Advances in Microbial Physiology Volume 47

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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Contents

CONTRIBUTORS

TO

VOLUME 47

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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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3. 4. 5. 6.

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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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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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4. Flavohaemoglobins. . . . 5. Evolution of Globins . . 6. Summary. . . . . . . . . . . Acknowledgements . . . . References . . . . . . . . . .

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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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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

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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

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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 <70% are not shown. The scale bar represents 0.1 substitutions per position.

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oligotrophic waters has also been demonstrated using 16S rDNA cladespecific oligonucleotides, which showed this clade to dominate an oligotrophic water column in the Gulf of Aqaba, Red Sea (Fuller et al., 2003). Such oligonucleotides will be useful in defining the in situ distributions of other members of the genus. Flow cytometry data have also shown general trends in the distribution of Synechococcus populations (see Partensky et al., 1999a for review). The different orange fluorescences of these populations is a reflection of the differences in the two chromophores, phycourobilin (PUB) [maxffi 495–500 nm] and phycoerythrobilin (PEB) [maxffi 540–570 nm] (Glazer, 1985) which attach to the light-harvesting pigment phycoerythrin (PE) (see Section 2). Thus, there is a general trend of PEB-rich populations dominating in mesotrophic or coastal waters and PUB-rich populations detectable only in oligotrophic waters, with PUB:PEB ratios also increasing with depth down a water column (Lantoine and Neveux, 1997). Since high and low PUB containing isolates can cluster phylogenetically (Toledo et al., 1999) and, more importantly, since some strains are capable of chromatic adaptation (Palenik, 2001), correlation of this flow cytometric data with the distribution of phylogenetically coherent clades is not straightforward. Nonetheless, the fact that flow cytometry can retrieve light scattering and fluorescence information on single cells whilst gathering data on many cells from many different samples, has contributed enormously to ecological studies of these organisms (Olson et al., 1988, 1990; Wood et al., 1998, 1999; Tarran et al., 1999; Collier, 2000; Shalapyonok et al., 2001).

2. LIGHT-HARVESTING APPARATUS The light environment is clearly a key parameter dictating the distribution and physiology of all photosynthetic organisms. Changes in light quality and intensity occur over spatial and temporal scales, and in aquatic systems light intensity is attenuated rapidly down the water column. Changes in light quality occur via absorption by water, dissolved yellow pigments, phytoplankton, and inorganic particulate matter, and this differs between coastal and open-ocean systems (Joint, 1990). Hence in coastal waters, blue and red light are significantly attenuated and green light shows the greatest transmission. Conversely short wavelength blue light tends to penetrate deepest in oceanic waters. These narrow windows of light quality obviously overlie the specific ecological niches occupied by coastal and open-ocean isolates, but have also dictated complementarity between these incident light

DIVERSITY AND ADAPTATION IN SYNECHOCOCCUS

9

wavelengths and the photosynthetic antenna pigments which absorb them. Thus, in the marine Synechococcus genus, and all other cyanobacteria, except Prochlorococcus, the light-harvesting accessory pigments that capture these wavelengths are contained within the phycobilisome, a large multi-protein complex comprising an allophycocyanin core (max 650 nm) and rods with phycocyanin (max 620 nm) and phycoerythrin (PE) (max 565 nm) as the major components (see Grossman et al., 1993; Sidler, 1994). Marine Synechococcus strains are unusual among cyanobacteria, however, in that two different PEs are present in the phycobilisomes. In comparison, less than half of the known freshwater cyanobacterial strains produce PE and these only produce one type. The two PE types of marine strains, which have been designated PE(II) and PE(I) respectively (Ong and Glazer 1988, 1991), are encoded by separate genes (Wilbanks et al., 1991; de Lorimier et al., 1992; Wilbanks and Glazer, 1993a,b; Newman et al., 1994) and encode proteins present in the phycobilisome at a relative molar ratio of approx. 3:1. PE(I) is typical of all known C-, B- and R-phycoerythrins containing five bilin groups, two attached to the a-subunit and three attached to the b-subunit. The main phycoerythrin of marine Synechococcus strains though is the PE(II) form, containing six bilin molecules, three each on the a and b subunits, and more PUB than the corresponding PE(I) form. However, the actual proportion of PEB and PUB within PE(II) differs between strains. Thus, Synechococcus sp. WH 8020 contains PE(II) with four PEBs and two PUBs whilst strain WH 8103 contains two PEBs and four PUBs. These differences are a function of the bilin content of the a subunit of PE(II) – strain WH 8020 contains a-75 (PUB), a-83 (PEB) and a140 (PEB), numbers referring to the sequence position of the cysteine residue(s) to which each bilin is attached, whilst that of strain WH 8103 contains PUB at all three of these sites (de Lorimier et al., 1992). It is the narrow absorbance band of PUB, centred around 490 nm, which complements excellently the light wavelengths of maximum transmittance through seawater, specifically the prevalent blue-green wavelengths (see Glazer, 1999). Thus, at this wavelength a PUB-rich phycoerythrin, typical of marine Synechococcus strains, would absorb around 4.5 times as much light as would a C-phycoerythrin typical of freshwater cyanobacteria (Ong and Glazer, 1988). The genetically determined differences in the forms of PE synthesised underlie variation in the fluorescence yield and fluorescence signature among different marine Synechococcus strains (Alberte et al., 1984; Wood et al., 1985, 1999; Ong and Glazer, 1988, 1991) and allow a fine-scale tuning of phycobilisome content to the natural light environment (Wood, 1985; Wood et al., 1998), and hence to the specific environmental niche occupied.

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Thus, in natural environments, in vivo PE emission studies from the Atlantic Ocean and Black Sea (Shalapenok and Shalapenok, 1997; Wood et al., 1998) and single-cell fluorescence excitation spectroscopy and flow cytometry (Campbell and Iturriaga, 1988; Olson et al., 1988, 1990) support a general predominance of PUB-lacking PEs in turbid waters (classified optically as case 2 waters), whilst PUB-containing PEs are more dominant in highly transparent waters (case 1 waters). More recently, Wood and colleagues (1999) used fluorescence excitation spectra to specifically distinguish between different PUB-containing PEs in situ in the Arabian Sea. Fluorescence excitation signatures varied in the relative excitation of PE emission by wavelengths absorbed by PUB ( 495 nm, ExPUB) and by wavelengths absorbed by PEB ( 550 nm, ExPEB). Differences in the ExPUB:ExPEB ratio showed a clear association of low PUB PEs and high PUB PEs with different water masses, the former occupying cooler water (24–27 C) of lower salinity (35.7–36.25 practical salinity units [psu]) but higher nitrate levels ( 10 mM), and the latter, warmer (27–29 C) saltier (36.16–36.4 psu) oligotrophic waters. Such categorisation of water masses into those dominated by PUB-lacking PEs or dominated by PUB-containing PEs has now been performed on a much larger scale using aircraftoperated lidar, which effectively allows global mapping of oceanic phycoerythrin types (Hoge et al., 1998). Relating this environmental pigment data back to the pigment content of the phylogenetically distinct clades described in Figure 1 is complicated. Although one clade (clade VI) comprises members containing no PUB (Rocap et al., 2002), and might be expected to dominate in coastal waters, other clades contain strains of variable PUB:PEB ratio. More importantly, we now know that some isolates are capable of increasing their PUB:PEB ratio when grown under blue light compared to growth under white light conditions (Palenik, 2001), a process traditionally called complementary chromatic adaptation (Grossman et al., 1994) – though usually comparing growth under green versus red light. This trait presents phylogenetic coherence in those non-motile strains studied so far, but members of clade III, which comprise only motile strains, contain both adapters and non-adapters. Only a complete survey of this capacity amongst representatives of all MC-A clades will allow the extent of this trait across the Synechococcus genus to be determined. Such data would then be extremely useful for ultimate comparison to geographic genotype distribution data based on the use of clade-specific 16S rDNA oligonucleotides since fluorescence-based measurements cannot distinguish between chromatic adaptation of a single strain or the presence of distinct strains with different PUB:PEB ratios. This is relevant to our understanding of Synechococcus

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population structure, both horizontally (coastal versus open-ocean systems, as we have already seen) and vertically (down a water column). The latter is indicated by fluorescence measurements of PUB:PEB ratios in the Arabian Sea and tropical north-eastern Atlantic which range from 0.53 in surface waters to 1.33 at 60 m (Lantoine and Neveux 1997; Wood et al., 1999), ratios similar to those found in strains capable of chromatic adaptation. Although most work on the acclimation of the light-harvesting accessory pigments of marine Synechococcus to the endogenous light environment has focused on the phycoerythrin component, it must be remembered that the phycocyanins of these organisms have the maximum content of green lightabsorbing bilin that is still compatible with retention of energy transfer function (Ong and Glazer, 1987). This is due to the presence of a novel type of phycocyanin, designated R-phycocyanin II. In marine Synechococcus strains WH 8103 (clade III), WH 8020 (clade I) and WH 7803 (clade V), this protein has absorbance maxima at 533, 554 and 615 nm and carries PEB at a-84 and b-155 and phycocyanobilin (PCB) at b-84. In freshwater strains, which possess C-phycocyanin, PCB is present in all three of these positions. Whether the presence of a phycocyanin containing two PEB and one PCB per ab monomer is typical of all marine Synechococcus MC-A, and indeed MC-B, isolates awaits further investigation. In fact, it is already known that the phycocyanin of Synechococcus WH 7805 (clade VI) possesses an unusual absorbance spectrum with a maximum at 555 nm and a shoulder at 590 nm, suggesting further subtle differences in phycocyanin chromophore content are possible. Indeed, the marine Synechocystis sp. WH 8501 phycocyanin contains a PUB chromophore on its a subunit, the only cyanobacterium known to possess this combination (Swanson et al., 1991). Certainly, thorough characterisation of the phycocyanin of MC-A and MC-B isolates and its significance to the ecological niche of MC-B strains, which lack phycoerythrin, will be illuminating!

3. C METABOLISM 3.1. Photosynthetic Physiology Unlike in the Prochlorococcus genus, where photosynthetic physiology in strains of specific clades is clearly a function of an evolutionary move towards occupation of surface (high light) or deep (low light) water column niches (Moore et al., 1995, 1998; Hess et al., 2001), a simple division of the marine Synechococcus genus into high light- and low light-adapted ecotypes

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is not likely, though this would not preclude the presence of one or more surface- or deep-adapted clades (see below). Indeed, the relative phylogenetic complexity of the Synechococcus genus compared to Prochlorococcus (Fig. 1) and the subtle variations of phycobiliprotein bilin composition that are possible at least in chromatically adapting strains, makes the picture even more complex. Certainly, the early prevailing hypothesis that marine Synechococcus strains required low light regimes for growth (Glover, 1985; Fogg, 1986; Barlow and Alberte, 1985, 1987) is no longer consistent with physiological information from pre-acclimated cultures (Kana and Glibert, 1987a,b; Moore et al., 1995) or ecological distribution data (see Partensky et al., 1999a), the latter pointing towards a general distribution restricted to the upper well-lit layer where fast growth rates have been reported (see Furnas and Crosbie, 1999). Thus, Synechococcus strain WH 7803 (clade V) is capable of growth over a wide range of irradiance (30–2000 mmol photons m2 s1) in nutrient-replete, continuous light, pre-acclimated batch cultures. Growth rate was observed to become light saturated around 200 mmol photons m2 s1, but remained maximal at irradiances ten-fold higher (Kana and Glibert, 1987a,b). This acclimation process involved changes in phycobiliprotein composition with high light acclimated cells appearing yellow due to a very reduced PE content. Interestingly, photo-inhibition of photosynthesis was observed only in light-limited cells exposed to light intensities ten-fold higher than the growth irradiance. These authors point out though that, since the time required for photo-acclimation was longer than the organism’s generation time, success of the genotype within a given natural light regime would be dependent on the stability of the water column. For another strain, Synechococcus WH 8103 (clade III), growth was again observed over a range of white light irradiance (10–450 mmol photons m2 s1) though in this case under a 14 h light:10 h dark regime (Moore et al., 1995). Interestingly, the light intensity at which growth reached a maximum (142  21 mmol photons m2 s1) is similar to that reported in the same study for Prochlorococcus sp. MED4, a high light-adapted strain, and much higher than that found for Prochlorococcus strain SS120 (37  8 mmol photons m2 s1), a low light-adapted strain (see Fig. 2). Moreover, photoinhibition of growth occurred only at the highest growth irradiance tested (450 mmol photons m2 s1) in both WH 8103 and MED4, considerably higher than that observed for the low light-adapted Prochlorococcus strain (37 mmol photons m2 s1). Thus, WH 8103 might be considered a high light-adapted strain. Recent molecular ecological studies have clearly shown that although several genetically defined Synechococcus clades can co-exist at the same site (Toledo and Palenik, 1997), one or more of these clades may contain surface-adapted organisms (Ferris and Palenik, 1998 and see

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Figure 2 Growth rate (day1) as a function of white light irradiance (mmol quanta m2 s1) for Prochlorococcus MED4 (^), Prochlorococcus SS120 (g) and Synechococcus WH 8103 (m). Data provided by Dr. L. Moore and Prof. S.W. Chisholm based on that presented in Moore et al. (1995).

Section 1.1) since i) rpoC1 sequences representing two distinct clades were detected only in clone libraries from surface waters and not deep waters from the same stratified water column, whilst ii) an isolate clustering with one of these clades shows good growth in high white light (100 mmol photons m2 s1) at 21 C, but not in low (15 mmol photons m2 s1) blue light at 19 C in comparison with other isolates from the same Californian current site (Ferris and Palenik, 1998). Certainly, finding any obvious division in photosynthetic physiology that is underpinned genetically, i.e. adaptive, will require cross-survey comparison of various photosynthetic parameters across the genus in several strains, and reference to genomic information, which ideally in the future will encompass many more genotypes than the already completed Synechococcus WH 8102. Thus far, work performed in this area has largely focused on comparison of photosynthetic or optical properties between the Prochlorococcus and Synechococcus genera (Morel et al., 1993; Moore et al., 1995; Shimada et al., 1996), rather than focusing solely on the latter genus. Further elucidation of photo-protective mechanisms across the genus is also required. The only reports so far addressing the role of carotenoids in this process show that cellular zeaxanthin concentrations do not vary appreciably in nutrient

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replete batch culture across a range of light intensity in Synechococcus strains WH 7803 and WH 8103 (Kana et al., 1988; Moore et al., 1995) with the proposal that zeaxanthin is located outside the thylakoid membrane i.e., either in the cytoplasmic or outer cell wall membrane consistent with a photo-protective function. In contrast, b-carotene in Synechococcus WH 7803 and WH 8103 appears to be associated with the photosynthetic apparatus, probably playing only a very minor role in light harvesting (Kana et al., 1988).

3.2. C Fixation: RubisCO and Carbon-Concentrating Mechanisms Ribulose 1,5-bisphosphate carboxylase/oxygenase (RubisCO), the ratelimiting catalyst for carbon fixation, is relatively inefficient with respect to the carboxylation reaction it performs and, as a result, most cyanobacteria have evolved an efficient carbon-concentrating mechanism (CCM) to improve this process (see Kaplan and Reinhold, 1999 for review). However, it is now becoming clear that marine Synechococcus, and indeed Prochlorococcus, are unusual with respect both to the type of RubisCO possessed, and the inorganic carbon-concentrating systems present, compared with other cyanobacteria. Thus, both these marine cyanobacterial genera possess a Form 1A type of RubisCO compared to the Form 1B typical of freshwater and halotolerant species, the latter exemplified by Synechococcus PCC 7002, see Table 2 (Shimada et al., 1995; Watson and Tabita, 1996). This Form 1A enzyme is most similar to those from b/g-purple bacteria such as Chromatium vinosum, Hydrogenovibrio marinus and Thiobacillus spp.; hence, it has been suggested that the genes encoding the enzyme have been acquired through a horizontal gene transfer event from a purple bacterium ancestor (Watson and Tabita, 1997; Hess et al., 2001). However, perhaps just as likely is an appearance in cyanobacteria preceding that in proteobacteria (Badger et al., 2002). Certainly, in the two marine Synechococcus strains, WH 7803 and WH 8102, where sequence information exists, the RubisCO genes rbcL and rbcS lie downstream of csoS1 encoding a component of the carboxysome. Carboxysomes are protein bodies surrounded by a protein shell and containing RubisCO. The presence of the csoS1 gene (and for strain WH 8102 other carboxysome components – see Table 2) has led to the proposal that marine Synechococcus possess a carboxysome significantly different from that of other cyanobacteria. Thus, Form 1A RubisCO is coincident with possession of an a-carboxysome whilst Form 1B containing cyanobacteria possess b-carboxysomes (Badger et al., 2002). These features

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15

of RubisCO and carboxysomes in marine Synechococcus are mirrored by other differences, compared to freshwater strains, in components of the C acquisition system that have become apparent through completion of the Synechococcus WH 8102 genome sequence. Thus, no carboxysomal carbonic anhydrase gene is found in this organism, although a b-carbonic anhydrase gene is present in the genome. Further, high affinity CO2 uptake components of the NAD(P)H dehydrogenase complex, which plays a role in transporting CO2 by supporting cyclic electron transport (Ohkawa et al., 2000), are absent, with genes encoding only low affinity CO2 uptake components of the complex being present. Finally, there also appears to be no high affinity bicarbonate transporter (Badger et al., 2002). The latter is also absent in Synechococcus PCC 7002, a halotolerant strain that contains all CO2 uptake-related NAD(P)H dehydrogenase components for both high and low affinity CO2 uptake. Whether the presence of the high affinity CO2 uptake components of the NDH-1 dehydrogenase complex gives a competitive advantage to this halotolerant species, compared to the MC-As in the estuarine niche it inhabits (i.e. a habitat subject to freshwater input), will require further analysis. Certainly for the MC-A cluster strain, WH 8102, and possibly for all MC-A isolates, the absence of high affinity components of the CCM probably correlates with the high inorganic carbon load of the oceanic environment. These genetic data may explain early physiological work suggesting the absence of a CCM in marine Synechococcus strains (Karagouni et al., 1990) and later work describing the presence of such a system (Hassidim et al., 1997). These latter workers document in Synechococcus WH 7803 a similar photosynthetic affinity for extracellular dissolved inorganic carbon (DIC) compared to Synechococcus sp. PCC 7942, a freshwater strain, but they also show that the marine strain had a substantially lower capacity to accumulate DIC internally than its freshwater counterpart. Such data, whilst indicating the absence of an active CO2 uptake system like that in freshwater strains, would be consistent with the presence of an as yet uncharacterised lower capacity system. It may also be consistent with the striking observation of net CO2 efflux during net photosynthesis which occurs in marine Synechococcus WH 7803 (Tchernov et al., 1997), a study which also provides evidence for HCO 3 utilisation from the external medium. The significance and ubiquity of this net CO2 evolution remains to be established, but it has been proposed that it may confer some protection against damage to the photosynthetic reaction centre at high light intensities by dissipating excess light energy by rapid consumption of ATP (Tchernov et al., 1997).

16

Table 2

CCM related genes in cyanobacterial genomes

Gene

Reference sequence

a-Cyanobacteria

b-Cyanobacteria

Synechocystis Anabaena Nostoc Synechococcus Synechococcus Synechococcus Prochlorococcus Prochlorococcus PCC 6803 PCC 7120 punctiforme* PCC 7002 PCC 7942 WH 8102* MED4* MIT9313* b-Carboxysomes ccmK 4 ccmL 1 ccmO 1 ccmM 1 rbcL activase n.f. ccmN 1 a-Carboxysomes csoS1 n.f. peptide A n.f. peptide B n.f. csoS2 n.f. csoS3 n.f. RuBisCO Forrn 1B Carboxysome CA 1 b-cCA NDH-1 genes ndhD1/D2 2 ndhD5/6 2 ndhF1 1 CO2 uptake ndhD3 1 ndhD4 1 ndhF3 1 ndhF4 1 chpY 1 chpX 1 Bicarbonate transporters cmpA 1

4 1 1 1 1 1

4 1 n.f. 1 1 1

1 1 n.f. 1 n.f. 1

1 1 1 1 n.f. 1

n.f. n.f. n.f. n.f. n.f. n.f.

n.f. n.f. n.f. n.f. n.f. n.f.

n.f. n.f. n.f. n.f. n.f. n.f.

SLL1028 SLL1030 SLR0436 SLL1031 PID:g296414 SLL1032

n.f. n.f n.f n.f n.f Form 1B ?

n.f. n.f. n.f. n.f. n.f. Form 1B 1 b-cCA

n.f. n.f. n.f. n.f. n.f. Form 1B 1 b-cCA

n.f. n.f. n.f. n.f. n.f. Form 1B 1 b-cCA

2 – 1 1 1 Form 1A n.f.

1 1 1 1 1 Form 1A n.f.

1 1 1 1 1 Form 1A n.f.

AAC24975.1 AAD30512.1 AAD30513.1 AAD30510.1 AAD30511.1 SLR0009 SLR1347

3 1 1

3 n.f. 1

2 2 1

– – 1

1 n.f. 1

2 n.f. 1

2 n.f. 2

SLR0331 SLR2007 SLR0844

1 1 1 1 1 1

1 1? 1 1 1 1

1 1 1 1 1 1

1 1 1 1 1 1

n.f. 1 n.f. 1 n.f. 1

n.f. n.f. n.f. n.f. n.f. n.f.

n.f. n.f. n.f. n.f. n.f. n.f.

SLL1733 SLL0027 SLL1732 SLL0026 SLL1734 SLR1302

1

1

n.f.

1

n.f.

n.f.

n.f.

SLR0040

cmpB cmpC cmpD slr1515 slr1512 glycolate metabolism PGP glcD glcE glcF Glycolate oxidase Carbonic anhydrases a-CAs b-CAs g-Cas

1 1 1 1 1

1 1 1 1 1

1 1 1 1 1?

n.f. n.f. n.f. 1 1

1 1 1 1 1?

n.f. n.f. n.f. 1 n.f.

n.f. n.f. n.f. n.f. n.f.

n.f. n.f. n.f. 1 n.f.

SLR0041 SLR0042 SLR0043 SLR1515 SLR1512

1 1 1 1 n.f.

1 1 1 1 1

1 1 1 1 1

1 1 1 1 n.f.

1 – – – –

n.f. 1 1 1 n.f.

n.f. n.f. n.f. n.f. n.f.

n.f. 1 1 1 n.f.

SLL1349 SLL0404 SLL1189 SLL1831 AAD25332.1

– ccA, ecaB

ecaA 1 b-CA

ecaA cCA

ecaA cCA

n.f. 1 b-CA

n.f. n.f.

n.f. n.f.

– –

ccmM, ferripyochelin

CcmM

– 5 b-CAs – cCA, ecaB ccmM

ccmM

ccmM

ferripyochelin

n.f.

n.f.

–

The following resources were used to make gene comparisons: for general information, Genbank protein and nucleotide sequence databases (http://www.ncbi.nlm.nih. gov/); for specific information on finished and unfinished cyanobacterial genomes, PEDANT – (http://pedant.gsf.de/index.html), DOE Microbial Genomics (http:// www.jgi.doe.gov/tempweb/JGI_microbial/html/index.html); for Synechocystis and Anabaena, Cyanobase (http://www.kazusa.or.jp/cyano/cyano.html). References are not given to each gene, however, the presence or absence of each gene was assessed by its protein homology to reference sequences for each gene. The protein references used are given. SLL and SLR numbers are from Cyanobase references for Synechocystis PCC 6803. –, genes not currently found, but genome not completely sequenced; n.f., genes not found in mostly completed genomes; *, species for which mostly completed genomes are currently available; ?, possible presence. Reproduced from Functional Plant Biology, Volume 29 (Badger, M.R., Hanson, D., and Price, G.D. 2002) with permission of CSIRO Publishing.

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4. NUTRIENT ACQUISITION Members of the marine Synechococcus genus are around one micron in size. This confers on them a high surface area to volume ratio which generally allows more efficient nutrient uptake particularly since diffusion gradients around a cell generally steepen as a cell’s linear dimension decreases. Even so, at the low nutrient concentrations often present in open-ocean oligotrophic areas and even in temperate regions in summer, there is a possibility of diffusion limitation of nutrient uptake (see Mierle, 1985; Fogg, 1987). In addition to the plethora of high affinity nutrient acquisition systems a marine Synechococcus may possess, other more subtle adaptations for nutrient acquisition in an oligotrophic environment may exist. For example, 42 ABC nutrient transport systems, based on the number of genes encoding the ATPase component, can be identified in the Synechococcus WH 8102 genome (see Scanlan and West, 2002). These might be in addition to more ‘general’ physiological adaptations for growth at low nutrient concentrations such as low elemental nutrient quotas or efficient mechanisms to prevent nutrient leakage. Such adaptive properties again must be put into the context of the specific niche occupied by members of an individual clade. In ecology it has been proposed that a niche implies a strategy for survival and growth (Koch, 2001), and this means that genotypes must acquire or even delete specific genes to suit their specific environmental requirements. Genetic adaptations of this type might allow utilisation of specific substrates (e.g. urea, cAMP see Sections 4.1 and 4.2 below) or conversely involve a commitment to utilise only a subset of the available nutrient sources. An excellent example of the latter has recently been strikingly demonstrated in the Prochlorococcus genus where all members of the HL and LL clades so far characterised show an inability to utilise nitrate as a N source, with a corresponding absence of the nitrate reductase gene from the genome (Moore et al., 2002). Certainly, given the obvious vertical and horizontal gradients in nutrient concentration that exist in natural systems, be it as a result of stratification in oligotrophic open-ocean systems or riverine input in coastal systems, adaptation to the specific nutrient environment will underlie much of the niche-adapted physiology of specific genotypes.

4.1. N Acquisition As for other phytoplankton, nitrogen (N) is an essential macroelement for Synechococcus and the availability of various nitrogenous nutrients, as

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dictated by the functioning of the marine N cycle (see Zehr and Ward, 2002), directly affects the growth and carbon fixation capacity of the genus. With respect to N availability, we can also include in this category the ability to store N and hence the accessibility of intracellular N reserves. As referred to in Section 2, phycoerythrin (PE) fluorescence appears to be related to irradiance – with fluorescence intensity increasing with depth. However, this precise function of PE fluorescence versus light intensity may be complicated by the fact that there is now good evidence that fluorescence is also a function of the nutrient history of the cell. Wyman and colleagues (Wyman et al., 1985; Heathcote et al., 1992) demonstrated that, under certain growth conditions, a proportion of the phycobiliprotein pool is not photosynthetically competent and functions primarily for N storage at least in strain WH 7803. There are some conflicting data from the same strain regarding whether phycobiliproteins are regulated independently from photosynthesis as a N storage pool (Yeh et al., 1986; Kana and Glibert, 1987a,b), which may reflect differences in growth rates of cultures and methodology between the studies. Nevertheless there are now several reports that show breakdown of phycobiliproteins does occur during periods of N depletion in marine Synechococcus isolates (Glibert et al., 1986; Lewis et al., 1986; Moyal et al., submitted), though the extent of this breakdown may differ between strains (Kana et al., 1992). This phycobiliprotein breakdown needs to be put into the context of a general nutrient stress response to limit the amount of energy reaching the photosystems during occasions when energy utilisation is retarded due to nutrient deficiency that may not be just N specific (Fig. 3). Regulation of phycobiliprotein degradation in marine Synechococcus strains awaits further investigation and lags far behind what is known for freshwater strains (see Collier and Grossman, 1994; Sauer et al., 1999; Luque et al., 2001; van Waasbergen et al., 2002). In contrast to this potential use of phycobiliproteins as a N store under conditions of N stress, there appears to be a general absence of the N reserve polymer cyanophycin (multi-L-arginyl-poly-[L-aspartic acid]) amongst marine MC-A Synechococcus strains (Newman et al., 1987). However, cyanophycin production has recently been demonstrated in a unicellular PE-containing marine cyanobacterium G2.1, isolated from the Arabian Sea (Wingard et al., 2002), with a phylogenetic affiliation relatively far removed from those Synechococcus strains described in Figure 1, the focus of this review. Marine Synechococcus strains appear to show a relatively broad  utilisation of N sources for growth. Ammonium ðNHþ 4 Þ, nitrate ðNO3 Þ,  nitrite ðNO2 Þ and urea are all capable of acting as the sole N source for growth in one or more strains (Glibert et al., 1986; Glibert and Ray, 1990; Lindell et al., 1998, 1999; Collier et al., 1999; Moyal et al., submitted). Since

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Figure 3 In vivo absorption spectra of three cyanobacterial clones (WH 7803, WH 7805, and WH 5701) grown under nutrient sufficient and nutrient depleted conditions. The difference spectra indicate the in vivo absorbance spectra of the pigments degraded following nutrient depletion, and corresponds to either phycoerythrin (WH 7803, WH 7805) or phycocyanin (WH 5701). Taken from Photosynthetic Picoplankton (Lewis, M.R., Warnock, R.E. and Platt, T. p. 245 [1986]), Canadian Bulletin of Fisheries and Aquatic Science 214, Fisheries and Oceans Canada, and reproduced with the permission of the Minister of Public Works and Government Services Canada, 2002.

strains WH 7803 and WH 8101 can assimilate amino acids (Kramer 1990; Kramer and Morris, 1990; Paerl, 1991), whilst others (e.g. WH 8102, WH 8103, WH 8011, WH 8112 and WH 7803) have been shown to express aminopeptidase activity (Martinez and Azam, 1993), it is likely that the ability to utilise amino acids is also widespread in the genus. However, some differences in N resource utilisation among strains are known. Although most strains are capable of consuming urea, a few exceptions have also been noted (Waterbury et al., 1986; Collier et al., 1999). For one Synechococcus WH 7803 (clade V), a portion of the ureC gene as well as a region of several kbp downstream of the urease gene cluster has been shown to be lacking (Collier et al., 1999). This eliminates urease activity in this strain, though whether this is a feature of other members of MC-A clade V is not known. Strain WH 7803 is, however, capable of ammonium, nitrate and nitrite utilisation (Glibert et al., 1986; Glibert and Ray, 1990; Post et al., submitted, and see Table 3, which includes a summary of physiological properties of strains, clustered with respect to their phylogenetic position in Fig. 1).

Table 3

Clade-specific physiological characteristics of marine Synechococcus strains. Phycobiliproteins

Phylogenetic clade & Strain No.

Isolation location

MC-A Clade I WH 8015

Woods Hole

WH 8016

Woods Hole Sargasso Sea (38 40.7’N, 69 19’W)

WH 9908 CC9311

Woods Hole California Current*

Almo 3

Mediterranean Sea (36 11’N, 1 51’W)

WH 8002

Phycoerythrin (PE)

Chromatic adaptation

Obligately marine (þ) or ()

Present (þ) Absent ()

PUB:PEB ratio

55.3

þ

0.44

55.5

þ

0.4

þ

0.78

þ

þ

1.16

þ

þ

0.4

þ



WH 8020

MC-A Clade II WH 6501

Mol % G þ C#

Utilisation of N sources

Capable of swimming motility (þ) or ()

References

(þ) or ()

þ

þ

nitrate (þ) ammonium (þ) nitrite (þ) nitrate (þ) ammonium (þ) nitrate (þ) ammonium (þ) nitrite (þ) nitrate (þ) ammonium (þ) urea (þ)

Tropical Atlantic (8 44’N, 50 50’W)

62.3

þ

0.43

þ

Gulf of Mexico (19 45’N, 92 25’W)

60.0

þ

0.48

þ



Waterbury et al., 1986 Moore et al., 2002



Waterbury et al., 1986



Waterbury et al., 1986 Moore et al., 2002 Palenik (2001) Rocap et al., 2002 Toledo & Palenik, 1997 Collier et al., 1999 Palenik (2001) Fuller et al., 2003





nitrate (þ) ammonium (þ) nitrite (þ) urea (þ) nitrate (þ) ammonium (þ) nitrite (þ) urea (þ)



Waterbury et al., 1986 Moore et al., 2002



Waterbury et al., 1986 Moore et al., 2002

21

(continued )

22

Table 3 Cont. Phycobiliproteins Phylogenetic clade & Strain No.

