate constant similar to that of GlbN (Tjepkema et al., 2002). The trHb from Synechocystis, a nonnitrogen fixing cyanobacterium, is thought unlikely to be involved in the facilitated transport of oxygen because, at 0.014 s1, its oxygen dissociation rate constant is extremely low (Hvitved et al., 2001). It is perhaps telling that the leprosy-causing bacterium Mycobacterium leprae, an organism whose genome has undergone extensive reductive evolution, has retained a trHb (Cole et al., 2001; Visca et al., 2002).
4. FLAVOHAEMOGLOBINS 4.1. Discovery of the Flavohaemoglobins Although the flavohaemoglobin, Hmp, of E. coli is now widely held to be the prototypical member of this class, it was not the first to be described. Almost thirty years ago, Oshino, Chance and collaborators purified a haemoglobin-like protein from yeast (Oshino et al., 1972, 1973a,b) and described a number of its most important properties. In particular, it was recognised as a ‘‘haemoglobin-reductase complex’’ (Oshino et al., 1972) and soon after it was recognised that two prosthetic groups (protohaem and FAD) coexisted in a single polypeptide. The first bacterial example to be described was the haemoglobin-like protein from R. eutropha; the protein was initially identified as a cytoplasmic b-type cytochrome capable of forming an oxygenated species (Probst and Schlegel, 1976). It was subsequently described as a flavohaemoprotein on the basis of analyses of the purified protein (Probst et al., 1979). The absorbance spectrum of the purified protein was similar to that of animal globins, with the addition of signals from the flavin. It was shown that the protein was reduced by NADH in aerobic solution, giving a transiently stable oxygenated complex with distinctive absorbance maxima. Furthermore, the protein catalysed the
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Figure 3 Serendipitous cloning of the E. coli hmp gene. This was described by Vasudevan et al. (1991). During the course of attempts to clone the dihydropteridine reductase (DHPR) gene, clones were obtained that were yellow (flavin) in colour and found to over-express a protein of 44 kDa. On sub-cloning and further characterisation, the gene responsible was found to be ‘‘gene X’’ now called hmp.
reduction of various dyes (dichlorophenol indophenol, methylene blue) and horse cytochrome c in a CO- and azide-insensitive manner. Studies of these proteins lapsed for many years, presumably because no physiological function for them could be found. The first bacterial flavohaemoglobin gene to be identified was hmp of E. coli. During the course of experiments designed to clone a dihydropteridine reductase from this bacterium, Vasudevan cloned, initially on a cosmid, a gene previously called gene X adjacent to glyA at 57.7 min on the E. coli chromosome (Fig. 3; Vasudevan et al., 1991). The gene was predicted to encode a protein of about 44 kDa, the N-terminal domain of which was clearly similar to globins. Furthermore, extracts of cells expressing the protein had spectral and ligand-binding features typical of a protohaem-containing protein, which was named nonetheless, with some circumspection, Hmp (haemoprotein) since no globin-like function was known. Shortly afterwards, Andrews et al. (1992) independently discovered Hmp (a ‘‘haemoglobin-like protein’’) in a search for ferric ion reductases and pointed to the homology of the C-terminal domain with similar domains, each resembling plant ferredoxin NADPþ reductase (FNR), in a number of proteins. Figure 4 illustrates the high level of identity of Vgb with the haem domains of
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Figure 4 ClustalW alignment (MacVector version 7.0, Oxford Molecular) of seven selected flavohaemoglobins and comparison with the single-domain bacterial haemoglobins from Vitreoscilla and C. jejuni (bottom). Identical or conserved replacements are in shadowed boxes.
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Phylogenetic tree of selected flavohaemoglobins.
selected flavohaemoglobins and also the conservation of sequences in the FNR domain, missing in Vgb. The discovery of a haemoglobin in E. coli was quite unexpected, but has led to the further identification and study of similar two-domain proteins in many organisms. Figure 5 illustrates the relationships between several of the 30 or so flavohaemoglobins currently in the databases. Several points are noteworthy. First, flavohaemoglobins are widely distributed in Gram-positive and Gram-negative bacteria, but not apparently in the Archaea. Many of these bacteria are animal or plant pathogens. Flavohaemoglobins are also found in several genera of yeasts. Second, some microbes (Streptomyces coelicolor A3 and Dictyostelium discoideum, for example) have two copies of an hmp-like gene. The physiological significance of this is unclear at present. Finally, Hu et al. (1999) reported the regulation of a gene that they described as encoding a flavohaemoglobin in M. tuberculosis H37Rv. However, the nucleotide sequence cited in that paper (EMBL-GenBank Z92774, encoding sequence nucleotides 17664 to 18741) with the gene name Rv3571 is predicted to encode a protein having the FAD- and NADH-binding sites of FNR, but not a haemoglobin-like domain. Therefore, this protein cannot be considered a homologue of the flavohaemoglobins. However, M. tuberculosis does
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possess another gene (Rv0385) which is predicted to encode a flavohaemoglobin that shares the same conserved haem domain as Hmp and this is shown in Figure 4. It should be noted that although claims have been made for the presence of a flavohaemoglobin in C. jejuni (Pesce et al., 2000; supplementary material in Wittenberg et al., 2002) our inspection of the sequence does not confirm this. Instead, we find a single-domain globin (included in Fig. 2), resembling that of Vitreoscilla (Section 2) and a truncated haemoglobin (Section 3).
4.2. Composition of Hmp and other Flavohaemoglobins The first flavohaemoglobin to be purified was that from yeast (Oshino et al., 1973a,b) over thirty years ago, and originally described as a ‘haemoglobinreductase complex’ (Oshino et al., 1972) Although the initial claim for the presence of non-haem iron was not substantiated, this work was the first to establish the association of a globin domain with a reductase. It is salutary to note, as did Oshino et al. (1972), that ferredoxin and NADPHferredoxin reductase were first discovered as ‘‘methaemoglobin-reducing factors’’. A similar protein described as an ‘oxygen-binding flavohaemprotein’ was later purified and characterised in Schlegel’s laboratory (Probst et al., 1979). The E. coli globin (Hmp), product of the hmp gene, is composed of 396 amino acids and its amino-terminal 144 residues have 45% sequence identity with Vgb (Fig. 4; Section 2). The carboxyterminal domain possesses an FAD- and NAD(P)H-binding domain (Fig. 1) and the sequence is clearly similar to other reductases (Andrews et al., 1992). An interesting feature of the extreme N-terminus is its overall hydrophobicity and thus close similarity with the corresponding region of Vgb. Of the first 30 residues, 16 are identical and a further four show conservative substitutions. Furthermore, in the case of Vgb, 16 residues have been shown by phoA fusions to specify export of Vgb to the periplasm (Khosla and Bailey, 1989) when this protein is expressed in E. coli; of these 16, 10 are identical in Hmp (Vasudevan et al., 1995). Consistent with this is the observation that, at least when over-expressed, about a third of the Hmp apoprotein is found in periplasmic fractions when assayed using an Hmpspecific antibody. However, no spectral signals attributable to the holoprotein could be detected in periplasmic fractions (Vasudevan et al., 1995). This may reflect failure to assemble the holoprotein with haem (or insensitivity of the spectral assay). Subsequent work has confirmed that
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Hmp can be detected in periplasmic fractions from both E. coli and Salmonella (C.E. Mills, S. Grogan and R.K. Poole, unpublished) but it is unclear whether this split location is an artefact arising from overexpression. The first purification of Hmp from E. coli and thereby confirmation of the redox centre complement was reported by Ioannidis et al. (1992). The protein contains flavin (FAD) and haem B (i.e. protohaem IX). Various reports of the stoichiometry of the redox centres in the pure protein have been published, notwithstanding the prediction from the sequence that there is one binding site for haem (in the N-terminal domain) and one each for flavin and NAD(P)H in the C-terminal portion. The later preparation described by Mills has about 0.6 mol haem and 0.6 mol FAD mol1 (Mills et al., 2001). Other reports give 0.1 and 0.01 mol of haem and FAD, respectively, in azide-treated preparations (Gardner et al., 1998a) and 0.28 and 0.42 mol mol1, respectively, in a later preparation (Gardner et al., 2000b). Catalytic activities of an Hmp preparation with equistoichiometric FAD and haem (c. 0.6 mol mol1) were stimulated by exogenous FAD (Mills et al., 2001). The affinity for O2 (in the absence of NO) was decreased from 80 to 15 mM with added FAD but was accompanied by cyanideinsensitive O2 consumption that might be attributed to direct electron transfer from the flavin. As purified, the protein is in the ferric state with absorbance maxima at 403.5, 540 (shoulder) and 627 nm (Ioannidis et al., 1992). The protein is readily reduced by dithionite to give a form that reacts with CO. These ferrous and carbonmonoxy states resemble those of other haemoglobins, with maxima at 431.5 and 558 nm, and 421, 542 and 566 nm, respectively. Aerobic addition of NADH to the ferric form generates a transiently stable oxygenated form with spectral maxima at 413, 544 and 580 nm. Addition of dithionite and nitrite to the ferric protein gives a nitrosyl complex, whose e.p.r. spectra indicated that the haem is attached to the protein via a His residue (Ioannidis et al., 1992) (see below).
4.3. Structure of the Flavohaemoglobins Fhp and Hmp The flavohaemoglobins from E. coli and R. eutropha are closely related (39% amino acid identity) and the haem domains are clearly homologous to Vitreoscilla Vgb (51% identity with the N-terminal globin portion of Hmp). Key amino acids in the haem domain are wholly conserved, particularly Tyr-B10 (in the distal pocket, i.e. on the ligand-binding ‘side’ of the haem plane), Phe-CD1, Leu-E11 and His-F8 (the haem-binding residue on the
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proximal side of the haem plane). Interestingly, the amino acid residue at E7, which is generally His in vertebrate haemoglobins and myoglobin, is generally a Gln residue in flavohaemoglobins and in Vgb. That His is the proximal ligand was originally suggested on the basis of sequence alignments (Vasudevan et al., 1991), and EPR spectroscopy (Ioannidis et al., 1992) was consistent with this view. Tyr-B10 and Gln-E7 on the distal side appear to be particularly important in defining the interactions of the protein with the haem-bound ligand. Resonance Raman spectroscopy of the purified E. coli Hmp has revealed that the active site is remarkably peroxidase-like (Mukai et al., 2001). At neutral pH, the ferric protein is five-coordinate and high-spin as in peroxidases. In the ferrous protein, an unusually strong iron-His stretching mode (244 cm1) suggests that His-F8 has imidazolate character. The FeHis-Glu triad appears analogous to the Fe-His-Asp arrangement in cytochrome c peroxidase. This strong hydrogen bonding interaction on the proximal side may provide a strong electronic ‘push’ for the activation of the O-O bond. Information on the distal side is provided by resonance Raman (Mukai et al., 2001) and infrared spectroscopy (Bonamore et al., 2001) of the CO complex. In the CO-bound form, open and closed conformations were detected. In the latter, the haem environment is highly polar, with the CO strongly interacting with Tyr-B10 and/or Gln-E7 (Mukai et al., 2001). The infrared stretching frequency of the ferric cyanide form (Bonamore et al., 2001) has an unusually high value (2136 cm1), again revealing a similarity in the active sites of Hmp and peroxidases. The distal environment may provide a strong electronic ‘pull’ for O-O bond activation, again as in peroxidases. Consistent with this heterogeneity, two phases are observed in the kinetics of CO recombination (Gardner et al., 2000a) The role of Tyr-B10 on the distal side is particularly noteworthy. In the trHbN (Section 3.1) from M. tuberculosis, mutation of Tyr-B10 causes loss of the closed conformation in the CO adduct and the O2 dissociation rate increases dramatically (Yeh et al., 2000a). Similar effects are seen on substituting Tyr-B10 in Hmp with Phe, which increases the O2 dissociation rate constant 80-fold and reduces the NO denitrosylase activity 30-fold demonstrating the importance of this distal residue (Gardner et al., 2000a). Two crystal structures have been reported for flavohaemoglobins. The first was that of the R. eutropha Fhp protein solved at a resolution of 1.75 A˚ (Ermler et al., 1995a,b). As anticipated from sequence analyses, Fhp comprises two fused modules: a globin domain and an FAD-binding oxidoreductase module, which adopts a fold like ferredoxin-NADPþ
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reductase (FNR). In the globin domain, the haem pocket appears to be enlarged relative to other globins by displacement of helix E. Electron transfer between FAD and haem may be facilitated by their proximity (6.3 A˚) and a predominantly polar environment. Very recently, the structure of E. coli Hmp has been described (Ilari et al., 2002). Despite the homology between Hmp and Fhp, the structures differ somewhat, due mainly to a rotation of the NAD-binding module and a substantial rearrangement of the E-helix. In the distal haem pocket, the position closest to the iron atom is occupied by the isopropyl sidechain of Leu-E11. It has been suggested that this might allow a ligandlinked conformational change in which rotation of the Leu-E11 sidechain to the haem edge allows Tyr-B10 to interact with the iron-bound ligand. In effect, Leu-E11 gates accommodation of the incoming ligand; only on displacement of the Leu-E11 sidechain does the Tyr-B10 hydroxyl, previously shown to be essential for O2 binding and catalysis (Gardner et al., 2000a), approach the iron-bound ligand. Indeed, resonance Raman and FTIR spectroscopy reveal participation of a Tyr hydroxyl group in distal coordination of CO in the ferrous protein (Bonamore et al., 2001; Mukai et al., 2001). On the proximal side of the haem, a hydrogen bonding network is seen, consistent with the imidazolate character of the proximal His described by Mukai et al. (2001). The spectral resemblance of the cyanide form of Hmp (Bonamore et al., 2001) and horseradish peroxidase provides additional evidence for similarities in the haem pocket. When the Fhp structure was first reported, an unidentified ligand was described at the haem active site (Ermler et al., 1995b). This was later identified (Ollesch et al., 1999) as a phospholipid, accommodation of which alters the positioning of the E-helix. Specifically, a cyclopropane ring of diacylglycerol-phosphatidic acid is located on top of the haem displacing Leu-E11 and moving the E-helix. The structure described for the E. coli flavohaemoglobin lacks a bound lipid but it is an intriguing possibility that both proteins may have a role in catalysing reactions involving lipids (see later, Section 4.8.3)
4.4. Enzymic Properties of Hmp Numerous catalytic activities have been ascribed to Hmp (Table 1). With the caveat that redox centres have not always been quantified in the enzyme preparations used, the data suggest a bewildering range of possible functions for Hmp. Although Hmp is now widely viewed as
Redox activities attributed to flavohaemoglobins.
