Advances in Microbial Physiology Volume 38

Advances in

MICROBIAL PHYSIOLOGY VOLUME 38

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Advances in

MICROBIAL PHYSIOLOGY Edited by

R. K. POOLE Department of Molecular Biology and Biotechnology The Krebs Institute f o r Biomolecular Research The University of Sheffield Firth Court, Western Bank Sheffield SlO 2TN, UK

Volume 38

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Contents

CONTRIBUTORS TO VOLUME 38

........................

vii

Hydrophobins: Proteins that Change the Nature of the Fungal Surface Joseph G . H.Wessels 1. 2. 3. 4. 5. 6. 7.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Identity of hydrophobins . . . . . . . . . . . . . . . . . . . . . . . . . Rodlets . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Surface activities of hydrophobins . . . . . . . . . . . . . . . . . . . . Formation of emergent structures . . . . . . . . . . . . . . . . . . . . . Technology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 4 10 13 19 34 36 36 36

Structure-function Analysis of the Bacterial Aromatic Ring-hydroxylating Dioxygenases Clive S . Butler and Jeremy R . Mason

................................ ........................... Structure of ring-hydroxylating dioxygenases . . . . . . . . . . . . . . Electron transport system . . . . . . . . . . . . . . . . . . . . . . . . . The catalytic terminal oxygenase component . . . . . . . . . . . . . . . Coordination of the iron-sulphur clusters . . . . . . . . . . . . . . . . . The catalytic non-haem iron centre . . . . . . . . . . . . . . . . . . . . Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction

2. Bacterial oxygenases

3.

4.

5. 6. 7.

8.

47 48 50 51 60 61 72 75 76 76

vi

CONTENTS

Thiol Template Peptide Synthesis Systems in Bacteria and Fungi Rainer Zocher and Ullrich Keller Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The peptide synthetase domain . . . . . . . . . . . . . . . . . . . . . . Enzymesystems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Peptide synthetases from fungi . . . . . . . . . . . . . . . . . . . . . . Prokaryotic peptide sythetase systems . . . . . . . . . . . . . . . . Future prospects of peptide synthetase research . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. 2. 3. 4. 5. 6.

. .

86 88 94 96 . 111 . 122 124 124

Microbial Dehalogenation of Halogenated Alkanoic Acids. Alcohols and Alkanes J . Howard Slater. Alan T. Bull and David J . Hardman 1. 2. 3. 4. 5.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 Dehalogenation of halogenated alkanoic acids . . . . . . . . . . . . . . 135 Dehalogenation of halogenated alcohols . . . . . . . . . . . . . . . . . 151 Dehalogenation of halogenated alkanes . . . . . . . . . . . . . . . . . .159 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166

Metal-Microbe Interactions: Contemporary Approaches T. J . Beveridge. M . N . Hughes. H . Lee. K . T. Leung. R . K . Poole. I . Savvaidis. S . Silver and J .T.Trevors Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 178 Microorganisms and metals: their essentialchemistry . . . . . . . . . . 179 Complexation of metal ions in media and cellular milieu . . . . . . . . 185 Analysis for total metal and for metal species . . . . . . . . . . . . . . 190 Spectroscopic techniques in the study of metals and microorganisnls . . 205 Molecular and genetic methods . . . . . . . . . . . . . . . . . . . . . . 210 214 7. Case studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231 8. Conclusions and outlook . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . 232 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 232

1. 2. 3. 4. 5. 6.

Author Index SubjectIndex

.................................. ..................................

245 263

Contributors to Volume 38

T. J. BEVERIDGE, Department of Microbiology, College of Biological Sciences, University of Guelph, Guelph, Canada NIG 2W1 Alan T. BULL,Research School of Biosciences, University of Kent at Canterbury, Canterbury, Kent CT2 7NJ, UK Clive S. BUTLER,School of Biological Sciences, Molecular and Microbiology Sector, University of East Anglia, Norwich NR4 7TJ, UK David J. HARDMAN,Carbury Heme Ltd, Research and Development Centre, Canterbury, Kent CT2 7PD, UK M. N. HUGHES,Department of Chemistry, King’s College London, Strand, London WC2R 2LS, UK Ullrich KELLER,Institut fur Biochemie und Molekulare Biologie, Technische Universitat Berlin, FranklinstraPe 29, D- 10587 Berlin-Charlottenburg, Germany H. LEE,Department of Environmental Biology, Ontario Agricultural College, University of Guelph, Guelph, Canada NIG 2W1

K. T. LEUNG,Department of Environmental Biology, Ontario Agricultural College, University of Guelph, Guelph, Canada N1G 2W1 Jeremy R. MASON,Division of Life Sciences, King’s College London, Campden Hill Road, London W8 7AH, UK R. K. POOLE,Department of Molecular Biology and Biotechnology, Krebs Institute for Biomolecular Research, The University of Sheffield, Firth Court, Western Bank, Sheffield S10 2TN, UK I. SAVVAIDIS, Department of Microbiology, University of Ioannina Medical School, Post Box 1186,45110 Ioannina, Greece

S . SILVER,College of MedicineDepartment of Microbiology and Immunology, University of Illinois, M-C 790,835 S. Wolcott Ave, Chicago, l L 60612, USA

viii

CONTRIBUTORS TO VOLUME 38

J. Howard SLATER,Molecular Ecology Research Unit, School of Pure and Applied Biology, University of Wales, PO Box 915, Cardiff CFI 3TL, UK

J. T.TREVORS, Department of Environmental Biology, Ontario Agricultural College, University of Guelph, Canada, NlG 2W1 Joseph G. H. WESSELS,Department of Plant Biology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Kerklaan 30, 975 1 NN Haren, The Netherlands Institut fur Biochemie und Molekulare Biologie, Technische Rainer ZOCHER, Universitat Berlin, FranklinstraPe 29, D- 10587 Berlin-Charlottenburg,Germany

Hydrophobins: Proteins that Change the Nature of the Fungal Surface Joseph G. H. Wessels Department of Plant Biology, Groningen Biomolecular Sciences and B i o t e c h l o g y Institute, University of Groningen, Kerklaan 30, 9751 NN Haren, The Netherlands 1. Introduction . . . . . . . . . . . . 2. Identity of hydrophobins . . . . . . 3. Rodlets . . . . . . . . . . . . . . 4. Surface activities of hydrophobins . 4.1. SC3 hydrophobin . . . . . . . 4.2. Cerato-ulmin . . . . . . . . . 5. Formation of emergent structures 5.1. Formation of aerial hyphae . . 5.2. Formation of fruit bodies . . . 5.3. Formation of conidia . . . . . 5.4. Pathogenesis. . . . . . . . . 5.5. Symbiosis . . . . . . . . . . 6. Technology . . . . . . . . . . . . 7. Conclusions . . . . . . . . . . . . Acknowledgements , . . . . . . . References . . . . . . . . . . . .