Isolation location

Mol % G þ C#

Phycoerythrin (PE)

Present (þ) Absent ()

PUB:PEB ratio

Chromatic adaptation

Obligately marine (þ) or ()

Gulf of Mexico (19 45’N, 92 25’W)

58.6

þ

0.44

WH 8012

Sargasso Sea (34 N, 65 W)

62.4

þ

0.4

WH 8109

Sargasso Sea (39 28.6’N, 70 27.72’W)

þ

0.89

CC9605 RS9902

California Current Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E)

þ

1.9

þ

þ

2.3

þ

RS9904 RS9907 RS9908 RS9910 RS9911

þ þ þ þ þ

2.1 0.5 0.5 0.5 1.0

Capable of swimming motility (þ) or ()

References

(þ) or ()

WH 8005

RS9903

Utilisation of N sources

þ

þ þ þ þ þ

nitrate (þ) ammonium (þ) nitrite (þ) urea (þ) nitrate (þ) ammonium (þ) nitrite (þ) nitrate (þ) ammonium (þ) nitrite (þ) nitrate (þ) ammonium nitrate (þ) ammonium nitrate (þ) ammonium nitrate (þ) ammonium nitrate (þ) ammonium nitrate (þ) ammonium nitrate (þ) ammonium



Waterbury et al., 1986 Moore et al., 2002



Waterbury et al., 1986 Moore et al., 2002



Waterbury et al., 1986 Moore et al., 2002

 

Toledo et al., 1999 Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003

(þ) (þ) (þ) (þ) (þ) (þ) (þ)

RS9912 RS9919 MC-A Clade III WH8102

Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E)

þ

0.5

þ

þ

1.9

þ

þ

nitrate (þ) ammonium (þ) nitrate (þ) ammonium (þ)



Fuller et al., 2003



Fuller et al., 2003

nitrate (þ) ammonium (þ) nitrite (þ) urea (þ) nitrate (þ) ammonium (þ) nitrite (þ) urea (þ)

þ

Waterbury et al., 1986

þ

nitrate (þ) ammonium (þ) nitrite (þ) urea (þ) nitrate (þ) ammonium (þ) urea (þ) nitrate (þ) ammonium (þ) nitrite (þ)

þ

Waterbury et al., 1986 Collier et al., 1999 Moore et al., 2002 Palenik (2001) Waterbury et al., 1986 Collier et al., 1999 Moore et al., 2002 Waterbury et al., 1986 Collier et al., 1999 Palenik (2001) Waterbury et al., 1986 Moore et al., 2002

Tropical Atlantic (22 29.7’N, 65 36’W)

60.4

þ

2.06

WH8103

Sargasso Sea (28 30’N, 67 23.5’W)

58.9

þ

2.40**

WH 8112

Sargasso Sea (36 N, 66 W)

59.8

þ

variable ***

WH 8113

Sargasso Sea (36 N, 66 W)

60.5

þ

variable***

WH 8406

Gulf of Mexico (30 N, 88 W)

þ

0.84

C8015 C129

Red Sea Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E) Mediterranean Sea (39 10’N, 6 10’E) Sargasso Sea (26 18’N, 63 26’W) Mediterranean Sea (34 0’N, 18 0’E)

þ þ

>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

þ

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

<1 <1

Chromatic adaptation

Obligately marine (þ) or ()

nitrate (þ) ammonium (þ) nitrite (þ) urea ()

þ

No PUB


þ

55.8

þ

No PUB


þ

WH 8017

Gulf of Mexico (19 45’N, 92 25’W) Woods Hole

54.5

þ

No PUB




WH 8018

Woods Hole

57.6

þ

No PUB


þ

1.7

þ

1.7

Oli31

Gulf of Aqaba (29 28’N, 34 55’E) Equatorial Pacific (5 60’S, 150 0’W)

References



Waterbury et al., 1986 Moore et al., 2002 Rocap et al., 2002 Rocap et al., 2002

59.7

MC-A Clade VII RS9920

Capable of swimming motility (þ) or ()

(þ) or ()

Sargasso Sea (33 44.8’N, 67 30’W)

WH 8008

Utilisation of N sources

þ

nitrate (þ) ammonium (þ) nitrite (þ) urea (þ) nitrate (þ) ammonium (þ) nitrate (þ) ammonium (þ) nitrate (þ) ammonium (þ)



Waterbury et al., 1986 Moore et al., 2002



Waterbury et al., 1986



Waterbury et al., 1986 Rocap et al., 2002 Waterbury et al., 1986



nitrate (þ) ammonium (þ) þ

Fuller et al., 2003 

Fuller et al., 2003

MIT S9220

Tropical Atlantic (21 2’N, 31 8’W) Equatorial Pacific

MC-A Clade VIII WH 8101

Woods Hole

Eum14

RS9906 RS9909 RS9913 RS9914 RS9917 RS9918 MC-A Clade IX RS9901 RS9916 RS9921 MC-A Clade X RCC309 Minos 11

þ

2.2

þ

Low



NA



(29 28’N,



NA



(29 28’N,



NA



(29 28’N,



NA



(29 28’N,



NA



(29 28’N,



NA



(29 28’N,



NA



Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E) Gulf of Aqaba (29 28’N, 34 55’E)

þ

0.7

þ

þ

0.7

þ

þ

0.7

þ

þ

0.7

þ

0.7

Gulf of Aqaba 34 55’E) Gulf of Aqaba 34 55’E) Gulf of Aqaba 34 55’E) Gulf of Aqaba 34 55’E) Gulf of Aqaba 34 55’E) Gulf of Aqaba 34 55’E)

63.9





Mediterranean Sea (39 10’N, 6 10’E) Mediterranean Sea (34 0’N, 18 0’E)

þ



Moore et al., 2002

nitrate () ammonium (þ) nitrite (þ) urea (þ) nitrate (þ) ammonium (þ) nitrite (þ) urea (þ) nitrate (þ) ammonium (þ) nitrate (þ) ammonium (þ) nitrate () ammonium (þ) nitrate () ammonium (þ) nitrate () ammonium (þ) nitrate (þ) ammonium (þ)

Fuller et al., 2003



Waterbury et al., 1986 Moore et al., 2002



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003



Fuller et al., 2003

þ



Fuller et al., 2003

þ



Fuller et al., 2003

nitrate (þ) ammonium (þ) nitrate (þ) ammonium (þ) nitrate (þ) ammonium (þ)

25

(continued )

26

Table 3 Cont. Phycobiliproteins Phylogenetic clade & Strain No.

MC-B WH 5701

Cyanobium sp. NS01

Isolation location

Long Island Sound

Central North Sea

Mol % G þ C#

65.8

Phycoerythrin (PE)

Chromatic adaptation

Obligately marine (þ) or ()

Utilisation of N sources

Capable of swimming motility (þ) or ()

References

Present (þ) Absent ()

PUB:PEB ratio

(þ) or ()



NA



nitrate (þ) ammonium (þ) nitrite (þ)



Waterbury et al., 1986 Moore et al., 2002



NA





Fuller et al., 2003

MC-A Clade IV is represented by 16S rDNA and ITS sequences obtained directly from environmental DNA; as yet, there are no cultured counterparts. RS ¼ Red Sea isolate WH ¼ Woods Hole RCC ¼ Roscoff culture collection # Mean DNA base composition *California Current station 83.110 of the CalCOFI (California Cooperative Oceanic Fisheries Investigations, cruise 9304) grid pattern, isolated at 95 m depth. **PUB:PEB values of 1.31 (Toledo and Palenik, 1997) and 1.3 (Fuller et al., 2003) have also been reported for this strain. ***These strains have phycoerythrins whose chromophore ratios vary with light intensity ****PUB:PEB values of 0.979 (Toledo and Palenik, 1997) and 0.5 (Fuller et al., 2003) have also been reported for this strain.
These strains contain C-phycoerythrin that lacks phycourobilin. NA, Not applicable. These strains lack phycoerythrin and hence have no PUB or PEB.

DIVERSITY AND ADAPTATION IN SYNECHOCOCCUS

27

Similarly, inactivation of ureC in Synechococcus WH 8102 and WH 7805 prevents growth of these strains on urea, but does not affect the ability of these strains to utilise nitrate or ammonium. Whether urease activity allows a more efficient internal recycling of N, which is beneficial in situ to some strains (Collier et al., 1999), or merely reflects a choice of resource utilisation that is lost in those strains occupying a niche with a rapid flux of the nutrient that makes utilisation impossible, and ultimately forces gene loss, remains to be determined. Recently, a gene cluster urtABCDE, was identified in the freshwater cyanobacterium Anabaena PCC 7120 to encode elements of an ABC-type permease involved in high affinity urea uptake (Valladares et al., 2002). Homologues of these genes are present in the completed genome of marine Synechococcus strain WH 8102, adjacent to the urease structural genes, suggesting the capacity for high affinity urea acquisition in this organism. Confirmation of their presence in other strains, or their absence in those strains that lack urease activity, requires further work or genome sequences. Utilisation of ammonium, nitrate and nitrite appears to be relatively widely distributed among the various Synechococcus clades (Table 3). This is in marked contrast to the Prochlorococcus genus in which utilisation of ammonium and nitrite N sources appears partitioned amongst closely related, but distinct, ecotypes, a feature correlating well to the environmental niche where these resources are available and the known niche of these ecotypes (Moore et al., 2002). As already mentioned in Section 4 above, nitrate utilisation among the Prochlorococcus strains presently in culture appears absent, with this lack of utilisation again correlating with actual gene loss. This compares with the relatively widespread ability to utilise nitrate amongst the marine Synechococcus genus, a feature agreeing both with in situ growth rate data which correlates with nitrate concentration (see Blanchot et al., 1992; Partensky et al., 1999a), and with observations of natural Synechococcus populations which have been shown to respond to periodic nitrate input (Glover et al., 1988; DuRand et al., 2001). The ability to retain N resource uptake capacity across both reduced and oxidised N forms may reflect a high cellular N requirement among marine Synechococcus strains as a result of the N-rich phycobilisome burden, a feature which has also been put forward to explain the more general surface distribution of the genus, since the low light levels at the base of the euphotic zone might not yield enough energy for reduction of oxidised N forms to ammonium. Even so, there is recent evidence that nitrate utilisation is not absolute across the genus with one strain MIT S9220 from the equatorial Pacific unable to utilise this source (Moore et al., 2002). In addition, clade VIII, whose members lack PE and which were originally

28

DAVID J. SCANLAN

designated MC-B, but which phylogenetically appear to be well supported within MC-A (see Fig. 1), contains strains, thus far non-axenic, that can and cannot utilise nitrate as sole N source (Fuller et al., 2003). Thus, although generalisations of N utilisation across the genus can be made, exceptions to this are now appearing. This assumes that the observed differences in utilisation do not merely reflect mutations that have arisen in strains during their time in culture. These differences in N nutrition physiology between strains may reflect adaptive responses of cells to the nutrient form(s) most frequently encountered by the organism in its own specific niche. However, it is possible there may be less obvious reasons why a particular N source is not utilised e.g., susceptibility to viral attack when expressing a particular cell surface-exposed N transporter, as a result of it acting as a phage receptor, or specific ‘consortial’ interaction with another phytoplankton species that can utilise the oxidised N source but then excreting the reduced form. Overlying obvious differences in N resource utilisation amongst different strains, are more subtle differences in N physiology that have also been documented and probably reflect the in situ niche of individual genotypes. Thus, Glibert et al. (1986) and Glibert and Ray (1990) report a differential response to N depletion and N uptake in two marine Synechococcus strains. WH 7803 (clade V) was able to maintain nitrate utilisation capacity even after two days of nitrogen starvation such that, although cell division was immediately arrested upon N depletion, growth and nitrate uptake occurred within 10 h of nitrate being re-supplied. In contrast, strain WH 8018 (clade VI), showed net growth throughout the 48 h N depletion period but had a 24 h lag in growth when nitrate was re-supplied. Such data have been reconciled with respect to ecological partitioning of the two strains, with a potential oceanic location for the former. This strain can readily respond to the intermittent nitrate supply which may occur due to only occasional physically driven turbulent flux across the nitracline or episodic pulses from water column mixing events. A more coastal location of strain WH 8018 would be consistent with its place of isolation and with the more continuous nitrate supply found in coastal waters, a result of both mixing and terrestrial run-off events, allowing growth at the expense of internal cellular reserves. Both strains have been shown to excrete little dissolved organic nitrogen during N starvation conditions, which suggests they can efficiently retain the N taken up during those circumstances (Bronk, 1999). A similar differential, though perhaps more dramatic, response to N deprivation among genotypes has been seen in freshwater Synechococcus isolates, characterised by PC-rich strains undergoing a rapid decrease in the phycobiliprotein:chlorophyll a ratio, but a concomitant increase in cellular carbohydrate and soluble

DIVERSITY AND ADAPTATION IN SYNECHOCOCCUS

29

extracellular polysaccharide. PE-rich strains show neither a change in pigment composition nor any accumulation of polysaccharide (Postius and Bo¨ger, 1998). Whilst further work would be required to define the molecular basis of the differing patterns of growth and N uptake in these two marine strains, for one of these, WH 7803, the mechanics and regulation of N uptake have recently been studied in detail (Lindell et al., 1998; Lindell and Post, 2001; A. F. Post, personal communication). What is immediately obvious from this work are the contrasting mechanisms and physiological properties of N acquisition between marine and freshwater Synechococcus strains. For example, an ABC-type transporter comprising the nrtABCD gene products appears to be the major route for nitrate transport in freshwater species (Omata, 1995), whereas a permease (NapA/NrtP) of the major facilitator super-family performs the same role in marine strains (Sakamoto et al., 1999; A. F. Post, pers. comm.). A nitrate permease of the NapA type has also been found in a filamentous marine nitrogen-fixing Trichodesmium strain (Wang et al., 2000). Interestingly, the transporter encoded by the nrtABCD genes appears also to transport nitrite (Luque et al., 1994), whilst recent mutagenesis of the napA gene in Synechococcus WH 7803 reveals a strain still capable of nitrite uptake, suggesting the presence of a separate nitrite transport system in this strain at least (A. F. Post, pers. comm.). Moreover, nitrite and nitrate appear to act as specific inducers for the synthesis of proteins required for nitrite uptake (Lindell et al., 1998). Another unusual property of N utilisation in WH 7803 is that ammonium appears to directly inhibit nitrate uptake in this strain, a feature different to that observed in all other cyanobacteria where the negative effect of ammonium on nitrate and nitrite uptake appears indirect, being prevented by the addition of the glutamine synthetase inhibitor L-methionine-DL-sulfoximine (MSX) (A. F. Post, pers. comm.). Ammonium, rather than nitrate or nitrite, is preferentially utilised by marine Synechococcus strains as their major N source (Glibert and Ray, 1990; Lindell et al., 1998), whilst during N starvation utilisation of phycobiliproteins as a source of amino acids for growth occurs (see above). This hierarchical utilisation of N substrates suggests a complex N regulatory network exists in these organisms, with the potential for subtle differences in regulation between strains. Key to the regulation of N metabolism in freshwater strains is the DNA binding protein NtcA (see Herrero et al., 2001) which, in the absence of ammonium, can act as a transcriptional activator of those genes required for utilisation of alternative N sources. Recently, ntcA was cloned from the marine Synechococcus strain WH 7803 (Lindell et al., 1998). This has allowed the construction and

30

DAVID J. SCANLAN

characterisation of a ntcA mutant strain (A. F. Post, pers. comm.). This mutant cannot assimilate, or grow on, nitrate or nitrite. In contrast, active ammonium transport is retained in the mutant. The mutant’s inability to assimilate or grow on nitrate coincides with an absence of napA expression, and indicates that NtcA regulates nitrate metabolism at the level of napA transcription (Post et al., submitted). This is supported by the finding of putative NtcA binding sequences, GTA N8 TAC..N22..T N4 T and GTA N8 AAC..N23..T N3 AT, at 225 and 132 bases respectively, upstream of the napA start codon (A. F. Post, pers. comm.), similar to reported consensus NtcA-binding sites (Luque et al., 1994). Conversely, the ability of the mutant to retain active ammonium transport suggests an absence of NtcA regulation in this process. Interestingly, in Prochlorococcus sp. PCC9511, a surface-adapted strain (Rippka et al., 2000), amt1, encoding a high affinity ammonium transporter, is expressed at high levels both in the presence of ammonium as well as during all but the severe stages of N-deprivation. This constitutive amt1 expression, and the absence of a typical NtcA binding site upstream of amt1, has led to the proposal that amt1 is not regulated by NtcA in this Prochlorococcus strain (Lindell et al., 2002). Whether constitutive amt1 expression occurs in Synechococcus WH 7803 and other marine Synechococcus strains awaits further investigation, but there is a recent report of constitutive expression of nirA, encoding nitrite reductase, in the marine Synechococcus strain WH 8103 (clade III), i.e. with little effect of ammonium on nirA transcription (Bird and Wyman, 2002). Such regulation contrasts with the enhanced expression of nirA and amt1 in the absence of ammonium in freshwater Synechococcus strains (see for example Vazquez-Bermudez et al., 2002) and highlights the regulatory constraints placed on these organisms in the N-poor marine ecosystem. This feature may well differ subtly between coastal and open-ocean strains. The critical role that N availability potentially plays in generally controlling phytoplankton growth rate and/or growth yield (Tyrell and Law, 1997), suggests that the availability of this resource is also vital to the ecological success of the Synechococcus genus. However, Synechococcus growth rates in situ are high despite low N (see Furnas and Crosbie, 1999 for review), perhaps a result of the rapid recycling of the N sources specifically used by members of this genus, so it is possible that these organisms do not encounter N stress in situ. To begin to address whether this is in fact the case, and to circumvent the problems of using oceanographic techniques which generally monitor bulk activities as a function of a complex phytoplankton assemblage making it difficult to assess the N status of a specific phytoplankton taxon, Lindell and Post (2001) have developed an assay based on ntcA gene expression as an indicator of the N status

DIVERSITY AND ADAPTATION IN SYNECHOCOCCUS

31

specifically of the marine Synechococcus genus. Using Synechococcus sp. WH 7803 as a model organism, ntcA expression was shown to be induced at ammonium levels below 1 mM, reaching maximal levels within 2 hours. Conversely, addition of >1 mM ammonium led to a rapid decline in ntcA mRNA. This negative effect of ammonium on ntcA expression could be prevented by addition of MSX or azaserine, inhibitors of ammonium assimilation. The highest ntcA transcript levels were, however, found in cells lacking a N source that could be used for growth. Thus, basal ntcA expression would indicate ammonium utilisation, whilst maximal expression would indicate N deprivation, levels of which could be induced experimentally using ammonium and MSX respectively, and compared with actual ntcA transcript levels from an in situ sample. Using such an assay, it was shown that natural Synechococcus populations expressed ntcA at a basal level at a nutrient-enriched site in the Gulf of Aqaba, Red Sea, suggesting they were utilising solely ammonium in this environment even though levels of nitrite and nitrate were measurable (Lindell and Post, 2001). Further application of this marker, particularly when used in tandem with molecular markers specific to other nutrients, e.g., phosphorus and trace metals (see Sections 4.2–4.3), will provide a much clearer picture of the role of particular nutrients in controlling the standing stock of Synechococcus in situ.

4.2. P Acquisition Inorganic orthophosphate (Pi) levels in the open ocean are often in the nanomolar range (see for example Bjo¨rkman et al., 2000) and there is now evidence from several ocean environments for phosphorus (P) limitation of microbial production (Zohary and Robarts, 1998; Wu et al., 2000; Karl et al., 2001a; San˜udo-Wilhelmy et al., 2001). The major consequence of a Plimited ocean is selection for microbes with high P acquisition efficiency, but low P quotas, a scenario that seems to be occurring now in the North Pacific subtropical gyre where Prochlorococcus tends to dominate numerically over Synechococcus (Karl et al., 2001b), presumably a function of the lower cell P quota and higher surface to volume ratio of the smaller celled Prochlorococcus genus (Cavender–Bares et al., 2001). Given the apparent importance of P controlling production in marine systems, we still know relatively little of the P physiology of either genus, though some insights pertaining to regulation of P acquisition and P source utilisation have recently been forthcoming in Prochlorococcus, a result of genome sequencing projects (http://www.jgi.doe.gov/JGI_microbial/html/index.html) and the

32

DAVID J. SCANLAN

availability of an axenic Prochlorococcus strain (Rippka et al., 2000; Scanlan and West, 2002). For the marine Synechococcus, work on P nutrition and regulation has focused on only a few strains (Scanlan et al., 1993, 1997a,b; Donald et al., 1997; Ikeya et al., 1997; reviewed in Scanlan and Wilson 1999). Although components of the P acquisition machinery are likely to be conserved across the genus, more data are required to assess the comparative utilisation of different P sources among genotypes, or whether open-ocean and coastal ecotypes possess different P quotas and/or P regulatory systems, for example. The differential bioavailability of organic P compounds in situ (Bjo¨rkman and Karl, 1994; Bjo¨rkman et al., 2000) suggests that different ecotypes might show specific preferences for meeting their P requirements for cell growth. For the most well studied marine Synechococcus strain WH 7803, a relatively broad utilisation of P sources has been described. This strain has been shown to utilise Pi, dCTP, p-nitrophenyl phosphate, glucose6-phosphate and glycerol phosphate, but not cAMP, as sole P source for growth (Donald et al., 1997). Interestingly, the incorporated Pi has been shown to accumulate largely in low molecular weight and RNA components in this strain, but with a general absence of phospholipid and little polyphosphate (Cuhel and Waterbury, 1984). It is possible that the ability to utilise cAMP as sole P source may be a clade-specific trait amongst marine Synechococcus strains; although strains WH 7803 (clade V) or WH 8012 (clade II) cannot utilise this substrate, cAMP will support the growth of strain WH 8103 (clade III) (L. Moore, personal communication). All three strains can utilise nucleotides as a P source, which suggests hydrolysis by alkaline phosphatase or the presence of a specific 50 -nucleotidase. Differences in alkaline phosphatase activity between strains WH 8103 and WH 8012 under P-deplete conditions (L. Moore, personal communication) hint at the potential different strategies of strains to utilise the available P pool. Some strains appear more capable of accessing the organic P pool than others, particularly if hydrolysis can be coupled to uptake. Utilisation of cAMP suggests the presence of a specific high affinity transport system for this substrate, a capacity which appears to be relatively rare among marine bacterioplankton (Ammerman and Azam, 1987). An in situ study of P uptake in natural picophytoplankton populations at the Bermuda Atlantic Time-Series (BATS) Station (Cotner et al., 1998) showed, however, no selectivity of uptake of inorganic (33P-Pi) over organic P (d32P-ATP) by Synechococcus or Prochlorococcus, though Synechococcus incorporated the majority of the labelled P sources (70% of the total). Little difference was observed in cell-specific incorporation of 14C-NaHCO 3

DIVERSITY AND ADAPTATION IN SYNECHOCOCCUS

33

suggesting that Synechococcus cells have a higher P requirement than Prochlorococcus and thus that Synechococcus spp. may be more susceptible to P limitation. However, there was a strong correlation between alkaline phosphatase activity and Synechococcus abundance in the Gulf of Aqaba, Red Sea, with the bulk of particulate alkaline phosphatase activity associated with the picoplanktonic fraction (Li et al., 1998) suggesting that this genus might be a significant contributor to removal of the bioavailable organic P in this region at least. It is important to highlight though, that this in situ alkaline phosphatase activity data needs to be put into the context of the contribution of individual genotypes to the total activity, a variable that may differ significantly between genotypes of specific Synechococcus clades. Depending on hydrographic conditions, it is possible to envisage times of relative abundance of Pi, e.g., after deep winter mixing events, compared to relative Pi scarcity when the external Pi concentration increases only transiently above the threshold for Pi incorporation, e.g., during summer stratification. The threshold concentration is the external Pi concentration at which incorporation of the nutrient stops due to energetic reasons. Any increase above this value is then utilised by the organism until the threshold is once again reached. It is obvious that survival in this special growth situation of fluctuating and limiting nutrient supply requires synthesis of a high-affinity Pi uptake system that senses changes in the external Pi status and is capable of ‘adapting’ to these changes rapidly in order to optimise the efficiency of nutrient incorporation. There has been much progress in understanding the kinetic mechanisms underlying Pi uptake during this ‘adaptive response’ to conditions of a pulsed P supply in a freshwater Synechococcus sp. (see Falkner et al., 1995; Wagner et al., 1995a, 2000), but our knowledge in marine strains is far less advanced. Kinetic data has confirmed the presence of a high affinity Pi uptake system in marine Synechococcus (Donald et al., 1997; Ikeya et al., 1997; F. Wagner and D.J. Scanlan unpublished data). Reported Km (Ks) values for this system range from 40–100 nM (Ikeya et al., 1997) in strain NIBB 1071, to 130 nM (F. Wagner and D.J. Scanlan, unpublished data) – 3 mM (Donald et al., 1997) in strain Synechococcus WH 7803. The wide range in Km value observed in the latter strain is likely to reflect the growth conditions used (semi-continuous in the former, and batch in the latter). This high affinity system was shown to be energy-dependent, the Km being reduced in the dark or on addition of the uncoupler FCCP, which inhibits ATP production (Donald et al., 1997). Consistent with this is that vanadate, a general inhibitor of P-type ATPases (Pedersen and Carafoli, 1987), also inhibits Pi uptake, suggesting translocation is coupled to the activity of an ATPase

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DAVID J. SCANLAN

(F. Wagner and D.J. Scanlan, unpublished data). Since uptake is specifically reduced by the addition of the photosynthetic electron transport chain inhibitors DCMU and DBMIB (Ikeya et al., 1997), it is likely that high affinity Pi uptake requires photophosphorylation carried out by both cyclic and non-cyclic electron flow. This high affinity system is also sensitive to osmotic shock (Ikeya et al., 1997), a property consistent with the presence of a periplasmic Pi binding component. Such a component, encoded by the pstS gene, has been well characterised in Synechococcus sp. WH 7803 (Scanlan et al., 1993) though in this, and other marine Synechococcus strains, the PstS protein appears to localise with the cell wall. The significance of this location awaits further work. Certainly the recent characterisation of a Synechococcus sp. WH 7803 pstS mutant is consistent with its role as a classical binding component of an ABC transport system, since the Km is unaltered in the mutant, whilst the Vmax decreases 2–3 fold (F. Wagner and D.J. Scanlan, unpublished data). This suggests that lack of PstS essentially influences the velocity of the overall transport process, a phenomenon which is coherent with PstS acting as a ‘simple’ harvesting protein that accelerates supply of the ABC membrane complex with substrate. PstS is induced in strain WH 7803 when external Pi levels drop below 50 nM (Scanlan et al., 1997b). Further, PstS expression under P depleted growth conditions is repressed by addition of all other utilisable P sources in this strain. These features of induction of PstS at Pi levels similar to those found in situ, and tight and specific repression by both inorganic and organic P sources, have led to use of the polypeptide as a molecular marker of the P status of natural Synechococcus populations (Scanlan et al., 1997b; reviewed in Scanlan and Wilson, 1999; Scanlan and West, 2002). This latter property of PstS repression is particularly important for its use as a reporter in situ where P sources other than that which can be chemically measured may be bioavailable (see Kolowith et al., 2001; Sundareshwar et al., 2001). The utility of such a protein as a molecular marker has been assessed in oligotrophic waters of the Sargasso Sea and Gulf of Aqaba (Red Sea) where expression appears to be both spatially and temporally regulated (Scanlan and Wilson 1999; N. Fuller, A. Post and D. J. Scanlan, unpublished data). Interestingly, temporal regulation of PstS expression at Station A in the Gulf of Aqaba coincides with the fall of a spring Synechococcus maximum (N.J. Fuller, D. Marie, A.F. Post and D.J. Scanlan, unpublished data) suggesting P control of production at that time. The same community structure data demonstrate that winter and summer Synechococcus populations differ markedly in genetic diversity, concomitant with organisms that may have markedly different strategies for P acquisition, or regulation of P metabolism.

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Regulation of the high affinity P acquisition system of marine Synechococcus is likely to require the functioning of the PhoB –PhoR two-component system (Watson et al., 1996) though confirmation of this by mutant construction is still needed. It is possible that the regulatory network controlling P acquisition may be more complicated than these ‘twocomponents’ since, immediately downstream of pstS in strain WH 7803, lies a putative transcriptional regulator (ptrA) (Scanlan et al., 1997a), of the cyclic AMP receptor protein (crp) and fumarate and nitrate reduction (fnr) family (Green et al., 2001), and similar to the cyanobacterial ntcA gene (see Section 4.1). Upstream of ptrA, an 18 nucleotide sequence representing a putative Pho box (Wanner, 1996) was found, suggesting regulation by the response regulator PhoB (Scanlan et al., 1997a). A similar gene arrangement of pstS immediately upstream of ptrA is found in the completed genome sequence of the marine Synechococcus strain WH 8102. This genome sequence hints at the potential complexity of P acquisition in this organism, since four copies of pstS, all potentially encoding Pi-binding proteins can be found. One of these copies shares greatest identity with sphX, a gene encoding a P-regulated polypeptide in the freshwater Synechococcus sp. PCC 7942 (Aiba and Mizuno, 1994). Mutation of the sphX gene in this freshwater Synechococcus strain does not affect the ability of the organism to utilise phosphate at nanomolar external concentrations, but it does lead to an increase in the Michaelis constant and the maximum velocity of the initial influx of 32P-Pi (Falkner et al., 1998). Moreover, the capacity of the wild type to adapt within a few minutes to a transitory increase in the external phosphate concentration in an energetically efficient way is lost in the mutant. As a result, the mutant can no longer attain pulse-adapted states that reflect in a characteristic way preceding exposures to higher phosphate concentrations. The role of sphX and the other pstS copies in the marine Synechococcus WH 8102 requires further work, though it could be that one of these copies transports a structurally related ion e.g. arsenate, a molecule known to be transported by marine Synechococcus (Takahashi et al., 2001), and also a molecule that can act as a competitive inhibitor of Pi transport (see Abedin et al., 2002). Interestingly, sphX is not found in the completed marine Prochlorococcus genomes, organisms which seem to show differential composition of their P acquisition and regulatory machinery between surface and deep ecotypes (see Scanlan and West, 2002). Finally, the presence in the Synechococcus WH 8102 genome of genes encoding components of a phosphonate or phosphite transport system reiterate the fact that these organisms may be able to access components of the in situ dissolved organic phosphorus (DOP) pool that are only just being recognised as important in marine systems (see Kolowith et al., 2001).