Protein
Electron acceptor
Km (mM)
Comments
Reference
E. coli Hmp
O2
90 (no added FAD) 16 (15 mM FAD) 2–4
Without removal of partial reduction products of O2 With removal of partial reduction products of O2 Activity stain on gel Stimulated by free flavins Fe(III)-hydroxamate K, CO-insensitive CO- and SOD-insensitive
Mills et al. (2001)
Fe(III) 104 Cytochrome c paraquat NO
R. eutropha Fhp
GSNO, nitrite dihydropteridine Azotobacter vinelandii NifL DCPIP, methylene blue, horse cytochrome c
6.8–7.3 7,300
Poole et al. (1996a) Andrews et al. (1992) Eschenbrenner et al. (1994) Poole et al. (1997)
CO- and SOD-insensitive
Poole et al. (1997) Anjum et al. (1998) Kim et al. (1999) Mills et al. (2001) Gardner and Gardner (2002) Vasudevan et al. (1991) Macheroux et al. (1998)
CO- and azide-insensitive
Probst et al. (1979)
N2O detected as product Cyanide-sensitive Measured anoxically
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Table 1
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a NO-detoxifying enzyme, particularly aerobically, other physiological functions cannot be ruled out and the physiological significance of some of the findings in Table 1 may yet be realised. NO ‘oxygenation’ (or denitrosylation) and NO reduction are thought to be catalysed exclusively at the haem (Poole and Hughes, 2000). Indeed, the haem pocket seems engineered to perform ligand chemistry (Mukai et al., 2001) rather than reversible ligand binding. Hmp binds and reduces O2 at the haem (Poole et al., 1994b; Membrillo-Herna´ndez et al., 1996; Poole et al., 1996a). Both NADH (Km 2 mM) and NADPH (Km 20 mM) are oxidised by Hmp (Anjum et al., 1998). The FAD is able to transfer electrons derived from NAD(P)H to a number of external acceptors. For example, purified Hmp reduces cytochrome c in a CO-insensitive manner, demonstrating that electrons exit at the level of FAD (see Table 1 for details and references). NADH oxidation proceeds in cyanide-treated Hmp but can be stimulated if exogenous FAD is added (Mills et al., 2001). The consumption of O2 by Hmp in the absence of NO is intriguing for several reasons. First, it suggests that Hmp might, in principle, and as claimed for Vitreoscilla globin (see Section 2.2.2), serve as an oxidase. However, at present there is evidence neither for association of Hmp with the cytoplasmic membrane nor for accepting electrons from the respiratory chain in a manner that would allow Hmp to intercept electron flow in a respiratory chain that was dysfunctional, for example, through mutation of both oxidases. Second, there is good evidence from experiments both with pure Hmp in vitro and using a (sodA-lacZ) reporter (i.e. a fusion of the sodA promoter to the gene encoding b-galactosidase), that Hmp reduces O2 to superoxide and peroxide (Membrillo-Herna´ndez et al., 1996; Mills et al., 2001). It is not known whether superoxide generation by Hmp in the presence of reductants and O2 is fortuitous or whether it has a physiological function, but the copy number of Hmp in vivo appears low, suggesting that this potentially deleterious activity is checked. Intriguingly, the hmp gene is up-regulated not only by NO and agents of nitrosative stress, but also by paraquat, a redoxcycling agent (Section 4.6). Might up-regulation of Hmp, a superoxidegenerating protein, by a superoxide-generating agent serve to amplify the signal perceived during oxidative stress? Third, O2 consumption by electron transfer from FAD to the bound ligand, coupled with the ability of the flavin to transfer electrons to other acceptors (although candidates in vivo have not been identified) is consistent with a possible role of Hmp in O2 sensing (Poole et al., 1994b). This model remains to be rigorously tested, but the presence of O2 at nearmicromolar concentrations does partially inhibit the transfer of electrons
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from NADH to cytochrome c via FAD, as predicted by the model (Poole et al., 1997). Hmp binds O2 strongly with a Kd of 12 nM (Gardner et al., 2000a), compared with a value of 857 nM for sperm whale myoglobin. Replacement of the Tyr-B10 with Phe increases the O2 dissociation 80-fold (Gardner et al., 2000a) demonstrating the importance of this residue in stabilising the haem-bound O2.
4.5. Functions of Hmp The pioneering work of the Oshinos and Chance failed to find a function for the yeast globin. Amongst proposals tested were that haemoglobin (a) acts as O2 store or buffer, (b) has enzymic activities, (c) represents a metabolic pool of prosthetic group(s) or (d) is involved in facilitated oxygen transport (Oshino et al., 1973a,b). The O2 affinity of the yeast haemoglobin, determined from simultaneous recordings of oxygenation-deoxygenation of globin within yeast cells and the oxygen-dependent luminescence of Photobacterium phosphoreum was estimated to about 0.02 mM and thus higher than the affinities of other ‘non-circulating’ globins in Paramecium and Ascaris (Oshino et al., 1971). To determine whether yeast haemoglobin might facilitate O2 transport, the Km value for O2 during respiration by intact cells was compared with cells in which the globin had been destroyed by ethyl hydrogen peroxide. The finding that the O2 concentration giving halfmaximal change in cytochrome a3 absorbance was similar in both types of cell (about 0.03 mM) suggests that respiration was not facilitated by yeast haemoglobin (Oshino et al., 1971). Although this work was inconclusive, it is worth noting that the Vitreoscilla globin has been suggested to facilitate O2 delivery in oxygen-depleted conditions (Section 2.2.2 and 2.2.6). Also considered was the possibility that the globin has a role in peroxide detoxification. Yeasts in which haemoglobin had been destroyed by ethyl hydrogen peroxide grew similarly to those with the globin (Oshino et al., 1973a). The presence of the globin domain in the bacterial flavohaemoglobins also hints at a role in O2 delivery/storage. However, in bacteria, high flavohaemoglobin levels are not generally observed in non-induced conditions, except after genetic manipulation of gene copy number. Therefore, the low abundance in vivo, together with the non-inducibility of hmp by limiting O2 availability (Poole et al., 1996b), makes a role in O2 storage or delivery unlikely. A clue that flavohaemoglobins may be involved primarily in the metabolism of nitrogen species was provided by the demonstration
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(Cramm et al., 1994) that a mutant of R. eutropha carrying a mutation in fhp (equivalent to hmp) was unable to generate nitrous oxide during denitrification. Although Cramm et al. suggested that Fhp is involved in gas metabolism during denitrification and that Fhp might possess NO reductase activity, they could not verify this through enzyme assays. The first direct evidence that a flavohaemoglobin was involved in protection against NO and its congeners was the finding that a single-copy chromosomal-borne fusion of the hmp promoter to lacZ, i.e. (hmplacZ), is up-regulated by NO, sodium nitroprusside (SNP), S-nitrosoglutathione (GSNO) – a nitrosating agent – and related species (Poole et al., 1996b; Membrillo-Herna´ndez et al., 1998). Subsequently, Gardner demonstrated that a mutant of E. coli selected in a screen for NO-resistance exhibited an O2-dependent NO-consuming activity referred to as ‘NO dioxygenase’ (Gardner et al., 1998a). This activity was attributed to Hmp. Results from a number of laboratories have lent support to the view that Hmp detoxifies NO. Hmp is part of a battery of measures employed by bacteria to resist stresses elicited by endogenously and exogenously generated NO and its congeners (Fig. 6). Amongst these clues are the following key observations. (a)
Null hmp mutants of Salmonella (Crawford and Goldberg, 1998a) and E. coli (Membrillo-Herna´ndez et al., 1999) are hyper-sensitive to killing by GSNO. (b) The haem of Hmp is readily reducible by physiological substrates (NAD(P)H) via electron transfer from FAD, and Hmp is capable of
Figure 6 Responses of a bacterium to NO and nitrosative stresses, typified by those occurring in E. coli.
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binding NO extremely rapidly to the Fe(II) form and of redox chemistry with NO and O2 at the haem (Poole et al., 1994b; Kim et al., 1999). (c) Hmp rapidly binds NO but this is reversible. Thus, under anoxic conditions in the presence of CO, the nitrosyl haem reverts to the carbonmonoxy form. This is consistent with the ability of Hmp to reduce NO to NO and subsequent formation of N2O (Kim et al., 1999). NO sequestering and NO reduction are plausible anaerobic functions for Hmp. (d) The level of resistance of cellular respiration to NO in E. coli is directly related to the level of Hmp. Cells pre-induced by treatment with SNP or paraquat are totally resistant to NO concentrations up to 50 mM, whereas respiration of an hmp mutant is highly sensitive to sub-micromolar NO (Stevanin et al., 2000). (e) Null hmp mutants of Salmonella (Stevanin et al., 2002) and E. coli (T.M. Stevanin, E. Demoncheux, R. Read and R.K. Poole, unpublished) are also hyper-sensitive to killing by human macrophages. The flavohaemoglobins of yeast (YHB1) (Liu et al., 2000; Gardner et al., 2000b) and Dictyostelium discoideum (Iijima et al., 2000) (see Section 4.9) have also been implicated in resistance to NO.
4.6. Regulation of hmp Transcription Understanding how and when gene transcription is up-regulated, or synthesis of a particular protein is increased, can give valuable clues to the function of the gene or protein. Thus, Probst et al. reported that the Ralstonia (Alcaligenes) flavohaemoglobin ‘‘concentration increases 20-fold in cells grown autotrophically under limited O2 supply. . . . . . . It might therefore be suggested that, under oxygen-limited growth conditions, the protein acts as an O2 store providing a constant O2 tension enabling the cell to maintain its rate of ATP production . . . . .’’(Probst et al., 1979).
4.6.1. E. coli Most work in this area has been done on the E. coli flavohaemoglobin. Expression of hmp is up-regulated by nitrosative stresses (see below), stationary phase of growth (involving the alternative sigma factor s) (Membrillo-Herna´ndez et al., 1997), anaerobiosis (partly as a result of Fnr
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regulation, see below), the Fe-chelator 2,20 -dipyridyl (Poole et al., 1996b) and the presence of paraquat (Membrillo-Herna´ndez et al., 1997; Anjum et al., 1998; Pomposiello et al., 2001). In marked contrast to the early results with the R. eutropha flavohaemoglobin, Hmp levels are not increased noticeably by limiting O2 supply (Poole et al., 1996b). Consistent with the proposed role of Hmp in NO detoxification, the flavohaemoglobinencoding gene of E. coli, hmp, is most dramatically up-regulated by NO and reactive nitrogen species; this appears not to involve SoxRS (Poole et al., 1996b). We have reported (Membrillo-Herna´ndez et al., 1998) a mechanism of hmp gene regulation that involves interaction between S-nitrosothiols and homocysteine. Intracellular homocysteine is an important co-regulator of several genes involved in methionine biosynthesis, via its effects on MetR, a LysR family DNA-binding protein. One gene activated by MetR with homocysteine as cofactor is glyA, which, in E. coli is adjacent to hmp and divergently transcribed from it. Elevated homocysteine (Hcy) levels, achieved either by exogenous Hcy or in certain met mutants, decrease hmp expression. Since Hcy has been shown to be nitrosated by GSNO (Membrillo-Herna´ndez et al., 1998), such nitrosating agents can deplete Hcy pool sizes, and are postulated to enhance MetR binding at a site proximal to hmp, and up-regulate hmp transcription. It is important to recognise that this mechanism does not readily explain hmp regulation by NO itself. First, NO induces hmp expression anoxically under which conditions NO will not nitrosate Hcy (although the presence of metal ions, for example, might allow NOþ formation from NO). Second, although the S-nitrosoHcy generated on reaction with GSNO breaks down to release NO, which could itself be the inducer, the reaction of Hcy with SNP (which also induces hmp) forms a more stable species from which NO is not released (Membrillo-Herna´ndez et al., 1998). Thus other mechanisms for hmp regulation by NO, particularly anoxically, must be present. Anaerobically, the global regulator Fnr (Kiley and Beinert, 1999; Green et al., 2001) is also involved in the regulation of hmp since an fnr mutation enhances hmp-lacZ expression (Poole et al., 1996b), but how Fnr contributes to NO-mediated regulation has remained obscure until very recently. The [4Fe-4S]2þ cluster of Fnr is oxygen-labile and controls protein dimerisation and site-specific DNA-binding. We have shown that NO also reacts anaerobically with the Fe-S cluster of purified Fnr generating changes in the optical absorbance and an EPR signal consistent with formation of a dinitrosyl-iron cysteine complex (Cruz-Ramos et al., 2002). The NOinactivated Fnr protein can be reconstituted, suggesting physiological
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relevance. Fnr binds at an Fnr box centred at þ5.5 within the hmp promoter (Phmp). Fnr samples inactivated by either O2 or NO bind specifically to Phmp, but with lower affinity. Dose-dependent up-regulation of Phmp in vivo by micromolar NO concentrations of pathophysiological relevance, provided by NOC-5 and NOC-7, is abolished by mutation of fnr, and NO also modulates expression from model Fnr-regulated promoters. Thus, Fnr can respond, not only to O2, but also to NO, with major implications for global gene regulation in bacteria. We have proposed a NO- and Fnr- mediated mechanism of hmp regulation by which E. coli responds to NO challenge (Cruz-Ramos et al., 2002). The hmp gene is also up-regulated by the redox-cycling agent methyl viologen (paraquat) (Membrillo-Herna´ndez et al., 1997; Anjum et al., 1998). Relatively high concentrations of paraquat are required (200 mM or more) and the effect is not shared by other redox-cycling agents (e.g. menadione, plumbagin or phenazine methosulphate). This specificity distinguishes hmp regulation from other genes that are regulated by paraquat as does the finding that the SoxRS two-component regulatory system is not required (Membrillo-Herna´ndez et al., 1997). However, a physiological significance for paraquat regulation is suggested by the sensitivity of hmp mutants to killing by this agent (Membrillo-Herna´ndez et al., 1999). Induction of hmp by paraquat in the stationary phase, but not in the exponential phase, is dependent on RpoS (MembrilloHerna´ndez et al., 1997). The underlying mechanism is at present obscure but the involvement of OxyR or Rob (known regulators of other stress regulons) has been ruled out. The iron chelator 20 20 -dipyridyl dramatically up-regulates hmp transcription (Poole et al., 1996b), both aerobically and anaerobically. Under similar anaerobic conditions, (frdA-lacZ) activity is down-regulated; both these observations are consistent with the view that iron chelation may act on hmp expression via inactivation of Fnr (Poole et al. 1996b).