. ... .. . . ... .. . . ... ... ....... . ... ... . . .. ... . . .. , . , . . . . .. , . . . . .. . . . . . , , . . . . . . . . . . . . .. . . . . . . , ,

. ..... . . ..... . . . .. .. . ....... . . . . .. . . , .. , . , , , , . . , . , , . , . . . . . . . . . . ,

,

,

,

.. .

. . . . . . . . .. . .. . ... , .. . . . . . . . . . . . . . . . . . . . .. .. , .. .. . . . . . . . . . . . . . . .

. . . . . . . . . . . 1 . . . . . . . . . . .4 . . . . . . . . . . . . 10 . . . . . . . . . . . . 13 . . . . . . . . . . . . 13 . . . , , . . . . . . 18

. . . .

,

. .. . . . .. . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . ..

. . . . .

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. . . . . . . . . . . . . .. . . . .. . . . ..

. . 19 . . 19 . . 22

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27

. 29 . 33 . 34

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. 36

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36

1. INTRODUCTION Although fungi, with the exception of yeasts, are multicellular, their mechanisms of growth and development are quite distinct from those of plants and animals. In the multicellular fungi, the fundamental growth unit is the hypha, which may or may not be divided into cells. The hypha is essentially a tubular structure containing all the typical eukaryotic cytoplasmic components surrounded by a rigid wall. Hyphae are highly polarized and grow only at one end, where a new wall is ADVANCES IN MICROBIALPHYSIOLOGY VOL 38 ISBN 0-12-027738-7

Copyright 0 1997 Academic Press Limited All rights of r e p d u d i o n in any form reserved

2

JOSEPH G.H.WESSELS

deposited in such a way that the tubular shape is generated, despite the presence of a high internal hydrostatic pressure (turgor) (Wessels, 1986,1990,1993a). Hyphae regularly branch and give rise to a mycelium that forms a colony. The colony grows radially at its periphery,whcre apical extcnsion and branching occur. The mycclium thus colonizes a substrate, maintaining a constant ratio between the total length of hyphae and the number of tips, a ratio known as the “hyphal growth unit” (Trinci. 1974; Trinci et nl., 1994). Another uscful concept developed by Trinci and co-workers (see Tritici el ul., 1994) is that extension of tips of individual hyphae is supported by a certain volume of protoplasm; this mycelial region involved in tip growth is called thc “peripheral growth zone” of the colony. In principle, hyphae appear designed for unlimited transport of water, nutrients and cytoplasmic components. In the zygomycetes the hyphae are not subdivided by septa so that the cytoplasm (with many nuclci) is essentially contained in a continuousbranched tubular system, making it difficult in these organisms to apply the concept of cellularity. In the second major group of the fungi, the ascomycetes, septa are present but these contain large pores that do not seem to obstruct movement of organelles such as nuclei and mitochondria. Only in thc third group. the basidiomycetes, do septa divide the hyphae into separate compartments. The small septa1 pore and the elabori~temembranous structures (parenthosomcs) that cover it (Moore. 1985) effectively prevent passage of nuclei and mitochondria, but apparently do allow for transport of water and nutrients. In fact. members of the basidiomycetes have a great capacity for long-rangetranslocation (Jennings, 1984). However, in situationswhere nuclci have to bc cxchangcd in these basidiomycetes, as in mating interactions, thc scpti are dissolved so that nuclei and other cytoplasmic components can move freely through the hyphal tubes (Raper, 1966; Wessels, 1978). Because hyphae not only grow but also sccrctc cnzymcs at their apices (Wosten cr ul., 1991). the fungal mycelium is ideally suited for growing into solid organic substrates and for degrading the constitucnt polymcrs from within (Wessels, 1993a). Thc colonization is Ihcilituted by the fact that, in the local absence of nutrients, hyphal growth can be sustained by the transport of water and nutrients from a food base (Jennings. 1984, 1994; Kayner et a[., 1995). In this way the mycelium can explore large arc~sthat conlain only isolated patches of nutrients. Another manifestation of the ability of the mycelium to transport water and assimilates to hyphae that are unable to acquire nutrients, is the Occurrence of emergent growth. For instance, hyphae may give up assimilatingnutrients and grow into the air, causing the “mouldy” appearance of many fungi. A function of the hydrophobic felt-like mats that arc often produced is not obvious but may be prevention of water loss from the substrate. Certain aerial hyphae, however, may differentiateinto spore-bearingstalksthat at their apices form sporangiacontaining sporangiospores (many zygomycetes) or sterigmnta that bud off conidiospores (many ascomycetes). Alternatively,aerial hyphae iit thcir apices may break up into oidiosporcs.