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4.3. Micro-nutrient Acquisition The bioactive trace metals Mn, Fe, Co, Cu, Ni and Zn are elements important for various metabolic functions (e.g. photosynthetic electron transport), and essential for phytoplankton growth. Much recent interest has centred on the role of these trace metals, and particularly iron, in controlling marine primary production (see Limnology and Oceanography [1991] 36, 1507–1970; Geider and La Roche, 1994; Wells et al., 1994, 1995; Zettler et al., 1996; Tortell et al., 1999; Sunda, 2000), though as well as availability constraints on production, toxicity effects are also important (see e.g. Robinson et al., 2000; Mann et al., 2002). Specifically with respect to the marine Synechococcus genus, most of these bioactive metals are likely to be essential for growth (Brand et al., 1983). Recent evidence suggests that cobalt is a specific requirement (Sunda and Huntsman, 1995; Saito and Moffett, 2002). This requirement for Co contrasts with data for Zn. Synechococcus bacillaris (CCMP1333 ¼ WH 5701) (MC-B) and Synechococcus WH 7803 (clade V) showed no difference in growth rate over a range of Zn concentrations (Brand et al., 1983; Sunda and Huntsman, 1995), with the former strain also showing no ability to metabolically replace Co with Zn (Sunda and Huntsman, 1995). A low Zn requirement by these organisms is suggested by (a)

the presence of a Cu/Zn superoxide dismutase in Synechococcus sp. WH 7803 (Chadd et al., 1996a), (b) the absence of a carboxysomal carbonic anhydrase, usually a zinc requiring metallo-enzyme (see Section 3.2) in Synechococcus sp. WH 8102, and (c) the apparent lack of a phoV gene homologue, which in a freshwater Synechococcus sp. PCC 7942 encodes a constitutively-expressed Zn(II)requiring alkaline phosphatase (Wagner et al., 1995b), an enzyme that might be considered advantageous in P-depleted oceanic environments.

This last point also demonstrates how cellular trace metal requirements could impact on acquisition and assimilation of the macro-nutrients C, N and P in this genus (see McKay et al., 2001 for a general review on this topic). Such a trace metal constraint on N metabolism is likely to be due to the Fe requirement of nitrate and nitrite reductase (though see Henley and Yin, 1998), and, for those strains that utilise urea, the Ni requirement for urease (Collier et al., 1999). This latter obligation may be obviated should some marine Synechococcus strains produce a Niindependent urea amidolyase similar to that found in green algae (see Antia et al., 1991).

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The ability of natural Synechococcus (and/or Prochlorococcus) populations to produce cobalt ligands (Saito and Moffett, 2001), whilst Cucomplexing ligands have been reported for marine Synechococcus cultures and in oceanic waters (Moffett, 1995; Moffett and Brand, 1996), demonstrates the importance of trace metal complexation in natural marine systems. Such complexation appears important in dictating trace metal speciation on the one hand, which may affect actual availability (Saito and Moffett, 2001), and also in limiting metal toxicity on the other (Moffett and Brand, 1996; Moffett et al., 1997). The observed Synechococcus sensitivity to Cu in coastal waters, and particularly polluted harbours, appears to result from an increase in free Cu2þ as a result of saturation of these organic ligands (Moffett et al., 1997). There is recent evidence to suggest that the Prochlorococcus genus is even more sensitive to Cu levels than its Synechococcus neighbour (Mann et al., 2002). Interestingly, there is a differential sensitivity to Cu in Prochlorococcus with the high light-adapted ecotype more Cu resistant than its low light counterpart, consistent with the higher free Cu2þ levels found in shallow mixed layers compared to deeper waters. Given the relative importance of trace metals in marine systems, little is known of the mechanisms of metal ion transport or physiological responses to specific trace metal deficiency in the marine Synechococcus genus, perhaps with the exception of iron, although even here most work relates to the halotolerant marine Synechococcus strain PCC 7002 (Trick and Wilhelm, 1995; Wilhelm 1995; Wilhelm and Trick, 1995a,b; Wilhelm et al., 1998), phylogenetically outside the MC-A/MC-B group. Genes encoding components of putative high affinity ‘ABC systems’ that might transport trace metals can, however, be found in the Synechococcus WH 8102 genome sequence. These include a potential iron-binding protein, an inner membrane channel and an ATPase that are predicted to comprise a complete periplasmic-binding component-dependent ABC transporter for Fe (Webb et al., 2001), although their specific functional characterisation via mutant construction awaits further work. One of these components, the iron binding protein, has recently been identified as an IdiA (Michel et al., 1996, 1999) or SfuA (Angerer et al., 1992) homologue, encoding a 39 kDa polypeptide specifically induced in Fe-depleted conditions in Synechococcus WH 8103, WH 8102 and WH 7803 and localised to the outer membrane (Webb et al., 2001). This polypeptide is probably synonymous with the 36 kDa Fe-induced polypeptide identified by Chadd and co-workers (Chadd et al., 1996b). Although some marine Synechococcus strains (e.g. WH 8101 and WH 7805) have been shown to produce siderophores – catechol or hydroxamate chelators with high affinity for Fe (Wilhelm and

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Trick, 1994) – other strains appear not to be capable of siderophore synthesis (e.g. WH 6501, WH 7803, WH8018 and WH 8102 [Rueter and Unsworth, 1991; Chadd et al., 1996b; Webb et al., 2001]), the last strain also lacking any known siderophore biosynthetic genes. Since WH 7803 can, however, incorporate Fe complexed to exogenous siderophore (Hutchins et al., 1999), it is possible that some strains are capable of siderophore ‘piracy’ – a trait also found in some marine heterotrophs (Granger and Price, 1999). Whether this ‘pirate’ behaviour is a clade-specific trait will require a thorough survey of siderophore production and utilisation across the genus. Further characterisation of Fe acquisition mechanisms among members of the genus is required, especially since strain WH 8102 also appears to lack genes encoding TonB, ExbB and ExbD, components required for Fe-siderophore transport (Webb et al., 2001). IdiA (Webb et al., 2001), together with the isiA gene (Geiß et al., 2001) encoding a chlorophyll a-binding protein that may protect the photosystems against excess light under iron-limiting conditions (Burnap et al., 1993), are currently being developed as potential diagnostic markers for assessing the Fe status of natural cyanobacterial populations. Use of these markers, rather than that of flavodoxin accumulation [which has been well characterised as a potential marker of Fe status in phytoplankton (La Roche et al., 1996)] has been necessitated by the reported absence of flavodoxin in the marine Synechococcus strain CCMP 1334 (¼ WH 7803) as determined by a sensitive HPLC assay (Erdner et al., 1999). This is consistent with an absence of isiB, encoding flavodoxin, in the marine Synechococcus WH 8102 genome, a gene notably present in both the Prochlorococcus MED4 and MIT9313 genomes. The physiological response of members of the marine Synechococcus genus to iron deficiency has been reported in only a few studies (Brand, 1991; Rueter and Unsworth, 1991; Chadd et al., 1996b; Kudo and Harrison, 1997; Henley and Yin, 1998) again mostly focusing on a single strain WH 7803. This strain shows an obvious decrease in growth rate as Fe levels fall below 100–20 nM, but there are conflicting data over whether ammonium, rather than nitrate, supports a higher growth rate under Fe limited conditions (Kudo and Harrison, 1997; Henley and Yin, 1998), as might be expected due to the Fe requirement of nitrate and nitrite reductase. Where comparative work has been performed in other marine Synechococcus strains, subtle differences in their responses to Fe limitation have been observed (Rueter and Unsworth, 1991; Henley and Yin, 1998) and this may underlie differences in Fe quotas between strains and/or the alternative strategies of Fe acquisition and utilisation dictated by their specific environmental niche.

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5. MOTILITY The non-flagellar-based swimming motility of marine Synechococcus (Waterbury et al., 1985; Pitta and Berg, 1995; Brahamsha, 1996a; Pitta et al., 1997) is one of the few phenotypic traits that thus far represents phylogenetic coherence within the genus (Toledo et al., 1999), with all motile strains currently falling within clade III (see Fig. 1; Rocap et al., 2002). This swimming motility is of a unique type characterised by the absence of flagella and of any other obvious motility organelle. The specific swimming mechanism remains elusive (see Brahamsha, 1999a for a review), although a protein SwmA, has been shown to be required for the generation of thrust, since swmA mutants lose the ability to translocate, although they can generate torque (Brahamsha, 1996a). Of specific interest here with respect to niche adaptation is that at least one motile strain, WH 8113, has been shown to be chemotactic towards a variety of nitrogenous sources including ammonia, nitrate, b-alanine, glycine and urea (Willey and Waterbury, 1989). Such positive chemotaxis probably allows motile strains to access local sources of nutrient enrichment, which may be particularly apparent around microaggregates (marine snow) or point source releases of nutrients, e.g., autolysis of a large microbe, the discharge of undigested organic matter and inorganic nutrients from food vacuoles or via zooplankton excretion. Such point sources of nutrients have been shown to spread into spherical patches a few millimetres in diameter and can sustain swarms of bacteria (Blackburn et al., 1998). Hence, across distances of millimetres to centimetres, motile Synechococcus strains appear to have a selective advantage over their non-motile counterparts particularly in oligotrophic open-ocean systems, an environment where motile strains appear to predominate. Such microscale structure to a water column imparts a three dimensional shape to any simplified niche system based solely on the vertical light and nutrient gradients, but ultimately will need to be considered with regard to niche adaptation. It will be interesting to resolve using molecular techniques whether motile Synechococcus strains show relatively high abundance on or around marine snow particles relative to their nonmotile counterparts.

6. CELL CYCLE The daily light:dark cycle of temperate and subtropical marine waters imparts a diel periodicity on natural microbial populations that can

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influence productivity, cell division, nutrient uptake processes and cellular optical properties (see e.g. Campbell and Carpenter, 1986b; Pre´zelin et al., 1986, 1987; Pre´zelin, 1992; Stramski et al., 1995; Jacquet et al., 1998; Kuipers et al., 2000; Binder and DuRand, 2002) as well as the expression of specific genes (Wyman, 1999). For photoautotrophs like Synechococcus, such periodicity imparts a naturally periodic energy supply to which cellular processes need to be aligned (Chisholm, 1981); the mechanisms by which this alignment is achieved may vary between genotypes. This phasing of cell division and other processes to the daily light:dark cycle may be a direct effect of light control of the cell cycle or be due to the existence of an endogenous circadian rhythm ( biological clock), both of which have been reported for marine Synechococcus (Armbrust et al., 1989; Sweeney and Borgese, 1989). In the freshwater Synechococcus strain PCC 7942, this circadian rhythm imparts a global influence on gene expression (see Nair et al., 2002), and hence presumably on the processes in which those genes function. Most physiological processes, excluding those of cell division, that have been characterised in the genus have been studied under continuous illumination thus circumventing such ‘circadian influence’, but this must be remembered when extending conclusions of culture work to natural populations. Analysis of the regulation of the cell cycle of marine Synechococcus strains has been explored in detail in the genus (Binder and Chisholm, 1995; Liu et al., 1999; Binder, 2000; Jacquet et al., 2001), particularly so since there is the potential to apply cell cycle analyses of division rate to estimate in situ growth rates of natural Synechococcus populations (see e.g. Jacquet et al., 1998). Two modes of cell cycle regulation have been reported for marine Synechococcus strains, a trait which appears to be strain-specific. For one strain, WH 7803, contrasting cell cycle data have been obtained, suggesting that culture conditions are also important or that specific genetic differences exist in strains of the same name (Binder and Chisholm, 1995; Liu et al., 1999; Jacquet et al., 2001). The first of these modes gives rise to bimodal DNA distributions (e.g. strains WH 8101, WH 8103 and WH 7805) in which cells arising with a single chromosome begin replication of that chromosome after a specific time interval before completing replication X min later and ultimately dividing Y min after this (Binder and Chisholm, 1995). These same workers also observed a second mode of cell cycle regulation (e.g. strain WH 7803) which gives rise to multimodal DNA distributions and probably involves the inheritance of multiple genome copies and the asynchronous replication of these copies. More detailed analysis of the cell cycle in Synechococcus WH 8101 during light and nitrogen limited

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growth, demonstrated by bimodal DNA distributions under all growth conditions, reiterates a relatively simple cell cycle behaviour in this strain (Binder, 2000). This study also showed that the timing of chromosome replication is dependent on cell mass (volume). Cell volume changes may in turn affect the non-linear relationship between growth rate and rRNA content in this strain (Binder and Liu, 1998), rRNA content potentially being another useful biochemical ‘index’ for assessing in situ growth rate. Differences in cell cycle regulation among strains may explain the relatively wide variation in the timing of division in the Synechococcus genus (see Table 4), compared to Prochlorococcus and picoeukaryotes (Jacquet et al., 2001). These investigators also provide data which hint at the potentially interesting observation that the timing of division may be different between low PUB:PEB strains and high PUB:PEB strains, with the former dividing during daylight and the latter at night. Variation in the timing of division among the low PUB:PEB isolates was also reported, possibly reflecting a differential response to their light environment and with individual strains requiring different integrated light doses to initiate the division process (Jacquet et al., 2001). Further, these low PUB strains stop dividing almost immediately in the dark suggesting that light is necessary to complete division. These culture studies agree well with field estimates of the timing of cell division, with coastal locations generally containing cells dividing earlier than openocean sites (see Table 4). This is consistent with the observed higher abundance of low PUB strains in coastal waters and of high PUB strains in more open-ocean systems, with the caveat that PUB : PEB ratio is polyphyletic in the genus and that chromatically adapting strains exist (see Section 2). The observed late division (i.e. after dusk) of natural Synechococcus populations in the Mediterranean Sea may be a result of P limitation of cell growth in that environment, particularly since P depletion was shown to retard DNA replication and hence delay cell division (Vaulot et al., 1996). However, it is also possible that Synechococcus cell division is controlled negatively by UV irradiation, rather than positively by nutrient supply (Jacquet et al., 1998). UV irradiation is an environmental variable that appears important in controlling the timing of DNA synthesis in natural Prochlorococcus populations (Vaulot et al., 1995). Even so, cell cycle variables do appear useful as potential molecular markers for assessing cellular nutrient status, particularly to monitor nutrient bioassays since nutrient deprived cells generally arrest at the beginning of their cell cycle (G1 phase), but initiate DNA synthesis (S phase) within a few hours of nutrient addition.

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Table 4

Timing of division cycle for marine Synechococcus under L:D in culture (A) and in the field (B). Strain

Phenotype

L:D

A – Culture studies Mitsui et al. (1986) Armbrust et al. (1989) Binder and Chisholm (1995) Binder and Chisholm (1995) Waterbury et al. (1986) Mori et al. (1996) Jacquet et al. (2001) Jacquet et al. (2001) Jacquet et al. (2001) Campbell and Carpenter (1986b) Campbell and Carpenter (1986b) Stramski et al. (1995) Sweeney and Borgese (1989) Campbell and Carpenter (1986b) Jacquet et al. (2001)

BG43511/12 WH 8101 WH 8101 WH 7803 WH 7803 PCC 7942 ROS 04 ALMO 03 WH 7803 WH 7803 WH 8107 WH 8103 WH 7803 WH 8012 WH 8103

n. a. No PE No PE Low PUB/PEB Low PUB/PEB No PE Low PUB/PEB Low PUB/PEB Low PUB/PEB Low PUB/PEB High PUB/PEB High PUB/PEB Low PUB/PEB Low PUB/PEB High PUB/PEB

12h:12h 14h:10h 14h:10h 14h:10h 14h:10h 12h:12h 12h:12h 12h:12h 12h:12h 14h:10h 14h:10h 12h:12h 12h:12h 14h:10h 12h:12h

Timing of division

DAVID J. SCANLAN

Author(s)

Populations

B – Field studies Waterbury et al. (1986) Affronti and Marshall (1994) Carpenter and Campbell (1988) Li and Dickie (1991) Sherry (1995) Xiuren and Vaulot (1992) Waterbury et al. (1986) Liu et al. (1998) Agawin and Agusti (1997) Neuer (1992) Olson et al. (1990) Olson et al. (1990) Kudoh et al. (1990) Vaulot and Marie (1999) Wyman (1999) S. Jacquet (unpub. data) Vaulot et al. (1996) Campbell and Carpenter (1986b) Pre´zelin et al. (1987) Dolan & Simek (1999) Jacquet et al. (1998)

Vineyard Sound Chesapeake Bay Long Island Sound N Atlantic N W Arabian Sea English Channel Sargasso Sea Arabian Sea NW Mediterranean Sea Sub. Arct. Pacific N Atlantic, Sargasso Sea Vineyard Sound N W Pacific Equat. Pacific N E Atlantic Alboran Sea NW Mediterranean Sea Sargasso Sea, Carribean Sea N W Atlantic N W Mediterranean Sea NW Mediterranean Sea

Timing of division

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Author(s)

Phenotype is indicated by the absence of phycoerythin (PE) or by the ratio of the chromophores Phycourobilin (PUB) to Phycoerythrobilin (PEB). n.a.: not available. As photoperiod varies with locations and culture conditions, the period of division is represented by dark symbols ( ) and located approximately over a daily 24 h ) for night and day, respectively. period without time precision but symbolised by dark and white bands ( Reproduced from Jacquet et al., 2001 with permission from the Journal of Phycology.

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7. GRAZING/VIRUSES Optimum growth of Synechococcus in situ generally occurs in the upper surface layers of the water column (see Partensky et al., 1999a), and identifying the controlling environmental influences on growth rate and growth yield is central to providing a robust understanding of pelagic food webs where Synechococcus plays a key role as a source of particulate organic material, as well as in nutrient utilisation and C transfer to higher trophic levels. Such factors important in controlling in situ growth dynamics of Synechococcus populations have been recently reviewed (Furnas and Crosbie, 1999), and include both abiotic and biotic factors. The former include nutrient (and light) availability, temperature fluctuations as well as seasonal mixing events, whilst the latter include grazing pressure (Campbell and Carpenter 1986a; Caron et al., 1991; Liu et al., 1995; Christaki et al., 1999; Dolan and Simek, 1999) and viral infection (see Suttle, 2000, N.H. Mann in press, for recent reviews). We have already considered the importance of abiotic factors in shaping the environmental niche occupied by specific Synechococcus genotypes, but these biotic factors also play an important role in shaping the success of niche occupancy, through selective grazing or genotype specific virus-infection. Selective grazing of marine Synechococcus versus Prochlorococcus populations has already been reported (Christaki et al., 1999) whilst selection within members of the genus is known for freshwater isolates. Thus, heterotrophic nanoflagellates isolated from Lake Constance show genotype-specific prey selection, with selection being a function of the ultrastructural morphology of the Synechococcus cell surface, strains containing a highly glycosylated, paracrystalline surface layer being ingested at lower rates. Moreover, a phycocyanin-rich strain was preferentially ingested by one of these nanoflagellates, a Paraphysomonas sp., even when it was just 10% of the prey, whilst another nanoflagellate, Bodo saltans, ignored the strain completely despite being the sole source of prey (Mu¨ller, 1996). Such prey selection, should it occur in marine systems, would have considerable influence on the population dynamics of individual Synechococcus genotypes, and it would be particularly interesting to see if motile strains, for example, are grazed at lower rates than their non-motile counterparts since the former may be capable of specific predator avoidance. Concomitant with a potential effect of grazing on dictating Synechococcus population structure, there is now good recent evidence that viruses can also play an important role in ordering the genetic structure

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of natural Synechococcus communities, as well as controlling their abundance. Thus, it is already known that specific marine cyanophage possess different host ranges (Suttle and Chan, 1993) with genetically distinct cyanophage in open-ocean versus coastal waters (Zhong et al., 2002). There is also evidence from in situ studies that Synechococcus genetic diversity co-occurs with viral diversity (M. Mu¨hling, N.J. Fuller, A. Millard, D.J. Scanlan, A.F. Post, W.H. Wilson, D. Marie and N.H. Mann, unpublished data). Viral infection, as well as affecting genetic diversity of the host population, may also have a direct influence on the physiological properties of the host cell. Thus, sequence information of phage SPM-2, a lytic phage capable of infecting Synechococcus WH 7803 (Fuller et al., 1998) with a genome size of 194 kb (Hambly et al., 2001), contains within its genome one open reading frame sharing sequence homology with CpeT (N.H. Mann, personal communication), a protein potentially involved in regulating PE expression in cyanobacteria (see Cobley et al., 2002). Should SPM-2 be maintained in the host as a pseudolysogen, phage-encoded genes like cpeT may confer a selective advantage on the host under specific environmental conditions. Examples of this so-called lysogenic or phage conversion have been most evident in studies on pathogenic bacteria where phage-encoded genes can contribute to virulence (Barondness and Beckwith, 1995; Pacheco et al., 1997). The gene encoding cholera toxin encoded on the filamentous bacteriophage CTXphi is a particularly striking example (Waldor and Mekalanos, 1996). As well as the possibility of the phage themselves imparting specific physiological properties on the host, there is also the likelihood that a specific physiological response of the host to its environment may then allow infection by a specific cyanophage. The most obvious scenario would be the induction of a high affinity nutrient acquisition system comprising an outer membrane component that subsequently acts as a phage receptor. Examples of outer membrane proteins acting as phage receptor proteins in Gram-negative bacteria include FhuA, a ferrichrome receptor and PhoE, a P-inducible anion selective pore (see Heller, 1992 for review). This scenario runs alongside that of the phage establishing (pseudo)lysogeny under these nutrient depleted conditions, which would be a direct mechanism for phage maintenance when host growth is impaired. Such a pseudolysogeny concept (i.e. a phage-host relationship in which a phage-infected cell grows and divides even though its virus is pursuing a lytic infection) is supported by experimental data using Synechococcus WH 7803 and cyanophage SPM-2 in which P-depleted, but not N-depleted growth of the host resulted in a 80% reduction in burst size but no difference in phage

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adsorption kinetics (Wilson et al., 1996). Such data support the concept that P status can dictate the outcome of phage-host interactions in this system.

8. CONCLUSIONS Although much is known of the physiology of the marine Synechococcus genus most work has focused on only a few strains and there await comparative analyses of physiological properties of representatives of different phylogenetic clades of this group. Such work should aim to clarify the light and nutrient physiology of the genus with respect to whether high and low light-adapted clades exist, as is the case for Prochlorococcus, as well as whether there is any obvious nutrient selection between members of the same clade, compared to other clades. It is always possible that only focusing on the abiotic factors mentioned above will look at the physiology of the genus too simplistically, and that there may be no real distinction in their nutrient and light physiologies with respect to phylogeny – a view that might be disputed given the rampant horizontal gene transfer that seems evident from currently sequenced microbial genomes (reviewed in Boucher et al., 2001). The fact that motility, and possibly chromatic adaptation in non-motile strains, shows phylogenetic coherence, however, suggests that some phenotypic traits are clade-specific. Even if some specific physiological traits are shared between clades, it would be expected that it is the total suite of traits that correlates with the environmental niche occupied. At the genome level this would be reflected in a genomic ‘core’ found in all strains and an auxillary set of genes found in different strain subsets ( a phylogenetic clade) required for occupation of that specific niche. Genomic microheterogeneity that could ultimately manifest itself in physiological diversity has already been reported for natural marine crenarchaeotes even though there was little 16S rRNA sequence variation (Be´ja`, et al., 2002). Subtleties in gene regulation networks also need to be studied in strains of different clades under standard conditions before some real grasp of the complexity and diversity of between-strain adaptation can be put into proper perspective. Such work reiterates the caveat that laboratory culture and selection must not have unrealistically ‘changed’ the physiology of such isolates, one of the major problems of modern microbiology. Further, astute interpretation of such physiological data is required for them to be correctly extrapolated to the in situ environment. Given such cautions, readily culturable aquatic microbes such as Synechococcus are relatively

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unique in allowing the investigator access to its in situ niche, and so permitting the coupling of physiology with the use of molecular ecological tools for defining the boundaries of genotype distribution in the water column. This really does allow comparative physiology versus ecological niche analysis to be realistically assessed. Moreover, since members of this genus are genetically amenable (Brahamsha, 1996b, 1999b) we have a system that also allows specific mutagenesis of niche-specific genes, as well as the potential to genetically shuffle clade-specific traits between ecotypes, which would allow competitive growth experiments to be performed. The latter would be useful since it is already known that differences in growth rate exist even among members of the same marine Synechococcus clade (Palenik, 1999), yet the underlying mechanism for such differences in clonal growth rate remain elusive. Although intrinsic cell properties alongside the environmental resources of light and nutrients available define these absolute growth rates, biological factors such as grazing and viral lysis ultimately dictate net growth rates. These latter biotic factors will also define the higher order level shaping of niches, a result of interactions which may be negative e.g. the virus and predator grazing just mentioned, or positive, e.g. with cooccurring photosynthetic picoeukaryotes or consortial bacteria. These positive interactions may include a potentially large array of biological signals that play a role in maintaining an environmental niche that is mapped into a three-dimensional shape by abiotic factors. Trying to dissect such a system would be a difficult challenge but it is encouraging that we are obtaining a clearer picture of the diversity of photosynthetic eukaryotes (see Moon-van der Staay et al., 2001), whilst also beginning to establish in culture marine bacteria once thought to be unculturable, with important biogeochemical function (Eguchi et al., 2001; Kolber et al., 2001; Zehr et al., 2001; Rappe´ et al., 2002). The future construction of a defined in vitro mesocosm or the like, containing selected components of the major photosynthetic and heterotrophic microbial ‘players’ in situ, might be possible for analyses of these interactions to be undertaken. As well as the marine environment, freshwater lake (see Postius and Ernst, 1999) and hot-spring environments (see Ward et al., 1998) are also yielding fascinating insights into the potential of Synechococcus genotypes to adapt to a distinct physiological niche, even if such niches might be separated by just a few hundred micrometres as is the case in the hot-spring mat system (Ramsing et al., 2000). Many surprises remain to be elucidated in these different systems but taken together they will certainly extend and promote the Synechococcus genus as a potentially excellent model to examine the physiology of microbes in their environment.

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ACKNOWLEDGEMENTS The author would like to thank Prof. N. H. Mann, Dr. L. Moore, Dr. B. Palenik and Prof. A. F. Post for communicating data prior to publication, and to N.H.M, A.F.P and Dr. F. Partensky for critical reading of the manuscript. The genome sequence data of Synechococcus WH 8102 are accessible via the DOE Joint Genome Institute at http://www.jgi.doe.gov/ JGI_microbial/html/index.html (data presented in this review exploiting this genomic information has been provided freely by the US DOE Joint Genome Institute for use in this publication only). Financial support provided by the Royal Society, NERC (NERC GR3/11606) and the European Union (PICODIV EVK2-1999-00064P, MARGENES QLRT2001-01226) is acknowledged for work in Warwick on the marine Synechococcus genus. D.J.S. is a Royal Society University Research Fellow.

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Adoption of the Transiently Non-culturable State – a Bacterial Survival Strategy? Galina V. Mukamolova1,2, Arseny S. Kaprelyants2, Douglas B. Kell1,3 and Michael Young1 1

Institute of Biological Sciences, University of Wales, Aberystwyth, Ceredigion SY23 3DD, UK 2 Bakh Institute of Biochemistry, Russian Academy of Sciences, Leninsky pr.33, 117071 Moscow, Russia 3 Present address: Department of Chemistry, UMIST, P.O. Box 88, Manchester M60 1QD, UK

ABSTRACT Microbial culturability can be ephemeral. Cells are not merely either dead or alive but can adopt physiological states in which they appear to be (transiently) non-culturable under conditions in which they are known normally to be able to grow and divide. The reacquisition of culturability from such states is referred to as resuscitation. We here develop the idea that this ‘‘transient non-culturability’’ is a consequence of a special survival strategy, and summarise the morphological, physiological and genetic evidence underpinning such behaviour and its adaptive significance.