4.6.2. Salmonella NO, provided in solution by spermine NONOate up-regulates expression of hmp in Salmonella in a SoxS- and OxyR-independent fashion. However, unlike the situation in E. coli, mutation of fnr does not affect either basal or NO-enhanced levels of hmp transcription; instead, a fur mutation derepresses hmp expression (Crawford and Goldberg, 1998b, 1999). An hmp mutant requires markedly lower NO concentrations to induce hmp, consistent with the view that Hmp is involved in NO removal.
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Very recently, changes in Salmonella hmp expression have been observed following macrophage infection (Eriksson et al., 2003). During replication in murine macrophage-like J774 cells, 919 of 4451 S. Typhimurium genes showed significant changes in transcription. A drastic repression of hmp expression was observed 4 h after infection, followed by induction at 8 h, coinciding with the induction of NO synthesis in J774 cells. In S. Typhimurium, hmp expression is repressed by Fur (the iron-responsive regulator); however, other facets of the gene expression profiling show that intracellular bacteria are not starved for iron, so hmp induction presumably reflects NO stress. It should be noted that changes in expression level in this work were given relative to free-living bacteria cultured in laboratory media: the initial drop in hmp expression on infection (above) probably reflects a change in environmental conditions other than NO or nitrosative stress. It might, for example, reflect a decrease in levels of superoxide anion in the macrophage relative to growth in highly aerated media containing excess oxidisable substrates.
4.6.3. Bacillus subtilis The hmp promoter in B. subtilis is activated by NO and a nitrosating agent (Nakano, 2002). The mechanism of up-regulation appears to be via the ResDE signal transduction system. ResD-dependent gene activation is heightened in vivo at limiting O2 concentrations, but full induction of the ResDE regulon requires both low O2 and nitrite (Lacelle et al., 1996). Nakano has shown that, in an hmp mutant, which might be anticipated to accumulate NO when grown in the presence of nitrite, the expression of both hmp and nasD (encoding nitrite reductase) was increased 20-fold. Furthermore, exogenously added SNP also up-regulated hmp and nasD, even in an hmp mutant. Mutants defective in ResD or ResE failed to show a response of nasD to SNP but some hmp transcription was observed. Therefore, an additional, unidentified, but kinetically distinct and ResDE-independent mechanism must induce hmp in response to SNP.
4.6.4. Other Bacteria The HmpX protein of the plant pathogenic enterobacterium Erwinia chrysanthemi has been described as a pathogenicity determinant, based on
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the finding that hmpX mutants produced only mild necrotic spots or no symptoms on plants (Favey et al., 1995). Using a gus gene fusion, hmpX transcription was shown to be induced in co-culture with tobacco cells, suggesting that the protein may be produced in response to factors released by the plant (Favey et al., 1995). Finally, although regulation of M. tuberculosis ‘‘hmp’’ by nitrosative and oxidative stresses has been claimed (Hu et al., 1999), the gene product is unlikely to be a flavohaemoglobin (Section 4.1). 4.7. Phenotypes of hmp Mutants Mutation of flavohaemoglobin-encoding genes compromises bacterial resistance to nitrosative stress. Some examples have been cited above. A Salmonella strain harbouring a deletion in hmp showed an increased sensitivity to acidified nitrite and S-nitrosothiols, both aerobically and anaerobically (Crawford and Goldberg, 1998a). Interestingly, the mutant was unaltered in its sensitivity to SIN-1, which generates superoxide and NO and is widely used as a convenient source of peroxynitrite, the product of the reaction between these two species (Poole and Hughes, 2000). Similar results have been obtained with a defined hmp knockout mutant of E. coli (Membrillo-Herna´ndez et al., 1999). This strain, RKP4545, was hypersensitive to paraquat, GSNO and SNP, consistent with the observed upregulation of the hmp promoter by these agents. A strain, RB9060, having a deletion that extends into hmp (but which also includes part of glyA) was also hypersensitive to NO (Gardner et al., 1998a), particularly during aerobic growth. The defined hmp E. coli mutant was shown to have a further interesting phenotype, namely the inability to reduce paraquat under anoxic conditions (Membrillo-Herna´ndez et al., 1999). This may reflect the broad spectrum of diaphorase activity of Hmp referred to in Section 4.4. 4.8. NO-Detoxifying Activities of Hmp and other Flavohaemoglobins At least two mechanisms appear to allow Hmp to play important roles in resisting the effects of NO. Aerobically, Hmp catalyses a reaction in which NO is transformed to give the relatively innocuous NO 3 ion. Anaerobically, Hmp reduces NO to N2O by a poorly understood mechanism. In both activities, the ability of the FAD moiety of Hmp to transfer electrons to haem B for ligand binding and reduction is probably crucial.
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4.8.1. NO Oxygenase or Denitrosylase Activity. Gardner et al. (1998a) first suggested that Hmp of E. coli might act as an enzyme catalysing the overall reaction: þ 2NO þ 2O2 þ NADðPÞH ! 2NO 3 þ NADðPÞ þ H
This proposal has its origins in the behaviour of a mutant isolated after treatment of strain AB1157 with the chemical mutagen N-methyl-N’nitro-N-nitrosoguanidine. NO-resistant mutants were selected as being able to grow in an atmosphere containing ‘‘960 ppm NO’’ and 10% air, balanced with nitrogen. One such mutant showed elevated NO consumption activity and from this strain was isolated an FAD-binding protein, the N-terminal sequence of which showed it to be Hmp. The nature of the mutation in this strain has not been described. A mutant RB9060 deficient in hmp (but with a deletion that extends into glyA) lacked the NO-consuming activity, whereas a strain carrying the hmpþ gene on a multicopy plasmid had 10-fold elevated activity. These results clearly demonstrated an NO consuming activity of Hmp that could explain the NO-sensitive phenotype of the hmp mutant and the NOresistance of the strain with elevated Hmp. Further, NO-grown cells exhibited an inducible, cyanide-sensitive and O2-dependent NO consumption that accompanied protection of aconitase (Gardner et al., 1998b). This has been assumed to be the activity of Hmp, but is not formally proven, for example through the use of mutants. Later, Hausladen et al. (1998) also reported the NO oxygenase activity of Hmp and showed that an hmp mutant (RB9060) was more severely inhibited in its growth by Snitrosocysteine than was a wild-type strain. Further studies of pure Hmp have confirmed that equistoichiometric O2 and NO are consumed during aerobic NO detoxification by Hmp (Mills et al., 2001). The reaction is not confined to the E. coli Hmp: the flavohaemoglobins of Saccharomyces cerevisiae, R. eutropha and E. coli have similar NO and CO binding kinetics and steady state ‘NO oxygenase’ kinetics (Gardner et al., 2000b). Under aerobic conditions, Hmp has a high affinity for NO, with a KM of 0.25 mM NO in the case of E. coli Hmp (Gardner et al., 2000b). Although the role of Hmp in protection from NO is widely accepted, based on the phenotypes of hmp mutants and the NO-inducibility of Hmp, the physiological significance of the direct NO-removing activities has been questioned. How effective is Hmp at NO detoxification at physiologically meaningful O2 concentrations? The intracellular O2
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concentration has not been reliably determined for any bacterium but it is instructive to look at one natural environment for E. coli – the gut. Here, it is generally accepted that the O2 concentration will be exceedingly low but not zero, particularly near capillaries (Savage, 1977). The O2 concentrations required for oxygenase activity have been addressed in several papers. E. coli cells grown with minimal aeration had reduced NO consumption activities relative to cells grown with vigorous shaking under air (Gardner et al., 1998a). Subsequently, an inducible NO-consuming activity that protects aconitase (an [Fe-S] protein) from NO was described (Gardner et al., 1998b) and attributed to Hmp. Aconitase was protected at O2 concentrations (about 17 mM), at which nitrate and nitrite formation in growing cultures was negligible. This suggests that the O2-mediated decomposition of NO that is anticipated to yield these ions (e.g. by Hmp action) cannot fully account for aconitase protection. Further evidence of the limited effectiveness of Hmp in protecting metabolic functions at low O2 tensions comes from the demonstration of enhanced sensitivity to NO of cell respiration under these conditions (Yu et al., 1997; Stevanin et al., 2000). A KM value for O2 of 60–100 mM has been determined using purified Hmp supplemented with free FAD when assayed with saturating NO (0.75 mM) and NAD(P)H (Gardner et al., 2000a,b). Sensitivity of the reaction to inhibition by NO at NO:O2 ratios of >1:100 makes the determination of Km values for O2 difficult (Gardner et al., 2000b) but in intact cells a value of 60 mM has been reported. Using a different approach, Mills et al. (2001) measured the dependence of NO consumption by pure Hmp (monitored directly using an NO electrode) as a function of O2 concentration and obtained a Km value of 47 mM (Mills et al., 2001). It is not clear whether other microorganisms that contain flavohaemoglobins have access to O2 concentrations that do not limit NO oxygenase activity. Nevertheless, it is clear that Hmp has robust denitrosylase activity: the Vmax is 670 s-1, and the Km values for O2 (with the caveats above), NO and NADH are c. 100, 0.28 and 4.8 mM, respectively at 37 C (Gardner et al., 2000a). The model first suggested for NO dioxygenase activity (Gardner et al., 1998a; Hausladen et al., 1998; Fig. 7) was re-examined by Hausladen et al. (2001). At biologically relevant O2 concentrations, Hmp preferentially binds NO, not O2, the bound species then reacting with O2 to form nitrate. Indeed, in the presence of NO and high O2, the predominant form is the ferric nitrosyl species. In effect the reaction involves the oxidation of the haem-bound nitroxyl (NO) equivalent and is properly called an O2 nitroxylase or denitrosylase reaction (Hausladen et al., 2001).
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Figure 7 Reactions at the haem iron of E. coli Hmp with NO (lower half) and other ligands (top half), both anaerobically (left) and aerobically (right). The Figure is modified from Kim et al. (1999) with the inclusion of newer information on the proposed denitrosylase reaction that leads to nitrate formation. 1. Hmp as prepared is in the Fe(III) form and is reducible by NADH or NADPH. 2. Fe(II) Hmp reacts with CO in a lightreversible fashion to yield the carbonmonoxy adduct. 3. Fe(II) Hmp reacts with oxygen to give the oxygenated species observable at room temperatures in the presence of reductant and O2. 4. Superoxide anion is releasable from the oxygenated species and appears in part as peroxide in solution. 5a. In the NO dioxygenase reaction scheme for nitrate formation (Gardner et al., 1998a; Hausladen et al., 1998) superoxide is attacked by NO to yield NO 3 regenerating Fe(III) Hmp. The peroxynitrite species (in square brackets) is a probable intermediate. 5b. In the later denitrosylase scheme (Hausladen et al., 2001) Hmp preferentially binds NO, not O2, which then reacts with high O2 concentrations to give nitrate. Fe(III) formed (5c) can rebind NO (8). The ferric nitrosyl form (within the oval) is the dominant form during steady state turnover at higher O2 concentrations. 6. Fe(II) Hmp reacts with NO to give a nitrosyl species. 7. Electron transfer from haem reduces NO to NO, which is released, regenerating Fe(III) Hmp. N2O formation (boxed) occurs possibly via a dimeric species (not shown). 8. Fe(III) Hmp reacts with NO to give a nitrosyl species, which is lost on exposure to air.