HYDROPHOBINS

3

All these structures serve vegetative reproduction. For sexual reproduction, many fungi form multihyphal fruit bodies within which meiosis occurs and meiospores are formed. As asexual spores, these can also be dispersed through the air or otherwise disseminated. Particularly in members of the basidiomycetes, the fruit bodies (mushrooms and brackets) can attain large sizes and their morphogenesis has attracted much attention both from a purely scientific and, for the edible species, from a commercial point of view (Wessels, 1993b). These large multicellular structures are not formed by cell divisions within meristems, as in plants, but they are formed by individual hyphae that grow at their tips and seem to “know” how to organize themselves into a distinct multihyphal structure. It is clear that, for the elaboration of these aerial fruit bodies, massive transport of water and assimilates from the assimilative substrate mycelium is required. When the substrate is exhausted, components of the substrate mycelium itself may be broken down and breakdown products reused for the construction of these emergent structures (Wessels and Sietsma, 1979; Wessels, 1993b). After spores have been dispersed, they must find a substrate in order to germinate and to produce a new mycelium. For saprotrophs this poses no problems, provided the spore lands in an area where dead organic material and enough moisture are available. For biotrophs, however, it is often necessary for a spore to attach to, and to germinate on, the bare surface of the host before infection structures can be formed and the host is penetrated to form an assimilative mycelium. This is particularly clear in parasitic relationships with plants and animals (Cole and Hoch, 1991), but is also evident in the mutually beneficial associations with plants, the mycorrhizas (Harley and Smith, 1983; Read, 1991). In addition, a large number of fungi, an estimated 20% of all species, have evolved as lichens, aerial structures in which the fungus obtains its assimilates from symbiotic algae and cyanobacteria (Honneger, 1993). This brief overview of fungal biology serves as an introduction to understanding the roles played by hydrophobins. These proteins were discovered while searching for genes expressed during emergent growth in Schizophyllurn cornmurre. AS in many homobasidiomycetes, the primary mycelium that grows from a meiospore forms aerial hyphae but, after mating of two compatible primary mycelia, a secondary mycelium is formed that produces fruit bodies in addition. The cDNAs of a number of abundant mRNAs appearing during emergence of the aerial structures of primary and secondary mycelia of S. commune were cloned (Mulder and Wessels, 1986). The most abundantly expressed genes were sequenced, revealing that at least four of the ten cloned genes encoded similar small cysteine-rich hydrophobic proteins (Schuren and Wessels, 1990; Wessels et al., 1995). At that time these proteins were totally unknown. Eventually, the product of one of these genes (SC3) was found in the walls of aerial hyphae, while the abundant product of another (SC4) was found in walls of hyphae that make up fruit bodies (Wessels et al., 1991a,b). The proteins were present in these walls as complexes, insoluble in a hot solution of 2% sodium dodecylsulfate (SDS), that

4

JOSEPH G. H. WESSELS

could be dissociated into monomers only by treatments with pure formic acid or trifluoroacetic acid, although monomers of these proteins were present in the medium of still cultures. Because of the abundance of hydrophobic residues and their presence in walls, we dubbed these proteins “hydrophobins”, a term used earlier to denote any substance conferring hydrophobicity to a microbial surface (Rosenberg and Kjelleberg, 1986). Around the same time, Stringer et al. (1991) found a gene in Aspergillus nidulans with homology to the S. cuinmune hydrophobin genes. Disruption of this gene caused a phenotype of wettable conidiospores from which the so-called rodlet layer was missing. This indicated hydrophobins as an essential component of hydrophobic rodlet layers, generally observed on fungal spores. We then showed that a single purified hydrophobin from S. cuininune (SC3) could form such a hydrophobic rodlet layer in vitm by self-assembly at a water-air interface (Wosten et ab, 1993), and that such a layer was formed at the surface of aerial hyphae (Wosten et al., 1994b).It was also found that this hydrophobin could mediate strong attachment of S. cuintnune hyphae to solid hydrophobic surfaces (Wosten et al., 1994a). In the meantime, hydrophobin-like proteins were found in all fungi examined (de Vries et al., 1993), while anonymous genes highly expressed in fungi during a variety of developmental processes turned out to encode proteins with clear homology to the S. cuminune hydrophobins (Fig. 1).

2. IDENTITY OF HYDROPHOBINS It is noteworthy that most of the hydrophobins listed i n Fig. 1 were found by sequencing cDNAs representing mRNAs abundantly expressed during certain stages of fungal development without knowing anything about the encoded proteins. Only ABHI, CoH1, cerato-ulmin and cryparin were first identified as proteins, and their genes then cloned by polymerase chain reaction (PCR) using degenerate primers based on determined N-terminal amino-acid sequences. In retrospect, the late discovery of these abundantly occurring proteins is understandable because many occur as SDS-insoluble complexes that can be dissociated into monomers only by using concentrated formic acid or trifluoroacetic acid (de Vries et af., 1993), agents not in common use for protein extraction. In principle, these proteins could have been seen when examining proteins present in media from standing cultures, but only after handling such media with special care because the hydrophobins easily aggregate upon exposure to air forming insoluble complexes. Precisely for this reason, cerato-ulmin (CU) and cryparin (CRYP) were detected early because, on shaking, these Class I1 hydrophobins formed a milky turbidity that could be dissolved in SDS. Yet, the fact that hydrophobin sequences are so readily found in screens for developmentally regulated sequences indicates that they are derived from the most abundantly expressed fungal genes. Indeed, the

5

HYDROPHOBINS

i t rl'

I'

Neumspm cmssa

Figure 1 Dendrogram of similarities between aligned hydrophobins obtained by the CLUSTAL programme of the PClGENE programs package, version 6.60 (Higgins and Sharp, 1988). Numbers in superscript indicate references where sequence information was gublished orrefer to unpublished data. 'hchuren and Wessels (1990);Wessels et al. (1991a). )Wessels et al. (1995). 3)S.A. Asgeirsd6ttir and L.A. Casselton (unpublished). 4)Martinet a1. (1995); Tagu et al. (1996). ')Lugones et at. (1996); de Groot et al. (1996). @StLeger et al. (1 992). 7)Talbotel at. (1 993). 8)Strjngerand Timberlake ( 1 995). 9)Strjngeret al. (1991); J. Rhodes and W.E. Timberlake, cited in Stringer and Timberlake (1995). '')Pam et a/. (1994); Thau et al. (1994). ")Bell-Pedersen et nl. (1992); Lauter el 01. (1992); Templeton et al. (1995). 12)Yaguchiet al. (1993); Bowden et at. (1993); Stringer and Timberlake (1993). 13)Loraet al. (1994). 14)Zhanget at. (1994); Carpenter et al. (1 992). 15)Nakari-Set;ilaet al. (1996). 16?. Nakari-Seala and M. Penttila (unpublished). Amino-acid sequences at the N-terminal end located before the first cysteine residue were omitted in the comparison since these include the signal sequence for secretion and in only eight cases (SC3, SC4, CoH1, ABHI, RodA, Eas, CU and CRYP) is the N-terminus of the mature protein known. However, aligning the whole protein sequence, including the signal sequence, results qualitatively in the same type of dendrogram, only the distances become larger. The overall identity of all sequences is only 4.3%, the overall similarity 1.7%.