Abbreviations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 2. Bacterial stress avoidance strategies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 2.1. Starvation survival . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 2.2. Formation of ‘‘non-culturable’’ cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 3. Are there specific chemical inducers of ‘‘non-culturability’’? . . . . . . . . . . . . . . . . . 92 4. Is ‘‘non-culturability’’ genetically controlled?. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94

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5. Resuscitation of ‘‘non-culturable’’ cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96 6. Social behaviour of bacterial populations and ‘‘non-culturability’’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 7. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 106 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109

ABBREVIATIONS BCG CFU CTC DVC GFP MPN NIC Rpf RT-PCR TNC VBNC

Bacille Calmette-Guerin Colony-forming units 5-Cyano-2,3-ditolyl tetrazolium chloride Direct viable count Green fluorescent protein Most probable numbers Not immediately culturable Resuscitation-promoting factor Reverse transcriptase polymerase chain reaction Transient non-culturability/Transiently non-culturable Viable but non-culturable

1. INTRODUCTION Bacteria in natural environments are exposed to a wide variety of stressful conditions such as nutrient deprivation, extremes of temperature or pH, changes in oxygen tension, the presence of toxic compounds, light, etc. (Edwards, 2000). Various stress avoidance strategies have evolved that enable bacteria to cope with fluctuations in natural environments (Kaprelyants et al., 1993; Neidhardt, 2002). Depending on the magnitude and combination of stresses experienced, different global stress responses ensure that cell integrity and the processes essential for viability (the ability to multiply) are maintained (Morita, 1990; Storz and Hengge-Aronis, 2000). Bacteria that live in fluctuating environments are endowed with an array of two-component systems (Hoch and Silhavy, 1995; Hoch, 2000) that detect environmental changes and initiate the appropriate responses to such changes. Bacterial two-component systems are made up of a sensor histidine

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kinase and a response regulator. The sensor kinase is activated by conformational change (that may result from ligand-binding), which leads to autophosphorylation (the g-phosphoryl group of ATP is transferred to a conserved histidine residue). The response regulator then catalyses the transfer of the phosphoryl group from the phosphoryl-histidine residue to a conserved aspartate within its own regulatory domain. Phosphorylated response regulators usually bind to specific DNA sequences and activate the expression of particular genes. Sensors have been described that detect changes associated with several stress responses such as changes in oxygen tension, redox potential and osmotic potential, and altered concentrations of toxic heavy metals and volatile fatty acids. Other sensors detect the presence or absence of specific nutrients (e.g. sources of phosphate, potassium, magnesium, carbon or nitrogen). Genome sequencing projects have uncovered many more two-component systems that respond to as yet unknown signals (e.g. Streptomyces coelicolor contains some 85 sensor kinases and 79 response regulators, including 53 kinase-regulator pairs (Bentley et al., 2002)). There are many variations on this basic theme, enabling different vital cellular processes to respond in an appropriate manner to a wide spectrum of environmental changes (Stock et al., 2000). Phosphorelays represent a further refinement, in which additional proteins are recruited to form phosphorylation cascades sensitive to multiple sensory inputs. Both chemotaxis and spore formation, which may be regarded as stress avoidance mechanisms, are under the control of dedicated phosphorelays (Falke et al., 1997; Hoch, 2000, 2002). Once a stress has been sensed, immediate action may be taken. Arguably, the simplest of these is the negative chemotactic response that enables bacteria to move away from localised regions in their immediate environment where conditions are unfavourable for growth and/or survival. However, flight is not always an option. Most stress avoidance mechanisms require well-orchestrated processes of up- and down-regulation of specific genes (Hengge-Aronis, 1999; Storz and Hengge-Aronis, 2000). Some of these genes feature in the responses to many different stresses (e.g. dnaK and groESL, which encode chaperonins involved in protein refolding). These processes consume energy and, depending on the severity of the stress, accumulated resources may have to be mobilised. As well as synthesis of chaperonins, responses to stressful conditions may involve the production of specific enzymes or specialised protective compounds. Examples of the former include catalase and superoxide dismutase, which are up-regulated in response to oxidative stress (Hengge-Aronis, 1993). Examples of the latter include low molecular weight osmo-protectants such as trehalose and betaine (Hengge-Aronis, 1993; Strom and Kaasen, 1993). Universal stress

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survival proteins play a significant role in the stress response and further survival of cells (Nystrom and Neidhardt, 1994). Synthesis of specialised DNA-binding (Dsp) proteins represents another important protective mechanism (Almiron et al., 1992; Gupta et al., 2002). These proteins co-crystallise with DNA in a metal ion-dependent process and protect it from degradation (Frenkiel-Krispin et al., 2001). After these relatively rapid responses have occurred, several different longer-term strategies may come into play, a common feature of which is a lowering of metabolic activity. If this lowering of metabolic activity is not very marked, nor very protracted, cells can emerge to re-initiate growth as soon as conditions become favourable again. However, if there is essentially metabolic shut-down, dormancy may ensue (Kaprelyants et al., 1993; Kell et al., 1998), with the production of highly specialised forms. Some bacteria produce spores in which metabolism is undetectable (Gould and Hurst, 1969). Others produce less specialised forms in which metabolism is very low (e.g. Kalakoutskii and Pouzharitskaja, 1973; Kaprelyants and Kell, 1993). In contrast to the first type of strategy, cells can only emerge from a dormant state after a protracted recovery period (in the case of spores, this corresponds to the period required for germination and outgrowth). In non-sporulating bacteria, metabolic shutdown is often accompanied by a loss of culturability (under conditions that support the growth of normal, viable cells). The process whereby culturability is restored is called resuscitation (Kaprelyants et al., 1993; Kell et al., 2003). Our present knowledge about dormancy in nonsporulating bacteria is very limited, although it is clear that ‘‘nonculturability’’ is only one of many specific properties of cells in this state. In this review we shall use the term ‘‘transiently non-culturable’’ (TNC) to describe this particular state, noting also that the definition of states such as this is purely operational (Kell et al., 1998). If the stress is extreme, or if the available energy supply is insufficient to permit avoidance, the cell will die. The death of some cells may permit the survival of others in the bacterial population, so death should not necessarily be viewed as a passive reaction to lethal stress (Postgate, 1976). Indeed, processes analogous to apoptosis (programmed cell death) in eukaryotic cells may also occur in prokaryotic cells. For example, it has been suggested that the death of the mother cell during endospore formation in Bacillus and the sacrifice of many cells during fruiting body formation in Myxococcus might be considered as examples of programmed cell death (Lewis, 2000). The plasmid-encoded toxin–anti-toxin systems found on bacterial plasmids provide another example (Yarmolinsky, 1995). Regulation of the expression of the genes encoding the toxin and the

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anti-toxin, and hence the ratio of these proteins in individual cells, will control the proportion of live cells in a population of bacteria harbouring the plasmid. Under certain conditions, cell death can be regarded as a complex multi-step mechanism aimed at sacrificing some bacteria in the population so as to release nutrients permitting the survival of others (Finkel and Kolter, 2001). This review will focus on the long-term strategies that bacteria employ to avoid stress, with special emphasis on those that involve the formation of (transiently) ‘‘non-culturable’’ cells. 2. BACTERIAL STRESS AVOIDANCE STRATEGIES 2.1. Starvation Survival 2.1.1. The Longevity of Bacteria The question of how long an individual organism can survive (according to any sensible definition of that verb, which in microbes implies the ability to replicate upon transfer to favourable conditions) is a very vexing one. It is of course of great importance to our Culture Collections, as this determines how often strains must be sub-cultured, the kinetics of die-off, the benefits of low water activity, of low oxygen tension and of suitable additives in maintaining culturability, and so on, but this is not the place to discuss these matters. However, it is appropriate to discuss briefly some of the issues. Given the comparative recency of microbiology as a discipline – let us say a century or two – it is obvious that no laboratory study can shed serious light on the question of whether microorganisms can in fact survive (to produce daughter cells) for centuries, millennia or even beyond. The current record claim (Vreeland et al., 2000) (see that article for references to earlier work) points at 250 million years for an organism isolated from a salt crystal. These authors also sequenced the rDNA of the organism with a view of establishing its lineage, which data in fact cast substantial doubt on such longevity (Nickle et al., 2002). Of course, the main issue in such studies (as in the isolation of other microbes from complex presentday samples (Schut et al., 1993) is to exclude contamination with modern organisms. This is extremely difficult (Keilin, 1959) (see other articles in the anthology edited by Crowe and Clegg (1973)), and the true (maximum) longevity of bacteria remains unknown. However, if we assume exponential die-off kinetics, a starting population of interest of 1010 cells, and a half-life of even 10 years, we lose 90 percent of

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the cells every 33 years; our 1010 cells have (obviously) then lost culturability in a rather modest 330 years. Even with this hefty inoculum of cells and a highly optimistic half-life of 100 years – well beyond anything ever measured in the laboratory – there are essentially no cells left after ca. 3000 years. The kinetics are simply against one here. The argument that extreme conditions in vitro may be tolerable under the unknown – and more organised – conditions in vivo certainly has merit, and the generally accepted record for thermophilic growth in the laboratory (at temperatures well above 100 C (Huber et al., 2000)) challenges the textbook notion that such temperatures would be expected to cause the hydrolysis of nucleic acids at an alarming rate, inconsistent with the doubling time of such organisms. Such arguments have indeed equivalently been made in the context of claims of a much higher temperature for microbial growth near ‘black smokers’ (Baross and Deming, 1983; Trent et al., 1984). Actually, a subtler issue of longevity as it pertains to individuals in a particular population hinges upon whether the causes of ‘death’ are entirely endogenous (e.g. thermally induced protein denaturation or hydrolysis) or have a more or less controlling exogenous cause (cosmic rays, reaction of the cells with oxygen radicals, etc.). In reality it must be the latter, since this determines the effective size of the inoculum considered as the starting population for the purposes of the kinetic argument. Our present view is that the likely reasonable survival half-life under optimal conditions (in the absence of nutrient influx, turnover and regeneration) of a microbe is between 101 and 102 years, but not greater. 2.1.2. Experimental Starvation Survival Leaving aside these considerations based on a quasi-geological timescale, it is nevertheless recognised from experiment that microorganisms can survive starvation for protracted periods (Kjelleberg et al., 1993), possibly with half-lives exceeding 30 years (Eisenstark et al., 1992). This specific strategy, called starvation survival (Morita, 1990, 1997), has been documented in many different bacteria including Escherichia coli (Kolter et al., 1993), Salmonella typhimurium (Spector, 1998), Vibrio spp. (Ostling et al., 1997), Listeria monocytogenes (Herbert and Foster, 2001), Staphylococcus aureus (Watson et al., 1998a) and Mycobacterium tuberculosis (Betts et al., 2002). Starvation survival shows several important physiological features that are equally applicable to survival of other kinds of stress:  Shut-down of metabolic processes related to cell multiplication and growth;

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 Use of specialised compounds and structures to enable cells to maintain their integrity and protect important cell components;  Maintenance of culturability (ability to grow in liquid and on solid media) so that cells can quickly recover and resume active growth as soon as nutrients become available or the stress is relieved. The transition from active metabolism (characteristic of dividing cells) to maintenance metabolism is an important feature of starvation survival. Valuable resources are conserved by the cessation of growth, but organisms retain a relatively high level of metabolic activity to support core metabolism and specific repair processes, commensurate with maintenance of cell integrity and culturability (Morita, 1990). When initially discovered, the stringent response was thought to be a specific attribute of the protein synthetic machinery, allowing the shutdown of protein and RNA synthesis when amino acids are not freely available (Gallant and Cashel, 1967; Cashel, 1975). The stringent response is activated when ppGpp is synthesised as a result of the binding of uncharged tRNA molecules to the ribosome. A ppGpp-independent mechanism that reduces the level of translation during amino acid starvation has also been reported (Christensen et al., 2001) and it is now accepted that ppGpp is a general stress alarmone, encompassing many diverse physiological processes. For example, ppGpp is produced when fatty acid synthesis is disrupted (Gong et al., 2002) and it controls the complex system of ‘‘competition’’ between different sigma factors, depending on the environment (Jishage et al., 2002). It also regulates the virulence of some bacteria (Bachman and Swanson, 2001) and is required for long-term survival of pathogenic mycobacteria under starvation conditions (Primm et al., 2000). The stress alarmone, ppGpp, also regulates the production of colicins under conditions of nutrient deprivation (Kuhar et al., 2001), which leads to death and lysis of some cells in the bacterial population, supplying others with nutrients. Another regulatory mechanism characteristic of starvation survival is a general increase in the half-life of mRNA (Takayama and Kjelleberg, 2000). Moreover, during extended starvation some transcripts remain stable but silent; they are immediately available for translation when a suitable substrate for growth becomes available (Marouga and Kjelleberg, 1996). Despite the fact that starvation survival consumes relatively little energy (as compared with the energy demands for growth and multiplication), prolonged starvation under these ‘‘low maintenance’’ conditions is or can be accompanied by the occurrence of irreversible changes to some cells, including some damage to important cellular components such as DNA,

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RNA, proteins and lipids (Hood et al., 1986). This may be the reason why the survival of some cells in a starving population is often accompanied by the death of others (so-called ‘cryptic growth’ (Postgate, 1967, 1976)). Nutrients released from dead bacteria can be used by other cells, permitting their survival. In a glycerol-limited culture of Klebsiella aerogenes, death by starvation of 50 individuals was necessary to permit the multiplication of one survivor in buffer lacking nutrients (Postgate and Hunter, 1962). In this study, it was shown that the ratio of multiplying/surviving/dying organisms depended on the particular conditions of starvation to which they were subjected. This population behaviour underlies the phenomenon of cryptic growth and may be considered as a kind of cannibalism (Postgate, 1976). Starvation survival must therefore be regarded as a temporary measure, at least under these conditions; owing to the gradual accumulation of damage, cells cannot remain in this state for protracted periods. When in suspension (as opposed to being dehydrated), they must either exit to resume active growth and multiplication, or they will die (i.e. lose culturability in an irreversible sense). Many laboratory models of nutrient starvation are based on the sudden transfer of copiotrophic organisms from a nutrient-rich environment, to one that is nutrient-poor. The organisms respond to this sudden stress using previously accumulated resources such as stored sources of energy (Dawes and Senior, 1973; Dietzler et al., 1979) and phosphate (Rao and Kornberg, 1999; Kim et al., 2002). In natural environments, most cell populations exist under the pressure of a ‘‘f(e)ast and famine’’ existence (Koch, 1971; Poindexter, 1981), with continuous exposure to environmental fluctuations and, frequently, multiple stresses. Cells in nature have adapted to a ‘‘low metabolic cost’’ existence, either genetically in the case of oligotrophs (Schut et al., 1993; Morita, 1997; Schut et al., 1997a,b), or physiologically in the case of copiotrophs. Both in the laboratory and in nature, organisms are preoccupied with accumulating resources for their next multiplication; this is generally more difficult in nature than in the laboratory. Moreover, under laboratory conditions the population density is usually very high, supporting cryptic growth or cannibalism, whereas when existing in a planktonic form in nature populations are either widely dispersed or, if localised, the effective population size may be comparatively small. In nature, there is therefore strong selective pressure for cells that have developed mechanisms permitting a low metabolic cost existence. Adoption of the TNC state in natural environments is easily understood within the conceptual framework of a low metabolic cost existence and may in fact be a necessary prerequisite for the successful colonisation of many natural habitats.

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Bacterial endospores represent an extreme adaptation to a low metabolic cost existence, i.e. profound dormancy (Gould and Hurst, 1969). The low metabolic cost existence adopted by non-sporulating organisms in natural environments may be viewed as a state akin to dormancy, which has been defined as ‘‘a reversible state of low metabolic activity, in which cells can persist for extended periods without division’’ (Kaprelyants et al., 1993). Both sporulation and dormancy are reversible states of metabolic shutdown, accompanied by loss of immediate culturability. We shall now focus on the TNC state.

2.2. Formation of ‘‘Non-culturable’’ Cells It is well known that the number of cells which can be observed microscopically in natural environments often exceeds by orders of magnitude the number which can be recovered and cultured (e.g. Amann et al., 1995; Torsvik et al., 1996; Head et al., 1998; Torsvik et al., 1998; Barer and Harwood, 1999; Rondon et al., 2000; Curtis et al., 2002; Kell et al., 2003). It is equally well documented that bacteria can enter states in which they have lost culturability but still retain measurable metabolic activity of some kind. These have been referred to as so-called ‘‘viable but non-culturable’’ (VBNC) forms (Roszak and Colwell, 1987). The terminology used to describe these forms is both confused and confusing; this, together with the heterogeneity of the cultures and the need for improved experimental designs – including in particular the use of ‘dilution to extinction’ methods – has been stressed before (Kell et al., 1998; Barer and Harwood, 1999). We shall adopt the terminology proposed by these authors, using the term ‘‘non-culturable’’ in an operational sense. A more precise term would be ‘‘not immediately culturable’’ (NIC) (Kell et al., 1998), but since the former is still in common use, we retain it here. The class of ‘‘non-culturable’’ bacteria is by now demonstrably a ‘‘rag-bag’’ of organisms displaying different physiological characteristics, which are discussed in turn in the following paragraphs. Cell integrity is the most obvious characteristic. If the cell envelope or any other major cell components are irreparably damaged, then despite their apparent integrity, such injured cells are manifestly not viable and therefore they cannot be cultured. Cell integrity can be investigated using a variety of conventional techniques such as electron microscopy or by staining with propidium iodide (Davey and Kell, 1996) and there are even so-called ‘‘live/dead’’ kits available for this specific purpose. Luna et al. (2002) reported that only 30% of apparently intact cells, isolated from

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sediment, had an intact permeability barrier. ‘‘Non-culturable’’ forms usually have an altered cell morphology, a reduced size, a coccoid shape and a thickened cell wall (Oliver et al., 1991; Turpin et al., 1993; Mukamolova et al., 1995c; Shleeva et al., 2002; Signoretto et al., 2000, 2002). The size reduction plausibly confers a reduced maintenance energy requirement. These morphological changes, and the biochemical changes that underlie them, contribute to the increased resistance of ‘‘nonculturable’’ cells to different kinds of stress. ‘‘VBNC’’ cells of Pseudomonas fluorescens were more resistant to heating than were actively growing cells (Evdokimova et al., 1994; van Overbeek et al., 1995), and ‘‘VBNC’’ cells of Vibrio vulnificus were less sensitive to sonication than actively growing cells (Weichart and Kjelleberg, 1996). However, sometimes the opposite (i.e. increased sensitivity to a challenge) has been observed (Biketov et al., 2000; Johnston and Brown, 2002). Moreover, it is not clear if ‘‘VBNC’’ cells really are more resistant to stress factors in natural environments, such as soil (Mascher et al., 2000). Another important criterion of the TNC state is that these cells are in an altered metabolic state with reduced respiratory activity. There are many controversial indications on this matter (Gribbon and Barer, 1995). It has been shown that ‘‘non-culturable’’ cells of Micrococcus luteus have a level of respiratory activity that is below the limit of detection using CTC (Kaprelyants and Kell, 1993). In the case of Pasteurella piscicida (Magarinos et al., 1994) and in Yersinia ruckeri (Romalde et al., 1994), the respiratory activity of ‘‘VBNC’’ cells was reduced by more than 80% compared with that associated with growing cells. However, in most reports concerning the (or a) ‘‘VBNC’’ state, detectable respiratory activity was used as a or even the criterion of ‘‘viability’’ (see, for example, Oliver and Wanucha, 1989; Garcia-Lara et al., 1993; Morgan et al., 1993; Boucher et al., 1994; Ramaiah et al., 2002). Since the products of CTC and tetrazolium reduction accumulate with time, these compounds are very sensitive indicators of respiratory activity. On the other hand, some cells referred to by the authors describing them as ‘‘VBNC’’ have metabolic activity similar to that of culturable bacteria and, in our opinion, such cells cannot be considered as being in a TNC state. Measurement of membrane energisation (often referred to as ‘‘the membrane potential’’) is another way to assess metabolic activity (Kaprelyants and Kell, 1992; Shapiro, 1995a; Davey and Kell, 1996; Davey et al., 1999; Shapiro, 2000). Sometimes, cells stained with CTC have a significantly reduced level of membrane energisation and adenylate energy charge (Tholozan et al., 1999). The potential to (re)activate metabolism has often been considered as one of the major criteria for viability, especially when assessing microbes

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in natural environments in situ where culturability is hard to observe. To this end, the direct viable count (DVC) method developed by Kogure and collaborators has been widely used to distinguish between viable and non-viable (dead) cells (Kogure et al., 1979). After incubation in medium with a low concentration of yeast extract in the presence of nalidixic acid (to prevent multiplication), cells are stained with acridine orange. Nonculturable cells activated by yeast extract are stained orange (because of RNA accumulation) and they increase their size, sometimes producing filaments. Dead cells do not respond. It is worth mentioning that this technique does not simply show the metabolic activity of ‘‘nonculturable’’ organisms. It demonstrates their ‘‘resuscitation’’ potential, i.e. their ability to restore metabolism, and recommence growth when stimulated by the addition of nutrients. Because of the presence of nalidixic acid it cannot of course tell us whether such cells would have multiplied in its absence. The DVC method, developed originally for use with Gram-negative bacteria, has been adapted for use with Grampositive bacteria. For example, ampicillin was used (instead of nalidixic acid) for M. luteus (Kaprelyants et al., 1994) and ciprofloxacin was used for L. monocytogenes (Besnard et al., 2000). The use of reporter gene fusions has also been proposed for estimating the ‘‘viability’’ of some ‘‘non-culturable’’ microorganisms (Bloemberg et al., 1997; Cho and Kim, 1999a,b). As shown by Lowder and colleagues, GFP was quite stable in starving cells of P. fluorescens and its fluorescence could be related to the number of so-called ‘‘VBNC’’ cells (Lowder et al., 2000). A modified, less stable version of GFP has also been used and a good correlation between metabolic activity and GFP fluorescence was noted (Lowder and Oliver, 2001). PCR has also been employed to detect ‘‘non-culturable’’ cells of many organisms (Domingue and Woody, 1997), including e.g., M. tuberculosis in vivo (de Wit et al., 1995; Herna´ndez-Pando et al., 2000), S. typhimurium under laboratory conditions (Romanova et al., 1996) and Campylobacter spp. in water samples (Moore et al., 2001). DNA stability is evidently one of the key criteria for viability. However, PCR products can be obtained from partially degraded DNA released from dead cells (Weichart et al., 1997) and this method must therefore be employed with great care. Since prokaryotic mRNA generally has a very short half-life (and certainly compared with that of DNA), RT-PCR represents a less equivocal method. It has been employed to detect ‘‘non-culturable’’ cells of E. faecalis (del Mar Lleo et al., 2000), Helicobacter pylori (Nilsson et al., 2002), M. tuberculosis (Herna´ndez-Pando et al., 2000; Pai et al., 2000) and V. vulnificus (Fischer-Le Saux et al., 2002).

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As is implied by their name, ‘‘non-culturable’’ organisms cannot be cultivated using conventional media and techniques. They should not be grouped together with injured organisms, which can grow under these conditions, but not in the presence of selective agents, such as detergents (Ray and Speck, 1973). An extremely important feature of (transiently) ‘‘non-culturable’’ cells, which serves to distinguish them from dead cells, is reversibility, i.e. their ability to convert to normally dividing cells under conditions permitting growth. This process, called ‘‘resuscitation’’ is discussed later. Finally, (transient) ‘‘non-culturability’’ as a survival strategy cannot simply be inferred from an inability to produce colonies or cultures. A convincing case can be made only when organisms are shown to conform to the various criteria mentioned above (i.e. cell integrity, metabolic shutdown, and reversibility/ability to resuscitate). ‘‘Non-culturable’’ bacteria can be found almost everywhere in nature (soil, air, sea, estuarine waters, etc.), in other living organisms and in foods. They can also be obtained in the laboratory. It has been suggested that the ‘‘non-culturable’’ forms of pathogenic bacteria that are found in natural environments are perfectly able to cause disease when re-introduced into their host (Oliver and Bockian, 1995; Colwell et al., 1996; Rahman et al., 1996; Chaiyanan et al., 2001) and this issue can obviously give cause for serious concern (Kogure and Ikemoto, 1997; Islam et al., 2001). However, all of these experiments have been done under conditions in which the ostensibly ‘‘non-culturable’’ organisms have in fact been a mixture of culturable and non-culturable cells. Only when the experiment is done under what amounts to MPN conditions of dilution to extinction (i.e. one pathogenic cell as the inoculum into the host) can this conclusion be considered unequivocal (Kell et al., 1998; Barer and Harwood, 1999). On the one occasion to date in which such an important experiment has been performed (Smith et al., 2002), the conclusion was that such ‘‘unculturable’’ cells did not retain pathogenicity. We now consider what may be learned about the TNC state from analyses of ‘‘non-culturable’’ forms in these different environments.

2.2.1. ‘‘Non-culturable’’ Cells in Natural Environments The vast majority of the microbial biota in natural environments has never been brought into culture, as revealed initially by the renaturation kinetics of DNA isolated from such environments (Torsvik et al., 1990) and subsequently by sequencing of PCR products obtained using primers corresponding to their 16S rRNA (Torsvik et al., 1990; Ward et al., 1990).

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For example, it has been estimated that the diversity of the total community in a particular soil was at least 200 times higher than the diversity of bacterial isolates cultured from the same soil (Torsvik et al., 1998). Many of the dominant organisms found in such studies belong to new species and even genera (Kalmbach et al., 1997). The reasons for this discrepancy are being widely discussed (Ward et al., 1990, 1998; Amann et al., 1995; Gobel, 1995; Kell et al., 2003). It has been suggested that appropriate cultivation conditions have not been found for these organisms. Very recently, significant strides have been made in several laboratories, using co-culture (Kaeberlein et al., 2002), gel microdroplets (Zengler et al., 2002), diluted media (Janssen et al., 2002) and polymeric growth substrates (Sait et al., 2002) to coax previously uncultured organisms into laboratory cultivation. In our view, the main difficulty is to understand whether (i) the ‘‘missing’’ organisms are fastidious, with highly specialised growth requirements, or (ii) they are simply surviving in a ‘‘non-culturable’’ state, occasionally ‘‘breaking’’ it to produce progeny when environmental conditions permit. Of course, both hypotheses are likely to contribute to the truth in different circumstances. The first hypothesis is supported by the existence of oligotrophs, whose cell form, size, metabolism and life cycle are adapted to extremely nutrientpoor conditions (Hirsch, 1986). They have specialised, high-affinity uptake systems for scavenging nutrients from environments in which they are extremely scarce, enzymes with extremely high affinity for their substrates, and a minimal complement of rrn operons and ribosomes (Fegatella et al., 1998). Oligotrophs are widespread and abundant in natural environments and can be found in water sources, soil and even within other living organisms (Wainwright et al., 1991; Tada et al., 1995). Because of their sheer abundance and their metabolic capabilities, they play a cardinal role in global recycling of nitrogen, sulphur and other inorganic materials (Wainwright et al., 1991). It has been suggested that some of the so-called ‘ultramicrobacteria’ (i.e. tiny bacteria that pass through 0.45 mm filter) may also be oligotrophic organisms (Anderson and Heffernan, 1965; Torrella and Morita, 1981). However, more recent evidence (Schut et al., 1993; Morita, 1997; Schut et al., 1997a,b; Fegatella and Cavicchioli, 2000) indicates that at least some such ‘ultramicrobacteria’ are not ‘small forms of normal (i.e. copiotrophic) bacteria’ but ‘normal forms of small bacteria’ (Kaprelyants et al., 1993). A recent investigation of one ultramicrobacterium showed that it had a very diverse protein complement, despite having a small-sized genome (only 1.5 Mb), suggesting that gene expression in these organisms may involve substantial co- and post-translational modifications of proteins (Fegatella and Cavicchioli, 2000). But in many cases,

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it is still not clear whether filterable cells are specialised forms of known bacteria, or a degenerate stage in their life cycle, or whether they represent a group of microorganisms that has not previously been described (Zweifel and Hangstrom, 1995; Haller et al., 2000). Finally, some copiotrophic organisms, such as Vibrio spp., can produce filterable cells under starvation conditions (Novitsky and Morita, 1976). However, there is also substantial support for the second hypothesis. There are many examples in which ‘‘non-culturable’’ forms of known copiotrophic bacteria have been found in natural environments (Magarinos et al., 1994; Kalmbach et al., 1997; Kozdroj and van Elsas, 2001). These cells could not be cultivated but showed reactivation in the presence of nalidixic acid and yeast extract (by DVC). Non-culturable cells of Yersinia pseudotuberculosis have also been detected in soil by PCR (Troitskaia et al., 1996). The mechanism of production of ‘‘non-culturable’’ cells in nature, and the factors influencing it, are still unknown. Like sporulation, the formation of ‘‘non-culturable’’ cells can be induced by various stress factors (pH, toxic compounds, lack of oxygen and nutrient limitation). Sometimes, a combination of different factors unique for a particular ecological niche determines the strategy of bacterial survival. For example, P. fluorescens cells produced ‘‘VBNC’’ forms in some soils, but not in others (van Overbeek et al., 1995). It is also possible that there is a seasonal dimension to the cycle of culturability/non-culturability. For example, it has been claimed that Vibrio cells produce only ‘‘non-culturable’’ cells when the water temperature in lakes and rivers drops below 10 C (see Oliver et al., 1995; Weichart and Kjelleberg, 1996). Complicated interactions between different species, including host–parasite interactions (Romalde et al., 1994), symbiotic associations (Lee and Ruby, 1995; Islam et al., 1999) or co-existence (Didenko et al., 2002), have great impact on bacterial survival strategies. Human activity can also influence significantly the diversity and physiological state of environmental bacteria. The introduction of heavy metals, disinfectants and other polluting compounds into natural environments inevitably influences bacterial culturability (Mascher et al., 2000; Pitonzo et al., 1999). In view of the uncertainty of the state of such complex environments, it is not possible to take into account all conceivable factors that may influence cell culturability in Nature. Therefore, defined laboratory models have been developed to simplify the problem and permit analysis of the relevance of different possible factors and mechanisms leading to ‘‘non-culturability’’. In the laboratory, pure bacterial cultures (or simple mixtures) are usually studied (although we note encouraging ‘‘largescale’’ developments in post-genomics experiments (Giaever et al., 1999)).

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This imposes a serious limitation, since pure cultures of individual microorganisms are seldom if ever found in nature. On the other hand it provides a distinct advantage, since individual organisms can be properly characterised physiologically.

2.2.2. ‘‘Non-culturable’’ Cells in Aqueous Laboratory Microcosms Microcosms have been widely used for studying environmental bacteria under laboratory conditions. In some cases, growth media have been formulated using a defined mixture of components to mimic as closely as possible the natural environment (artificial microcosm). In others, water is simply taken from natural environments (and in the better experiments is filter-sterilised). Autoclaving of the natural substrates, which may result in the release of readily available nutrients, often prevents the formation of ‘‘VBNC’’ cells (Garcia-Lara et al., 1993). A similar effect has also been observed in relation to sporulation of streptomycetes in soil (Mayfield et al., 1972). A very important feature of these microcosm experiments is that a low inoculum of organisms should be employed, effectively excluding cryptic growth. Many different conditions have been used to produce ‘‘non-culturable’’ organisms. In many cases, starvation per se in a microcosm does not cause loss of culturability on a timescale of weeks. For example, in V. vulnificus, incubation temperature is the most important factor. Starving cells do not lose culturability at 24 C but similar cells incubated in the same buffer at 4 C lose viability rapidly, and produce an ostensibly homogeneous population of ‘‘VBNC’’ cells in 3 weeks (Weichart et al., 1992). Similar results have been reported for many organisms, including Vibrio anguillarum (Eguchi et al., 2000), E. faecalis (Lleo et al., 2001), Aeromonas hydrophila (Wai et al., 2000; Mary et al., 2002), Salmonella enteritidis (Chmielewski and Frank, 1995), H. pylori (Shahamat et al., 1993) and Campylobacter spp. (Thomas et al., 2002). Remarkably, in the case of V. vulnificus, temperature but not starvation was the factor that ‘‘triggered’’ the transition to non-culturability (Oliver and Wanucha, 1989). E. coli cells incubated in a microcosm exposed to visible light also produced ‘‘VBNC’’ forms (Arana et al., 1992). A combination of desiccating conditions and starvation induced the formation of ‘‘VBNC’’ forms in S. typhimurium (Lesn et al., 2000). In some cases, mild stress did not cause a transition to the ‘‘non-culturable’’ state, whereas a stronger stress did. For example, ‘‘non-culturable’’ cells of P. fluorescens were produced when they were either exposed to a combination of low redox potential and oxygen limitation or a high

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level (1.5 M but not 0.7 M) of NaCl (Mascher et al., 2000). Addition of CuSO4 to starving bacteria was also claimed to have induced a ‘‘VBNC’’ state (Ghezzi and Steck, 1999; Grey and Steck, 2001a), but (given the wellknown bactericidal effects of heavy metals) it is more likely that the cells were simply killed, as has been found in other cases where mixtures of cells in different physiological states are studied indiscriminately (Bogosian et al., 1998, 2000; Kell et al., 1998; Nystrom, 2001). We remain puzzled by the apparent unwillingness of experimenters to perform these studies of culturability under conditions of dilution to extinction. As will be clear from the foregoing, it is not possible to define a universal procedure for induction of the ‘‘non-culturable’’ state. The precise conditions required for its establishment vary from one organism to another, presumably reflecting their different genetic, biochemical and physiological characteristics. Generally speaking, bacteria seem to lose culturability under conditions when they cannot initiate a starvation survival program. This may occur if a severe stress is encountered, or if they are simultaneously subjected to a combination of several different stresses, or even as a result of prolonged exposure to conditions that are not ideal for bacterial survival. Moreover, if the starvation survival program has already been initiated (for example, by pre-starvation at 24 C before ‘‘cold induction’’, in the case of V. vulnificus), cells do not then lose culturability (Weichart et al., 1992). We still do not know if the examples described above represent specialised forms or simply dying cells. Exposure to a sudden stress without pre-adaptation, poor recoverability and, often, a lack of metabolic reorganisation, may suggest that ‘‘VBNC’’ (or dead) forms frequently arise as a result of a process of gradual cellular degradation.