4.8.2. NO Reductase Activity The ability of Hmp to remove NO from solution anoxically was briefly noted by Stamler’s group (Hausladen et al., 1998). More detailed studies (Kim et al., 1999) of NO binding to Fe(III) Hmp revealed a second order rate constant of 7.5 105 M1 s1 and generation of a nitrosyl adduct that was stable anoxically but decayed in the presence of air to reform the Fe(III) protein. NO displaced CO bound to dithionite-reduced Hmp but,
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remarkably, CO recombined after only 2 s at room temperature indicative of NO reduction and dissociation from the haem. Addition of NO to anoxic NADH-reduced Hmp also generated a nitrosyl species, which persisted while NADH was oxidised. These results were consistent with direct demonstration by membrane-inlet mass spectrometry of NO consumption and nitrous oxide (N2O) production during anoxic incubation of NADHreduced Hmp (Kim et al., 1999). Subsequently, we demonstrated the anoxic NO removing activity by incubating pure Hmp in an O2 electrode chamber and injecting a solution of NO after anoxia had been achieved (Mills et al., 2001). Simultaneous measurements with an NO electrode detected the NO addition and its subsequent removal. In addition to a cyanide-insensitive component, an Hmp-dependent activity that was sensitive to 0.1 mM cyanide was measurable having an initial rate of NO removal, expressed as a turnover number, of 0.15 s1. This is in reasonable agreement with the rate (0.25 s1) determined by membrane-inlet mass spectrometry at higher NO concentrations (Kim et al., 1999) and very similar to that obtained by Gardner et al. (2000a) (0.14 s1). Although these rates are at best 2% of the aerobic NOconsuming activity, they suggest that flavohaemoglobin has an anoxic function. This is consistent with (1) anoxic up-regulation of Hmp synthesis (Section 4.6), (2) the anaerobic phenotype of an fhp mutant of R. eutropha (Cramm et al., 1994), (3) the protection that hmp has been observed to confer in E. coli (Gardner et al., 1998a) and Salmonella (Crawford and Goldberg, 1998a). However, very recently, Gardner et al. (2002) have questioned the ability of Hmp to function significantly under anaerobic conditions and have identified a flavorubredoxin with higher rates of NO removal (Gardener and Gardner 2002), which is outside the scope of this review.
4.8.3. Other Roles for Flavohaemoglobins in Protection from Nitrosative and Oxidative Stresses? Although attention has focussed on the NO-reducing activity of Hmp, new evidence has suggested that this may be of low activity at physiological O2 concentrations (see above). Other roles in protection from nitrosative stress might be envisaged, however. Hmp might, for example, have a role in sequestering NO (as the formation of an anaerobically stable nitrosyl species might suggest (Kim et al., 1999). In this respect, Hmp would resemble the ‘green’ cytochrome b of Bacillus halodenitrificans (Denariaz et al., 1994) which spectrally resembles cytochrome c’. The latter, interestingly, can function as an NO reductase, generating N2O even though the protein
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does not possess a reductase domain (Cross et al., 2000, 2001). Another possibility is that Hmp functions aerobically or anaerobically to repair damage inflicted by nitrosative stress. This might occur whether or not the source of the stress (i.e. NO or a reactive nitrogen species) is directly removed by the protein. In this context, it is notable (a) that the Hmp haem pocket is configured to effect peroxidase-like transformations (Mukai et al., 2001) and (b) that bound lipid, at least in the case of the R. eutropha protein, may modulate enzyme activity. The presence of lipid at the active site of this protein suggests that phospholipids may gain access to act as modulators of function or indeed as substrates (Ollesch et al., 1999; Ilari et al., 2002). Globins from C. jejuni and Vitreoscilla conferred resistance to SNP when over-expressed in E. coli and such cells had increased NO-consuming activities (Frey et al., 2002). However, globins from E. coli, Klebsiella pneumoniae, Deinococcus radiodurans and Pseudomonas aeruginosa conferred resistance to oxidative stress when expressed in E. coli. The molecular and physiological mechanisms underlying these results are not known, and it may be imprudent to assign such functions to the globins when expressed at normal levels in their respective bacterial hosts.
4.9. Flavohaemoglobins of Yeasts and Fungi For many years after its discovery, the yeast flavohaemoglobin could not be assigned a clear function. Gene regulation studies led to the view in 1995 that, in marked contrast to Vgb and the bacterial flavohaemoglobins, the S. cerevisiae globin is up-regulated in logarithmic phase of growth and under oxygen-replete conditions (Crawford et al., 1995). Transcription was shown to be regulated by haem availability via the action of haem-activated proteins (HAP), although anaerobically a low-level HAP-independent induction was detectable. Disruption of the flavohaemoglobin gene did not alter cell viability or growth in a variety of growth conditions in which oxygen availability and carbon source were altered. Subsequent work showed that flavohaemoglobin levels were elevated (from the very low levels extant in actively respiring cells) under conditions in which mitochondrial respiration was compromised either by mutation or by inhibitors of respiration such as antimycin A (Zhao et al., 1996). Other conditions that elevated globin levels were the absence through mutation of superoxide dismutase or the presence of thiol conditions. On the basis of these data it was suggested that the function of flavohaemoglobin in yeast is related to tolerance of oxidative stress. Note that this is reminiscent of the effects on
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E. coli hmp expression of paraquat, which dramatically up-regulates transcription although the transcriptional regulator(s) has not been identified. Mutational analysis (Zhao et al., 1996) showed that loss of flavohaemoglobin (yhb1) did not affect growth in either rho0 or rhoþ strains (i.e. strains without and with, respectively, functional respiratory systems). Thus the globin does not appear to be able to act as a functional oxidase. Supporting the idea that the protein plays a role in oxidative stress tolerance was the observation that yhb1 deletions resulted in increased sensitivity to oxidative stress. Further complications in assigning function arose from subsequent work (Buisson and Labbe-Bois, 1998) that failed to support the view that YHB1 synthesis is increased under conditions of respiratory deficiency, or the oxidative stresses incurred on exposure to H2O2, diothiothreitol and other agents. Recently, the Stamler laboratory has presented persuasive evidence that YHB1 protects S. cerevisiae from nitrosative stress, bringing studies on this eukaryotic flavohaemoglobin into line with the large body of evidence on bacterial flavohaemoglobins. Deletion of the YHB1 gene abolished the NOconsuming activity of cells and resulted in higher levels of protein nitrosylation in the presence of NO donors. Nitrosative stress exerted a more severe effect on mutant cells than on the wild-type. Interestingly, the phenotypes were observed under both aerobic and anaerobic growth conditions suggesting that YHB1 can detoxify NO and related species irrespective of the presence of O2 and fuelling further discussion of the relative efficacy of bacterial flavohemoglobins in anoxic NO removal. Kinetic evidence for NO-consuming activity of YHB1 was subsequently presented by Gardner et al. (2000b). Studies of the globin of Candida norvengesis have been resurrected after more than ten years (Kobayashi et al., 2002), in the light of new data on other flavohaemoglobins. During purification on butyl-Toyopearl resin, the FAD is partly lost, as has been observed in the purification of other flavohaemoglobins. Loss of the flavin renders the protein readily autoxidisable to the met-form. It is suggested that, unlike bacterial flavohaemoglobins, the C. norvegensis globin might serve as an O2 storage protein in aerobic conditions. Although such a role has been discounted in many other cases on the basis of low abundance of the globin in vivo, it is worth noting that, in this yeast, intracellular levels are relatively high (Oshino et al., 1973a). The flavohaemoglobin from Fusarium oxysporum has been introduced into Pseudomonas stutzeri (Takaya and Shoun, 2002), resulting in a strain that emits less nitrous oxide. The basis of this phenomenon is poorly understood.
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5. Evolution of Globins The finding that globins occur in plants, animals, lower eukaryotes and bacteria has fuelled speculation on the evolution of this ubiquitous superfamily (Perutz, 1986). Although it has sometimes been suggested (Takagi, 1993) that the truncated globins described in Section 3 might have arisen from a different ancestor, these proteins have all the essential determinants of the globin fold and are considered, along with the other bacterial globins, to be ‘‘true’’ globins (Moens et al., 1996). Keilin long ago suggested that globins may have their origins in primitive haem proteins and Runnegar has proposed that globins may derive from cytochromes of the b5 type (for references, see Hardison, 1996). Moens et al. (1996) have suggested that all globins evolved from an ancestral haem protein of about 17 kDa that displayed the globin fold (characterised by the eight helices A to H), which functioned as a redox protein. The identification of haemoglobins in Deinococcus and Aquifex, regarded as ‘primitive’, suggests that they existed about 2 109 years ago, i.e. before O2 existed in Earth’s atmosphere. Under such conditions we may speculate that NO transformations and detoxification were a/the primary function. Since NO toxicity is potentiated by O2, Hausladen et al. (2001) suggest that the NO/O2 (denitrosylase) reaction evolved to detoxify the two ligands, while retaining the preference for binding NO. An increasing concentration of O2 on Earth, as a result of microbial photosynthesis, may have led to the evolution of O2 reactivity as a primary function. Moens et al. (1996) further suggest that formation of the chimeric flavohaemoglobins was an additional and distinct evolutionary trend. An evolutionary tree reveals three apparently discrete clusters, one containing the globins of metazoans and plants, another the ciliates, Chlamydomonas and Nostoc, and a third the yeasts and the proteobacteria Escherichia, Ralstonia and Vitreoscilla (Moens et al., 1996). On the other hand, rRNA analyses and the generally accepted view that the eukaryotic/prokaryotic split was decisive, some 1,800 million years ago, argue that all bacteria should be grouped together. To reconcile these opposing lines of evidence, Moens et al. (1996) suggest that horizontal globin gene transfer occurred from an ancient ancestor, common to the yeasts Saccharomyces and Candida, to a bacterial ancestor of Escherichia, Ralstonia and Vitreoscilla. The flavoprotein domain was then lost from the line leading to the Vitreoscilla globin. If we accept that the prime function of the flavohaemoglobins is aerobic, as in NO oxygenase
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(denitrosylase) activity, it might be speculated that this gene transfer followed O2 appearance. The finding that NO detoxifying activity is conserved among the flavohaemoglobins suggests that reaction of NO with O2 may have been an early function for these proteins and possibly their prime function.
6. SUMMARY In the past ten years, it has become clear that the functions of many, but not all, classes of microbial globins relate to tolerance to NO or other forms of nitrosative stress. This is a marked shift in perception from earlier work of twenty or thirty years ago, in which speculations on possible function were dominated by the ‘classical’ view of the globins of higher organisms, namely that O2 storage and/or transport were pre-eminent. Interestingly our views on those globins has altered also and haemoglobin is sometimes considered an ‘honorary enzyme’. Although it is widely accepted that some microbial globins function in NO stress tolerance, and NO-consuming activities have often been demonstrated in whole cells, extracts and for the purified proteins, there is in many cases a lack of sound physiological and genetic evidence that NO tolerance is the sole or dominant function. Only in a very small number of cases has the globin been (a) shown to be up-regulated in response to NO, (2) deleted by mutation with clear phenotypic consequences, and (c) shown to exhibit NO-detoxifying activities commensurate with the rates of NO removal required to alleviate stress and under conditions (e.g. of O2 supply) that are likely to prevail in the microbe’s natural habitat. Conclusions regarding true physiological function should not be drawn solely based on studies with the purified protein or on the effects of massive overexpression of the globin in either the organism in which the globin is naturally found or, worse, in heterologous hosts. Until such criteria are met, it would be prudent to continue to consider other roles, including participation in O2 metabolism and perhaps hitherto unknown enzymic roles. It should be remembered that although the major function of mammalian haemoglobin is O2 delivery, it has a subsidiary role in NO transport and conservation or in NO consumption (see Joshi et al., 2002 and references therein). Whatever the outcome of these studies, the interdisciplinary study of microbial globins will remain a challenging area for study at the crossroads of physiology, biochemistry, molecular biology and genetics.
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ACKNOWLEDGEMENTS Work in RKP’s laboratory has been generously supported by the Biotechnology and Biological Sciences Research Council (BBSRC, UK) and The Royal Society. We are grateful to our many colleagues and collaborators, especially Professor Martin Hughes (Kings College London) for his guidance in nitric oxide chemistry.