SC3 and SC4 genes of S. coininune were shown to produce 1% and 3.596, respectively, of the mRNA mass at the tiine of emergent growth [Mulder and Wessels, 1986), while the record is probably set by the mRNA for cryparin that amounted to 25% of the mRNA mass (Zhang ef nl., 1994). The sequence diversity of hydrophobins (Fig. 1) means that isolation of

hydrophobin genes on the basis of sequence homology is mostly impossible. For instance, the four hydrophobin genes cloned from S. coininune do not crosshybridize (Mulder and Wessels, 1986), even under non-stringent conditions. Only in the case of related species has nucleic acid homology been used to isolate a hydrophobin gene that fulfils a similar function: the hydrophobin gene that is responsible for formation of rodlets on conidia of Aspergillus fumigatus was isolated on the basis of its homology to the KodA gene of A. nidulans (Parta et al.,

6

JOSEPH G. H. WESSELS

1994; Thau et al., 1994). This state of affairs means that it is generally unknown how many hydrophobin genes exist in a given fungal species, but the identification of multiple genes in species, such as S. cornrnune,I? tinctorius and A. nidulans, just by screening cDNA libraries, indicates that, in most studied species, only the most abundantly expressed hydrophobin genes may have been identified. Of the (putative) hydrophobins listed in Fig. 1, only SC3 (Wosten et al., 1993) and SC4 (this laboratory, unpublished data) from S. commune, ABHl from Agaricus bisporus (Lugones et al., 1996), CoHl from Coprinus cinereus (S.A. Asgeirsd6ttir and L.A. Casselton, unpublished data), CU from Ophiostorna ulmi (Takai and Richards, 1978; Russo et al., 1982) and CRYP from Cryphonectria parasitica (Carpenter et al., 1992) have been physically isolated and their properties studied. Wessels (1992) noted that the remarkable property of interfacial self-assembly exhibited by the SC3 hydrophobin of S. commune (see below) was earlier observed with CU (Takai and Richards, 1978; Russo etal., 1982)andCRYP (Carpenter et al., 1992). When the amino-acid sequence of CU became available (Yaguchi et al., 1993), Stringer and Timberlake (1993) noted the sequence homology to known hydrophobins. However, whereas interfacial self-assembly of, for instance, SC3, SC4 and ABHl hydrophobins results in aggregates that are highly insoluble in water, organic solvents and 2% SDS, the aggregates formed by CU and CRYP were found to be unstable in water, and soluble in aqueous ethanol and 2% SDS. In addition, they display a hydropathy pattern that is clearly different from that of hydrophobins like SC3 (Fig. 2). Therefore, Wessels (1994) proposed a distinction between Class I hydrophobins that form highly insoluble assemblages and Class I1 hydrophobins that form less stable assemblages (e.g. soluble in 60% ethanol or 2% SDS), a distinction supported by the alignment dendrogram shown in Fig. 1. In the Class I hydrophobins, the cysteine doublets are followed by a stretch of hydrophilic amino acids whereas, in Class TI hydrophobins, hydrophobic residues immediately follow the cysteine doublets (Fig. 2). Also, fewer amino acids separate the third and fourth cysteine residue in Class I1 hydrophobins than in Class I hydrophobins. Whether this grouping is correct can only be decided after isolation and characterization of all the listed hydrophobins. However, because most of the hydrophobins tabulated in Fig. 1 have not yet been physically isolated, they can be only tentatively grouped as Class I and Class I1 hydrophobins on the basis of similarities in hydropathy patterns and solubility characteristics of assemblages. It would not be surprising if some of these hydrophobins exhibit solubility characteristics intermediate between the two classes now distinguished. On the basis of the available information on hydrophobins, they would seem to have the following characteristics:

1. Hydrophobins are small proteins (100 k 25 amino acids) that are moderately hydrophobic. The hydrophobicity indices (Kyte and Doolittle, 1982) for mature proteins vary from 0.01 (RodA) to 0.60 (SC3). The overall hydrophobicity thus varies widely.

HYDROPHOBINS

7

ichi:ophvllum cummune SC3”

irhizophvl/um commune SCI”

Werorhizium onisupliue SSCA*’

Veuruspora cmssa Eaa”’

bfagnuyorrhe griseu MPG 1 ’I

Figure 2 Comparison of hydropathy patterns of selected hydrophobins (SC3, SCI, SC4, SSGA, Eas, RodA, MPGI, CU and CRYP) (for references, see Fig. 1). The patterns were determined using the parameters of Kyte and Doolittle (1982). A six amino-acid window was used and plotted against position in the deduced amino-acid sequence. The hydropathy patterns were then aligned around the first and second cysteine doublet, and around the fourth and eighth cysteine residue leaving gaps in the sequences where the hydrophobic regions (above the lines) alternate with hydrophilic regions. The hydrophobic amino-terminal sequences serve as signal sequences for secretion. The amino termini for the mature hydrophobins, when known, are indicated by arrows. Note that the first seven hydrophobins (Class I) have similar hydropathy patterns, which deviate from those of the last two hydrophobins (Class 11). (Modified from Wessels, 1994, with permission from the publisher.)

JOSEPH G. H.WESSELS

8 2.

3.

4.

5.