2.2.3. Stationary Phase Culture as a Model for Induction of ‘‘Non-culturability’’ Compared with the microcosms discussed above, the stationary phase of batch culture in the laboratory is a very different model for studying the physiological processes that induce ‘‘non-culturability’’. It allows bacteria to be studied under complex conditions in a multicellular community, in which cell–cell interactions occur. The stationary phase in laboratory batch culture does not have a real counterpart in natural environments because in the laboratory, homogeneous populations of cells of a single species reach a comparatively high population density (often under conditions of nutrient excess). Many environmental factors (oxidative stress, low or high pH, accumulation of toxic compounds, lack of nutrients) are changing as

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bacteria enter stationary phase, to which they are forced to adapt simultaneously. Most of these stresses arise as an indirect result of the metabolic activity of the cells themselves. The ‘‘artificial’’ stationary phase laboratory model makes it possible to explore the complete adaptive potential of different organisms. Entry into stationary phase is therefore a gradual process of adaptation to an increasingly hostile environment. The so called ‘‘transition state’’ from exponential to stationary phase has been extensively characterised in model organisms, such as Bacillus subtilis and E. coli (Kolter et al., 1993; Siegele and Kolter, 1993; Strauch and Hoch, 1993), from which it is clear that it represents a highly regulated, genetically programmed, developmental process during which the organisms change from one physiological state to another. By definition, bacteria cease to multiply when they enter stationary phase; however, as we shall see, the reason for this is not always obvious. In some cases, growth arrest is an irreversible process and cells cannot divide normally when re-inoculated into fresh medium (e.g. Weichart and Kjelleberg, 1996; Bogosian et al., 1998; Kell et al., 1998; McDougald et al., 1998; Bogosian et al., 2000) and this too remains largely enigmatic. We shall restrict our further consideration of stationary phase to this phenomenon of (ostensible) ‘‘non-culturability’’. As is our custom (Kaprelyants et al., 1993; Kell et al., 1998; Kell and Young, 2000), as well as the longstanding convention in microbiology (e.g. Postgate, 1969, 1976), we again equate the terms ‘viability’ and ‘culturability’ (the ability to multiply). We also recognise that these are not innate properties of the microbes themselves but a conflation of such properties (which are themselves inaccessible) with the experiment that is actually performed to assess them (Kell et al., 1998), a phenomenon equivalent to the ‘‘Schro¨dinger’s Cat’’ paradox in quantum mechanics (Primas, 1981). The phrase ‘‘viable but non-culturable’’ is thus an oxymoron (Barer et al., 1993; Barer and Harwood, 1999). During stationary phase, the proportion of surviving cells (which may or may not retain their culturability) and of lysing organisms, depends on the medium used and on the culture conditions. In E. coli, the richer the growth medium, the more rapidly the cells lose their ability to produce colonies on plates (Vulic and Kolter, 2002). Oxygen availability is also an important factor. For instance, a sigE sigS double mutant of S. typhimurium lost culturability (measured by cfu) very quickly when it was cultured with shaking, but not when it was cultured without shaking (Testerman et al., 2002). The simplest explanation for this behaviour is the damaging influence of oxygen in shaking culture, but it is also possible that cells need oxygen for initiation of the ‘‘death’’ program and in its absence,

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they use a different survival strategy. Unfortunately, in many cases it is difficult to understand whether cells are dying or whether they are surviving as ‘‘non-culturable’’ cells. One of the approaches frequently used is to study the stationary phase that is provoked by starvation for a known substance. In this type of model, cells are inoculated in medium containing a limiting amount of a particular nutrient (usually a carbon, phosphate or nitrogen source). Cells grow until they almost exhaust the limiting compound and they then enter stationary phase. Cells sense that the nutrient is about to become limiting before its complete exhaustion (Smeulders et al., 1999), so adaptation to starvation is a more or less gradual process. Depending on the nature of the limitation, cells can adopt different survival strategies, using different metabolic pathways. In a nitrate-limited culture of P. fluorescens, dormant (i.e. reversibly ‘‘non-culturable’’) cells were observed, whereas they did not arise when the organism was carbon- or phosphate-limited (Evdokimova et al., 1994). In the case of Mycobacterium smegmatis, significant differences in population behaviour under carbon-, phosphorus- or nitrogen-starvation were not observed (Smeulders et al., 1999). Unfortunately, these authors did not specifically study the ‘‘non-culturable’’ cells (which were the majority class in their mixed populations) and the reason for their lack of culturability was not established. The fact that bacteria are simultaneously exposed to multiple stresses and have to adapt to the gradual environmental changes that occur during stationary phase, may explain the dramatic difference observed in population behaviour in stationary phase as compared with that observed during starvation survival in buffer. For example, ‘‘non-culturable’’ coccoid forms were produced when H. pylori cells were exposed to starvation in either phosphate-buffered saline at 8  C, or under prolonged incubation in stationary phase (Nilsson et al., 2002). Moreover, in both cases, transcription of some genes (vacA, ureA, tsaA) could be detected by RT-PCR. But in contrast to cold-starved bacteria, a significant proportion of stationary phase cells revealed ‘‘degenerative’’ changes according to electron microscopy. Moreover, they also showed a more rapid decrease of their ATP pool. The authors concluded that cells subjected to cold starvation can survive in a ‘‘non-culturable’’ state for a longer period than those exposed to prolonged stationary phase (at least 3 months in the first case and up to 40 days in the second). Production of any such ‘‘non-culturable’’ cells in stationary phase is presently not known to occur readily in other species. For example, neither S. typhimurium (Turner et al., 2000) nor E. coli (Weichart and Kell, 2001) nor S. aureus (Watson et al., 1998b) produced ‘‘non-culturable’’ forms in prolonged stationary phase. Interestingly,

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Ericsson and colleagues (Ericsson et al., 2000) reported that fewer than 10% of the total number of cells in a stationary phase culture of E. coli maintained cell integrity. However, as was mentioned previously, both species can produce ‘‘non-culturable’’ forms under other conditions. A difference in adaptation strategy for cells incubated in stationary phase, as compared with cells suddenly transferred to phosphate buffer, has also been reported for M. tuberculosis (Betts et al., 2002). Under conditions of buffer starvation, activity of the respiratory chain was reduced dramatically over the first 96 hours, whereas it continued unchanged over the same period in stationary phase. These experiments showed that the ‘‘sudden’’ stress occasioned by transfer to phosphate buffer led to the adoption of a starvation survival strategy, in which cells did not lose culturability. In this altered metabolic state there was a slowdown of the transcription apparatus, energy metabolism, lipid biosynthesis and cell division, in addition to induction of the stringent response. These cells could survive in the apparent absence of major nutrients (i.e. they adopted a typical starvation survival strategy) and they were not sensitive to rifampicin. In contrast, in the so-called ‘‘dormancy’’ model of M. tuberculosis, developed by Wayne and defined by him as adoption of a state of ‘‘non-replicating persistence’’ (Wayne and Sohaskey, 2001), the state of the cell population is determined by oxygen unavailability (Wayne, 1976). Cells are cultivated in sealed tubes filled with medium (almost no head space). After gradual consumption of nearly all the available oxygen, cells transit to a synchronised non-dividing state, i.e. stationary phase (Wayne, 1977). They become resistant to rifampicin, but sensitive to metronidazole (Wayne and Sramek, 1994), indicating that they develop an ‘‘anaerobic’’ type of metabolism, using the glyoxylate pathway and lipids as source of carbon for energy production. Protein synthesis is also shut down in these cells (Hu et al., 1998). However, they do not lose culturability on plates and resume growth immediately after oxygen input. In contrast, cells ‘‘suddenly’’ depleted of oxygen do not develop this ‘‘anaerobic’’ persistence strategy, but simply die (Wayne and Diaz, 1967). This shows again that cells need time and adequate resources successfully to implement a survival strategy. M. tuberculosis populations incubated in stationary phase for prolonged periods of time can also enter a state of ‘‘non-culturability’’. After growth without apparent limitation of oxygen or nutrients, a significant proportion of cells lose the ability to form colonies on plates. Some of these cells may be dead/lysed; others can be recovered after incubation in liquid medium in the presence of supernatant obtained from early stationary phase cultures (Zhang et al., 2001; Shleeva et al., 2002). A diagrammatic representation of the culturability of these bacteria in different physiological states is shown in Figure 1.

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Figure 1 A model depicting the transition of actively growing, viable M. tuberculosis to dormant forms during prolonged stationary phase. The intensity of shading of the arrows reflects the intensity of metabolic activity; small arrows indicate reversibility of transitions. The term ‘non-culturable’ is operational (Barer et al., 1998) and refers, in this figure, to the inability of the bacteria to form colonies. Reproduced from Shleeva et al. (2002), with the kind permission of the publishers.

Most bacterial populations probably contain a mixture of culturable/nonculturable and viable/dead cells and the proportions of the different types will vary according to the culture conditions. This may depend on a single factor or on a combination of several different factors. For example, cultures of M. tuberculosis and Mycobacterium bovis (BCG) incubated in prolonged stationary phase (two months) in a sealed flask contained at least 99% of ‘‘non-culturable’’ cells, whereas similar cultures in ‘‘unsealed’’ flasks contained 99% of culturable cells. Variations in pH, medium composition, and airspace/liquid ratio again influenced the proportion of culturable versus ‘‘non-culturable’’ cells (Shleeva et al., 2002; O.A. Turapov and D.I. Young, unpublished data). In summary, stationary phase is an example of a dynamic environment in which many complex stresses are being generated and responded to over an extended period of time. We suggest that this constellation of conditions favours a survival strategy that results in a loss of immediate culturability of surviving cells. Another very important question is how stable are these ‘‘non-culturable’’ cells and how long can they exist in a reversible, ‘‘non-culturable’’ state? The answer depends on the organism under consideration. Some bacteria can survive in a ‘‘non-culturable’’ state for weeks or months, others for just a

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few hours. This may be determined, in part, by the life style of the organism. Thus M. tuberculosis has a generation time of ca. 20 h and can survive in a non-culturable state for months (Sun and Zhang, 1999; Shleeva et al., 2002) and possibly even years (Lillebaek et al., 2002). However, the period of ‘‘non-culturability’’ is time-limited; after surviving for some critical period, at least in these planktonic conditions, the cell must eventually divide or die. It is possible that the fluctuations of culturability sometimes observed in stationary phase cultures (e.g. Ekweozor et al., 1998; Shleeva et al., 2002, and many unpublished observations) are determined by this necessity for division to occur after the survival time has reached its limit. In the case of M. tuberculosis after incubation for 4 months in sealed tubes, the population consisted entirely of ‘‘non-culturable’’ cells, but during the next few months, CFU numbers increased by up to 5–6 orders of magnitude (Shleeva et al., 2002), usually accompanied by the lysis of some cells. The cells then started to lose culturability once again. This cyclical behaviour may permit populations to survive without added nutrients for many years (Corper and Cohn, 1933). Any attempt to make general conclusions about the conditions that lead to the formation of ‘‘non-culturable’’ cells in stationary phase faces obvious problems. Indeed, it seems that each species probably represents a unique case and many conditions need to be explored to find those that are suitable. This can be illustrated by consideration of some detailed studies of the stationary phase behaviour of three fast growing Gram-positive bacteria, viz. M. luteus, Rhodococcus sp. and M. smegmatis (Kaprelyants and Kell, 1993; Mukamolova et al., 1995b; Shleeva et al., 2002). These organisms did not produce significant numbers of ‘‘non-culturable’’ cells in stationary phase under optimum conditions of cultivation (temperature, medium composition, aeration, etc.). However, if cells of Rhodococcus sp. were grown in a sub-optimal minimal medium, they transited quite rapidly to a ‘‘non-culturable’’ state in stationary phase. This was maintained for a period lasting only a few hours that occurred some 90 hours post-inoculation. During this period, the bacteria could not produce colonies or grow in liquid medium unless it was supplemented with culture supernatant obtained from actively growing cells. Several hours later, culturability was spontaneously restored. Inoculum age, medium composition, speed of flask agitation and culture volume in relation to the flask capacity significantly influenced the formation of non-culturable bacteria. For each culture, there was a window of several hours in stationary phase, during which ca. 99.9% of the bacteria lost culturability (Shleeva et al., 2002). Similar results were obtained for M. smegmatis cells where a narrow window of ‘‘non-culturability’’, sensitive to medium composition and other parameters, has also been observed

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(M. Shleeva and A.S. Kaprelyants, unpublished data). In contrast to M. smegmatis and Rhodococcus sp. cultures, where ‘‘non-culturability’’ was a transient phenomenon in stationary phase, the majority of cells of M. luteus persisted in a ‘‘non-culturable’’ state during a prolonged stationary phase lasting several months (Mukamolova et al., 1998b; Kaprelyants and Kell, 1993). This timescale is intermediate between those of M. tuberculosis and Rhodococcus sp. ‘‘Non-culturability’’ in M. luteus is a particularly wellcharacterised example, in which some limited information is now available concerning the molecular basis of the loss and gain of culturability (see later). Again, like the two examples already mentioned, induction of the ‘‘non-culturable’’ state was achieved if cells were grown under sub-optimal conditions (in a lactate minimal medium). ‘‘Non-culturable’’ cells were not formed under starvation conditions (whether nitrogen-, carbon- or phosphate-starvation) for any of the three species tested. Possibly, cells grown in sub-optimal media have some sort of pre-adaptation to a ‘‘low cost metabolism’’, and at some stage adaptation reaches its extreme in the production of ‘‘non-culturable’’ cells. It is not yet clear whether lysis of some M. smegmatis or Rhodococcus sp. cells occurs during the phase of minimal culturability, which might serve as a signal for the observed spontaneous ‘‘resuscitation’’, or whether there are some other triggers that control the remarkable behaviour of these cells. It is also possible that ‘‘nonculturability’’ is a direct consequence of their continued existence under very poor growth conditions, i.e. the available nutrients are insufficient to support the normal metabolism and multiplication of all cells in the population. This situation can be modelled in chemostat culture. Loss of culturability has been reported, for example, in chemostat cultures of Cytophaga johnsonae (Hofle, 1983) and Aerobacter aerogenes (Tempest et al., 1967). Under such conditions there is a tendency for cultures to ‘differentiate’, i.e. the more metabolically active cells are increasingly more effective at scavenging nutrients as they enter the chemostat such that the chemostat effectively becomes heterogeneous. Significantly, cells lost culturability much more quickly at a very low dilution rate. Similarly, in lactate-limited chemostat cultures of M. luteus, only 40% of cells were culturable, at a very low dilution rate of 0.01 h1 (Kaprelyants and Kell, 1992). The inability to produce ‘‘non-culturable’’ cells in a rich medium can be explained by an imbalance between anabolism and catabolism (see Fig. 2). Cells in very rich medium grow at a maximal rate, supported by a maximal rate of metabolic activity. In a growing culture, catabolism and anabolism are balanced. When cells reach stationary phase, they shut down their main anabolic processes, mainly due to imposition of the

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Figure 2 Different mechanisms of loss of culturability of microbial cells operate under conditions that normally support slow (a) versus rapid growth (b). For detailed discussion, see the text.

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stringent response, and induction of the RpoS-regulated global network (Cashel et al., 1996). But these cells cannot reduce catabolic processes to the same extent as anabolic processes (Nystrom et al., 1996), leading to the intracellular production of free radicals (Bloomfield et al., 1998). If this unbalanced state is not rapidly rectified, free-radical scavenging mechanisms cannot cope adequately and cell structure deteriorates (Dukan and Nystrom, 1999; Nystrom, 2001), leading in extreme cases to lysis. (There is, perhaps, a parallel here with the phenomenon of ‘‘acid crash’’ that is seen when solventogenic clostridia growing in a rich medium fail to activate the metabolic switch from acidogenic to solventogenic metabolism, which would permit their long-term survival via the production of dormant endospores (Schuster et al., 2001).) Such cells are ‘‘nonculturable’’ but metabolically active. Their ‘‘non-culturability’’ is a consequence of cell injury but not a low-metabolic cost existence, and they cannot therefore be considered to be in a TNC state. The requirement for a medium that is not optimised for growth in order for a particular species to achieve a TNC state may explain why some bacteria do not produce ‘‘non-culturable’’ cells in stationary phase models. Such bacteria (e.g. E. coli and Vibrio spp., which can grow in almost any poor or rich medium) may have an extremely flexible metabolic potential, being able to adjust their metabolism to many different types of medium. In these organisms, low temperature serves to induce ‘‘non-culturability’’. The requirement for growth under sub-optimal conditions for transition to the ‘‘non-culturable’’ state could explain why, in some cases, mutant strains (e.g. auxotrophs) cultivated under conditions appropriate for the wild type, can display temporary ‘‘non-culturability’’. For instance, purF mutants of M. smegmatis, defective in stationary phase survival, revealed a transient decrease in culturability followed by an increase (Keer et al., 2000; Keer et al., 2001). The authors suggested two possible explanations of this effect: (i) re-growth (cryptic growth) of viable cells and (ii) formation of ‘‘non-culturable’’ (and possibly dormant) bacteria in stationary phase and their subsequent resuscitation. According to our studies, the second suggestion is correct, as such cells could be resuscitated (M. Shleeva and A.S. Kaprelyants, unpublished results). M. smegmatis is therefore a good example of an organism in which different experimental conditions result in the formation of the same ‘‘non-culturable’’ phenotype. Indeed, it is possible that an essentially identical mechanism underlies the production of ‘‘non-culturable’’ cells in all these cases. However, more work is required to verify this hypothesis (see below). Finally, another important point should be considered. Populations in stationary phase are very heterogeneous, often containing a mixture of

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different cells doing different things. Some may be in a state of starvation survival and others may be in a TNC state; some may be actively growing and others may be dying. This makes it difficult to design incisive experiments and is a source of considerable confusion in the scientific literature. Flow cytometric cell sorting of subpopulations (Davey and Kell, 1996) coupled to, for instance, microarray measurements (Schena, 2000; Betts et al., 2002), should soon help to set out the magnitude of this type of problem, i.e. how differentiated such cultures actually are.

2.2.4. ‘‘Non-culturable’’ Cells in Vivo ‘‘Non-culturability’’ is probably one of the major strategies of bacterial survival in vivo. The environments encountered by bacteria in vivo are very complex and they face several lines of host defence. These include general stresses like exposure to low pH and oxidising agents, as well as the influence of specific antibacterial compounds produced by specialised cells (Deretic and Fratti, 1999; Flynn and Chan, 2001). Lack of nutrients and low oxygen availability are additional challenges that they must face and overcome if they are to survive (Wayne and Sohaskey, 2001). There are many examples of latency, in which bacteria adopt a ‘‘non-culturable’’ state in vivo. The mechanism of this latency has been discussed extensively, but remains poorly understood (Young and Duncan, 1995; Parrish et al., 1998; Honer zu Bentrup and Russell, 2001; Wayne and Sohaskey, 2001). The organisms present in latent infections represent a reservoir of bacteria that could, in principle, reactivate and hence they have important implications for infectious disease and public health. It is still not clear if these persisting bacteria in latent infections are specialised forms, or if they are injured, or if they are simply ‘‘normal’’ bacteria that have been suppressed by the host immune system. The available evidence is controversial. The best-known example is afforded by the ‘‘non-culturable’’ forms of M. tuberculosis that have been found in lung lesions, sometimes in substantial numbers, but there is no evidence that these forms could be converted to normal multiplying cells (Wayne, 1960). Another important factor is the response of the bacteria following antibiotic therapy. The Cornell model of latency is based on the treatment of M. tuberculosis-infected mice with a mixture of antibiotics. After treatment, mice reach a so-called ‘‘sterile’’ condition during which no culturable bacteria are detectable. However, during the next 3 months without antibiotics, about one third of the mice go on to develop tuberculosis (McCune et al., 1966a, b). There is substantial variation on this model, depending on the precise experimental procedure

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that is employed (Scanga et al., 1999). During the ‘‘sterile’’ state, when culturable organisms are absent, DNA equivalent to ca. 105 bacteria per organ (lung or spleen) can be detected using PCR (de Wit et al., 1995). This might represent residual DNA resulting from the lysis of dead cells, but recently the presence of ‘‘non-culturable’’ organisms has been confirmed by RT-PCR (Pai et al., 2000). This is the first evidence that the ‘‘nonculturable’’ cells of M. tuberculosis present in the tissues are actively expressing genes and are thus metabolically active (and note too that even ostensibly ‘‘normal’’ lung tissue may contain mycobacterial DNA (Herna´ndez-Pando et al., 2000)). Since metronidazole had no activity, either in the initial sterilising phase or in the subsequent sterile state in the Cornell model (Dhillon et al., 1998; Brooks et al., 1999), the bacteria present (or at least some of them) do not have an anaerobic type of metabolism. Reactivation of M. tuberculosis was prevented by treatment with a combined course of isoniazid, pyrazinamide and rifampin (Dhillon et al., 1996), but not by rifapentine monotherapy (Miyazaki et al., 1999). It is possible that tuberculosis bacilli in the Cornell model are injured cells or L-forms, resulting from the specific action of the antibiotics used in this model, as has been suggested for human patients undergoing anti-tubercular chemotherapy (Khomenko et al., 1980). These cells may require special recovery conditions. In fact, the presence of L-forms and non acid-fast bacilli in vivo has been reported (Khomenko and Muratov, 1993). Interestingly, cells of M. tuberculosis from late stationary phase treated with rifampin showed zero culturability on agar plates and about 102 cells per ml in liquid culture (Hu et al., 2000). It also has been shown that some M. tuberculosis cells recovered from macrophages were extremely sensitive to freezing/thawing and treatment with detergents (Biketov et al., 2000). The current practice of decontamination for recovery of bacteria from sputum samples, involves harsh alkali treatment. Many culturable bacteria may become ‘‘nonculturable’’ as a result of injury during this procedure and they may need special stimuli to restore culturability. Significantly, the addition of a resuscitation-promoting factor increased the number of culturable cells by one or two orders of magnitude (Biketov et al., 2000). There are other examples of an apparent association between ‘‘nonculturable’’ bacteria and disease states. H. pylori, the causative agent of stomach ulcers (Marshall and Warren, 1984; Decross and Marshall, 1993), is detectable by PCR of biopsy samples, but not by other methods (culture, urease test, histology), for a substantial period of time after patients have been treated with antibiotics (Loginov et al., 1999). Moreover, BALB/c mice orally infected with coccoid (non-culturable) forms of H. pylori developed gastric inflammation with the same severity as animals infected with spiral

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(culturable) forms of this organism (Wang et al., 1997). ‘‘VBNC’’ uropathogenic bacteria have also been found in the mouse urinary tract after chemotherapy (Rivers and Steck, 2001) although, again, the important ‘‘dilution to extinction’’ tests (see Kell et al., 1998) were not performed. At all events, these types of reports suggest that some bacteria can survive in vivo as ‘‘non-culturable’’ cells. Moreover, there is the additional complication posed by the existence of some infectious bacteria that have never been cultured in vitro. Apart from the well-known case of M. leprae, with its degenerated genome (Cole et al., 2001), Tropheryma whipplei (or whippelii) which has never been cultured in vitro, is now considered as the causative agent of Whipple’s disease (Lynch et al., 1997; Fredricks and Relman, 2001; Maiwald et al., 2001). Even for organisms that have been cultured from some sources, there are diseases in which such organisms have been implicated but from which they have not been brought into culture. Sarcoidosis may be related to infection by M. tuberculosis (Khomenko et al., 1994) or Propionibacterium (Eishi et al., 2002) and Eales disease may be caused by M. tuberculosis (Madhavan et al., 2002). The main problem in all of these diseases is an inability to isolate the infectious agent in pure culture from patients and, hence, Koch’s postulates cannot be fulfilled (Fredricks and Relman, 1996). The presence of bacteria has been indicated by PCR but, except in the case of Whipple’s disease, there is the attendant risk of sample contamination. Another problem is the possibility of secondary infection. In some cases, the microorganisms isolated from patients appeared to be very specialised forms. For example, ‘‘dense bodies’’ were isolated from a patient with interstitial cystitis (Domingue et al., 1993). These ‘‘bodies’’ have a unique morphology with multilayer protective envelopes. The authors suggested that these structures could be resting forms of bacteria. L-form bacteria have also been reported as the causative agents of severe urological diseases (Domingue and Woody, 1997). It is still not clear if some of these organisms are truly specialised forms, or if they are simply injured cells suppressed by the host’s immune system. Nevertheless, two clear facts show the importance of atypical/‘‘non-culturable’’ bacteria in medicine. One is that the causative agent of infection can still be found in patients even after strong chemotherapy and the apparent clearance of infection (Velayati et al., 2002). The other is the occurrence of reactivation disease, which is well known in the case of tuberculosis (Dubrow, 1976; Kuznetsova, 1985; Flynn and Chan, 2001; Lillebaek et al., 2002) and other diseases (Domingue and Woody, 1997). Despite ongoing controversy, it seems very likely that adoption of a TNC state does truly represent a survival strategy exploited by several pathogenic bacteria in vivo. It effectively makes them ‘‘invisible’’ to the immune system, resistant to

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most (probably all) antibiotics, and since it is a low energy cost strategy, bacteria can survive for protracted periods in this state.

3. ARE THERE SPECIFIC CHEMICAL INDUCERS OF ‘‘NON-CULTURABILITY’’? In the preceding discussion we have developed the idea that ‘‘nonculturability’’ represents a survival strategy employed by bacteria as they gradually adapt to various stressful environments. By analogy with spore formation in bacilli, streptomycetes and myxobacteria (Losick and Kaiser, 1997), we may surmise that ‘‘non-culturability’’ is induced in response to the perception of specific chemical stimuli generated by cells as a result of the prevailing environmental conditions. For example, specific microbially produced peptides regulate the development of competence for transformation and sporulation in bacilli (Kleerebezem et al., 1997; Lazazzera and Grossman, 1998; Kleerebezem and Quadri, 2001; Lazazzera, 2001; Morrison, 2002) and N-acyl-homoserine lactones are involved in densitydependent signalling in Gram-negative bacteria (e.g. Salmond et al., 1995; Fuqua et al., 2001; Whitehead et al., 2002). Such signals act as pheromones (Stephens, 1986; Kell et al., 1995). They induce a wide spectrum of different physiological processes including, for example, bioluminescence in Vibrio harveyii (Cao and Meighen, 1989; Meighen, 1994), virulence in Pseudomonas aeruginosa (Winson et al., 1995), antibiotic production in Erwinia (McGowan et al., 1995), biofilm development in P. aeruginosa (Davies et al., 1998) and the cell division cycle in E. coli (Huisman and Kolter, 1994; Garcia-Lara et al., 1996; Sitnikov et al., 1996). It has been proposed that an N-acyl homoserine lactone may serve as a signalling molecule under starvation conditions in E. coli, leading to activation of the sigS gene (Huisman and Kolter, 1994). Later it was reported that the level of sigS expression in E. coli is positively controlled by a homocysteine thiolactone during stationary phase (Goodrich-Blair and Kolter, 2000), providing a possible link between production of this molecule and the growth arrest that occurs under these conditions. Homoserine lactones may activate the starvation-survival response in Vibrio sp. (Srinivasan et al., 1998). These authors showed that halogenated furanones, which are putative antagonists of acylated homoserine lactones (Manefield et al., 1999, 2002; Hentzer et al., 2002), inhibited the synthesis of proteins specifically induced upon carbon starvation. Of course many other compounds, for which no signalling role has yet been clearly established, are known to be excreted from

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microbial cells, especially under conditions of slow or zero growth and multiplication rates. These include uracil and xanthine (Rinas et al., 1995), furanones (Slaughter, 1999) and the numerous bioactive molecules of both ‘‘primary’’ (e.g. Wittmann and Heinzle, 2001) and ‘‘secondary’’ (e.g. Abel et al., 1999) metabolism, which are often of applied interest (Devlin, 1997). Indeed, the ‘‘global’’ measurement of culture supernatants using modern mass spectrometric procedures shows that a vast and largely uncharacterised panoply of substances are normally excreted, and the application of numerical deconvolution methods to such data allows one to discriminate strains which differ only in the presence of a single gene (Allen et al., 2002). There is as yet no really direct evidence for the existence of chemical inducers of ‘‘non-culturability’’, but there are indications that such molecules may exist. The accumulation of compounds in the external environment that inhibit bacterial growth during the transition to the ‘‘non-culturable state’’ has been reported (Weichart et al., 1992) (as have the presence of molecules stimulating regrowth following transfer of E. coli cells from stationary phase to growth medium (Weichart and Kell, 2001)). Accumulation of toxic compounds could also be responsible for the transition to ‘‘non-culturability’’ in the case of M. tuberculosis (Sun and Zhang, 1999; Zhang et al., 2001; Shleeva et al., 2002), Rhodococcus sp. (Shleeva et al., 2002) and M. luteus (Mukamolova et al., 1995a). Indeed, so-called ‘staling factors’ have been known for many years (e.g. Barker et al., 1983). These toxic compounds may be implicated in adoption of the TNC state. In M. xanthus, production of bactericidal derivatives of unsaturated fatty acids causes lysis of the majority of cells in the population during the formation of fruiting bodies and the release of spores (Varon et al., 1984). Accumulation of some specific low molecular weight compounds is considered responsible for the ‘‘programmed’’ loss of culturability of E. coli cells in stationary phase (Vulic and Kolter, 2002). It is interesting that ethanol interferes with this process, since it also interferes with endospore formation in B. subtilis (Vulic and Kolter, 2002). The accumulation of low molecular weight compounds that suppress growth in stationary phase has also been reported in S. typhimurium (Barrow et al., 1996). Alkylresorcinols are involved in cyst formation in Azotobacter vinelandii, suggesting that these compounds may act as regulators of ‘‘non-culturability’’ in this organism (Reusch and Sadoff, 1979). The addition of alkylresorcinols to growing cells of Bacillus cereus caused the formation of ‘‘refractile’’ spore-like ‘‘non-culturable’’ forms (Duda et al., 1982). However, there is no clear evidence that these ‘‘non-culturable’’ forms can be resuscitated.

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Alternatively, the toxic compounds listed above may provoke a ‘‘nonspecific’’ form of chemical stress (like exposure to heavy metals, for example), which is known to cause the formation of ‘‘non-culturable’’ (or dead) cells (Grey and Steck, 2001a, b). These effects are observed only in high-density cultures. There is no evidence of accumulation of inhibitory compounds or of inducers of ‘‘non-culturability’’ in the diluted microcosm experiments discussed previously (although we note that non-planktonic bacteria in nature may co-exist at concentrations that are locally (effectively) very high). It is not clear whether these substances are natural physiological ‘‘waste’’ products of overflow metabolism (Neijssel and Tempest, 1976) or specific regulators of the culturability to ‘‘non-culturability’’ transition. Therefore, the important question of chemical induction of ‘‘non-culturability’’ – which has substantial implications for public health – remains unanswered, and more experimental research is required.