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Author Index
Page numbers in italics indicate where a reference is given in full. Names beginning de, van and von have been listed under their respective alphabets. Abedin, M.J., 35, 48 Abel, C.B.L., 93, 109 Adams, M.W., 203, 240 Affronti, L.F., 43, 48 Agar, J.N., 203, 240 Agawin, N.S.R., 43, 48 Agusti, S., 43, 48 Aiba, H., 35, 48 Akhtar, M., 136, 176 Alami, M., 191, 222, 240, 243 Alberte, R.S., 9, 12, 48, 49 Alexander, K., 136, 176 Allen, J.K., 93, 109 Allen, S.C., 230, 232, 240 Almiron, M., 68, 109 Amann, R.I., 73, 77, 109 Ammerman, J.W., 32, 48 Andersen, C., 236, 240 Andersen, J.F., 143, 176 Anderson, J.I., 77, 109 Andersson, C.I., 267, 300 Andersson, C.R., 260, 300 Andrews, S.C., 279, 283, 300 Angelini, S., 216, 240 Angerer, A., 37, 48 Anjum, M.F., 283, 284, 288, 289, 300 Antia, N.J., 3, 36, 48, 60 Aoyama, Y., 136, 176, 186 Appleby, C.A., 134, 176, 260, 300 Arana, I., 79, 109 Armbrust, E.V., 40, 42, 48 Asakura, H., 99, 110 Ascenzi, P., 258, 300, 303 Attar, R.M., 164, 176
August, P.R., 143, 176 Azam, F., 20, 32, 48, 56 Be´ja`, O., 3, 46, 49 Bo¨ck, A., 205, 241, 245, 247, 249, 253 Bo¨ger, P., 29, 58 Bachman, M.A., 71, 110 Badger, M.R., 14, 15, 17, 48 Bailey, J.E., 263, 265, 266, 267, 300, 304 Baitsch, D., 198, 240 Barer, M.R., 73, 74, 76, 81, 84, 107, 110, 114, 115, 118 Barlow, R.G., 12, 48, 49 Barondness, J.J., 45, 49 Baross, J.A., 70, 110 Barrow, P.A., 93, 110 Batchelor, S.E., 105, 110 Bateman, A., 103, 110 Beckwith, J., 45, 49, 210, 250 Beinert, H., 288, 304 Bell, S.G., 175, 176 Bellamine, A., 137, 154, 158, 177 Bentley, S.D., 67, 102, 110, 216, 240 Berg, B.L., 236, 237, 240 Berg, H.C., 39, 58 Berger, A.S., 97, 110 Berks, B.C., 191, 192, 195, 200, 201, 206, 208, 212, 215, 216, 223, 225, 227, 230, 233, 234, 235, 237, 240, 241 Berlin, D.L., 99, 110 Bernhard, M., 207, 237, 241 Bernstein, H.D., 195, 246 Berrie, J.R., 143, 177 Besnard, V., 75, 110
312 Betanzos, M., 236, 241 Betts, J.C., 70, 83, 89, 96, 110 Biketov, S., 74, 90, 97, 103, 104, 110 Binder, B., 40, 41, 49 Binder, B.J., 40, 41, 42, 49 Binder, U., 205, 241 Binnerup, S.J., 100, 111 Bird, C., 30, 49 Bird, C.W., 154, 177 Bjo¨rkman, K., 31, 32, 49 Black, S.D., 135, 168, 177 Blackburn, N., 39, 49 Blanchot, J., 27, 49 Blasco, F., 201, 241 Bloemberg, G.V., 75, 111 Blokesch, M., 205, 206, 241, 245 Bloomfield, S.F., 88, 97, 111 Bockian, R., 76, 99, 123 Bogosian, G., 80, 81, 97, 98, 100, 105, 111 Bogsch, E., 191, 192, 206, 219, 241 Bolhuis, A., 221, 222, 225, 241 Bollinger, C.J., 267, 300 Bolognesi, M., 264, 266, 300 Bonamore, A., 281, 282, 300 Boon, C., 96, 111 Borgese, M.B., 40, 42, 60 Boscott, P.E., 174, 177 Boucher, S.N., 74, 111 Boucher, Y., 46, 49 Bouvier, P., 154, 177 Bovill, R.A., 97, 105, 111 Boxer, D.H., 205, 206, 251, 253 Brahamsha, B., 39, 47, 49, 50 Brand, L.E., 36, 37, 38, 50, 56 Breskvar, K., 164, 177 Breyton, C., 189, 241 Briza, P., 163, 165, 177 Brock, I.W., 233, 241 Bronk, D.A., 28, 50 Brooks, J.V., 90, 111 Brown, K., 209, 242 Brown, M.H., 74, 117 Brown, S., 74, 110 Brunori, M., 258, 300 Buchanan, G., 195, 196, 198, 220, 230, 231, 232, 242, 247, 252 Burmester, T., 271, 301 Burnap, R.L., 38, 50
AUTHOR INDEX Bussmann, I., 97, 111 Bycroft, M., 103, 110 Caipo, M.L., 106, 111 Calcott, P.H., 97, 111 Campbell, L., 2, 6, 10, 40, 42, 43, 44, 50, 53, 58 Cao, J.G., 92, 111 Cappelier, J.M., 99, 111, 127 Carafoli, E., 33, 58 Caron, D.A., 44, 50 Carpenter, E.J., 2, 6, 40, 42, 43, 44, 50 Carr, N.G., 3, 6, 50, 51, 57 Casalot, L., 207, 242 Casamayor, E.O., 6, 50 Cashel, M., 71, 88, 111, 115 Casula, G., 99, 112 Cavicchioli, R., 77, 114 Chadd, H.E., 36, 38, 50 Chaiyanan, S., 76, 112 Chan, A.M., 45, 60 Chan, J., 89, 91, 114 Chanal, A., 219, 242, 251 Chen, L., 210, 211, 242 Chen, L.-Y., 208, 242, 247 Chen, Z.W., 211, 242 Chisholm, S.W., 4, 40, 42, 49, 51, 56, 57, 61 Chistoserdov, A.Y., 210, 211, 242 Chmielewski, R.A., 79, 112 Cho, J.C., 75, 112 Christaki, U., 44, 51 Christensen, S.K., 71, 112 Clark, S.A., 192, 242 Clegg, J.S., 69, 112 Cleverley, R.M., 195, 242 Cline, K., 191, 221, 222, 223, 225, 235, 236, 242, 247, 248, 252 Coates, A.R., 96, 117 Cobley, J.G., 45, 51 Cohn, M.L., 85, 112 Cole, J.A., 200, 250 Cole, S.T., 91, 102, 112, 275, 301 Collier, J.L., 8, 19, 20, 21, 22, 27, 36, 51, 53 Colwell, R.R., 73, 76, 99, 107, 112, 124, 125 Contreras, I., 219, 243 Cook, G.M., 260, 274, 307 Coon, M.J., 135, 168, 177 Corper, H.J., 85, 112 Cosme, J., 174, 178 Cotner, J.B., 32, 51
313
AUTHOR INDEX Couture, M., 270, 271, 272, 273, 275, 301, 307, 309 Cramm, R., 286, 295, 301 Crawford, M.J., 286, 289, 291, 295, 296, 301 Cristo´bal, S., 192, 194, 195, 196, 214, 238, 243 Crooke, H., 219, 243 Crosbie, N.D., 12, 44, 52 Cross, R., 296, 301 Crowe, J.H., 69, 112 Cruz-Ramos, H., 288, 289, 301 Cuhel, R.L., 32, 51 Curtis, T.P., 73, 112 Cutting, S.M., 99, 112 Danese, P.N., 104, 112 Darby, C., 218, 243 Das, T.K., 270, 271, 272, 275, 301, 302 Davey, H.M., 73, 74, 89, 95, 112, 113 Davies, D.G., 92, 113 Davies, R., 92, 112 Dawes, E.A., 72, 113 Decross, A.J., 90, 113 de Gier, J.-W., 189, 243–247 de Leeuw, E., 219, 220, 221, 222, 223, 224, 229, 235, 242, 243, 250, 251 DeLisa, M.P., 195, 196, 243 del Mar Lleo, M., 75, 113, 126 de Lorimier, R., 9, 51 Delye, C., 171, 173, 178 Deming, J.W., 70, 110 Denariaz, G., 295, 302 de Pina, K., 204, 243 Deretic, V., 89, 113 Devlin, J.P., 93, 113 de Wit, D., 75, 90, 113 Dhillon, J., 90, 113 Diaz, G.A., 83, 128 Dickie, P.M., 43, 55 Didenko, L.V., 78, 99, 113 Dietzler, D.N., 72, 113 Dikshit, K.L., 261, 264, 265, 266, 267, 302, 304 Dikshit, R.P., 265, 266, 302 Dobbek, H., 198, 243 Dolan, J.R., 43, 44, 51 Dolata, M.M., 201, 243 Domingue, G.J., 75, 91, 113 Donald, K.M., 32, 33, 51
Douglas, S.E., 6, 51 Dreusch, A., 195, 209, 243 Drew, D., 230, 243 Driessen, A.J., 189, 247, 252 DuRand, M.D., 27, 40, 49, 51 Dubini, A., 206, 207, 237, 244 Dubrow, E.L., 91, 113 Duda, V.I., 93, 113 Dukan, S., 88, 100, 113, 114 Duncan, K., 89, 129 Duport, C., 175, 178 Eaves, D.J., 198, 244 Edwards, C., 66, 114, 127 Eguchi, M., 47, 51, 79, 100, 114 Eisenstark, A., 70, 114 Eishi, Y., 91, 114 Eitinger, T., 204, 244 Ekweozor, C.C., 85, 114 Ensign, J.C., 99, 116 Erdner, D.L., 38, 51 Ericsson, M., 83, 100, 114 Eriksson, S., 290, 302 Ermler, U., 281, 282, 302 Ernst, A., 47, 58 Ernst, S., 99, 120 Eschenbrenner, M., 283, 302 Evdokimova, N.V., 74, 82, 98, 114 Falke, J.J., 67, 114 Falkner, G., 33, 35, 52 Falkner, R., 33, 35, 52 Falzone, C.J., 269, 271, 302, 308 Favey, S., 291, 302 Favre, B., 173, 178 Fegatella, F., 77, 114 Ferguson, S.J., 211, 249 Ferris, M.J., 6, 12, 13, 52, 62 Field, K.G., 3, 52 Finkel, S.E., 69, 114, 115 Fischer-Le Saux, M., 75, 114 Flynn, J.L., 89, 91, 114 Fogg, G.E., 3, 12, 18, 52 Fogler, H.S., 104, 118 Foster, S.J., 70, 116 Frau´sto da Silva, J.J.R., 207, 244 Frank, J.F., 79, 112 Franz, R., 171, 172, 173, 178 Fratti, R.A., 89, 113
314 Frazzon, J., 202, 244 Fredricks, D.N., 91, 114, 120 Freeman, R., 103, 114 Frenkiel-Krispin, D., 68, 115 Frey, A.D., 267, 296, 302 Freymann, D.M., 195, 244 Fu, C., 204, 244 Fulco, A.J., 140, 141, 180, 182, 184, 186 Fuller, N.J., 4, 6, 8, 21, 22, 23, 24, 25, 26, 28, 45, 52 Fuqua, C., 92, 115 Furnas, M., 12, 44, 52 Gagne´, G., 275, 303 Gallagher, J.C., 3, 52 Gallant, J., 71, 115 Garcia-Lara, J., 74, 79, 92, 115 Gardner, A.M., 265, 280, 281, 282, 283, 285, 286, 287, 291, 292, 293, 295, 297, 303 Gardner, M.J., 162, 178 Gardner, P.R., 265, 280, 281, 282, 283, 285, 286, 287, 291, 292, 293, 295, 297, 303 Geider, R.J., 36, 52 Geller, B., 192, 244 Ghezzi, J.I., 80, 115 Giaever, G., 78, 115 Giangiacomo, L., 263, 264, 303 Gierasch, L.M., 195, 242 Gilbert, P., 104, 115 Giovannoni, S.J., 3, 52 Glazer, A.N., 8, 9, 11, 52, 57, 60, 62 Glibert, P.M., 12, 19, 20, 29, 52, 54 Glockner, A.B., 210, 244 Glover, H.E., 3, 12, 27, 53 Gobel, U.B., 77, 115 Godden, J.W., 207, 244 Goffeau, A., 163, 178 Goldberg, D., 289, 301 Goldberg, D.E., 286, 289, 291, 295, 301, 306 Gong, L., 71, 115 Goodrich-Blair, H., 92, 115 Gouffi, K., 230, 231, 244 Gould, G.W., 68, 73, 115 Graham, S.E., 133, 140, 178 Granger, J., 38, 53, 60 Grant, G.H., 174, 177 Green, J., 35, 53, 288, 303 Grey, B., 80, 94, 100, 115 Grey, B.E., 94, 99, 115
AUTHOR INDEX Gribbon, L.T., 74, 115 Griesbeck, C., 211, 244 Gross, R., 195, 201, 237, 238, 244, 252 Grossman, A.D., 92, 119 Grossman, A.R., 9, 10, 19, 51, 53 Guengerich, F.P., 132, 178 Guertin, M., 275, 303 Guest, J., 267, 308 Gunsalus, I.C., 139, 178 Gupta, R., 202, 244 Gupta, S., 68, 115 Halbig, D., 195, 214, 244 Haller, C.M., 78, 116 Hamamoto, H., 171, 178 Hambly, E., 45, 53 Han, Y.N., 165, 178 Hangstrom, A., 78, 129 Hanson, D., 17, 48 Hardison, R.C., 298, 303 Hargrove, M.S., 271, 308 Harrison, A.P., 105, 116 Harrison, P.J., 38, 55 Harwood, C.R., 73, 76, 81, 107, 110 Hasona, A., 198, 245 Hassidim, M., 15, 53, 60 Hata, S., 163, 178, 179 Hatchikian, C.E., 203, 249 Hatchikian, E.C., 203, 245 Hausladen, A., 293, 294, 303 Hawkes, D.B., 141, 179 Hayflick, L., 101, 116 Head, I.M., 73, 116 Heathcote, P., 19, 53 Heffernan, W.P., 77, 109 Heidrich, C., 217, 218, 245 Heikkila¨, M.P., 208, 209, 210, 214, 245 Heim, S., 95, 116 Heinzinger, N.K., 213, 245 Heinzle, E., 93, 129 Heller, K.J., 45, 53 Henderson, B., 99, 116 Hengge-Aronis, R., 66, 67, 116, 126 Henley, W.J., 38, 53 Hensel, M., 213, 245 Hentzer, M., 92, 116 Herbert, K.C., 70, 116 Herdman, M., 4, 5, 53 Herna´ndez-Pando, R., 75, 90, 116
315