6.

Hydrophobins are all secreted as suggested by the presence of signal sequences. This was actually shown for those hydrophobins in which the amino terminus of the mature protein was determined (arrows in Fig. 2). Hydrophobins have a conserved spacing of eight cysteine residues: X2-38-C-X5-9-C-C-X 1~-~9-c-x~~2~-c-x~~~c-c-x~~~ g-C-X*-13 in which X signifies any other amino acid, except for tryptophan, which has been reported only in HydPtl, while methionine has been found only in HydPtl, HydPt2, Eas, SSGA and MPG1. Asparagine mostly follows the first cysteine doublet. Of course, the numbers of amino acids that separate the cysteine residues may change as more hydrophobins are sequenced but the recurrent hydropathy patterns around the sequence C-X5-9-C-€ in the amino-terminal and carboxy-terminal halves of the molecule are remarkable (Fig. 2). (Note that in the putative QID3 protein listed in Fig. 1, serine substitutes for the second cysteine residue.) Hydrophobins have poor amino-acid homology. For instance, the SC1, SC3 and SC4 hydrophobins, all produced by S. cornmune, are only 39% identical. However, many of the differences concern conservative substitutions so that the similarity between these hydrophobins becomes 80%. If the RodA hydrophobin ofA. nidulaiis and the Eas hydrophobin of N. c r a m are also taken into account, the identity between the five hydrophobins drops to 11% and the similarity to 34%. The similarities between the hydrophobins therefore become most clear when both the conserved spacings of cysteine residues and the hydropathy patterns are compared (Fig. 2). Hydrophobins have the capacity to assemble into an amphipathic protein film when confronted with a hydrophilic-hydrophobic interface, such as between water and air. As indicated above for the Class I hydrophobins, this was shown only for the hydrophobins SC3, SC4, CoHl and ABHl. However, the hydrophobins Eas (Templeton et al., 1995), MPGl (Talbot et al., 1993), RodA(Stringeret al., 1991) and HYPl (Partaetal., 1994; Thau et al., 1994) have all been shown to be part of, or constitute, the hot SDS-insoluble hydrophobic rodlet layer on conidiospores and thus most probably had gone through the interfacial self-assembly process. For the putative Class I1 hydrophobins, interfacial self-assembly has clearly been established for CU (Takai and Richards, 1978;Russo et al., 1982; Richards, 1993), CRYP (Carpenter etal., 1992) and HFBl (Nakari-Seda et af., 1996). As far as is known, all hydrophobins are present as assemblages on the surfaces of emergent hyphal structures.

These criteria delimit the hydrophobins from other cysteine-rich proteins of fungal or other origins. It has been suggested (St Leger el al., 1992; Templeton et al., 1994) that hydrophobins may be related to proteins exhibiting the so-called toxin-agglutinin fold (Drenth et al., 1980; Andersen et al., 1993). For the agglutinins belonging to the chitin-binding family (Raikhel et al., 1993) disulphide

HYDROPHOBINS

9

bridges occur between C1-424, C2-C5, C 3 4 6 and C7 ser mutated enzymes. Biochemistry 31,46024612, Pang, C.-P., Chakravarti, B., Adlington, R.M., Ting, H.-H., White, R.L., Jayatilake, G.S., Baldwin, J.E. and Abraham, E.P. (1984) Purification of isopenicillin N synthase. Biochem. J. 222,789-795. Pember, S.O., Johnson, K.A., Villafranca,J.J. and Benkovic, S.J.(1989) Mechanistic studies on phenylalanine hydroxylase from Chromobacterium violaceum. Evidence for the formation of an enzyme-oxygen complex. Biochemistry 28,2124-2130. Pfefferkom, B. and Meyer, H.E. (1986) N-terminal amino-acid sequence of the Rieske iron-sulfur protein from the cytochrome bd'complex of spinach thylakoids. FEBS Lett. 206,233-237. Que, L. Jr, Widom, J. and Crawford, R.L. (1981) 3.4-dihydroxyphenylacetate 2,3dioxygenase: a manganese(I1) dioxygenase from Bacillus brevis. J. Uiol. Chem. 256, 10941-10944. Raag, R. and Poulos, T.L. (1989) Crystal structure of the carbon monoxide-substratecytochrome P-450CAM ternary complex. Biochemistry 28,7586-7592.