4. IS ‘‘NON-CULTURABILITY’’ GENETICALLY CONTROLLED? We now consider whether adoption of the TNC state is associated with the expression of specific genes. Currently all that we can (or tend to) measure is the end result, i.e. ‘‘non-culturability’’. Until the processes that underpin the transition to, the maintenance of, and the exit from, the ‘‘non-culturable’’ state are better understood, we shall remain unable to provide a definitive answer to the question. Nevertheless, the simple fact that ‘‘non-culturable’’ cells show altered morphology, biochemistry and physiology, suggests that the TNC state is genetically controlled. During the alternative process of starvation survival (see above) some genes are switched on and others are switched off, suggestive of control by a general starvation-induced program (Kjelleberg et al., 1993) and it has also been argued that cell death can be genetically controlled (Vulic and Kolter, 2002). In addition, the adoption of several other physiological states is also genetically controlled (transition to a non-dividing state in stationary phase or under starvation conditions, development of resistance to different stresses). For example, an arcA mutant of E. coli was unable to reduce respiration and total metabolic activity in stationary phase (Nystrom et al., 1996) and, as a result, it was severely impaired in its ability to survive prolonged periods of carbon starvation. There are many reports showing induction of the synthesis of specific proteins depending on conditions of starvation. For example, cultures of V. vulnificus pre-starved at 24 C and then shifted to 5 C maintained

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culturability at low temperature in a starvation-condition-dependent manner (Paludan-Muller et al., 1996). This is probably related to initiation of the starvation survival program. A null mutation in the smcR (luxR) gene prevented starvation survival under these conditions (McDougald et al., 2001). The physiological age of the cells also influenced the transition to ‘‘non-culturability’’ (Oliver et al., 1991). There are many examples of changed biochemical and morphological characteristics of ‘‘non-culturable’’ cells. Moreover, it has been shown that some ‘‘non-culturable’’ cells can take up methionine and synthesize proteins (Rahman et al., 1994). These processes presumably reflect the expression of particular genes. Additional evidence in favour of this interpretation has been obtained by Boshnakov et al. (2001) who identified several genes that are preferentially expressed in ‘‘VBNC’’ forms of S. typhimurium, using differential display technology. Very recently, different protein expression profiles have been found in starved vs. ‘‘VBNC’’ cells of E. faecalis (Heim et al., 2002). Further evidence for genetic control of ‘‘non-culturability’’ comes from analysis of other mutant strains. A mutation has been described that speeds up the transition of V. cholerae to some kind of ‘‘VBNC’’ state (Ravel et al., 1994), although it is not clear what particular gene was disrupted in this mutant. In the case of M. smegmatis, a purF mutant lost culturability in stationary phase, whereas the wild type control did not (Keer et al., 2001). The mutant cells subsequently regained culturability and it was established that this was not due to reversion. The intriguing behaviour of this mutant is not currently understood. Several mutants with an altered ‘‘VBNC’’ response have also been isolated in S. typhimurium (Romanova et al., 1996). For example, an lpf mutant did not produce ‘‘VBNC’’ forms when incubated under conditions that normally lead to the adoption of a ‘‘VBNC’’ state. Disruption of glgC (glycogen metabolism) increased the period of survival in the ‘‘VBNC’’ state, whereas disruption of pqi, a paraquat-inducible gene that is induced under stress and controlled by the soxRS locus, and another gene of unknown function, altered the timing of the period of transition to the ‘‘VBNC’’ state. One of the most difficult problems to be overcome in any study of gene expression in ‘‘non-culturable’’ cells is population heterogeneity. Cultures frequently contain mixtures of culturable and ‘‘non-culturable’’ cells, which can make it very difficult to ascribe quantitative changes to a particular cell type with any degree of reliability unless careful and quantitative arguments are made. Flow cytometry can be very helpful in this regard (Davey and Kell, 1996) but this technology is presently only applicable in cases where the genes under investigation are expressed at a comparatively high level. Homogeneous cultures (100% of cells in the population in the same

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physiological state) are the best model for this type of study, but they are not readily obtained. One example of such a nominally ‘‘homogeneous’’ culture is the so-called Wayne model of ‘‘dormancy’’ for M. tuberculosis, induced by gradual oxygen depletion in vitro, that was mentioned previously. Bacteria stop DNA replication and cell division in a synchronised fashion. However, they remain ‘‘culturable’’ because they resume synchronous division and growth immediately after oxygen input. These cells have an active but altered ‘‘anaerobic’’ type of metabolism, based on use of the glyoxylate pathway with fatty acids as carbon source. Microarray and proteomic analyses have confirmed that this ‘‘dormancy’’ model is more correctly viewed as an adaptive response to oxygen depletion. In cells from the Wayne model, there is strong up-regulation of genes encoding an -crystallin homologue thought to be a chaperonin, a putative response regulator Rv3133c, and two conserved hypothetical proteins Rv2623 and Rv2626c (Boon et al., 2001). A similar set of genes is up-regulated during oxygen shift-down in actively growing cultures (Sherman et al., 2001). The Wayne model therefore represents a clear example of bacterial adaptation to unfavourable growth conditions. M. tuberculosis cells in a starvation survival model showed a completely different profile of gene expression (Betts et al., 2002). It has been suggested that during prolonged stationary phase in vitro, sigJregulated gene expression determines the survival strategy (Hu and Coates, 2001). It would not be surprising if yet another completely distinct strategy were involved in the survival of M. tuberculosis in vivo. Unfortunately, little is known about mycobacterial gene expression in the Cornell model of latency. Apart from the heterogeneity of cells, analysis is hampered by the poor availability of material. It is equally important to understand the molecular basis of cell recovery/ resuscitation from the ‘‘non-culturable’’ state. Recovery of E. coli after starvation was dependent on the expression of some specific genes (Siegele and Kolter, 1993) but, as yet, there is no clear evidence supporting the presumed existence of genes specifically required for restoration of culturability in other organisms. 5. RESUSCITATION OF ‘‘NON-CULTURABLE’’ CELLS The important distinction between cells in the TNC state and cells that are ‘‘dead’’ is that only the former can be brought back into a culturable form, by a process called resuscitation. The distinction between cells that are ‘‘non-culturable’’ and those that are dead is, as with higher organisms

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(Watson, 1987; Berger, 1992), purely operational (Kell et al., 1998); there are undoubtedly many living organisms that we do not yet know how cultivate. (In these cases it is the microbiologist, rather than the organism, that would more correctly be regarded as ‘‘uncultured’’.) The main criterion for the presence of ‘‘non-culturable’’ cells is that under specialised conditions they can be coaxed back into a state where growth and division are resumed. The ability to resuscitate thus serves unequivocally to distinguish ‘‘non-culturable’’ cells from dead cells. Indirect criteria (metabolic activity, cell integrity, etc.) are not sufficient proof of viability (the ultimate ability to resume growth), and such measurements are a source of much confusion, especially when carried out in cultures consisting of mixtures of cells in different physiological states (Kell et al., 1998). Resuscitation is almost certainly a very complex process, requiring restoration of many cellular functions, leading finally to the reacquisition of culturability (Kaprelyants and Kell, 1993), and the ability to restore one particular function does not necessarily imply competence to perform them all. We thus concentrate here on the end result of resuscitation, viz. the restoration of culturability. The essential problem – assuming that the objects of study are not merely dead – is to find conditions that are suitable for resuscitation. Different physiological processes have to be taken in account when the medium is selected. First of all, the cells may be ‘‘metabolically injured’’ (Postgate, 1976); one or more components of a medium that is harmless for normal cells can be toxic for recovering cells. Substrate-accelerated death is a welldocumented example (Calcott and Postgate, 1972). For isolation of bacteria from natural environments, a diluted broth is normally used (MacDonell and Hood, 1982). The optimal conditions for the recovery of ‘‘nonculturable’’ bacteria from lake water were very low substrate concentrations and an incubation atmosphere of 21% oxygen at 16 C for 4 weeks (Bussmann et al., 2001). Consistent with this, a very rich medium did not permit the resuscitation of ‘‘non-culturable’’ cells of the copiotroph M. luteus (Mukamolova et al., 1998b). It has been reported that addition of catalase or sodium pyruvate can significantly improve the recovery of some, but not all, ‘‘non-culturable’’ cells found in nature or obtained under laboratory conditions (Bogosian et al., 2000). This observation supports the old idea that some cases of ‘‘non-culturability’’ are a direct consequence of the damaging action of free radicals (Bloomfield et al., 1998). In some cases, restoration of culturability was observed when cells were plated in soft agar (Weichart and Kjelleberg, 1996). Sometimes cells can be cultivated successfully in liquid but not on solid media (Bovill and Mackey, 1997; Biketov et al., 2000; Hu et al., 2000; Shleeva et al., 2002). A combination of

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liquid medium and catalase was the best procedure for resuscitation of ‘‘non-culturable’’ cells of A. hydrophila (Wai et al., 2000). It is also possible that ‘‘non-culturable’’ cells cannot produce some important compounds required for the initiation of division. They have to be supplied in the medium. In the case of M. luteus and Rhodococcus sp., small amounts of yeast extract improved resuscitation dramatically and, moreover, the optimal concentration of yeast extract depended greatly on the age of the ‘‘non-culturable’’ cells (Mukamolova et al., 1998b; Shleeva et al., 2002). Improved recovery of cells of C. jejuni and Campylobacter coli has also been observed in media containing yeast extract (Moore, 2000). Addition of certain phospholipids in agar increased the number of CFU in populations of ‘‘non-culturable’’ M. tuberculosis cells by several orders of magnitude (Zhang et al., 2001). These agents are known to promote the repair of damaged membranes (Ray and Speck, 1973) suggesting perhaps that the cells in these experiments were ‘‘non-culturable’’ as a result of injury. However, other explanations should not be ruled out. In the case of M. luteus, ‘‘non-culturable’’ cells pre-treated with cerulenin (an inhibitor of fatty acid synthase) did not resuscitate (Kaprelyants and Kell, 1993). Moreover, during the process of resuscitation of ‘‘non-culturable’’ cells of this organism, the disappearance of lysophospholipids correlated with restoration of the permeability barrier (de novo protein synthesis was not required) and permitted resuscitation of the majority of ‘‘non-culturable’’ cells in the population (Mukamolova et al., 1995b). The detailed cultivation regime (shaking speed, temperature, pH, etc.) can be an important factor for resuscitation. Sometimes a transition from one type of starvation to another can be helpful. For example, when ‘‘nonculturable’’ cells of P. fluorescens obtained under conditions of nitrogen starvation were transferred to complete medium, there was no resuscitation (substrate-accelerated death?) whereas they were resuscitated successfully when transferred to conditions of carbon (and energy) starvation before plating (Evdokimova et al., 1994). The authors noted that metabolic activity was restored before culturability. In some cases, pre-incubation of ‘‘nonculturable’’ cells in liquid medium in the presence of antibiotics improved resuscitation dramatically (Mukamolova et al., 1995b). However, in other cases, cells do not need special manipulation and culturability is restored after the stress has been relieved. For example, Nilsson and colleagues (Nilsson et al., 1991) reported that ‘‘VBNC’’ cells of V. vulnificus, obtained by starvation at 4 C, could be resuscitated by simple temperature upshift. However, it has been pointed out by others that re-growth of a small number of culturable cells present in these populations could be responsible for the increased culturability that they observed (Weichart et al., 1992; Bogosian

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et al., 2000). Similarly, Ohtomo and Saito (2001) reported the rapid resuscitation of saline-stressed cells of E. coli following a reduction in salt concentration. Some attempts have also been made to resuscitate cells by physical treatments. For example, sonication and heating in the case of Streptomyces (Hirsch and Ensign, 1976; Miguelez et al., 1993), or high pressure and heating in the case of Bacillus (Paidhungat et al., 2002), promotes spore germination. In two reported cases, heating of VBNC forms stimulated the restoration of culturability (Wai et al., 1996; Kurokawa et al., 1999), but high pressure did not similarly affect VBNC cells of Vibrio (Berlin et al., 1999). Some ‘‘non-culturable’’ cells may require highly specialised environments for resuscitation. This may apply in particular to certain pathogenic bacteria, ‘‘non-culturable’’ forms of which can resuscitate and multiply only in vivo, or in artificial conditions that are very similar to those encountered in vivo. Restoration of culturability and virulence in vivo to previously ‘‘nonculturable’’ cells has been reported for V. vulnificus (Oliver and Bockian, 1995), V. cholerae (Colwell et al., 1996), Salmonella enterica (Asakura et al., 2002), Ralstonia solanacearum (Grey and Steck, 2001b) and C. jejuni (Cappelier et al., 1999), though in none of these studies were the estimates of culturability done under conditions of dilution to extinction. Resuscitation of ‘‘non-culturable’’ cells of Legionella pneumophila within Acanthameoba castellanii has also been reported (Steinert et al., 1997). ‘‘Non-culturable’’ cells of Y. pseudotuberculosis could be resuscitated by passage through an axenic culture of Tetrahymena pyriformis (Didenko et al., 2002). Interestingly, Bacillus spores could germinate and colonise the mouse intestine for brief periods, indicating that this environment supports spore germination (usually requiring a specific chemical germinant) and multiplication (Casula and Cutting, 2002). It is possible that some specific compounds in vivo stimulate resuscitation of ‘‘non-culturable’’ forms. For example, growth of some bacteria is stimulated by hormones or lymphokines from higher organisms (reviewed by Kaprelyants and Kell, 1996) – and vice versa (Henderson et al., 1996) – and Williams, Lyte and colleagues have shown the significance of mammalian neurotransmitters in the former regard (Lyte, 1992, 1993; Lyte and Ernst, 1992, 1993; Freestone et al., 1999, 2000; Neal et al., 2001). VBNC cells of H. pylori showed metabolic reactivation (increase of ATP level and transcription of some genes) when human erythrocyte lysates were added to the recovery medium (Nilsson et al., 2002). A crucial problem in any resuscitation experiment is distinguishing between resuscitation of ‘‘non-culturable’’ cells and re-growth of culturable cells, even if the latter represents only a very small proportion of the total

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cells present. Sometimes a mixture of re-growth and resuscitation occurs (Dukan et al., 1997). Several procedures have been employed to infer the occurrence of resuscitation, e.g. measurement of the kinetics of growth/ resuscitation, following the ratio of ‘‘non-culturable’’/culturable cells, etc. However, it is very difficult, if not impossible, to determine the percentage of resuscitating cells using these methods. For quantitative analysis, Most Probable Numbers (MPN, i.e. counts based on dilution to extinction) should be determined. Different dilutions of cells are inoculated into several replicate tubes containing liquid medium. By scoring positive/negative tubes and using statistical tables, it is possible to back-calculate the most probable number of culturable cells in the original sample. An important feature of this simple methodology is that the behaviour of ‘‘non-culturable’’ cells can be monitored at low dilutions where, statistically speaking, culturable cells are absent. This method is especially useful when different stimulatory (or indeed inhibitory) compounds are added to or present in the growth medium. If a substance stimulates the growth of culturable cells in the population, changing their growth rate, the kinetic argument (see above) is invalidated. For example, it was found that some low molecular weight compound stimulated re-growth but not resuscitation of starved E. coli cells (Weichart and Kell, 2001). If these experiments had not been done using the MPN method it would not have been possible to distinguish the re-growth observed by these authors from resuscitation. Unfortunately, there are very few unequivocal demonstrations of the resuscitation of ‘‘non-culturable’’ cells (Kell et al., 1998). In many cases, no resuscitation was observed under any of the conditions tested (Bogosian et al., 1996; Eguchi et al., 2000; Kolling and Matthews, 2001; Mary et al., 2002). In others, resuscitation was possible only over a limited period of time following adoption of the TNC state (Roszak et al., 1984; Lleo et al., 2001; Grey and Steck, 2001a). There are many discrepancies and complications in the published literature describing the resuscitation of ‘‘non-culturable’’ cells. A further feature is that in some experiments ‘‘nonculturable’’ cells inoculated in fresh medium were able to make limited cycles of division, and then stopped. For example, VBNC forms of P. fluorescens, obtained from a soil microcosm, could only produce microcolonies resulting from two or three divisions when resuscitated (Binnerup et al., 1993). The reason for such limited divisions is not known. For M. luteus, accumulation of toxic compounds has been observed during resuscitation in fresh medium and under such conditions, cells also made only a limited number of divisions (Mukamolova et al., 1995a). Stationary phase populations of E. coli contain some cells capable of making just one division (Ericsson et al., 2000). In a few cases, only one of the two daughter

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cells could divide subsequently. The authors proposed that this behaviour might result from some kind of localised deterioration or damage, inherited by only one daughter cell. The phenomenon of limited divisions in eukaryotic cells is of course very well documented (Hayflick and Moorehead, 1961; Rubin, 2002). Finally, as previously mentioned, many of the VBNC cells described in the literature may be dying or metabolically injured. By definition, such cells cannot be ‘‘resuscitated’’. One of the most intriguing questions is whether special resuscitationinducing factors exist. Addition of some compounds (so-called germinants, such as alanine or adenosine) dramatically increases the rate and proportion of spores of various Bacillus spp. that germinate (Moir and Smith, 1990). Germinants bind to specific Ger proteins and stimulate ion transport, leading to the activation of lytic enzymes that degrade the spore envelopes, to rehydration of the spore core, and to the breaking of dormancy (Paidhungat and Setlow, 2000). Some components in resuscitation medium may have a similar effect to that of germinants. A change of growth medium may switch on the early stages of resuscitation, i.e. restoration of membrane energisation and respiratory activity, but in some cases this is not sufficient for restoration of culturability (Votyakova et al., 1994). In the case of M. luteus, the addition of sterile culture supernatant was necessary for the restoration of culturability of ‘‘non-culturable’’ cells, suggesting that active cells could secrete compounds that promote resuscitation (Kaprelyants et al., 1994). Interestingly, the resuscitation of undiluted populations containing small numbers of culturable cells did not require the addition of any specific factors (Kaprelyants and Kell, 1993; Votyakova et al., 1994). The resuscitation-promoting factor (Rpf ) in the sterile culture supernatant of M. luteus was non-dialysable, heat-labile and trypsin-sensitive, suggesting that it was a protein and this was subsequently confirmed (Mukamolova et al., 1998a). The presence of Rpf was necessary for the restoration of culturability to ‘‘non-culturable’’ (dormant) cells of M. luteus, although their metabolic activity could be restored in its absence (Votyakova et al., 1994). In addition to promoting the resuscitation of ‘‘non-culturable’’ cells, Rpf caused a reduction in the apparent lag phase of cultures growing from small inocula. An additional assay was developed using washed vegetative cells, whose growth was dependent on supplementation with Rpf, which indicates that Rpf has a more general role as a growth factor in M. luteus (Mukamolova et al., 1998a; Kaprelyants et al., 1999) (Figure 3). Rpf is a secreted protein, active at picomolar concentrations, suggesting that it may represent a signalling molecule. Disruption of the rpf gene was not possible in M. luteus in the absence of a second functional copy, indicating essentiality (Mukamolova et al., 2002a). Moreover, the protein

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Figure 3 M. luteus requires Rpf for growth. Growing cells make Rpf. During a protracted stationary phase, bacteria may enter a state of dormancy during which they are ‘‘non-culturable’’. Culturability is restored by the exogenous provision of Rpf. Normally, the source of Rpf is an actively growing neighbouring cell(s) but recombinant Rpf made in E. coli also suffices (Mukamolova et al., 1998a). In the laboratory, actively growing cells can be made Rpf-dependent by extensively washing them, which removes pre-synthesised protein molecules from the cell surface, and inoculating them at low density into a minimal medium.

must be secreted (or provided exogenously) to be biologically active (Mukamolova et al., 2002a). However, its mechanism of action remains unknown. Our working hypothesis is that Rpf, or a peptide derived from it, is the ligand for a surface-located receptor, but until the predicted receptor has been characterised this speculation is supported only by the extreme potency of the molecule – a truncated derivative encompassing only the Rpf domain (see below) was active at femtomolar concentrations (Mukamolova et al., 2002a). It still remains entirely possible that Rpf has a catalytic activity of some kind. The schematic diagram in Figure 4 shows the relationship between Rpf production and the growth cycle of M. luteus in laboratory batch culture. Rpf is the founder member of an extended protein family that is found throughout (and, so far as is known, exclusively within) the high G þ C cohort of Gram-positive bacteria. There are several representatives in most organisms and the gene family now contains in excess of 30 members (Kell and Young, 2000; Matsuda et al., 2001; Jang et al., 2002). For example, M. tuberculosis (Cole et al., 1998) and S. coelicolor (Bentley et al., 2002) both contain five rpf-like genes. It has recently been established that recombinant versions of all five proteins from M. tuberculosis have similar

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Figure 4 The distribution of Rpf during different stages of growth of M. luteus in laboratory batch culture. Bacteria make and secrete Rpf during lag phase and early log phase (Mukamolova et al., 2002a). Most molecules remain associated with the cell surface. As growth continues and clumps of cells start to break up, Rpf is released into the culture supernatant. In stationary phase, Rpf molecules in the supernatant lose biological activity, and eventually disappear (degradation). If cells spend a protracted period in stationary phase they enter the state of ‘‘transient non-culturability’’.

biological activity to that of M. luteus Rpf (Mukamolova et al., 2002b). Moreover, Rpf-like biological activity has also been observed using recombinant proteins from Corynebacterium glutamicum (Jang et al., 2002) and M. smegmatis (D.I. Young and M. Shleeva, unpublished data) and in culture supernatant from Rhodococcus sp. (Shleeva et al., 2002). Most members of the protein family are predicted to be secreted, or membraneanchored, and lysM modules (Bateman and Bycroft, 2000) promoting association with the cell wall peptidoglycan are a common feature of the Rpf-like proteins from the non-mycolate actinomycetes. Although there is some evidence for selectivity (see below), all of the Rpf-like proteins so far tested show cross-species activity (Mukamolova et al., 1998a, 2002b; Biketov et al., 2000; Shleeva et al., 2002; A. Davies, unpublished data). In addition, M. luteus culture supernatant (presumably containing Rpf ) has been reported to enhance the isolation of mycobacteria in liquid medium (Freeman et al., 2002).

6. SOCIAL BEHAVIOUR OF BACTERIAL POPULATIONS AND ‘‘NON-CULTURABILITY’’ Over the last decade it has become abundantly clear that bacteria do not live in isolation one from another. Rather, they form structured communities in

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which individual cells communicate with each other via signalling molecules (reviewed by Kaiser and Losick, 1993; Losick and Kaiser, 1997; O’Toole et al., 2000; Shapiro, 1995b). Social behaviour of cells is undoubtedly of cardinal importance for many different cellular processes associated with multiplication, differentiation, survival in fluctuating environments and death. In this section we consider how multicellular bacterial communities respond to stress. One good example is presented by the behaviour of bacteria growing in biofilm communities. Biofilms are complex structures (Rittman, 1999; Stoodley et al., 2002) resulting from the expression of genetic information according to a defined developmental programme (Danese et al., 2001; O’Toole et al., 2000). They provide a niche in which growth conditions are optimised for communal living (Gilbert et al., 2002) and polysaccharide production confers substantial resistance to several different stresses (Li et al., 2001; Mah and O’Toole, 2001), including starvation (Kim and Fogler, 1999). In response to nutrient depletion, cells in the biofilm can adopt two alternative survival strategies. Organisms may enter a starvation survival programme in situ, or the biofilm may dissociate as its members adopt a motile free-living (planktonic) existence. Failure to find food in the latter case can lead to adoption of a TNC state. There is an interesting parallel here with cells that normally grow as clumps or aggregates. During prolonged incubation of cells of M. tuberculosis or M. bovis (BCG) in stationary phase, a heterogeneous population arose containing ‘‘aggregates’’ composed of dead lysing cells and single short rods that were ‘‘non-culturable’’ and could be resuscitated (Shleeva et al., 2002; O.A. Turapov and D.I. Young, unpublished data). Persistent cells inside murine macrophages are also disaggregated (Biketov et al., 2000). In denser populations, cells often survive using a strategy that has been likened to ‘‘cannibalism’’ (Postgate, 1976). Generally this allows some members of the population to survive and generate progeny in the future at the expense of the others and, hence, it may be regarded as an example of cooperative or social behaviour, permitting species maintenance. In stationary phase cultures of E. coli mutants arise showing an ‘‘evolutionary cheating’’ behaviour (Vulic and Kolter, 2001). These cells have rpoS mutations and cannot initiate a proper ‘‘stationary’’ phase response. They continue to multiply in conditions of stationary phase due to the development of amino acid-dependent catabolism (Zinser and Kolter, 2000). Moreover, they have a growth advantage in mixed culture with wild type bacteria and tend to overgrow them (Zambrano and Kolter, 1996). The survival advantage of the mutant is inversely dependent on its frequency in

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the population, and a balance of normal cells and ‘‘cheaters’’ can be maintained. Moreover, these cells are more sensitive to various stresses than is the wild type, A similar phenomenon has also been described for M. smegmatis cells incubated in prolonged stationary phase (Smeulders et al., 1999). Cells producing colonies with altered morphology had a ‘‘growth advantage in stationary phase’’ but not under normal growth conditions. There is some evidence that cells can produce or release compounds that help other cells to survive under conditions of stress (Harrison, 1960; Rowbury, 2001) and adapt to new environments (Nikolaev et al., 2000). These compounds are found in the culture supernatant of many different species and, apparently, there is no specificity in their action. Usually, the population of surviving cells in a starving culture is very heterogeneous. In many organisms it consists of a mixture of dead (lysing) cells and survivors; the latter may include viable cells that will produce CFU and may even be slowly multiplying, and ‘‘non-culturable’’ cells, which may be dormant. This may result from the ‘‘initial’’ heterogeneity of the population due to unsynchronised growth. In the case of M. luteus, the surviving population always contained some residual culturable cells (103–105 cells/ml in populations containing 109–1010 cells/ml). These culturable cells promote the resuscitation and growth of the majority of ‘‘non-culturable’’ cells when inoculated into fresh medium (Votyakova et al., 1994). This can be regarded as a particularly efficient survival strategy since it permits survival of the majority of the cells in the original bacterial population and is a clear example of the cooperative behaviour of surviving cells. In the case of V. vulnificus, a population of ‘‘non-culturable’’ cells, starved in buffer at 4 C, were restored to culturability when they were transferred to 20 C. However, this was not possible when the population was diluted 10- or 100-fold. The simplest explanation is that in this case, ‘‘resuscitation’’ is in fact re-growth of initially undetectable culturable cells. But it is also possible that culturability was restored to some cells more quickly than to others and that the cells that became culturable early on, secreted a growth-promoting substance. The experimental data on ‘‘cooperative’’ resuscitation of Vibrio cells are quite controversial: in some experiments active cells, mixed with VBNC cells, helped the latter to restore culturability (Whitesides and Oliver, 1997), whereas in other experiments no growth-promoting effect of active cells was detected (Bogosian et al., 1998). The resuscitation of ‘‘non-culturable’’ cells by compounds accumulated in culture supernatant was first observed in M. luteus (Kaprelyants et al., 1994). It has also been observed in C. jejuni (Bovill and Mackey, 1997), Nitrosomonas europaeae (Batchelor et al., 1997), M. tuberculosis (Sun and Zhang, 1999), Rhodococcus sp. (Shleeva et al., 2002) and M. smegmatis

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(M. Shleeva and A.S. Kaprelyants, unpublished). It is clear that cooperative behaviour is an important feature not only in populations during the transition to a non-dividing state but also during the reverse process of recovery or resuscitation. Interestingly, large inocula of Bacillus megaterium spores germinated at a significantly higher rate (in percentage terms) than did small inocula, suggesting that germinating spores may produce something that stimulates the germination of their immediate neighbours (Caipo et al., 2002). Similar effects have been observed in Clostridium botulinum (Wong et al., 1988). In contrast, inhibition of Streptomyces spore germination was observed at very high population densities (Polyanskaya and Triger, 1990; Triger and Polyanskaya, 1991). The discovery of endogenous germination inhibitors in Streptomyces may be significant in this context (Petersen et al., 1993). The resuscitation of ‘‘non-culturable’’ cells occurs in response to the perception of external factors and it is now becoming apparent that some of these factors can be produced by neighbouring cells. We have suggested that, in the high G þ C cohort of Gram-positive bacteria, the active molecules are, or include, proteins of the Rpf family (Kaprelyants et al., 1999). It has been argued elsewhere (Postgate, 1995) that adoption of the ‘‘non-culturable’’ state by the majority of cells in a population is of adaptive significance, since a small number of ‘‘sentinel’’ organisms can make best use of scarce resources during periods of starvation and then signal the onset of conditions favourable for growth to their genetically identical but quiescent neighbours. Should the entire population lose culturability, all is not lost, since the resuscitation-promoting factors show cross-species activity (Mukamolova et al., 1998a, 2002b). Finally, it has recently been demonstrated that rpf is an essential gene in M. luteus (Mukamolova et al., 2002a). Why does this organism (and possibly others that make Rpf-like proteins) secrete a protein that is essential for its growth? Why have bacteria evolved such a precarious system for growth regulation? This is most readily understood if we adopt the view that the organism is not in fact the individual bacterial cell, but the entire bacterial population. The bacteria that we grow as isolated planktonic forms in laboratory shake flasks probably form structured communities in nature where they enjoy all the benefits of a multicellular existence. 7. CONCLUSION There has been much discussion and controversy concerning the nature and biological significance of ‘‘non-culturable’’ cells. Nevertheless, it is generally

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agreed that ‘‘non-culturable’’ cells are abundant in Nature and that they can be obtained under laboratory conditions and, therefore, they have importance for ecology (Roszak and Colwell, 1987), medicine (Barer et al., 1993; Barer and Harwood, 1999) and industry (Millet and LonvaudFunel, 2000; Nebe-von-Caron et al., 2000). In this review, we have developed the hypothesis that transient ‘‘non-culturability’’ is a consequence of a special survival strategy. The fact that ‘‘non-culturable’’ cells have altered morphology and are adapted for survival under stressful conditions suggest that they show some resemblance to the more specialised survival structures produced by sporulating bacteria. We have emphasised that only cells that satisfy several different criteria, i.e. integrity of all cellular components, low metabolic activity and, most importantly, the ability to be resuscitated – preferably in the absence of other culturable cells – should be considered as having adopted the TNC state. Unfortunately, in many cases in which ‘‘non-culturability’’ has been inferred, only one criterion (e.g. staining with CTC or tetrazolium) has been applied. Moreover, we still do not know whether VBNC cells, obtained under cold starvation, persisting cells in latent infections in vivo and ‘‘non-culturable’’ cells, arising in stationary-phase/chemostat culture, are different manifestations of the same underlying process, i.e. whether they represent similar surviving forms of different bacteria. We exclude ‘‘non-culturable’’ metabolically active cells from the category of special surviving forms. In order to adopt the TNC state, bacteria must first adapt to a low metabolic cost existence and as a consequence of this they then lose culturability. Initial loss of culturability, which may be followed by gradual metabolic decline, is indicative of injury, leading to death, and is quite distinct from entry into a TNC state. One of the main potential criticisms of the idea that ‘‘non-culturable’’ cells are specialised resting forms, is that their state of ‘‘non-culturability’’ is temporary, and that it can have two very different outcomes, resulting in resuscitation on the one hand and death on the other. It could therefore be argued that it simply represents an intermediate state in a process of gradual decline towards death, rather then a programme of adaptation to unfavourable conditions (Barer and Harwood, 1999). Admittedly, the ‘‘non-culturable’’ forms under discussion here cannot survive for years, as is the case for profoundly dormant forms like bacterial endospores. However, Streptomyces spores (generally accepted as specialised survival forms) also have limited longevity, and are much more sensitive to environmental fluctuations than are the highly resistant bacterial endospores. In fact, actively growing cells, ‘‘non-culturable’’ forms, Streptomyces spores and endospores can be considered to represent different points on a continuum, where survival of inimical conditions becomes favoured as metabolic activity

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is gradually reduced (see Fig. 1). Long-term survival of endospores is dependent on a complete shutdown of metabolic activity, resulting from the adoption of a state of low water activity (in the laboratory a similar state is achieved artificially by lyophilisation). The limited duration of the TNC state is not therefore a compelling argument against the view we have taken, which is that the TNC state should be regarded as a survival strategy. Another point of general concern to people working in the field is the low reproducibility/high variability of experiments. We suggest that this is mainly related to the fact that there is very substantial heterogeneity of cells in bacterial populations. From our own experience, we know that a slight change of culture conditions can cause dramatic changes in population behaviour (Shleeva et al., 2002). Sometimes these changes are very difficult to control. The best experimental models are uniform populations of ‘‘non-culturable’’ cells. Unfortunately, this ideal is seldom realised; ‘‘non-culturable’’ cells very often represent less than 10% of the total population. In our opinion, this fact does not challenge their existence, nor does it undermine their biological significance, but it certainly complicates attempts to investigate them. The most severe criticism of the significance of the phenomenon as a general survival strategy (if such it is) is the fact that only in a relatively few instances has resuscitation been convincingly demonstrated. There can be many reasons why attempted resuscitation has failed. In some instances the organisms under investigation may be dead, whereas in others inappropriate conditions may have been employed. Finding the right conditions still remains one of the major challenges, since they vary from one organism to another and no generalisations can currently be made. ‘‘Non-culturable’’ cells are very different from culturable cells and finding the appropriate conditions for resuscitation is every bit as difficult as finding the appropriate conditions to support the growth of any previously unknown organism. Like endospores, ‘‘non-culturable’’ cells may be able to resuscitate only in the presence of specific chemical signals. Another parallel is that spore germination and resuscitation from the ‘‘non-culturable’’ state are both stochastic processes depending on many different factors. The production of such specialised forms does not automatically mean that the organism will survive and give rise to progeny at some time in the future; it merely increases the probability that this will happen. The main factor that has to be overcome in order to clarify the status of ‘‘non-culturable’’ cells is ignorance. We still do not know how and why bacteria adopt a ‘‘non-culturable’’ form. Also, we do not know what is happening at the molecular level during the transition to ‘‘nonculturability’’, in the period of ‘‘non-culturability’’ itself, and during the

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subsequent process of resuscitation. Unless we have a much fuller understanding of these matters we cannot decide whether ‘‘non-culturable’’ cells are dormant or whether they are forms of life that are merely undergoing a more or less graceful degradation. In spite of the fact that the phenomenon was first described 20 years ago (Xu et al., 1982) and many experiments have been done since then, we still remain woefully ignorant. The application of the new powerful -omics methods (transcriptomics, proteomics and metabolomics) to organisms of known macro-physiological state may be expected to provide significant new insights in the future.