AUTHOR INDEX Herrero, A., 29, 53 Hess, W.R., 3, 11, 14, 53 Hicks, M.G., 227, 228, 245 Hill, D.R., 274, 303 Hill, R.D., 260, 306 Hilton, J.C., 191, 245 Hinsley, A.P., 196, 202, 245 Hirsch, C.F., 99, 116 Hirsch, P., 77, 116 Hoch, J.A., 66, 67, 81, 116, 126 Hofle, M., 86, 116 Hoge, F.E., 10, 53 Hohn, T.M., 165, 179 Honda, D., 4, 53 Honer zu Bentrup, K., 89, 116 Hood, M., 97, 120 Hood, M.A., 72, 117 Hopwood, D.A., 143, 179 Hu, Y., 90, 96, 97, 117, 278, 291, 303 Hu, Y.M., 83, 113, 117 Hube, M., 205, 245 Huber, H., 70, 117 Huber, R., 70, 117 Hughes, M.N., 258, 273, 284, 291, 307, 308 Huisman, G.W., 92, 117 Hunt, J.F, 195, 245 Hunter, J.R., 72, 124 Huntsman, S.A., 36, 60 Hurst, A., 68, 73, 115 Hutchins, D.A., 38, 53 Hutchinson, C.R., 143, 176 Hvitved, A.N., 271, 272, 275, 303, 309 Hynds, P.J., 192, 245 Ignatova, Z., 216, 245 Iida, T., 164, 179, 183 Iijima, M., 287, 303 Ikemoto, E., 76, 119 Ikeya, T., 32, 33, 34, 54 Ilari, A., 296, 304 Imlay, J.A., 208, 245 Imlay, K.R., 208, 245 Ioannidis, N., 280, 281, 304 Islam, M.S., 76, 78, 117 Iturriaga, R., 10, 50 Ize, B., 195, 196, 216, 217, 245, 246 Jack, R.L., 220, 246 Jackson, C.J., 138, 154, 155, 158, 179
Jacquet, S., 40, 41, 42, 43, 54 Jang, J.W., 102, 103, 117 Janssen, P.H., 77, 117, 125 Ji, H.T., 174, 179 Jiang, Y., 236, 246 Jishage, M., 71, 117 Johnson, E.F., 174, 178 Johnston, M.D., 74, 117 Joint, I., 8, 54 Jongbloed, J.D., 216, 246 Jongbloed, J.D.H., 216, 246 Joo, H., 175, 179 Jormakka, M., 213, 236, 237, 246 Joshi, M., 266, 267, 304 Kaasen, I., 67, 126 Kaeberlein, T., 77, 117 Kahn, R.A., 136, 179 Kaiser, D., 92, 104, 117, 120 Kakeya, H., 173, 179 Kalakoutskii, L.V., 68, 117 Kalb, V.F., 136, 179 Kallio, P.T., 267, 304 Kalmbach, S., 77, 78, 117 Kana, T.M., 12, 14, 19, 54 Kaprelyants, A.S., 66, 68, 73, 74, 75, 77, 81, 85, 86, 97, 98, 99, 101, 105, 106, 112, 118, 127 Karagouni, A.D., 15, 54 Karl, D.M., 31, 32, 49, 54 Katagiri, M., 139, 179 Kaur, R., 261, 265, 267, 304 Kebir, M.O., 195, 246 Keenan, R.J., 195, 246 Keer, J., 88, 95, 118 Keilin, D., 69, 118, 257, 304 Kell, D.B., 68, 73, 74, 76, 77, 80, 81, 82, 85, 86, 89, 91, 92, 93, 95, 97, 98, 99, 100, 101, 102, 110, 113, 118, 129 Kelly, D.E., 133, 136, 158, 163, 169, 170, 172, 173, 179, 180, 181, 184 Kelly, S.L., 133, 136, 158, 163, 169, 170, 172, 173, 179, 180, 181, 182, 184, 185 Kendall, D.A., 195, 246 Khomenko, A.G., 90, 91, 118 Khosla, C., 263, 265, 266, 267, 304 Kieser, T., 143, 180 Kiley, P.J., 288, 304 Kim, C., 208, 246
316 Kim, D.S., 104, 118 Kim, K.S., 72, 118 Kim, S.J., 72, 75, 112, 117 Kim, S.O., 283, 287, 294, 295, 304 King, D.J., 136, 180 Kingston, R.L., 201, 246 Kitazume, T., 165, 180 Kjelleberg, S., 70, 71, 74, 78, 81, 94, 97, 119, 121, 123, 126, 128 Klabunde, T., 208, 246 Kleerebezem, M., 92, 119 Klein, M.L., 141, 180 Klingenberg, M., 134, 180 Klipp, W., 201, 247 Kobayashi, G., 297, 305 Koch, A.L., 18, 54, 72, 119 Kogure, K., 75, 76, 119 Kolber, Z.S., 47, 54 Kolling, G.L., 100, 119 Kolowith, L.C., 34, 35, 54 Kolter, R., 69, 70, 81, 92, 93, 94, 96, 104, 112, 114, 115, 117, 119, 123, 126, 128, 129 Kornberg, A., 72, 124 Kozdroj, J., 78, 119 Krafft, T., 201, 213, 246 Kramer, J.G., 20, 54 Krebs, C., 203, 246 Kroneck, P.M.H., 209, 254 Kudo, I., 38, 55 Kudoh, S., 43, 55 Kuhar, I., 71, 119 Kuipers, B., 40, 55 Kullman, S.W., 166, 180 Kurokawa, M., 99, 119 Kuznetsova, Z.A., 91, 119 La Roche, J., 36, 38, 52, 55 Lacelle, M., 290, 305 Lake, M.W., 198, 246 Lamb, D.C., 133, 137, 138, 143, 145, 150, 154, 168, 171, 172, 173, 174, 179, 180, 181, 185 Lantoine, F., 8, 11, 55 Larson, L.J., 135, 181 Law, C.S., 30, 61 Lazazzera, B.A., 92, 119 Leatham, T., 3, 63 Lebeault, J.M., 164, 181 Lebioda, L., 258, 305
AUTHOR INDEX Lecomte, J.T.J., 271, 305 Lee, A., 228, 246 Lee, H.C., 195, 246 Lee, K.-H., 78, 119 Lee, P.A., 228, 247 Lee, Y.-H.W., 208, 247, 253 Leimkuhler, S., 201, 247 Lentz, O., 175, 181 Lesn, J., 79, 119 Leu, W.-M., 195, 208, 242 Leu, W.M., 195, 208, 247 Lewis, K., 68, 119 Lewis, M.R., 19, 55 Li, H., 33, 55, 175, 186 Li, H.Y., 141, 181 Li, Q.S., 175, 181 Li, R., 104, 111 Li, W.K.W., 2, 43, 55 Li, Y.H., 104, 119 Lidstrom, M.E., 210, 242 Lillebaek, T., 85, 91, 119 Lindberg, R.L.P., 145, 181 Lindell, D., 19, 29, 30, 31, 55 Lindenmayer, A., 134, 182 Liu, H., 40, 41, 43, 44, 55 Liu, L.M., 287, 305 Liu, M.T., 198, 247, 254 Liu, Y.C., 41, 43, 49 Lleo, M.M., 79, 100, 119 Loeffler, J., 172, 173, 182 Loginov, A.S., 90, 120 Lonvaud-Funel, A., 107, 121 Loper, J.C., 164, 168, 184, 185 Losick, R., 92, 104, 117, 120 Lowder, M., 75, 120 Luirink, J., 189, 243, 247 Lupetti, L., 172, 182 Luque, I., 19, 29, 30, 55, 56 Lynch, T., 91, 120 Lyte, M., 99, 120 Mu¨ller, H., 44, 57 Ma, X., 222, 225, 247 MacDonell, M.T., 97, 120 MacMicking, J.D., 274, 305 Macheroux, P., 283, 305 Mackey, B.M., 97, 105, 111 Madhavan, H.N., 91, 120 Madueno, F., 202, 247
317
AUTHOR INDEX Magalon, A., 198, 205, 247 Magarinos, B., 74, 78, 120, 124 Magnani, P., 237, 247 Mah, T.F., 104, 120 Maier, R.J., 204, 205, 219, 244, 249 Maier, T., 205, 247 Maiwald, M., 91, 120 Mandrand, M.-A., 237, 254 Mandrand-Berthelot, M.A., 200, 204, 244, 247 Manefield, M., 92, 120 Mann, E.L., 36, 37, 56 Mann, N.H., 3, 50, 57 Manting, E.H., 189, 247 Marichal, P., 173, 182 Marie, D., 43, 61 Marouga, R., 71, 121 Marshall, B.J., 90, 113, 121 Marshall, H.G., 43, 48 Martinez, J., 20, 56 Mary, P., 79, 100, 121 Masaphy, S., 164, 166, 182 Mascher, F., 74, 78, 80, 121 Matsuda, M., 102, 121 Matsumura, F., 166, 180 Matsuoka, T., 143, 184 Matthews, K.R., 100, 119 Mayfield, C.I., 79, 121 McCune, R.M., 89, 121 McDevitt, C.A., 201, 247 McDougald, D., 81, 95, 121 McGowan, S., 92, 121 McGuirl, M.A., 209, 248 McLean, K.J., 158, 182 Meighen, E.A., 92, 111, 121 Mekalanos, J.J., 45, 61 Mellado, E., 169, 182 Membrillo-Herna´ndez, J., 286, 288, 289, 291, 305 Menendez, C., 197, 248 Menon, N.K., 206, 207, 248 Michel, K.P., 37, 56 Mierle, G., 18, 56 Miguelez, E.M., 99, 121 Mihara, H., 203, 248 Milani, M., 269, 272, 273, 305, 306 Miller, E., 212, 248 Millet, V., 107, 121 Mills, C.E., 264, 280, 283, 284, 293, 295, 306 Mingot, J.M., 165, 182
Minning, D.M., 258, 306 Mitsui, A., 42, 56 Miyazaki, E., 90, 121 Mizuno, T., 35, 48 Mizutani, M., 168, 182 Moens, L., 260, 298, 306 Moffett, J.W., 36, 37, 56, 59, 62 Moir, A., 101, 121 Molik, S., 196, 202, 216, 248 Moore, J., 75, 121 Moore, J.E., 75, 98, 121 Moore, L.R., 3, 6, 11, 12, 13, 14, 18, 21, 22, 24, 25, 27, 56 Moorehead, P.S., 101, 116 Morel, A., 13, 56 Morgan, J.A., 74, 121 Mori, H, 220, 248 Mori, H., 191, 220, 221, 222, 223, 225, 235, 236, 242, 248 Mori, J., 223, 235, 248 Mori, T., 42, 56 Morita, R., 77, 127 Morita, R.Y., 66, 70, 71, 72, 77, 78, 122, 123 Morris, I., 20, 54 Morrison, D.A., 92, 122 Mould, R.M., 191, 248 Mukai, M., 265, 270, 272, 273, 274, 281, 282, 284, 296, 306 Mukamolova, G.V., 74, 85, 86, 93, 97, 98, 100, 101, 102, 103, 106, 122 Munro, A.W., 141, 182 Muratov, V.V., 90, 118 Musser, S.M., 191, 222, 233, 248 Nagy, I., 140, 182 Nair, U., 40, 57 Nakano, M.M., 290, 306 Narhi, L.O., 140, 182 Navarro, C., 204, 248 Neal, C.P., 99, 122 Nebe-von-Caron, G., 107, 122 Negishi, M., 145, 181 Neidhardt, F.C., 66, 68, 122, 123 Neijssel, O.M., 94, 122 Nelson, D.R., 133, 136, 183 Neuer, S., 43, 57 Neumann, A., 212, 248 Neveux, J., 8, 11, 55 Newman, J., 9, 19, 57
318 Nichols, J., 198, 249 Nickle, D.C., 69, 122 Nicolet, Y., 203, 245, 249 Nie, X., 260, 306 Nikolaev, I.A., 105, 122 Nilsson, H.O., 75, 82, 99, 123 Nilsson, L., 98, 123 Novitsky, J.A., 78, 123 Nurizzo, D., 202, 249 Nystrom, T., 68, 80, 88, 94, 114, 123 O’Keefe, D.P., 140, 146, 183 O’Toole, G., 104, 123 O’Toole, G.A., 104, 120 Ochsner, U.A., 218, 249 Ohkawa, H., 15, 57 Ohkuma, M., 164, 183 Ohta, A., 168, 179, 183 Ohta, D., 168, 182 Ohtomo, R., 99, 123 Oliver, J.D., 74, 75, 76, 78, 79, 95, 99, 105, 120, 123, 129 Oliver, K., 75, 112 Ollesch, G., 261, 282, 296, 306 Olson, J.W., 204, 205, 219, 249 Olson, R.J., 8, 10, 43, 51, 57 Omata, T., 29, 57 Omura, S., 143, 183 Omura, T., 134, 183 Ong, L.J., 9, 11, 57, 60 Oresnik, I.J., 200, 249 Orii, Y., 264, 306 Ortiz de Montellano, P.R., 132, 183 Oshino, N., 275, 279, 285, 297, 306 Oshino, R., 275, 279, 285, 297, 306 Ostling, J., 70, 123 Ostroff, R.M., 218, 249 Ouellet, H., 274, 306 Ouellet, Y., 274, 306 Pacheco, S.V., 45, 57 Paerl, H.W., 20, 57 Page, M.D., 211, 249 Pai, S.R., 75, 90, 123 Paidhungat, M., 99, 101, 123 Palenik, B., 5, 6, 8, 10, 12, 13, 21, 22, 47, 51, 52, 57, 60 Paludan-Muller, C., 95, 123 Park, K.W., 265, 268, 306
AUTHOR INDEX Park, S.Y., 165, 183 Parrish, N.M., 89, 123 Partensky, F., 3, 4, 8, 12, 27, 44, 57, 58 Paschos, A., 204, 249 Pathania, R., 269, 272, 274, 306 Pedersen, P.L., 33, 58 Peltier, J.-B., 192, 195, 249 Perozo, E., 236, 249 Perutz, M., 298, 307 Pesce, A., 208, 249, 269, 272, 273, 279, 307 Petersen, F., 106, 123 Peterson, J.A., 133, 140, 178, 183, 184 Pitonzo, B.J., 78, 124 Pitta, T.P., 39, 58 Platt, T., 20, 55 Podust, L.M., 137, 158, 174, 183 Poindexter, J., 72, 124 Polyanskaya, L.M., 106, 124, 127 Pomar, M.L.C.A., 6, 58 Pommier, J., 200, 250 Pomposiello, P.J., 288, 307 Poock, S.R., 219, 250 Poole, R.K., 258, 260, 263, 273, 274, 283, 284, 285, 286, 287, 288, 289, 291, 301, 305, 307, 308 Porcelli, I., 221, 225, 229, 235, 243, 250 Post, A.F., 31, 55 Postgate, J.R., 68, 72, 81, 97, 104, 106, 111, 124 Postius, C., 3, 29, 47, 58 Potter, L.C., 200, 250 Potts, M., 274, 307 Poulos, T.L., 139, 141, 181, 183 Poupin, P., 140, 183 Pouzharitskaja, L.M., 68, 117 Pre´zelin, B.B., 40, 43, 58 Price, G.D., 17, 48 Price, N.M., 38, 51, 53, 60 Primas, H., 81, 124 Primm, T.P., 71, 124 Probst, I., 275, 279, 283, 287, 307 Pugsley, A.P., 189, 250 Quadri, L.E., 92, 119 Rahman, I., 76, 95, 124 Rahme, L.G., 218, 250 Rain, J.C., 204, 250 Rajagopalan, K., 191, 215, 254