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Riedel, A., Rutherford, A.W., Hauska, G., Muller, A. and Nitschke, W. (1991) Chloroplast Rieske protein-EPR study on its spectral characteristics, relaxation and orientation properties. J. Biol. Chem. 266, 17838-17844. Rieske, J.S., Zaugg, W.S. and Hansen, R.E. (1964) Properties of a new oxidation-reduction component of the respiratory chain as studied by electron paramagnetic resonance spectroscopy. J. Biol. Chein 239,3017-3022. Romanov, V. and Hausinger, R.P. (1994) Pseudomonas aeruginosa 142 uses a threecomponent ortho-halobenzoate 1,2-dioxygenase for the metabolism of 2.4-dichloro- and 2-chlorobenzoate. J. Bacleriol. 176, 3368-3374. Rosche, B., Tshisuaka, B., Fetzner, S. and Lingens, F. (1995) 2-0~0-1,2-dihydroquinoline 8-monooxygenase, a two-component enzyme system from Pseudomonas putida 86. J. Biol. Chem. 270,17836-17842. Rosenzweig, A.C., Frederick, C.A., Lippard, S.J. and Nordlund, P. (1993) Crystal structure of a bacterial non-haem iron hydroxylase that catalyses the biological oxidation of methane. Ncrture 366,537-543. Salemo, J.C., Blum, H. and Ohnishi, T. (1979) The orientation of iron-sulphur clusters and a spin-coupled ubiquinone pair in the mitochondria1 membrane. Biochim. Biophys. Acta 547,270-281. Sauber, K., Frohner, C., Rosenberg, G, Eberspocher, I . and Lingens, F. (1977) Purification and properties of pyrazon dioxygenase from pyrazon-degrading bacteria. E m J. Biochem. 74,89-97. Schlafli, H.R., Baker, D.P., Leisinger, T. and Cook, A.M. (1995) Stereospecificity of hydride removal from NADH by reductases of multi-component non-heme iron oxygenase systems. J. Bacteriol. 177, 831-834. Schlafli, H.R., Weiss, M.A., Leisinger, T. and Cook, A.M. (1994) Terephthalate 1.2dioxygenase system from Comamonas teslosteroni T-2: purification and some properties of the oxygenase component. J. Bncteriol. 176.66444652. Schweizer, D., Markus, A., Seez,M., Ruf, H.H. and Lingens, F. (1987) Purificationand some properties of component B of the 4-chlorophenylacetate 3.4-dioxygenase from Pseudornonns species strain CBS3. J. Biol. Chem. 262,9340-9346. Sheldrake, G.N. ( I 992) Biologically derived arene cis-dihydrodiols as synthetic building blocks. In: Chircrlify in Industry (A.N. Collins, G.N. Sheldrake and J. Crosby, eds), pp. 127-1 66. Wiley, Chichester. Shergill, J.K. and Cammack, R. (1994) ESEEM and ENDOR studies of the Rieske iron-sulphur protein in bovine heart mitochondria1 membranes. Biochim. Biophys. Acta 1185,3542. Shergill, J.K., Joannou, C.L.. Bratt, P.J., Mason, J.R. arid Cammack, R. (1995) Coordination of the Rieske-type [2Fe-2S] cluster of the terminal iron-sulphur protein of Pseudomoms pufida benzene 1,2-dioxygenase, studied by one- and two-dimensional electron spinecho envelope modulation spectroscopy. Biochemistry 34,16533-16542. Simon, M.J., Osslund,T.D., Saunders, R., Ensley, B.D., Suggs, S.,Harcourt, A., Suen, W.-C., Cmden, D.L., Gibson, D.T. and Zylstra, G.J. (1993) Sequences of the genes encoding naphthalene dioxygenase in Pseudomonas putida strains G7 and NCIB 9816-4. Gene 127,31-37. Spain, J.C., Zylstra, G.J., Blake, C.K. and Gibson, D.T. (1989) Monohydroxylation of phenol and 2,5-dichlol-ophenol by toluene dioxygenase in Pseudornonas putida F1. Appf. Environ. Microbiol. 55,2648-2652. Subramanian, V., Te-Ning, L., Yeh, W.K. and Gibson, D.T. (1979) Toluene dioxygenase: purification of an iron-sulfur protein by affinity chromatography. Biochem. Biophys. Res. Commnn. 91, 1131-1139. Subramanian, V., Liu,T.-N., Yeh, W.-K., Narro, M. and Gibson, D.T. (1981) Purification and

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properties of NADH-ferredoxinToLreductase. A component of toluene dioxygenase from Pseudomonas purida. J. Biol. &hem. 256,2723-2730. Subramanian, V., Liu,T.-N., Yeh, W.-K., Serdar,C.M., Wackett, L.P. andGibson, D.T. (1985) Purification and properties of ferredoxinmL. A component of toluene dioxygenase from Pseudoinonas putida F1. J. Biol. Chein. 260,2355-2363. Suemori, A., Kurane, R. and Tomizuka, N. (1993) Purification and properties of phthalate oxygenase from Rhodococcus eryrhropolis S-1 . Biosci. Biotechnol. Biochem. 57, 1482-1 486. Suen, W.-C. and Gibson, D.T. (1993) Isolation and preliminary characterization of the subunits of the terminal component of naphthalene dioxygenase from Pseudomonas putida NCIB 9816-4. J. Bacferiol. 175,5877-5881. Suen, W.-C. and Gibson, D.T. (1994) Recombinant Escherichia coli strains synthesize active forms of naphthalene dioxygenase and its individual a and p subunits. Gene 143,67-71, Taira, K., Hirose, J., iiayashida, S. and Furukawa, K. (1992) Analysis of bph operon from the polychlorinated biphenyl-degrading strain of Pseudomonas pseudoalcaligenes KF707. J. Biol. Chem. 267,48444853. Takizawa, N., Kaida, N., Torigoe, S., Moritani, T., Sawada, T., Satoh, S . and Kiyohara, H. (1994) Identification and characterization of genes encoding polycyclic aromatic hydrocarbon dioxygenase and polycyclic aromatic hydrocarbon dihydrodiol dehydrogenase in Pseudotnonus putida OUS82. J. Bacteriol. 176,2444-2449. Tan, H.-M. and Mason, J.R. (1990) Cloning and expression of the plasmid-encoded benzene dioxygenase genes from Pseuhnonasputida ML2. FEMS Microbiol. Lett. 72,259-264. Tan, H.-M. (1991) The benzene dioxygenase genes from Pseudomonasputida ML2: cloning, expression and identification of functional domains. Ph.D. thesis, University of London. Tan,H.-M., Tang, H.-Y., Joannou, C.L., Abdel-Wahab, N.H. and Mason, J.R. (1993) The Pseudomonas putida ML2 plasmid-encoded genes for benzene dioxygenase are unusual in codon usage and low in G + C content. Gene 130,33-39. Tan, H.-M. and Cheong, C.-M. (1994) Substitution of the ISP a subunit of biphenyl dioxygenase from Pseudoinonas results in a modification of the enzyme activity. Biochem. Riophys. Res. Coininun. 204,912-917. Tan H.-M., Joannou, C.L., Cooper, C.E., Butler, C.S., Cammack, R. and Mason, J.R. (1994) The effect of fenedoxinBED over-expression on benzene dioxygenase activity in Pseudornonns puiida ML2. J. Bacieriol. 176,2507-25 12. Trumpower, B.L. (1990) Cytochrome bcl complexes of microorganisms. Microbiol. Rev. 54,101-129. Tsang, H.-Y., Batie, C.J., Ballou, D.P. and Penner-Hahn, J.E. (1989) X-ray absorption spectroscopy of the [2Fe-2S] Rieske clusters in Pseudoinonas cepaciu phthalate dioxygenase. Determination of core dimensions and iron ligation. Biochemistry 28, 7233-7240. Twilfer, H., Bernhardt. F.-H. and Gersonde, K. (1981) An electron-spin-resonance study on the redox-active centers of the 4-methoxybenzoate monooxygenase from Pseudomonas putida. Eu,: J. Biochem. 119,595-602. Twilfer, H., Bernhardt, F.H. and Gersonde, K. (1985) Dioxygen-activating iron centre in putidamonooxin. Electron spin resonance investigation of the nitrosylated putidamonooxin. Eu,: J. Biochern. 147, 171-176. Wackett, L.P. (1990) Tvluene dioxygenase from Psedomonas putida FI. In: Methods in Enzymology, V01188, Hydrocarbons nndMethylotrophy (M.E. Lidstrom. ed.), pp. 3 9 4 6 . Academic Press, New York. Wackett, L.P., Kwart, L.D. and Gibson, D.T. (1988) Benzylic monooxygenation catalysed by toluene dioxygenase from Pseudornonas putida. Biochemistry 27. 1360-1 367. Wang, Y., Gamon, J., Labbe, D., Bergeron, H. and Lau, P.C.K. (1995) Sequence and