ACKNOWLEDGEMENTS We thank Obolbek Turapov, Margarita Shleeva, Jim Clegg and Danielle Young for stimulating discussions and access to their unpublished data. We also gratefully acknowledge financial support from the UK BBSRC, the Russian Foundation for Basic Research (grant 00-04-48691), the WHO Global Programme for Vaccines and Immunization, and the Wellcome Trust.

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The Biodiversity of Microbial Cytochromes P450 Steven L. Kelly, David C. Lamb, Colin J. Jackson, Andrew G.S. Warrilow and Diane E. Kelly Wolfson Laboratory of P450 Biodiversity, Institute of Biological Sciences, University of Wales Aberystwyth, Aberystwyth, Wales, UK

ABSTRACT The cytochrome P450 (CYP) superfamily of genes and proteins are well known for their involvement in pharmacology and toxicology, but also increasingly for their importance and diversity in microbes. The extent of diversity has only recently become apparent with the emergence of data from whole genome sequencing projects and the coming years will reveal even more information on the diversity in microbial eukaryotes. This review seeks to describe the historical development of these studies and to highlight the importance of the genes and proteins. CYPs are deeply involved in the development of strategies for deterrence and attraction as well as detoxification. As such, there is intense interest in pathways of secondary metabolism that include CYPs in oxidative tailoring of antibiotics, sometimes influencing potency as bioactive compounds. Further to this is interest in CYPs in metabolism of xenobiotics for use as carbon sources for microbial growth and as biotransformation agents or in bioremediation. CYPs are also current and potential drug targets; compounds inhibiting CYP are antifungal and anti-protozoan agents, and potentially similar compounds may be useful against some bacterial diseases such as tuberculosis. Of note is the diversity of CYP

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requirements within an organism, ranging from Escherichia coli that has no CYPs as in many bacteria, to Mycobacterium smegmatis that has 40 representing 1% of coding genes. The basidiomycete fungus Phanerochaete chrysosporium surprised all when it was found to contain a hundred or more CYPs. The functional genomic investigation of these orphan CYPs is a major challenge for the future.

Abbreviations. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 132 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 132 1.1. Discovery of CYPs in microbial systems . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 1.2. CYP51 (sterol 14a-demethylase), an ancestral CYP form . . . . . . . . . . . . . . . 136 1.3. Bacterial CYPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 1.4. CYP superfamily in Streptomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 143 1.5. CYP superfamily in mycobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150 1.6. CYPs in other bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158 1.7. CYPs in archaebacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 1.8. CYPs in protists . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 1.9. Fungal CYPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163 1.10. Future perspectives for microbial CYP research . . . . . . . . . . . . . . . . . . . . . 174 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 176 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 176

ABBREVIATIONS MTB ER

Mycobacterium tuberculosis endoplasmic reticulum

1. INTRODUCTION Cytochromes P450-dependent monooxygenases (CYPs) constitute a superfamily of haem-containing enzymes that are involved in a wide array of NADPH/NADH-dependent reactions (Guengerich, 1991; Ortiz de Montellano, 1995). Their role in the metabolism of xenobiotic drugs and toxic chemicals has been firmly established in animals where a wealth of information exists. However, we also see an ever-increasing biodiversity of these genes/proteins within microbes. Predominantly, these CYPs are ‘‘orphans’’ with no known function. CYPs also play pivotal roles in the biosynthesis of endogenous compounds such as sterols, steroids, fatty acids

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and prostaglandins and, in microbes, functions are known both in primary and secondary metabolism. Many of these are discussed in this review. The extraordinary biodiversity of the CYP superfamily of enzymes within both prokaryotes and eukaryotes has been revealed from the many completed genome sequencing projects (Nelson, 1999). For example, humans possess 57 CYP genes, the nematode Caenorhabditis elegans, 80, the fruitfly Drosophila melanogaster, 90, and the plant Arabidopsis thaliana, 275. Within the microbial world, CYPs also show variations in distribution: the yeast Saccharomyces cerevisiae, has 3 CYP genes, the white-rot fungus Phanerochaete chrysosporium, 100, the bacterial pathogen, Mycobacterium tuberculosis, 20, whilst Escherichia coli has none (Kelly et al., 2001). The last observation is of interest because genes for cytochrome P450 are not needed in all organisms. Many bacteria and the primitive eukaryote Giardia lamblia appear to have no CYPs based on annotated genomes and probing raw data using the conserved CYP haem-binding domain signature. Furthermore, examination of the primary amino-acid sequence of all CYPs reveals the diverse nature and low sequence homology from one sequence to another. Structural studies do, however, reveal a conservation in the overall fold of the proteins in the superfamily to give a triangular prism structure in which variability is due to features of substrate recognition and binding as well as redox partner binding, (Graham and Peterson, 1999). The uniquely conserved cysteine ligand to the prosthetic haem and flanking conserved residues allow most CYP sequences to be identified (Nelson et al., 1996). An EXXR domain in the K-helix is also observed except for some recent variants in Streptomyces coelicolor (Lamb et al., 2002a). Besides these three defining amino acids, a frequently observed motif involves a threonine in the I-helix associated with oxygen activation and electron transfer. There is a nomenclature for the CYP superfamily that is documented (Nelson et al., 1993, 1996) and family assignments are made by Dr David Nelson. Essentially, CYPs fall into the same family (e.g. CYP52) if they have >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 <20% amino acid identity to CYP101 and CYP102A1.

1.4.2. Roles of CYPs in Streptomycetes sp. The genus Streptomyces produces approximately two-thirds of naturally occurring antibiotics and a wide array of other secondary metabolites. These include numerous antibacterial agents (e.g. erythromycin, tetracycline), anticancer agents (doxorubicin, enediyenes), immunosuppressants (FK506, rapamycin), antiparasitic agents (avermectin, nemadectin), antifungals (amphotericin, griseofulvin, nystatin), cardiovascular agents (lovastatin, compactin) and veterinary products (monensin, tylosin). Such compounds are often referred to as polyketides, natural products containing multiple carbonyl and/or hydroxyl groups each separated by a methylene carbon. Although the final structures of the polyketides are diverse, these molecules share common features in their biosynthetic pathways, and within intermediates formed within these pathways. The impetus for the discovery of new polyketides and enhancing the activity of existing drugs has gathered momentum since infectious diseases continue to be a growing and serious health problem. This is due to a number of reasons including the changing spectrum of the pathogens, drug resistance, immunocompromising treatments and immunodeficiency diseases. The understanding of the molecular basis for the biosynthesis of many complex polyketide antibiotics has only been gained over the past decade and has provided an explanation for the basis of the extensive structural diversity seen amongst this class of naturally occurring molecules. Polyketide diversity is generated by variations in (a) use of different starter and extender units, (b) degrees of reduction, (c) polyketide length, and (d) post-polyketide modification (e.g. hydroxylations, methylations, etc.). In this latter step, it is now generally recognised that enzymes belonging to

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the CYP superfamily play an instrumental role in determining antibiotic potency and efficacy. These steps are stereo- and regiospecific oxidative tailoring of the antibiotics e.g. as classically described for macrolides, such as in the biosynthesis of erythromycin, tylosin, oleadomycin and methymycin. Such CYP reactions occur in the late stages of biosynthesis after formation of the macrolactone ring by polyketide synthase. The biological importance of the resulting hydroxyl and/or epoxide substituents to the biological activity of the compounds is often represented by a significant increase in antibiotic potency following CYP biotransformation. Although cytochromes P450 genes are common in macrolide gene clusters, there are considerable variations in the substrates utilised and the position of oxidation catalysed by these enzymes. This should come as no surprise given the known effect of even a single amino acid change in determining the position of steroid hydroxylation with mammalian CYP forms (Lindberg and Negishi, 1989). The majority of the antibiotics produced by Streptomyces spp. are encoded by clustered genes located contiguously on the linear chromosome of the individual streptomycete species. To date, CYPs have been described in numerous different species of Streptomyces (Table 2), and in the majority of these, the individual CYP described is directly involved in the biosynthesis of the antibiotic produced by that organism.

1.4.3. The CYP Superfamily in the Model Actinomycete, S. coelicolor A3(2) Streptomyces coelicolor has been used extensively for the study of morphological and physiological development of streptomycetes and for the investigation of the genetic control of antibiotic production using the array of molecular tools that have been developed. Recently, eighteen CYP genes dispersed throughout the linear chromosome have been described (Lamb et al., 2002a) (Table 3). In assessing putative function, it is useful to consider sequence comparison to CYPs of known function as well as considering the genetic locus of a CYP. As discussed earlier, we observed that a low-level similarity exists for a S. coelicolor CYP51-like protein (encoded by SC 7E4.20) and we have observed sterol 14-demethylase activity for the protein after heterologous expression (Lamb et al., 2002b). CYP102B1 has closest amino-acid identity with CYP102A1 from Bacillus megaterium, although the latter is fused to a reductase domain. CYP102B1 is the longest bacterial

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Table 2 Streptomycete cytochromes P450 described in databases. Many of the CYPs have been shown to be involved in the biosynthesis of antibiotic. CYP names are annotated at website: www.drnelson.utmem.edu/P450.family.list.html. Streptomyces sp.

CYP identification

S. S. S. S. S. S. S. S. S. S. S. S. S. S. S. S. S. S. S. S. S.

CYP105D1, 105D2, 107F1 CYP105A1, 105B1, 105C1 CYP105A3 CYP105D3 CYP105D4 CYP107C1 CYP105F1, 107N1, 160A1 CYP105K1, 162A1 CYP105L1, 113B1, 154B1 CYP105M1 PikC (PicK) CYP107A1, 107B1 CYP107G1, 122A2, 122A3 CYP107D1 CYP163A1 CYP107R1 CYP129A2, 131A1, 131A2 CYP105H1, 161A1 Orf1, Orf2 TxtC CYP171A1

griseus griseolus carbophilus scerotialus lividans thermotolerans lavendulae tendae fradiae clavuligerus venezuelae erythraea hygroscopius antibioticus spheroides maritimus peucetius noursei nodosus acidscabies avermitilis

Antibiotic produced

Function

complestatin nikkomycin tylosin

Anti-HIV Insecticidal Promotant

pikromycin erythromycin rapamycin oleandomycin novobiocin

Antibacterial Antibacterial Antibacterial Antibacterial Antibacterial

daunorubicin nystatin amphotericin thaxtomin avermectin

Antitumor Antifungal Antifungal Phytotoxin Antiparasitic

CYP encountered so far possessing 527 amino acids whereas most bacterial CYPs are approximately 45kDa (less than 500 amino acids in length). The gene encoding CYP105D5 and surrounding genes had the closest homology to the S. lividans AUD4 gene cluster, a region of DNA that could be amplified within the organism (Schmid et al., 1999). A S. griseus protein homologue, CYP105D1, is associated with the metabolism of a vast array of xenobiotics (Taylor et al., 1999) as were other CYP105s (O’Keefe et al., 1994). As with CYP105D1 of S. griseus and CYP105D4 of S. lividans, the CYP105D5 of S. coelicolor is adjacent to a ferredoxin in the chromosomal locus, but there is not a ferredoxin reductase gene in this operon that is needed to complete CYP catalysis. This CYP is particularly interesting from the viewpoint of a detoxifying role in the soil environment where the bacteria may be exposed to toxic secondary metabolites from live organisms and decaying organic material. Furthermore, CYP105 proteins are important in biotransformations and bioremediation, including their use in manipulating plant herbicide tolerance (O’Keefe et al., 1994).

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Table 3 (a) The CYPome of Streptomyces coelicolor A3(2) listed with CYP family assignments and gene names. The conserved motifs of the CYPs are shown including the region often containing Thr in the I-helix of CYPs as well as the EXXR motif in the Khelix and the haem-binding domain containing the cysteine that acts as the fifth haem ligand of CYPs. (b) The 18 CYPs of S. coelicolor A3(2) together with a list of their closest homologues and the extent of amino-acid identity. (a) Gene name1

CYP name2

I-helix

K-helix

SC7E4.20 SCF43.12 3SCF60.06c SC4C2.21 SCH10.14c SCH63.17 SCE41.08c SCE6.21 SC6D11.13c SC6D11.40c SCE6.20 SCIF3.12 SC6D11.14c SCF55.08c SCI41.09c SC8F11.24c 2SCG58.07 SCF55.07

CYP51 CYP102B1 CYP105D5 CYP105N1 CYP107P1 CYP107T1 CYP107U1 CYP154A1 CYP154C1 CYP155A1 CYP156A1 CYP156B1 CYP157A1 CYP157B1 CYP157C1 CYP158A1 CYP158A2 CYP159A1

274

330

PGSET278 AGHET305 251 AGHET255 249 AGRET253 253 AGHEA257 231 AGHET235 250 AGFET254 242 AGYET246 242 AGHET246 250 AGMVT254 241 AGIEP245 248 AGADP252 240 AGHQP244 255 AQQPT259 259 AAYEA263 245 GGEAV249 242 GGEAV246 234 AGGET238 301

Haem binding motif

EAMR333 ESLR361 290 ELMR293 288 ELLR291 292 ELMR295 270 ESLR273 294 ELLR297 281 ETLR284 281 ETLR284 290 ELLR293 272 STVR275 269 EYAR272 279 EVLW282 293 EVLW296 297 EQSLW301 283 ELLR286 280 ELLR283 273 ETLR276 358

403

FSAGKRKCPS412 FGTGARACIG442 354 FGFGIHQCLG363 353 FGYGVHQCVG362 356 FSAGIHYCIG365 335 FGHGPHHCLG344 359 YGHGIHYCLG368 347 FGHGVHFCLG356 348 FGHGPHVCPG357 361 FGDGPHRCPG370 344 WSTGPHTCPA353 344 WSAGPHACPS353 342 FGHGEHRCPF351 355 FSNGEHRCPY364 361 FGGGPHECPG370 349 YGNGHHFCTG358 346 FGFGPHYCPG355 347 FALGRHFCVG356 433

Match in the database3

(b) Gene name1

CYP name2

Amino acid, no.

Species

CYP name

Protein identifier

% identity

Amino acid overlap

SC7E4.20

CYP51

461

Streptomyces avermitilis Mycobacterium tuberculosis Bacillus megaterium Bacillus subtilis Streptomyces lividans Streptomyces griseus Streptomyces griseolus Streptomyces lividans

CYP171A1

Q9S0R5

32.4

429

CYP51

P77901

26.8

421

CYP102A1

P14779

42.8

488

– CYP105D4

gi|16077792 O85697

42.2 97.5

502 405

CYP105D1

P26911

67.5

412

CYP105A1

P18326

45.5

407

CYP105D4

O85697

44.3

411

SC F43.12

CYP102B1

527

SCF60.06c

CYP105D5

412

SC4C2.21

CYP105N1

411

(continued )

148 Table 3

STEVEN L. KELLY et al. Cont. Match in the database3

Gene name1

CYP name2

Amino acid, no.

Species

CYP name

Protein identifier

% identity

Amino acid overlap

SCH10.14c

CYP107P1

411

Streptomyces coelicolor Streptomyces erythraeus Streptomyces venezuelae Streptomyces erythraeus Streptomyces coelicolor Actinomadura hibisca Streptomyces fradiae Streptomyces coelicolor Streptomyces coelicolor Streptomyces erythraeus Brevibacterium linens Streptomyces tendae Streptomyces coelicolor Streptomyces griseus Streptomyces coelicolor Streptomyces griseus Streptomyces coelicolor Streptomyces griseus Streptomyces coelicolor Streptomyces griseus Streptomyces griseus Streptomyces coelicolor

CYP157C1

Q9RJ75

40.3

426

CYP107B1

P33271

39.2

406

CYP107L1

O87605

51.5

394

CYP107B1

P33271

46.9

373

CYP107P1

Q9X8Q3

40.4

426

CYP107M1

O32460

40.0

434

CYP154B1

Q9XCC6

50.9

409

CYP154C1

Q9L142

42.1

411

CYP154A1

Q9KZR7

42.1

411

CYP107B1

P33271

38.4

409

–

Q9KK77

38.8

408

CYP162A1

Q9L465

27.9

394

CYP156B1

gi|13872774

43.2

410

ORF5

Q9AJP0

31.3

403

CYP156A1

Q9KZR8

43.2

410

ORF5

Q9AJP0

34.7

452

CYP157B1

Q9RJQ6

51.7

404

ORF5

Q9AJP0

41.4

406

CYP157A1

Q9L141

51.7

404

ORF5

Q9AJP0

41.3

421

ORF5

Q9AJP0

50.8

519

CYP157A1

Q9L141

39.8

400

SCH63.17

SCE41.08c

SCE6.21

SC6D11.13c

SC6D11.40c

SCE6.20

SCIF3.12

SC6D11.14c

SCF55.08c

SCI41.09c

CYP107T1

CYP107U1

CYP154A1

CYP154C1

CYP155A1

CYP156A1

CYP156B1

CYP157A1

CYP157B1

CYP157C1

394

433

408

407

423

410

447

413

420

498

149

THE BIODIVERSITY OF MICROBIAL CYTOCHROMES P450 Table 3

Cont. Match in the database3

Gene name1

CYP name2

Amino acid, no.

Species

CYP name

Protein identifier

% identity

Amino acid overlap

SC8F11.24c

CYP158A1

407

Streptomyces coelicolor Streptomyces avermitilis Streptomyces avermitilis Streptomyces coelicolor Bacillus subtilis M. tuberculosis

CYP158A2

Q9FCA6

61.2

407

–

gi|15824133

58.7

407

–

gi|15824133

81.4

404

CYP158A1

Q9KZF5

61.2

407

CYPX CYP123

O34926 P77902

41.3 33.3

407 403

2SCG58.07

SCF55.07

CYP158A2

CYP159A1

404

407

1

Gene name as annotated at website: http://www.sanger.ac.uk/Projects/S_coelicolor/ CYP names as annotated at website: http://drnelson.utmem.edu/CytochromeP450.html 3 Database search at FASTA website: http://fasta.birch.virginia.edu 2

CYP107U1, CYP107P1 and CYP107T1 are three CYP107 genes in the S. coelicolor genome. Amino-acid sequence comparisons reveal the first has homology to CYP113 and the latter to cytochrome P450 EryF, an enzyme in antibiotic production. Consequently, these proteins may also participate in secondary metabolism. CYP155A1 has only low-level homology to other CYPs. The genes encoding CYP156A1 and CYP154A1 are adjacent to each other on the same DNA strand with only 12 bp separation. Such close association points to a related function with polycistronic translation. Homology to CYPs with known function, such as in tylosin biosynthesis, points again to an important secondary metabolism function. The genes CYP154C1 and CYP157A1 also lie adjacent to each other on the chromosome and may reflect a gene duplication event and a putative operon involved in secondary metabolism. As with CYP154A1 and CYP156A1, the putative operon structure is conserved such that it consists of genes for a putative sensor kinase/membrane protein, two orphan ORFs, a putative ATP binding protein and the two CYPs. This structure has been called a conservon (Bentley et al., 2002). CYP157C1 has homology to known CYP proteins that are orphans with little clue as to function, but again is associated with a conservon as is CYP157B1. Many of the S. coelicolor CYPs have closest identity with other CYPs of S. coelicolor A3(2) reflecting the recent evolution of forms by gene duplication events. Genes encoding CYP159A1 and CYP157B1 are also

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adjacent, but lie on opposite strands. Comparison of protein sequences indicates different functions for each. CYP158A2 has 81.4% identity to a CYP from S. avermitilis and the genetic organisation at the corresponding gene locus is similar to the S. avermitilis chromosome. Both loci include an adjacent Type III polyketide synthase suggesting a likely role in secondary metabolism. CYP158A1 is also 58.7% identical to this S. avermitilis CYP. CYP105N1 is similar to other CYP105s involved in xenobiotic metabolism, but is encoded in a likely operon of secondary metabolism. This includes ORFs for probable nonribosomal peptide synthase, a probable thioesterase and probable ABC transporter and so divergence of function within CYP105s may occur, with xenobiotic and biosynthetic functions for different forms. The identification of eighteen novel CYPs in the genome of S. coelicolor A3(2) represented the first systematic approach to production of all the CYPs of an organism (the CYPome) for biochemical, structural and genetic characterisation (Lamb et al., 2002a). As we have emphasised here, CYPs are remarkably diverse oxygenation catalysts found throughout nature and have the potential to be manipulated for the production of new antibiotics. Deciphering the biology of the CYPs of S. coelicolor can be considered as a suitable model for understanding the role of CYPs in the biosynthesis of such important medicinal compounds within the genus.

1.5. CYP Superfamily in Mycobacteria The order Actinomycetales consists largely of saprophytic, soil-inhabiting gram-positive bacteria that have a high DNA G þ C content and often show a branched or mycelial-like growth habit. Besides the streptomycetes dealt with above, four other genera, Actinomyces, Corynebacterium, Mycobacterium and Nocardia are of primary medical or commercial importance. In the genus Mycobacterium, complete genome sequence data are available for M. bovis and M. leprae and for two strains of M. tuberculosis. In addition, sequencing is in progress for the genomes of M. smegmatis, M. avium, M. avium subsp. paratuberculosis and M. marinum. These resources have provided a unique opportunity to compare the CYP systems of diverse members of the same taxonomic order. Perhaps the most surprising observation is the enormous CYP biodiversity in the mycobacteria. The numbers of CYPs present in individual mycobacterial species varies widely, with relatively few CYPs apparently conserved between species or between genera. M. tuberculosis and S. coelicolor have 20 and 18 CYPs respectively, none of which seem to be highly conserved between

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Table 4 The CYPome of M. tuberculosis compared to another strain with genomic information and the presence or absence of these forms in M. bovis, M. avium, M. smegmatis. Also shown is whether a pseudogene for the M. tuberculosis CYP exists in M. leprae. ML0447 and ML2159 (M. leprae ORF identifier) are identical pseudogenes in M. leprae that resemble S. coelicolor A3(2) CYP102B1 and ML 2088* is the only putative functional CYP of M. leprae. P450

CYP number

H37Rv CDC1441,210

M. bovis

M. avium

M. smegmatis

M. leprae

Rv0136 Rv0327c Rv0568 Rv0764c Rv0766c Rv0778 Rv1256c Rv1394c Rv1666c Rv1777 Rv1785c Rv1880c Rv2276 Rv2268c Rv2276 Rv3059 Rv3121 Rv3518c Rv3545c Rv3685c

CYP138 CYP135A1 CYP135B1 CYP51 CYP123 CYP126 CYP130 CYP132 CYP139 CYP144 CYP143 CYP140 CYP124 CYP128 CYP121 CYP136 CYP141 CYP142 CYP125 CYP137 CYP102 CYP102 CYP164

þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ

þ þ þ þ þ þ

þ

þ

ML2684

þ þ þ þ

þ þ þ þ þ

ML2229 ML1102

þ þ þ þ þ þ þ þ þ þ þ þ

þ þ þ þ þ

þ þ

ML1237 ML1185 ML1542 ML2033 ML1787

þ

þ

ML1742

þ þ

þ þ

ML2024

þ

ML0447 ML2159 ML2088*

þ

these species. Only 12 of the 20 M. tuberculosis CYPs have potential functional homologues in M. smegmatis, an organism that has forty putative CYPs and at least two CYP pseudogenes (Table 4, Fig. 6). The related species M. avium and M. avium ssp paratuberculosis similarly appear to have approximately forty CYPs, and again, only some of these are present in other mycobacteria (Jackson C.J. and Kelly S.L., unpublished observation). These observations are based on the incomplete genome sequence but, with approximately 3800 genes in M. tuberculosis, these mycobacterial CYPomes are likely to represent 1% of the genes of some mycobacteria. M. leprae has only a single CYP and, remarkably, a homologue of this putative protein is not present in M. tuberculosis, but is found in the more distantly related M. smegmatis (60% identity, 75% similarity over the whole

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Figure 6 A phylogram of CYPs from mycobacteria including the twenty named CYPs of M. tuberculosis and the one CYP of M. leprae. The closest homologue of the last is in M. smegmatis with 67% identity over 415 amino acids. The M. smegmatis CYPs are unclassified and given our assignment in roman numerals except where they fall into a known family. Mtb: M. tubercutosis; Mlep: M. leprae; Msmeg: M. smegmatis.

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protein). The closest homologue in M. tuberculosis is CYP140 with 41% identity over only a part of the protein (334 amino acids) with twelve gaps. The leprosy bacillus also has eleven degenerate CYP pseudogenes, and counterparts of nine of these are present as entire genes in M. tuberculosis (MTB). Intriguingly, the two non-MTB pseudogenes are exact direct repeats of each other (ML0447 and ML2159 in the M. leprae annotation). The predicted translation product of these identical pseudogenes is most similar to CYP102B1 of S. coelicolor and to CYP102A1, the P450 BM-3 protein of B. megaterium. This CYP was, therefore, probably present in the last common ancestor of mycobacteria and streptomycetes.

1.5.1. Eukaryote-like CYPs in Mycobacteria The 20 MTB CYPs have been categorised by Nelson (1999) as being either ‘eukaryote-like’ (5 CYPs ) or ‘prokaryote-like’ (15 CYPs). The former are CYP138 (gene locus Rv0136); CYP51 (locus Rv0764c); CYP132 (locus Rv1394c); CYP139 (locus Rv1666c) and CYP136 (locus Rv3059). The first of these, CYP138, belongs to a cluster of four MTB CYPs that are related and share a relatively recent common ancestor (CYPs 135A1, CYP135B1 and CYP137). Although these genes are an outgroup in the general prokaryote CYP tree (see Nelson above), CYP138 certainly belongs with it. CYP132, CYP136, CYP139 and CYP51 share a common branch in the tree of MTB CYPs (Fig. 6), and are also similar to an unusual S. coelicolor CYP51. Interestingly, among the CYP51-like genes is included S. coelicolor CYP102B1. Differentiation of CYP51 and CYP102 enzymes with distinct functions may have occurred before the divergence of Streptomyces spp. and Mycobacterium spp. The latter (CYP102B1) has been lost in mycobacteria studied so far with the exception of M. leprae where two pseudogenes can be detected. The CYP51 homologue may have undergone duplication/loss and refinement in MTB and other mycobacteria. Of course, horizontal transfer and convergent evolution may also play a role here.

1.5.2. Evolutionary scenario for CYP51-like CYPs in mycobacteria We suggest above that a CYP51-like P450 was present in an ancestral actinomycete, and that this is related to CYP102B1. In MTB, CYP132

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and CYP139 cluster most closely with the S. coelicolor CYP51 and CYP102, and have probably arisen by duplication events at different stages in the evolution of individual mycobacterial species. The same duplication process has probably also produced CYP136. CYP136 is present in all mycobacteria, including M. leprae (as a pseudogene) whilst CYP132 is only present in the highly related MTB and M. bovis. CYP139 is absent from the fast-growing M. smegmatis, but is present in all slowgrowing mycobacteria (Table 3).

1.5.3. MTB CYP51 May Have Arisen by Lateral Transfer From an Ancestor of Methylococcus capsulatus MTB CYP51 shows a number of unique characteristics (Bellamine et al., 1999) but, nevertheless, is a eukaryotic CYP51 homologue, most similar to plant obtusifoliol 14a-demethylase. However, MTB CYP51 forms part of a six gene operon, which is universally present in both fast- and slow-growing mycobacteria with the exception of M. leprae. A second MTB CYP, CYP136, is similar to MTB CYP51 and is also present in all mycobacteria, including as a pseudogene in M. leprae (ML1742). It appears that in leprosy, the entire CYP51 operon has been lost by a deletion event, as neither the second CYP gene in the operon, CYP123 (Rv766c) or any of the other operon genes have M. leprae pseudogenes. Immediately downstream of MTB CYP51 is a ferredoxin gene. We have recently discovered that the gram-negative proteobacterium and methylotroph, Methylococcus capsulatus, contains a CYP51 fused to a ferredoxin (MCCYP51FX, Fig. 7), and that both domains of the protein show very high identity with separate counterparts in MTB that are encoded by adjacent genes on the chromosome. M. capsulatus produces sterols (Bird et al., 1971; Bouvier et al., 1976), and the genome contains other homologues of genes from the eukaryotic sterol biosynthesis pathway in addition to CYP51 (Jackson C.J. and Kelly S.L., personal observation). M. capsulatus CYP51FX does not form part of an operon and logically its role in this organism would be as the 14a-demethylase component of the sterol biosynthesis pathway. Indeed such an activity has been confirmed (Jackson et al., 2002). There are some other open-reading frames in M. tuberculosis containing domains with significant identity/similarity to sterol biosynthesis proteins including to a late stage enzyme, sterol C5-desaturase (Cole et al., 1998; Lamb et al., 1998a), but their roles are not defined. We are evaluating

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Figure 7 Diagram of the genetic locus containing the M. capsulatus CYP51 that has a ferredoxin domain fused at the C-terminus and depicting the alanine-rich linker region between the CYP and ferredoxin domain.

Figure 8 The predicted conserved operon containing M. tuberculosis CYP51 together with the gene assignments and predicted ORF functions.

their functions, but they have so far failed to genetically complement yeast sterol mutants and no known sterol has been detected in M. smegmatis from minimal medium, lacking trace sterol (Jackson C.J., et al., manuscript in preparation). The presence of MTB CYP51 in an operon with other genes that have no known role in sterol production (Fig. 8) further suggests that the function of MTB CYP51 is different from that of MCCYP51FX. A possible role for the MTB CYP51 operon could be in maintaining the structural architecture of the mycobacterial cell wall or sterol or steroid modifications. The leprosy bacillus is an obligate intracellular parasite, in which many of the genes involved in envelope biogenesis in MTB are absent. It has been suggested that many of these

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deleted genes, encoding biosynthetic routes to polyketides, lipids and aromatic compounds, produce substrates for cytochromes P450 (Cole et al., 2001). The loss of the MTB CYP51 operon and many other MTB CYPs in M. leprae may, therefore, be a function of the coincident deletion of genes required for envelope-related compounds. We propose that an ancestral copy of the genes now encoding the CYP51/ferredoxin fusion (MCCYP51FX) identified in M. capsulatus has been transferred horizontally to a mycobacterial ancestor. The evidence for this transfer is the identical gene order and the high degree of amino acid sequence similarity encoded by the two genes from these very different species. There is 49% identity for the CYP51 (Fig. 9) and 42% for the ferredoxin and each have the strongest similarities to each other. The alternative possibility, namely that similar contiguous CYP51/FX genes of presumably different function arose independently in such evolutionarily diverse species, seems highly unlikely. It is implied that, during evolution of the form observed now in M. capsulatus, there was loss of the termination triplet of the CYP51 and read-through and evolution of a fusion polypeptide encoding also the ferredoxin domain. The transfer of CYP51 from M. capsulatus to mycobacteria would imply that MCCYP51FX could represent an ‘ancestral’ CYP51 domain, and that CYP51 originated during the evolution of a prokaryote sterol biosynthesis pathway. This pathway was maintained in the early stages of eukaryotic evolution, although some primitive eukaryotes may have lost this biosynthetic pathway as it also seems they lost mitochondria (Roger and Silberman, 2002).