AUTHOR INDEX Rajagopalan, K.V., 191, 198, 215, 246, 247, 249, 251, 252, 253, 254 Ramaiah, N., 74, 124 Ramsing, N.B., 47, 58 Rao, N.N., 72, 124 Rappe´, M., 47, 58 Rasmussen, T., 209, 250 Ravel, J., 95, 124 Ravichandran, K.G., 141, 183 Ray, B., 76, 98, 124 Ray, R.T., 19, 20, 29, 52 Relman, D.A., 91, 114, 120 Reusch, R.N., 93, 124 Rheinwald, J.G., 139, 184 Riester, J., 209, 250 Rinas, U., 93, 124 Rippka, R., 4, 5, 30, 32, 58, 62 Rittman, B., 104, 124 Ritz, D., 210, 250 Rivers, B., 91, 124 Roberts, G.A., 161, 184 Roberts, S.A., 208, 250 Robertson, B.R., 4, 58 Robinson, C., 191, 248 Robinson, N.J., 36, 58 Rocap, G., 6, 10, 21, 22, 23, 39, 56, 58, 60 Rodrigue, A., 191, 250 Roger, A.J., 156, 184 Romalde, J.L., 74, 78, 120, 124 Romanova, I.M., 75, 95, 125 Rondon, M.R., 73, 125 Roos, V., 267, 307 Rosano, C., 204, 250 Rosazza, J.P., 143, 184 Rose, R.W., 195, 216, 250 Roszak, D.B., 73, 100, 107, 125 Rousset, M., 207, 242 Rowbury, R.J., 105, 125 Rubin, H., 101, 125 Ruby, E.G., 78, 119 Rudolph, M.J., 198, 251 Rueter, J.G., 38, 59 Ruettinger, R.T., 140, 184 Russell, D.G., 89, 116 Ryley, J.F., 257, 304 Sadoff, H.L., 93, 124 Saito, M., 99, 123 Saito, M.A., 36, 37, 59
319 Salmond, G.P., 92, 121, 125, 129 Samuelson, J.C., 190, 251 Sanders, C., 192, 214, 251 Sanglard, D., 164, 172, 173, 184 Santana, M.A., 260, 308 Santini, C.-L., 191, 192, 196, 200, 217, 240, 242, 251 Santini, C.L., 191, 200, 241 Sargent, F., 190, 191, 196, 200, 206, 213, 216, 219, 220, 221, 225, 233, 234, 236, 237, 238, 242, 243, 244, 246, 251, 252 Sariaslani, F.S., 143, 184 Sartori, G., 260, 308 Sasaki, S., 196, 251 Sato, R., 134, 183 Sauer, J., 19, 59 Saunders, N.F., 209, 251 Savage, D.C., 293, 308 Sawers, R.G., 206, 251 Scanga, C.A., 90, 125 Scanlan, D.J., 3, 4, 18, 32, 34, 35, 55, 59, 62 Schaerlaekens, K., 208, 251 Schena, M., 89, 125 Schlegel, H.G., 275, 307 Schlichting, I., 140, 184 Schmid, B., 215, 251 Schmid, E., 146, 184 Schmidt, M., 271, 308 Schuster, K.C., 88, 125 Schut, F., 69, 72, 77, 125 Schutz, M., 201, 252 Scott, C., 271, 301 Scott, N.L., 271, 302, 308 Scotti, P.A., 190, 252 Senior, P.J., 72, 113 Serizawa, N., 143, 184 Setlow, P., 101, 123 Settles, A.M., 220, 225, 252 Sevrioukova, I.F., 141, 184 Shahamat, M., 79, 125 Shalapenok, A., 10, 59 Shalapenok, L.S., 10, 59 Shalapyonok, A., 8, 59 Shalapyonok, L.S., 8, 59 Shapiro, H.M., 74, 125 Shapiro, J.A., 104, 125 Sherman, D.R., 96, 125 Sherry, N.D., 43, 59 Shimada, A., 13, 14, 59
320 Shleeva, M.O., 74, 83, 84, 85, 93, 97, 98, 103, 104, 105, 108, 126 Shoun, H., 297, 308 Shyadehi, A.Z., 172, 184 Sidler, W.A., 9, 60 Sieburth, J. McN., 2, 54 Siegele, D.A., 81, 96, 119, 126 Signoretto, C., 74, 113, 126 Sijbrandi, R., 190, 252 Silberman, J.D., 156, 184 Silhavy, T.J., 66, 116 Simek, K., 43, 44, 51 Simon, J., 201, 244, 252 Sitnikov, D.M., 92, 126 Skaggs, B.A., 163, 184 Slaughter, J.C., 93, 126 Sligar, S.G., 139, 185 Smeulders, M.J., 82, 126 Smith, D.A., 101, 121 Smith, K.E., 164, 185 Smith, L., 134, 182 Smith, R.J., 76, 126 Sohaskey, C.D., 83, 89, 128 Solomon, P.S., 197, 252 Speck, M.L., 76, 98, 124 Spector, M.P., 70, 126 Spiro, S., 267, 308 Sramek, H.A., 83, 128 Srinivasan, S., 92, 126 Stanley, N.R., 195, 196, 200, 208, 214, 216, 217, 237, 242, 245, 247, 251, 252, 254 Steck, T.R., 80, 91, 94, 99, 100, 115, 124 Steinert, M., 99, 126 Stephens, K., 92, 126 Stevanin, T., 287, 308 Stevanin, T.M., 287, 293, 308 Stevenson, C.E., 198, 252 Stiefel, E.I., 203, 240 Stock, A.M., 67, 126 Stockner, J.G., 3, 60 Stoodley, P., 104, 126 Storz, G., 66, 67, 126 Stramski, D., 40, 42, 60 Strauch, M.A., 81, 126 Strom, A.R., 67, 126 Summer, E.J., 238, 239, 252 Sun, Z., 85, 93, 105, 126 Sunda, W., 36, 63 Sunda, W.G., 36, 60
AUTHOR INDEX Sundareshwar, P.V., 34, 60 Sutter, T.C., 168, 185 Suttle, C.A., 45, 60 Suyama, A., 212, 252 Swanson, M.S., 71, 110 Swanson, R.V., 11, 60 Sweeney, B.M., 40, 42, 60 Tabita, F.R., 14, 62 Tada, Y., 77, 126 Tajkhorshid, E., 235, 252 Takagi, T., 260, 298, 308 Takahashi, A., 35, 60 Takahashi, Y., 202, 252 Takaya, N., 297, 308 Takayama, K., 71, 126 Tang, L., 142, 185 Tarran, G.A., 8, 60 Tarricone, C., 261, 265, 266, 308 Taylor, M., 143, 146, 185 Tchernov, D., 15, 60 Tempest, D.W., 94, 122 Temple, C.A., 191, 215, 246, 252 Testerman, T.L., 81, 127 Teter, S.A., 233, 253 Theg, S.M., 191, 192, 222, 233, 242, 248, 253 Theodoratou, E., 205, 253 Tholozan, J.L., 74, 127 Thomas, C., 79, 127 Thomas, J.D., 192, 196, 217, 241, 253 Thorsteinsson, M.V., 274, 303, 308 Ting, C.S., 3, 60 Tissieres, A., 257, 304 Tjepkema, J.D., 275, 308 Tokumoto, U., 202, 252 Toledo, G., 6, 8, 39, 60 Torrella, F., 77, 127 Torsvik, V., 73, 76, 77, 127 Tortell, P.D., 36, 60 Townsend, D., 6, 63 Tranier, S., 200, 253 Trent, J.D., 70, 127 Trent, J.T., 271, 308 Trick, C.G., 37, 61, 63 Triger, Y.G., 106, 124, 127 Troitskaia, V.V., 78, 127 Truglio, J.J., 198, 253 Trzaskos, J.M., 136, 185 Tsai, P.S., 267, 308
AUTHOR INDEX Tsai, T.-Y., 208, 253 Tudzynski, B., 165, 185 Turner, K., 82, 127 Turpin, P.E., 74, 127 Tyree, B., 263, 264, 309 Tyrell, T., 30, 61 Tyson, C.A., 139, 185 Unsworth, N.L., 38, 59 Urbach, E., 6, 61 Urban, P., 166, 185 Valent, Q.A., 195, 253 Valladares, A., 27, 61 van de Pas, B.A., 212, 253 van Elsas, J.D., 78, 119 van Overbeek, L.S., 74, 78, 127 van Waasbergen, L.G., 19, 61 Varon, M., 93, 127 Vasudevan, S.G., 276, 279, 281, 283, 309 Vaulot, D., 41, 43, 51, 61, 63 Velayati, A.A., 91, 127 Venkateswarlu, K., 168, 172, 185 Venrick, E.L., 3, 61 Vignais, P.M., 204, 253 Villee, C.A., 137, 141, 186 Vinogradov, S.N., 260, 309 Visca, P., 275, 309 von Heijne, G., 192, 225, 253 Voordouw, G., 191, 253 Votyakova, T.V., 101, 105, 127 Voulhoux, R., 216, 218, 253 Vreeland, R.H., 69, 128 Vulic, M., 81, 93, 94, 104, 128 Wagner, F., 33, 61 Wagner, G.C., 139, 178 Wagner, K-U., 36, 61 Wai, S.N., 79, 98, 99, 128 Wainwright, M., 77, 128 Wakabayashi, S., 261, 263, 309 Waldor, M.K., 45, 61 Walkenhorst, H.M., 198, 253 Walker, M.B., 220, 253 Wang, Q., 29, 61 Wang, X., 91, 128 Wanner, B.L., 35, 61 Wanucha, D., 74, 79, 123
321 Ward, B.B., 19, 64 Ward, D.M., 47, 62, 76, 77, 128 Warren, J.R., 90, 121 Warrilow, A.G.S., 166, 179, 185 Waterbury, J.B., 2, 3, 4, 5, 20, 21, 22, 23, 24, 25, 32, 39, 42, 43, 51, 62, 63 Watson, G.M.F., 14, 35, 62 Watson, L., 97, 128 Watson, P.F., 169, 185 Watson, S.P., 70, 82, 128 Watts, R.A., 260, 309 Waugh, R., 205, 253 Wayne, L.G., 83, 89, 128 Webb, E.A., 37, 38, 62 Weber, R.E., 260, 309 Webster, D.A., 260, 261, 263, 264, 266, 302, 306, 309 Weichart, D., 74, 75, 78, 79, 80, 81, 93, 97, 98, 123, 128, 129 Weichart, D.H., 82, 93, 100, 129 Weiner, J.H., 190, 219, 227, 254 Wen, L.P., 140, 186 West, N.J., 3, 4, 18, 32, 34, 35, 59, 62 Wexler, M., 219, 254 White, T.C., 171, 172, 173, 186 Whitehead, N.A., 92, 129 Whitesides, D.M., 105, 129 Wickner, W.T., 191, 222, 254 Wickramashighe, R.H., 137, 141, 186 Wilbanks, S.M., 9, 60, 62 Wilhelm, S.W., 37, 61, 62, 63 Willey, J.M., 39, 63 Williams, P.A., 174, 186 Williams, R.J.P., 207, 244 Wilson, W.H., 32, 34, 46, 59, 63 Wingard, L.L., 19, 63 Winson, M.K., 92, 129 Wittenberg, B.A., 265, 268, 269, 270, 273, 279, 306, 309 Wittenberg, J.B., 265, 268, 269, 270, 273, 279, 306, 309 Wittmann, C., 93, 129 Wodara, C., 213, 254 Wong, D.M., 106, 129 Wood, A.M., 3, 6, 8, 9, 10, 11, 63 Woody, Sr., H.B., 75, 91, 113 Wu, L.-F., 237, 248, 254 Wu, L.F., 204, 254 Wyman, M., 19, 30, 40, 43, 49, 63
322 Xiang, S., 198, 254 Xiuren, N., 43, 63 Xu, H.S., 109, 129 Yahr, T.L., 191, 222, 254 Yang, J., 272, 309 Yano, J.K., 161, 186 Yarmolinsky, M.B., 68, 129 Yeh, D.C., 271, 309 Yeh, M., 19, 64 Yeh, S.R., 272, 281, 309 Yen, M.-R., 191, 216, 218, 220, 225, 230, 254 Yeom, H., 141, 186 Yin, Y., 38, 53 Yoshida, Y., 136, 176, 186 Young, C., 165, 186 Young, D.B., 89, 129
AUTHOR INDEX Young, M., 81, 102, 110, 118 Yu, H., 293, 310 Yu, J., 165, 186 Yu, J.J., 165, 186 Yu, T-W., 165, 176 Zambrano, M.M., 104, 129 Zehr, J.P., 19, 47, 64 Zengler, K., 77, 129 Zettler, E.R., 36, 64 Zhang, Y., 83, 85, 93, 98, 105, 126, 129, 160, 186 Zhao, X.-J., 296, 297, 310 Zheng, L., 203, 254 Zhong, Y., 45, 64 Zinser, E.R., 104, 129 Zumft, W.G., 209, 210, 243, 244, 254 Zweifel, U.L., 78, 129
Subject Index
actinomycetes, CYPs 142–3 alarmones, stress 71 alkylresorcinols, TNC 93 amidase puzzle, Tat protein translocation pathway 217–18 ammonium, nitrogen acquisition 20–31 anabolism/catabolism, non-culturable cells 86–9 ancestral forms, CYPs 136–9, 156–8 antibiotics biosynthesis CYP107A1 (P450 eryF) 143–4 CYPs 143–50 archaebacteria, CYPs 161–2 azole inhibition, CYPs 158, 159, 160, 169–74 Bermuda Atlantic Time-Series Station (BATS) 32–3 biofilms, TNC 104 cAMP, phosphorus acquisition 32 Candida, CYPs 164, 169–74 cannibalism, TNC 104–5 carbon-concentrating mechanisms (CCM) 14–17 carbon metabolism carbon fixation 14–17 photosynthetic physiology 11–14 Synechococcus 11–17 CCM see carbon-concentrating mechanisms cell cycle, Synechococcus 39–43
cell death TNC 68–9 see also longevity, bacterial cell wall biosynthesis, Tat protein translocation pathway 217–18 chemical inducers, TNC 92–4 chemotaxis Synechococcus 39 TNC 67 cobalamin cofactors, Tat protein translocation pathway 211–12 consensus tree, Synechococcus 7 cooperative behaviour, TNC 105–6 copper cofactors, Tat protein translocation pathway 207–10 Corynebacterium, CYPs 150 cryptic growth, starvation survival 71–2 cyanobacterial genomes, CCM related genes 14–17 cytochromes P450 (CYPs) actinomycetes 142–3 ancestral forms 136–9, 156–8 anti-mycobacterial activity 158, 159 antibiotics biosynthesis 143–50 archaebacteria 161–2 azole inhibition 158, 159, 160, 169–74 bacterial 139–43, 158–62 biodiversity 131–86 Candida 164, 169–74 carbon monoxide difference spectrum 134–5 catalytic cycle 135–6