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expression of the bpdClC2BADE genes involved in the initial steps of biphenylkhlorobiphenyl degradation by Rhodococcus sp M5. Gene 164,117-122. Wende, I?, Pfleger, K. and Bernhardt, EH. (1982) Dioxygen activation by putidamonooxin: substrate-modulated reaction of activated dioxygen. Biochem. Biophys. Res. Commun. 104,527-532. Wierenga, R.K., Terpstra, P. and Hol, W.G.J. (1986) Prediction of the occurrence of the ADP-binding pap-fold in proteins, using an amino-acid sequence fingerprint. J. Mol. Biol. 187. 101-107. Yamaguchi, M. and Fujisawa, H. (1978) Characterization of NADH-cytochrome creductase, a component in benzoate 1,2-dioxygenase system from Pseudomonas arvilla C-1 .J. Biol. Chem. 253,8848-8853. Yamaguchi, M. and Fujisawa, H. (1980) Purification and characterization of an oxygenase component in benzoate 1,2-dioxygenase system from Pseudomonas arvilla C-1, J. Biol. Chem. 255,5058-5063. Yamaguchi, M . and Fujisawa, H. (1982) Subunit structure of oxygenase component in benzoate-l,2-dioxygenasesystem from Pseudomonns arvilln C-1. J. Biol. Chem. 257, 12497-12502. Yang, Y., Chen, R.F. and Shiaris, M.P. (1994) Metabolism of naphthalene, fluorene and phenanthrene: preliminary characterization of a cloned gene cluster from Pseudoinonas pu/ida NCIB 9816. J. Bacteriol. 176,2158-2164. Zamanian, M. and Mason, J.R. (1987) Benzene dioxygenase in Pseudomonas putida. Subunit composition and immuno-cross-reactivity with other aromatic dioxygenases. Biochem. J. 244,611616. Ziffer, H., Kabuto, K., Gibson, D.T., Kobal, V.M. and Jerina, D.M. (1977) The absolute stereochemistry of several cis-dihydrodiols microbially produced from substituted benzenes. Tetrahedron 33,2491-2496. Zylstra, G.J. and Gibson, D.T. (1989) Toluene degradation by Pseudomonas putida F1, Nucleotide sequence of todCl C2BADE genes and their expression in Escherichia coli. J. Biol. Chern. 264, 14940-14946. Zylstra, G.J., McCombie, W.R., Gibson, D.T. and Finctte, B.A. (1988) Toluene degradation by Pseudomonas putida F1: Genetic organization of the lod operon. Appl. Environ. Microbiol. 54, 1498-1503.

Thiol Template Peptide Synthesis Systems in Bacteria and Fungi Rainer Zocher and Ullrich Keller lnstitut fur Biocheinie und Molekulare Biologie. Technische Universitiit Berlin. Franklinstrafle 29. 0-10587Berlin.Charlottenburg. Germany

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86 2. The peptide synthetase domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88 2.1. Peptide synthetases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 88 2.2. Motifs of the carboxyl-adenylate-formingdomain . . . . . . . . . . . . . . . . 90 2.3. Modules in the activation domain . . . . . . . . . . . . . . . . . . . . . . . . 90 2.4. The N-methylation module . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 2.5. Acyltransfer and epimerization modules . . . . . . . . . . . . . . . . . . . . . 91 2.6. Thioesterase modules in peptide synthetase genes . . . . . . . . . . . . . . . 92 2.7. Properties of amino-acid activating domains . . . . . . . . . . . . . . . . . . . 93 3 . Enzymesystems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94 3.1. Organization of activation domains in prokaryotes and eukaryotes . . . . . . . 94 4. Peptide synthetases from fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . 96 96 4.1 . 6(La-Aminoadipyl)-cysteinyl-o-valine . . . . . . . . . . . . . . . . . . . . . . 4.2. Enniatins and beauvericin . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 4.3. Cyclosporin and related peptides . . . . . . . . . . . . . . . . . . . . . . . 105 4.4. Ergot peptide alkaloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108 5 . Prokaryotic peptide synthetase systems . . . . . . . . . . . . . . . . . . . . . . 111 111 5.1, Acyi peptide lactones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2. Surfactin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 5.3. Bialaphos . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 6. Future prospects of peptide synthetase research . . . . . . . . . . . . . . . . . . 122 6.1. Domain exchange in thiol template peptide synthesis systems . . . . . . . . 123 6.2. Combinatorial approaches in future peptide synthesis development . . . . . 123 Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 124 124 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Abbreviations: Abu. or-aminobutyric acid; ACMS. actinomycin synthetase; ACP. acyl carrier protein; ACV. &(L-a-aminoadipyl)-cysteinyl-D-valine; ACVS. 6-(~-aaminoadipy1)-cysteinyl-D-valinesynthetase; AdoHCy. S-adenosyl-L-homocysteine; ADVANCES IN MICROBIAL PHYSIOLOGY VOL 38 ISBN 0-12-027738-7