1.5.4. Ancestral CYP Forms in Mycobacteria Another possible ancestral CYP within actinomycetes is CYP107. Homologues of a CYP107-like CYP (Rv1880c) are present in most mycobacteria and as pseudogenes in M. leprae. These genes cluster with CYP107P1, CYP107T1 and CYP107U1 of S. coelicolor. This family of CYPs is diverse, merging with the CYP105 family and others. Functional data to date suggest that the CYP107s are involved in secondary metabolism, but only a small fraction of them have had their role revealed. In the consideration of ancestral CYP function in actinomycetes, we recently also detected a CYP125 in both mycobacteria and Streptomyces avermitilis and from the genetic locus, this appears to be involved in lipid metabolism (Lamb, D.C., Ikeda, H., Waterman, M.R. and Kelly, S.L., unpublished).

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Figure 9 Amino acid alignment of the P450 domain of the M. capsulatus (Mcap) CYP51 with other CYP51s from M. tuberculosis (MTB), Arabidopsis thaliana (Atha), human and from Candida albicans. The bacterial forms are truncated, lacking a N-terminal membrane anchor and the cysteine forming the fifth ligand with haem is at 394 in the Mcap sequence.

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1.5.5. CYPs and Anti-Mycobacterial Activity The development of therapeutic azole inhibitors began with the antifungal compounds that target CYP51. Such compounds were made for use in agriculture and medicine and new compounds continue to be developed and are discussed in the section below on fungi. Given the diversity of CYPs including CYP51 in mycobacteria it would be extraordinary if some CYPs proved not to be good drug targets for the treatment of M. tuberculosis. The study of the sensitivity of mycobacteria to azole antifungals has been undertaken and many of the topical antifungal drugs have potent activity against M. smegmatis at concentrations that were effective against Candida albicans (Jackson et al., 2000). A number of drugs have been evaluated including topical mycotics, such as miconazole and econazole, and systemic drugs such as ketoconazole and fluconazole (Fig. 10). Unfortunately, the systemic drugs, especially the best current antifungal fluconazole, had low activity against M. bovis BCG (Kelly et al., 2001), a close relation of M. tuberculosis. High activity has been seen against some other bacteria including Mycobacterium chelonei that causes skin infections that are recalcitrant to current treatment (Table 5). Binding of azoles to mycobacterial CYPs has been observed, including to CYP51 (Bellamine et al., 1999) whose azole-bound structure has been determined (Podust et al., 2001). The mode of CYP inhibition by azole compounds involves co-ordination to the haem as a sixth ligand via N-3 of imidazole or N-4 of triazole compounds and binding of the varied N1 substituent moieties to the protein active site in a way that differentiates between CYPs of different structure. Interaction of azoles with other CYP forms such as CYP125 (Perally S., et al., unpublished) and CYP121 (McLean et al., 2002) has also been reported. As with antifungal compounds, agents specific for M. tuberculosis will have to target essential CYP function and give a selective effect against that CYP, without serious drug/drug interactions or deleterious effect on human CYP activity.

1.6. CYPs in Other Bacteria A large number of other bacterial CYPs have been described. Of particular interest are CYPs that are natural fusion proteins with their redox partners. Such proteins will provide excellent models for probing P450 electron transfer, substrate specificity and catalysis, as well as offering the prospect of a basis for improved biotechnological processes.

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Figure 10 Chemical structures of azole antifungals including the triazole drug fluconazole and tebuconazole (an agrochemical) and the imidazoles drugs miconazole, econazole, clotrimazole and ketoconazole. Imidazole is also shown and requires N-1 substituent groups to enable effective binding to CYP51 although the azole group itself is responsible for forming the sixth ligand with haem.

This is because coupling of CYP and reductase partners may be limiting in the catalytic turnover. Two recent examples of such proteins (see below) should provide major advances in understanding CYP/reductase structure/ function.

1.6.1. The First Naturally Occurring Bacterial CYP/Ferredoxin Fusion Protein is Isolated From M. capsulatus Examination of the genome sequence data for the gram-negative bacterium M. capsulatus, a bacterial species known to produce sterol, revealed the

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Table 5 The minimum inhibitory concentrations (MICs), measured by a broth dilution method, for several azole compounds plus amphotericin B against C. albicans and Mycobacterium spp. Clotrimazole, econazole and miconazole were highly inhibitory against M. smegmatis, and also showed good activity against pathogenic mycobacteria such as M. chelonei and M. fortuitum. Skin infections due to these two species are often resistant to treatment with standard antimycobacterial agents. Less activity was seen with these compounds against the slow growing species M. bovis BCG, although some growth reduction was evident. ND denote no assay information was available. Fluconazole and imidazole have no useful activity against mycobacteria, whilst ketoconazole and tebuconazole are only weakly effective in reducing growth. MIC mg/ml Species Strain

Azole Clotrimazole Econazole Fluconazole Ketoconazole Imidazole Miconazole Tebuconazole Amphotericin B

C. albicans 505

M. smegmatis 700084

M. smegmatis 13116

M. fortuitum 01/2341

M. chelonei LRT 6680

M. bovis BCG

<2 <2 8 <2 > 256 4 4 <2

<2 <2 > 256 16 > 256 <2 32 > 256

<2 <2 > 256 16 > 256 <2 32 > 256

4 8 > 256 > 256 ND 8 64 ND

<2 <2 > 256 8 ND <2 ND ND

16 8 > 256 32 ND 8 ND ND

presence of a single CYP with significant similarity in the amino acid sequence to CYP51, particularly to a form in M. tuberculosis. Interestingly, this M. capsulatus CYP51 protein represents a new class of CYP consisting of the CYP domain naturally fused to a ferredoxin domain at the Cterminus via an alanine-rich linker (Fig. 7). Expression of the M. capsulatus MCCYP51FX fusion in E. coli yielded a P450, which, when purified to homogeneity, had the predicted molecular mass, ca 62 kDa. Sterol 14ademethylase activity was demonstrated (0.24 nmol lanosterol metabolised/ min/nmol MCCYP51FX fusion) and this was dependent upon the presence of ferredoxin reductase and NADPH. The requirement for sterol biosynthesis in bacteria and the evolutionary origin of this CYP, normally associated with eukaryotes, is open to much conjecture as described above. The flanking genes for MCCYP51Fx are, upsteam, a potential thiC-type gene, which in B. subtilis, is involved in thiamine biosynthesis (Zhang et al., 1997) and, downstream, an orphan ORF. Neither flanking genes are found adjacent to CYP51 in the genome of

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M. tuberculosis. However, conservation of synteny from gram-negative to gram-positive bacteria might not be expected.

1.6.2. A Fusion CYP Protein from Rhodococcus Another novel CYP protein recently discovered in a Rhodococcus sp. consisted of a reductase domain at the C-terminus similar to dioxygenasereductase proteins (Roberts et al., 2002). Here, the reductase domain occurred after the CYP domain and contained flavin mononucleotide and NADH-binding domains with a C-terminal [2Fe2S] ferredoxin center at the end of the polypeptide. This catalytically self-sufficient CYP has many properties that could be useful to develop and enhance as a basis for biocatalytic processes. These new classes of CYPs described here are represented in Figure 11 and, together with P450nor (see below), represent five classes/ types of CYPs found in microbes to date.

1.7. CYPs in Archaebacteria CYPs have also been discovered in some archaebacteria. Whether these were acquired later in evolution (since CYPs generally require oxygen for catalytic function) or whether some ‘‘electron transfer’’ or reductive function might be detected in these organisms is not clear. What is of applied interest is the ability of such CYPs to maintain structural and functional integrity under conditions that could be useful in biotechnological processes such as operation at high temperature. The structure of one archaebacterial CYP119 from Sulfobolus solfataricus has been obtained and the temperature stability of the protein has been ascribed to extensive clustering of aromatic residues (Yano et al., 2000). The structure of a second thermophilic cytochrome P450, CYP175A1 from the thermophilic bacterium Thermus thermophilus HB27, has recently been solved to 1.8A˚ resolution. The CYP175A1 structure lacks the large aromatic network found in CYP119. The primary difference between CYP175A1 and its mesophile counterparts is the investment of charged residues into salt-link networks at the expense of single charge–charge interactions. Additional factors involved in the increase of thermal stability are a decrease in the overall size, especially shortening of loops and connecting regions, and

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Figure 11 The different classes of microbial CYPs are depicted including typical bacterial and mitochondrial three-component systems (class I); typical eukaryotic ERassociated two-component systems (class IIa), P450 BM3 fusion protein and a similar membrane-bound fungal CYP505 form (classIIb); P450 cin, a similar, but threecomponent enzyme system (class IIc); a soluble single-component fungal CYP55 (class III); fusion protein from Rhodococcus exhibiting a fusion protein of CYP, reductase and ferredoxin domains (class IV) and fusion protein consisting of CYP with a ferredoxin fusion at the C-terminus as found in M. capsulatus CYP51 (class V).

a decrease in the number of labile residues such as Asn, Gln and Cys (Yano et al., 2002).

1.8. CYPs in Protists For those interested in CYP evolution, the protists are a focus of growing attention, but information is still lacking. These eukaryotic micro-organisms are very diverse, distinct from the plant, animal and fungal kingdoms and yield clues to early evolution of eukaryotes and eukaryotic CYPs. Given the diversity of CYPs, many of these organisms will contain many tens of families and others seemingly none, as in the bacteria. The ability to synthesise sterols requires CYP51 and many will contain this. Given the potential for use of CYP51 inhibitors against pathogenic protists, such as trypanosomes and Leishmania, this will be an area of close scrutiny. The completed genome of Plasmodium falciparum contained no CYPs (Gardner et al., 2002) and so is a parasitic sterol auxotroph. The number of CYPs in many protists may be large, as is already indicated by the genome project on Dictyostelium discoidum (>42 CYPs so far, http://drnelson.utmem.edu/nelsonhomepage.html).

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This multiplicity probably reflects adaptation to niches and the use of CYPs in biochemical mechanisms of deterrence, attraction and detoxification. Many protists, like fungi and some algae, contain ergosterol which is likely to confer osmotic resilience on unicellular eukaryotes (Kelly et al., 1997a). As ergosterol biosynthesis requires the further activity of CYP61, a sterol 22desaturase originally found in yeast (Kelly et al., 1995a), this CYP may also be present in different kingdoms of life as with CYP51. Another form, CYP98, has been identified in both diatoms and plants (http://drnelson.utmem.edu/nelsonhomepage.html), but the function of this CYP is not known. Within the protists there will be many new CYPs involved in secondary metabolism, CYPs representing drug targets and resistance factors. This area is only just beginning to be unraveled, but it is certain it will be a focus of much future research.

1.9. Fungal CYPs Fungal genome sequences will invigorate fungal CYP studies. Fungal CYPs are generally class II forms, located in the endoplasmic reticulum. The first eukaryotic genome sequenced was that of S. cerevisiae (Goffeau et al., 1996). Besides the known target of the azole antifungals, CYP51, the genome was also known to contain a CYP producing dityrosine for the spore wall of the meiotic tetrads (CYP56, Briza et al., 1994). In mitotic cells growing vegetatively, the presence of a second CYP protein was suspected, encoding sterol
22-desaturase. This was based on inhibitor studies (Hata et al., 1981, 1983) and indeed CYP was still detectable in a CYP51-disrupted strain (Kelly et al., 1993). Using amino-acid sequence information of purified protein the first proteomic study of orphan CYP function was possible linking this information and known activity to a gene sequence on yeast chromosome XIII (Kelly et al., 1995a, 1997b). This gene called CYP61 was also isolated using traditional complementation (Skaggs et al., 1996). The total CYPome of yeast comprised only three CYPs while the fission yeast Schizosaccharomyces pombe was even more minimalist, containing just CYP51 and CYP61. Besides the biotechnological importance of ergosterol, and hence CYP51 and CYP61, in conferring osmotolerance during fermentation, other biotechnological importance for fungal CYPs has been discovered. Studies on fungal CYPs began (unknowingly) with some of the earliest biotransformations for the production of corticosteroids. Rumours during the Second World War that the Luftwaffe was administering corticosteroids to pilots stimulated steroid research and the therapeutic value of cortisone

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became apparent (Smith et al., 1993). The ability of fungi to perform stereo-and regiospecific hydroxylations of steroids is now well recognised and the importance of the steroid 11-hydroxylation was a costly step, overcome in 1952 by biotransformations by a number of species including Rhizopus nigricans and Aspergillus niger (for review Smith et al., 1993). Purification of fungal steroid hydroxylase CYPs has been reported (for example, Breskvar et al., 1987), but CYP purification from filamentous fungi is fraught with low yield and protein instability problems, as well as the likely problems of CYP multiplicity and co-purification. It is surprising that the CYP genes responsible for filamentous fungal steroid biotransformations are still unidentified as are the forms responsible for efficient metabolism of polycyclic aromatic hydrocarbons that are of potential value in bioremediation (Masaphy et al., 1999). The ability of some yeasts to grow on alkanes as single carbon sources is of biotechnological interest. Indeed, British Petroleum had set up an industrial scale facility to convert oil to single-cell protein utilizing Candida tropicalis. However, with increased oil prices this became uneconomical. Biochemical data had indicated that CYP was responsible for the initial oxidation (Lebeault et al., 1971) and molecular cloning of the CYP concerned in this and other yeasts showed that a new family, CYP52, was responsible (Sanglard and Loper, 1989). This family is very diverse with numerous forms present within a single species and CYP52 forms have been identified in many yeast including Candida maltosa (Ohkuma et al., 1998) and Yarrowia lipolytica (Iida et al., 1998). In yeast genetic nomenclature, these CYPs are encoded by ALK genes. In the study by Ohkuma et al., (1998), the effect of knocking out the structurally related ALK genes was investigated. Eight were detected in C. maltosa, but loss of four prevented growth on n-alkane. The reason for such functional redundancy remains unclear. Besides the forms isolated by traditional molecular genetic approaches, there is also the newly emerging information from the uncompleted annotation of the C. albicans genome (http://www-sequence.stanford.edu/group/candida/). Surprisingly, this pathogenic yeast is not associated with an environmental niche requiring alkane utilisation yet has numerous ALK genes, as does C. tropicalis, which can also be pathogenic. Retention of these genes suggests a role in lipid metabolism/utilisation from the host or an unrecognised alternative environmental niche requiring lipid/alkane metabolism. The third CYP family to be identified in fungi was from Aspergillus niger. The new family, designated CYP53, was detected through studies on the para-hydroxylation of benzoate (Van Gorcom et al., 1990). A further family, CYP54, has an unknown function, but was cloned as a cycloheximide-inducible protein from Neurospora crassa (Attar et al., 1989).

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CYP55 (P450nor) is an extremely interesting soluble CYP form from Fusarium oxysporum as it undertakes nitrate/nitrite reduction without the need for a CYP-reductase (CPR) or a requirement for molecular oxygen. As such, it fell into a new class of CYP, the first to be discovered since the CYP102A1 (P450 BM3) fusion protein. Its activity independent of CPR is similar to the allene oxide synthase of plants and the structure of CYP55A1 was the first eukaryotic CYP structure to be determined (Park et al., 1997). CYP56 has already been described above (Briza et al., 1994) as a form generating dityrosine for the spore wall of S. cerevisiae and is so far restricted to this yeast. CYP57 is a particularly interesting CYP in the context of showing the roles of CYPs in biochemical deterrence and/or attraction. It was identified in Nectria heamatococca (Han et al., 2001) as a gene on a supernumary chromosome and had activity in the demethylation (detoxification) of pisatin produced by the plant host for this pathogen. CYP57 is one of six pea pathogenicity (PEP) genes located in a gene cluster. Other fungal CYPs have been assigned family numbers and a novel Fusarium oxysporum CYP similar in structure to CYP102A1 has been found consisting of a CYP domain fused to a CPR. We would place this CYP in class IIb of the nomenclature as it resembles the classical class II eukaryotic forms along with CYP102A1. This form, CYP505, is membrane-bound and also metabolises fatty acids (Kitazume et al., 2000). Numerous other forms have no known role and have emerged from genome projects e.g. CYP501 from Candida albicans and others have emerged from experimental studies such as those revealing the importance of CYP in controlling penicillin production in the presence of phenylacetate (Mingot et al., 1999). Once again, genetic loci may give vital clues to function. For example, some CYPs have been found to play roles in mycotoxin and aflatoxin biosynthesis in a cluster of functionally related genes (CYP 58,59,64; Hohn et al., 1995; Yu et al., 1998, 2000). This has also been found for CYPs implicated in giberellin biosynthesis in Giberella fujikuroi (CYP68, Tudzynski et al., 2001) and for CYP involved in the biosynthesis of the indole-diterpene paxilline by Penicillium paxilli (Young et al., 2001). With regard to identifying function, it is likely that many forms will be integrated into the secondary metabolic pathways present in fungi and these will be systematically revealed within the genomes that are beginning to emerge.

1.9.1. Phanerochaete Chrysosporium – Unexpected Levels of CYP Biodiversity Despite S. cerevisiae containing only three CYPs, it was always anticipated that filamentous fungi would contain more CYPs due to the array of

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secondary metabolism and due to their capabilities in biotransformations and bioremediation. However, the genome of Neurospora crassa revealed a CYPome with 38 forms (http://drnelson.utmem.edu/nelsonhomepage.html). An even bigger surprise was the greater biodiversity found within the genome of the basidiomycete higher fungus P. chrysosporium. P. chrysosporium is a white-rot fungus found in the environment growing on dead wood and amongst leaf litter. It is the most widely studied fungus in relation to bioremediation of organic pollutants and CYP has been implicated in that process, besides the laccase and peroxidase enzymes that have received most focus (Masaphy et al., 1996; Kullman and Matsumura, 1996). The genome of P. chrysosporium was recently sequenced by the U.S. Department of Energy Joint Genome Institute (http:// www.jgi.doe.gov/programs/whiterot.htm). Although the genome has not yet been annotated, Dr David Nelson (http://drnelson.utmem.edu/CytochromeP450.html) has searched the released DNA scaffolds using the conserved CYP haem-binding motif. The P. chrysosporium genome was found to contain over 123 CYP haem motifs and one CPR gene corresponding to that describe earlier by Yadav and Loper (2000). The CPR has been expressed and preliminary characterisation undertaken (Warrilow et al., 2002). 78 full-length CYP genes were identified from the P. chrysosporium genome, but only five of these CYP genes can currently be ascribed names and only two function. These are CYP51, CYP61, CYP63A1, CYP63A2 and CYP63A3 where the function of CYP63s is unclear (Kullman and Matsumura, 1997). Considerable interest lies in how a single CYP reductase can interact with such a large number of CYP forms representing approximately 1% of genes. In plants, the reductase (CPR) is present in more than one form (Urban et al., 1997) and the relatedness of the P. chrysosporium CPR to other eukaryotic enzymes is shown in Figure 12. P. chrysosporium CPR resembles most closely CPRs from Coriolus vesicolor (Q8X1W0), Rhodotorula minuta (Q9C498), Rhizopus stolonifer (Q9HFV3), Aspergillus niger (Q00141) and Cunninghamella elegans (Q9P4E2) with identities of 80, 53, 45, 43 and 43%, respectively. P. chrysosporium CPR also had identities of 34, 36, 36 and 32% with rat (P00388), house fly (Q07994), yeast (P16603) and A. thaliana ATR1 (3) CPRs, respectively. Wang et al. (1997), identified three FMN-binding regions in rat CPR which were the phosphate moiety-binding region, 79IIVFYGSQTGTAEEFANRLS, the FMN ring (re-face)-binding site, 133VFCMATYGEGDPTDNAQDFYDWL and the FMN ring (si-face)-binding site, 163TGVKFAVFGLGNKTYEHFNAMGKYVDQRL. Four FADbinding sites were identified; these were the FAD ring (si-face)-binding

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Figure 12 Phylogram of NADPH-cytochrome P450 reductases. A phylogenic tree was constructed from previously published CPR sequences using Clustal X 1.8 and TreeView 1.6.1. The sequences used were Candida maltosa (P50126), Candida tropicalis (P37201), Cunninghamella elegans (JC7192), human (P16435), pig (P04175), rat (AAA41683), Schizosaccharomyces pombe (CAA22429), yeast (AAC09468), Aspergillus niger (S38427), Phanerochaete chrysosporium (AAG31349), Catharanthus roseus (Q05001), fruit fly (CAA63639), house fly (Q07994), mung bean (A47298), pea (AAC09468), parsley (T14903), Douglas fir (CAA89837), poplar (AAK15260), wheat (AAG17471), Rhizopus nigricans (AAG23833), silk moth (BAA95684), Arabidopsis thaliana (S21530), Caenorhabditis elegans (NP_498103), Coriolus versicolor (BAB83588) and Rhodotorula minuta (BAB21543).

site 450LQARYYSIASSSKVHPNSVH, the adenine-binding site 470ICAV484 VNKGVAVEYEAKSGR, the pyrophosphate-binding site 670 ATSWLRAKE and the FAD ring (re-face)-binding site RYSLDVWS. Two cytochrome c-binding sites were identified, these were

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TGVKFAVFGLGNKTYEHFNAMGKYVDQRL and 205GDDDGN. Two NADP(H)-binding sites were identified, which were the pyrophosphate-binding site 527VIMVGPGTGIAPFMGFIQER and the NADP(H) adenine-binding site 599AHKVYVQHLLKRDREHLWKLIHEGGAHIYVCGDARNMAKDV. The similarities between these binding sites on rat CPR (P00388) and CPRs from yeast (P16603), house fly (Q07994), P. chrysosporium (Q9HDG2) and A. thaliana ATR1 (Mizutani and Ohta, 1998) were compared. All three FMN-binding regions are highly conserved amongst the five CPR enzymes. Of the four FAD-binding regions, only two are highly conserved with the adenine- and pyrophosphate-binding regions being poorly conserved. This suggests that the FAD-binding sites differ between CPR enzymes from different species. The two cytochrome cbinding regions are highly conserved. This contrasts with the two NADP(H)-binding regions, where only the pyrophosphate-binding region is highly conserved, whilst the NADP(H) adenine-binding site is poorly conserved, except for the last 13 amino acid residues. This again suggests that the binding of NADP(H) differs between the five CPR enzymes. Whether these differences in the co-factor- and substrate-binding regions would lead to differences in kinetic properties requires further investigation. In general, studies on CPR have been largely restricted to mammalian forms. It has been suggested that CPR must have an intact membrane anchor for function. It has also been indicated that for supply of the two electrons for the CYP catalytic cycle CPR supplies the first and can also provide the second of these (Black and Coon, 1982). Differences between CPRs have been observed in fungi; yeast CPR was not found to be an essential gene as might be expected from mammalian studies and in contrast to studies on yeast CYP51 (Sutter and Loper, 1989). This suggested a different mechanism for providing both electrons to yeast CYP and it has been observed that the cytochrome b5 and NADHcytochrome b5 reductase system can substitute for yeast CPR and support normal activity (Lamb et al., 1999b). Yeast CPR can also drive the CYP catalytic cycle when present as an engineered soluble form lacking an N-terminal membrane anchor (Venkateswarlu et al., 1998; Lamb et al., 1999d). These differences indicate structural and mechanistic biodiversity between organisms that requires further investigation. We have taken this further to the X-ray crystal structure of the functional soluble yeast CPR in collaboration with Professor M. Waterman and Dr L. Podust at Vanderbilt University (Podust et al., unpublished).

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Figure 13 A molecular model of fluconazole bound in the active site of Candida albicans CYP51 (described in Boscott and Grant, 1994).

1.9.2. Fungal CYP as a Drug Target – Evolution of Resistant CYP51 As has been mentioned earlier one of the main stimuli for work on fungal CYP, specifically CYP51, came from the realisation that it was the target for azole antifungals. The azole compounds, and some other nitrogen-containing cyclic compounds for agriculture, bind to the haem of CYP51 as a sixth ligand and N-1 substituent groups of the compounds interact with the protein to give potent inhibition (Fig. 13). Azole compounds also bind to other fungal CYPs, but at the moment only CYP51 has been identified as essential for growth. Inhibition of CYP51 causes growth arrest through the accumulation of fungistatic 14-methylated sterols (Watson et al., 1989), but it is a particular 14-methyl diol sterol that is central to the mode of action (Kelly et al., 1995b). An interesting recent observation is the presence of two CYP51s in A. fumigatus (Mellado et al., 2001), but the significance of this finding for azole antifungal susceptibility remains unclear. Through study of resistance to azoles and suppressor mutations of CYP51 disrupted mutants, the role of sterol
5,6-desaturase (Erg3p) in the mode of action of these important drugs was revealed. The product of metabolism of 14-methylated sterol by this enzyme was a 6-hydroxy sterol that arrested growth. In a
5,6-desaturase mutant strain this sterol, 14methylergosta-8, 24(28)-dien-3b,6a-diol, was not made but instead 14methylfecosterol was produced that sustained growth (Watson et al., 1989;

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Figure 14 A simplified biosynthetic route to ergosterol showing the effect of blockage of 14-demethylation on sterol accumulation. Instead of proceeding via the sterol
5,6desaturation (mediated by the protein Erg3p) to ergosterol the 4-methylation steps still occur, but Erg3p activity results in the formation of 14-methylergosta-8,24(28)-dien-3b, 6a-diol production and growth arrest. The dashed lines reflect the simplified nature of the scheme i.e. other enzymes participate in the metabolism between the sterol structures. The continuous line is a single step mediated by Erg3p we showed is responsible for the mode of action of the azole drugs.

Kelly et al., 1995b, Fig. 14 ). Similar fluconazole-resistant C. albicans strains were observed in the clinic (Kelly et al., 1996, 1997c). Unfortunately, this mechanism of resistance involves mutant cells lacking ergosterol and, as a consequence, the strains are cross-resistant to amphotericin, the other main antifungal drug available for treatment that works by binding to ergosterol and disrupting cell membranes (Fig. 14).

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The importance of azole antifungals in therapy is growing and new compounds continue to emerge for treating systemic and superficial infection including skin and nail disease. The rapid increase in fungal infections has meant that they represent one of the top five infectious diseases in the world today with the immunocompromised being particularly susceptible to infection. Patients undergoing organ transplantation, cancer chemotherapy, intensive care and HIV positive patients are all at risk. The types of fungal infections vary geographically, but the main pathogens are various species of Candida, particularly C. albicans. However, particularly under azole treatment, more intrinsically resistant forms such as C. krusei can be selected. Other important pathogens include A. fumigatus where infection is seriously life-threatening and C. neoformans, Histoplasma capsulatum and Penicillium marnefii. With opportunistic infections, fungi associated with plant disease have also been observed such as Fusarium spp. and even S. cerevisiae. This applied relevance in part explains the large number of fungal CYP51s in the sequence databases that includes sequences of genes/proteins from medical pathogens, but also from phytopathogens such as Ustilago maydis (Lamb et al., 1998b) and mildews (Delye et al., 1997). The CYP51 sequences represent an interesting source with which to begin comparing conserved residues across the superfamily and some of these have been associated with changed resistance to azoles. Within the fungi there are known to be differences in the sensitivity of the target enzyme, but using purified protein from human and C. albicans there was only a small reduction is sensitivity of the human CYP51 representing less than five-fold change in IC50 in the case of ketoconazole and itraconazole (Lamb et al., 1999c). Possibly the pool of human CYPs can protect the CYP51 and a similar finding was made in plant/fungal comparative studies (Lamb et al., 2001). In contrast to these findings are the observations that point mutations in CYP51 can cause resistance in the clinic and the field. In medicine, the prophylactic and prolonged use of azole therapy, sometimes extending for years, provides strong evolutionary pressure for the selection of resistant forms. Overexpression of CYP51 may also cause resistance, but the evidence for this is less strong. Among the strongest evidence for this resistance mechanism was the report that increased CYP51 mRNA has sometimes been seen in resistant strains (e.g. White, 1997; Franz et al., 1998) and that Penicillium digitanum resistant strains exhibited five repeats of a 126 bp sequence in the CYP51 promoter in contrast to one in sensitive strains (Hamamoto et al., 2000). Although we deal in this review with resistance associated with alterations in CYP51, it is important to note as well that this

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is not the only cause of resistance. In addition, there is resistance through defects in sterol
5,6-desaturase that change the sterol accumulating under treatment (Kelly et al., 1996) and there is an involvement in C. albicans azole resistance of major facilitator proteins like MDR1p and ABC transporters like CDR1p and CDR2p. Most of the work published on resistance is with C. albicans and multiple mechanisms of resistance can operate in the same cells (Sanglard et al., 1995; Venkateswarlu et al., 1995; Kelly et al., 1997c; White, 1997; Franz et al., 1998; Sanglard et al., 1998; Lamb et al., 1999a; Lupetti et al., 2002). In a sequence analysis of the CYP51 gene from 19 fluconazole-resistant isolates from C. albicans, alterations were evident that were not present in a similar sized control set of sequences from sensitive isolates (Loeffler et al., 1997). In the resistant strains, the following amino acid substitutions were observed: five had F105L, five had E266D, one had K287R, one had G448E, three had G464S and one V488I. None of the mutations were where molecular modelling or laboratory-based site-directed mutation studies had indicated resistance could arise (Lamb et al., 1997a,b). The changes included mutations in different regions and distinguishing true resistance mutations from CYP51 polymorphisms is ongoing, but G464S appears the most common of these changes and has been demonstrated to cause resistance (see below). Work with matched sets of strains from individual patients has proved valuable, as changes in sensitivity can be correlated with molecular changes with greater confidence that the genetic background was not altered, thus contributing to changed sensitivity. Such studies first indicated the importance of R467K in fluconazole resistance where cellfree extracts of this mutant sustained ergosterol biosynthesis at a higher concentration of fluconazole (White, 1997). The use of S. cerevisiae as a host for expression of C. albicans CYP51 and other CYP51s has been important in that heterologous CYP51 expression can confirm an in vivo role for genetic change in causing microbiological resistance through tests of cellular sensitivity. Such host strains can contain a knock-out of the endogenous CYP51 (Kelly et al., 1995; Sanglard et al., 1998) or be selected host strains with a low endogenous CYP content (Lamb et al., 1997a). Furthermore, using the GAL10 promoter on YEp51, mmol/L quantities of C. albicans CYP51 have been obtained for in vitro studies using pure protein to investigate the catalytic reaction mechanism and probing of the role of the Thr315 residue of CYP51 (Shyadehi et al., 1996; Lamb et al., 1997a). Many further mutations have been shown to cause resistance using heterologous expression systems in yeast (Lamb et al., 1997a; Sanglard et al.,

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1998; Favre et al., 1999), or by in vitro sterol biosynthesis studies (Franz et al., 1998; Favre et al., 1999; Marichal et al., 1999). The residues are not restricted to a single, or few, residue(s), and it appears that many changes can result in resistance. We are currently assessing almost 60 residues in the 530 amino acid C. albicans CYP51 polypeptide. Among the more commonly observed changes are Y132H, G464S and R467K. The latter two have been investigated at the level of protein and shown to have reduced affinity for fluconazole (Kelly et al., 1999; Lamb et al., 2000). As these residues lie beneath the plane of the haem in the active site of CYP51, it is possible that they may give rise to resistance by changing the position of haem and, thereby, displacing the fluconazole bound above it in relation to important binding sites within the active site. As with all such resistance mutations, it is required that catalytic activity is maintained in the protein for viability (fitness) of the fungus. Within the C. albicans cysteine-containing haembinding domain 464GGGRHRCIGEQF, 464G>S (Loeffler et al., 1997; Kelly et al., 1999), 465G>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.

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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

262 GUANGHUI WU, LAURA M. WAINWRIGHT AND ROBERT K. POOLE

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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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