324 cytochromes P450 (CYPs) (Continued ) classes 161, 162 Corynebacterium 150 CYP101 (P450cam) 139–40 CYP102A1 (P450 BM-3) 140–1 CYP105D5 146 CYP107A1 (P450 eryF) 143–4 CYP107s 156 CYP158A2 150 CYP51 136–9, 156–8 discovery 134–6 drug resistance 169–74 ergosterol 170–1 eukaryote-like 153 evolutionary scenario 153–4 ferredoxin fusion protein 159–61 fungal 163–74 fusion proteins 159–61 gene transfer, horizontal 138–9 genomes, bacterial 141–2 MTB CYP51 154–6 mycobacteria 150–8 Nocardia 150 nomenclature 133–4 oil-protein conversion 164 phylogenetic tree 138 phylograms 136–7, 152 protists 162–3 roles 143, 144–5 Streptomycetes 143–50
disease states, non-culturable cells 89–92 diversity, genetic, Synechococcus 6 diversity, physiological, Synechococcus 1–64 division cycle Prochlorococcus 41–3 Synechococcus 41–3 drug resistance, CYPs 169–74 drug target, fungal CYPs 169–74
environments fluctuating 66–7 natural 76–9 non-culturable cells 76–9, 99 ergosterol, CYPs 170–1
SUBJECT INDEX erythromycin biosynthesis, CYP107A1 (P450 eryF) 143–4 eukaryote-like CYPs 153 ferredoxin fusion protein, CYPs 159–61 flavohaemoglobins 275–97 clustalW alignment 277 composition 279–80 crystal structures 281–2 discovery 275–9 fungi 296–7 NO-detoxifying activities 291–6 redox activities 282–4 cf. single-domain globins 277 structure 280–2 yeasts 296–7 see also Hmp fungal CYPs 163–74 drug target 169–74 oil-protein conversion 164 Phanerochaete chrysosporium 165–9 gene transfer, horizontal, CYPs 138–9 genetic control, TNC 94–6 genomes, bacterial, CYPs 141–2 GFOR see glucose-fructose oxidoreductase globins classical view 257–60 definition 257–60 enzymes? 258 evolution 298–9 haemoglobin 257–8 myoglobin 257–8 single-domain globins 258–68, 277 see also microbial globins glucose-fructose oxidoreductase (GFOR), Tat protein translocation pathway 201–2 grazing, Synechococcus 44–6 growth irradiance Prochlorococcus 12–14 Synechococcus 12–14 haem coordination, truncated globins 270–1 haemoglobin 257–8 heterogeneity, population, TNC 95–6
SUBJECT INDEX Hmp composition 279–80 discovery 275–9 enzymic properties 282–5 functions 285–7 mutation 291 NO-detoxifying activities 291–6 transcription regulation 287–91 hydrogenases, Tat protein translocation pathway 203–7 imidazole antifungals 158, 159, 160 iron-binding protein 37–8 iron deficiency, Synechococcus 38 iron-sulphur clusters, Tat protein translocation pathway 202–3 latency, non-culturable cells 89–90 ligand binding, truncated globins 271–3 light-harvesting apparatus, Synechococcus 8–11 longevity, bacterial 69–70 see also cell death membrane protein biosynthesis, Tat protein translocation pathway 236–9 metabolic activity, lowering, TNC 68 metabolically injured non-culturable cells 97–8 Methylococcus capsulatus, CYP/ferredoxin fusion protein 159–61 MGD see molybdopterin guanine dinucleotide micro-nutrient acquisition iron-binding protein 37–8 macro-nutrients 36 Prochlorococcus 36–8 Synechococcus 36–8 microbial globins 255–310 classification 258–60 distribution 258–60 flavohaemoglobins see flavohaemoglobins single-domain globins 258–68, 277 truncated globins see truncated globins Vgb 258–68 molybdopterin guanine dinucleotide (MGD), Tat protein translocation pathway 198–201
325 molybdopterin (MPT) cofactors, Tat protein translocation pathway 197–201 motility, Synechococcus 39 MPT see molybdopterin mRNA, starvation survival 71–2 MTB CYP51 154–6 Mtt pathway see Tat protein translocation pathway mycobacteria CYPs 150–8 CYPs, anti-mycobacterial activity 158, 159 myoglobin 257–8 NADPH-CYP reductase, phylogram 167 niche adaptation, Synechococcus 1–64 niche partitioning, Prochlorococcus 2 nitrate, nitrogen acquisition 20–31 nitrite, nitrogen acquisition 20–31 nitrogen acquisition Prochlorococcus 27–8, 30 Synechococcus 18–31 nitrous oxide reductase, Tat protein translocation pathway 209–10 NO-detoxifying activities, flavohaemoglobins 291–6 Nocardia, CYPs 150 non-culturable cells acid crash 88 anabolism/catabolism 86–9 cultivation regime 98–9 disease states 89–92 division initiation 98 environments 99 environments, fluctuating 66–7 environments, natural 76–9 laboratory microcosms 79–80 latency 89–90 metabolically injured 97–8 oligotrophs 77 oxygen 83 resuscitation 96–103 stability 84–5 stationary phase culture 80–9 TNC 73–92 transition 84 tuberculosis 89, 91 in vivo 89–92 see also transient non-culturability
326 nutrient acquisition micro-nutrient acquisition 36–8 nitrogen acquisition 18–31 phosphorus acquisition 31–5 Synechococcus 18–38 oil-protein conversion, CYPs 164 oligomeric protein biogenesis, Tat protein translocation pathway 199, 212–13 oligotrophs, non-culturable cells 77 pathogenicity, Tat protein translocation pathway 218–19 PCB see phycocyanobilin PE see phycoerythrin PEB see phycoerythrobilin Phanerochaete chrysosporium 165–9 phospholipid bilayer, Tat protein translocation pathway 233 phosphorus acquisition Prochlorococcus 31–5 sphX gene 35 Synechococcus 31–5 temporal factors 34 photosynthetic physiology, carbon metabolism 11–14 phycobiliproteins characteristics, clade-specific physiological 20–7 nitrogen acquisition 19 phycocyanobilin (PCB), Synechococcus 11 phycoerythrin (PE) characteristics, clade-specific physiological 20–7 fluorescence 19 Synechococcus 9–10 phycoerythrobilin (PEB) cell cycle 41–3 Synechococcus 9–11 phycourobilin (PUB) cell cycle 41–3 Synechococcus 9–11 phylogenetic tree, CYPs 138 phylogeny, Synechococcus 4–8 phylograms CYPs 136–7, 152 NADPH-CYP reductase 167 physiological diversity, Synechococcus 1–64
SUBJECT INDEX populations distribution, Synechococcus 8 heterogeneity, TNC 95–6 and social behaviour 103–6 ppGpp 71 Prochlorococcus division cycle 41–3 growth irradiance 12–14 micro-nutrient acquisition 36–8 niche partitioning 2 nitrogen acquisition 27–8, 30 phosphorus acquisition 31–5 proofreading properties, Tat protein translocation pathway 213–15 proteins CYP/ferredoxin fusion protein 159–61 iron-binding 37–8 membrane protein biosynthesis, Tat protein translocation pathway 236–9 oil conversion, CYPs 164 TatA/B protein family 224–30 TatC protein family 230–2 without cofactors, Tat protein translocation pathway 215–19 protists, CYPs 162–3 PUB see phycourobilin response regulator, TNC 67 resuscitation non-culturable cells 96–103 TNC 76 resuscitation-promoting factor (Rpf) 101–3, 106 ribulose 1,5-bisphosphate carboxylase/ oxygenase (RubisCO), carbon fixation 14–17 Rpf see resuscitation-promoting factor RubisCO see ribulose 1,5-bisphosphate carboxylase/oxygenase Sec pathway, cf. Tat protein translocation pathway 189–90, 192–5 sensor histidine kinase, TNC 67 single-domain globins 258–68 cf. flavohaemoglobins 277 smcR (luxR) gene, TNC 95 social behaviour, TNC 103–6 sphX gene, phosphorus acquisition 35
SUBJECT INDEX starvation survival cryptic growth 71–2 experimental 70–3 mRNA 71–2 physiological features 70–1 stress avoidance 69–73 stringent response 71 stationary phase culture, non-culturable cells 80–9 Streptomycetes, CYPs 143–50 stress avoidance starvation survival 69–73 TNC 69–92 stringent response, starvation survival 71 Synechococcus carbon fixation 2 carbon metabolism 11–17 cell cycle 39–43 characteristics 4–6 characteristics, clade-specific physiological 20–7 chemotaxis 39 consensus tree 7 diversity, genetic 6 diversity, physiological 1–64 division cycle 41–3 grazing 44–6 growth irradiance 12–14 iron deficiency 38 light-harvesting apparatus 8–11 marine clusters characteristics 4–6 micro-nutrient acquisition 36–8 motility 39 niche adaptation 1–64 nutrient acquisition 18–38 PCB 11 PE 9–10 PEB 9–11 phosphorus acquisition 31–5 phylogeny 4–8 populations distribution 8 PUB 9–11 viruses 44–6 Tat protein translocation pathway 187–254 amidase puzzle 217–18 cell wall biosynthesis 217–18 cobalamin cofactors 211–12 components 219–22
327 copper cofactors 207–10 GFOR 201–2 hydrogenases 203–7 iron-sulphur clusters 202–3 mechanism 232–6 membrane protein biosynthesis 236–9 MGD 198–201 mis-targeting mechanisms 192–5 MPT cofactors 197–201 nitrous oxide reductase 209–10 oligomeric protein biogenesis 199, 212–13 organisation 219–22 pathogenicity 218–19 phospholipid bilayer 233 proofreading properties 213–15 proteins without cofactors 215–19 routing 195–7 cf. Sec pathway 189–90, 192–5 substrate biogenesis 192–5 substrate diversity 197–215 substrate transport preparation 197–215 TatA/B protein family 224–30 TatC protein family 230–2 transport cycle 222–4 TTQ cofactor 210–11 virulence attenuation 219 temporal factors, phosphorus acquisition 34 TNC see transient non-culturability trace metals see micro-nutrient acquisition transcription regulation, Hmp 287–91 transient non-culturability (TNC) 65–129 alkylresorcinols 93 biofilms 104 cannibalism 104–5 cell death 68–9 chemical inducers 92–4 chemotaxis 67 cooperative behaviour 105–6 environments 99 environments, fluctuating 66–7 environments, natural 76–9 genetic control 94–6 metabolic activity, lowering 68
328 transient non-culturability (Continued ) non-culturable cells 73–92 population heterogeneity 95–6 response regulator 67 resuscitation 76, 96–103 sensor histidine kinase 67 smcR (luxR) gene 95 social behaviour 103–6 stress avoidance 69–92 transition 84 see also non-culturable cells triazole antifungals 158, 159, 160 truncated globins 268–75 conserved residues 270 function 273–5 haem coordination 270–1 ligand binding 271–3 two-over-two a-helical fold 269–70
SUBJECT INDEX Tryptophyl Tryptophanquinone (TTQ) cofactor, Tat protein translocation pathway 210–11 tuberculosis, non-culturable cells 89, 91 twin arginine translocation (Tat) see Tat protein translocation pathway Vgb see Vitreoscilla haemoglobin virulence attenuation, Tat protein translocation pathway 219 viruses, Synechococcus 44–6 Vitreoscilla haemoglobin (Vgb) 258–68 biochemical characterisation 264–5 biotechnological implications 267–8 crystal structures 265–6 function 264–5 gene expression regulation 266–7 heterologous expression 267–8 reductase 266