CopyrightQ 1997 Academic Press Limited All rights of reproduction in any form reserved

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AdoMet, S-adenosyl-L-methionine; Bmt, (4R)-4-[(E)-2-butenyl]-4-methyl-~threonine; Cysyn, cyclosporin synthetase; D-Hiv, D-2-hydroxyisovaleric acid; DMPT, desmethylphosphinothcin; Esyn, enniatin synthetase; FAS, fatty acid synthase; GSI, gramicidin S synthetase I; GSII, gramicidin S synthetase 11; MeLeu, methyl leucine; 4-MHA, 4-methyl-3-hydroxyanthranilicacid; Orf, open reading frame; PKSI, polyketide synthetase type I; PKSII, polyketide synthetase type 11; Pt, phosphinothricin; Ptt, phosphinothricin tripeptide; Sar, sarcosine; TE, thioesterase; TYI, tyrocidine synthetase I.

1. INTRODUCTION

Bacteria and fungi produce numerous peptides as secondary metabolites, which are valuable as antibiotics, cytostatics. immunosuppressants, enzyme inhibitors and effectors acting on various cellular targets. It was speculated rather early that the mechanism of synthesis must differ from that of protein synthesis, since the vast majority of microbial peptides often contain unusual amino acids that are not found in proteins. Many of the non-proteinogenic amino acids have unique structures with respect to their carbon skeletons, presence of double bonds, unusual functional groups, aromatic character, D-configuration at a-C or presence of methyl groups at the amino groups involved in peptide-bond formation (Kurahashi, 1974; Kleinkauf and von Dohren, 1987, 1990). Also, the fact that such peptides in the producing organisms are very often formed as homologous series instead of single compounds indicates the broad specificity of enzymes responsible for incorporation of the relevant amino acids into peptides, which is in contrast to the specificity observed in protein synthesis. Nevertheless, the normal proteinogenic amino acids are present in most of the peptides of microbial origin, which indicates that the process of microbial peptide synthesis may have evolved in parallel to that of protein synthesis. The discovery of the activation with ATP of both acetate and amino acids led Lipmann, in the early 1950s, to postulate arelationship of peptide/protein synthesis with fatty acid synthesis, which have in common the mechanism of activation of building blocks and the direction of chain growth (head growth). In this model, peptide synthesis takes place on a template characterized by various condensation domains (Lipmann, 1954). The deciphering of the genetic code and later the elucidation of the ribosomal system eventually made it clear that protein synthesis mechanistically is very different from this model, especially in the nature of the template. However, the discovery that the cyclodecapeptide antibiotic gramicidin S is synthesized by enzymes instead of ribosomes independent of RNA (Berg et a]., 1965) confirmed the idea of a protein template directing the incorporation of amino acids into polymers (Lipmann, 1971, 1973). The first such templates with analogy to fatty acid synthase were found to be gramicidin S synthetase and later

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THIOL TEMPLATE PEPTIDE SYNTHESIS SYSTEMS IN BACTERIA AND FUNGI

GS I I

L-Om

L-Pro

L-Val

L-Pro

Activation

ESH+ aa + 2)

E.S.H.~~-AMP

MZ+

ATP -T-C

E.s .H.aa-~~~ 4 PPi

E

~ +- AMP ~

~

Figure 1 Structure of gramicidin S and its assembly by the protein thiol templates gramicidin synthetase I and I1 (GSI and GSII). Each of the five domains of the multifunctional enzyme system activates its corresponding amino acid as adenylate, subsequently binds it to a specific thiol in thioester linkage as shown in lower part of the figure. GSI epimerizes thioester bound Phe to D-Phe prior to transfer of this residue to GSII where condensation of the amino acids takes place. Two pentapeptide chains are finally dimerized in head-to-tail condensation.

tyrocidine synthetase, which polymerize amino acids in a stepwise fashion into covalently bound peptidyl intermediates (Gevers et al., 1969; Roskoski et al., 1970). Gramicidin S synthetase I (GS1)-at that time with an estimated molecular mass of 100 kDa-activates the starter amino acid L-phenylalanine as adenylate and binds it in thioester linkage. Similarly, the 280 kDa gramicidin S synthetase I1 (GSII) recruits the residual four amino acids of gramicidin S as thioesters (Fig. 1). After epimerization of phenylalanine, the various amino acids are polymerized into a pentapeptide chain of the sequential order as shown in Fig. 1. Head-to-tail condensation of two of these chains yields gramicidin S. The same mechanism of formation was also shown with tyrocidine, which is structurally similar to gramicidin S (Fig. l), but has more different amino-acid positions and therefore requires more enzymes for its assembly (Lipmann, 1973). The presence of 4'-phospho- pantetheine in these enzymes strongly supported the thiol template model in which the enzyme is composed of activating domains rather than of subunits, with each responsible for activation of one single amino acid. Each domain has an Mr equivalent of more than 70 000, which reflects that GSII is able to activate four amino acids in a defined sequence (Lipmann, 1973). An essential

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feature in every domain is the presence of a specific (peripheral) thiol group to which the corresponding amino acid becomes attached in thioester linkage. A central sulfhydryl was proposed to reside on the 4’-phosphopantetheine group, which, like the swinging arm of acyl carrier protein (ACP) in fatty acid synthesis, was to carry the growing peptide chain from one reaction centre to the next (Lynen, 1972). Thus, the analogy to fatty acid synthase (FAS), which acts upon acyl-coenzyme A (CoA) thioesters with subsequent covalent substrate binding is apparent except that the peptide synthetases consist of an array of similar domains each directing the incorporation of its cognate amino acid into the growing peptide chain. In contrast, in fatty acid synthesis, it is always the same extender unit (ie. malonyl coenzyme A) that contributes to fatty acyl chain growth. Accordingly, FAS has only two thiol groups, one on the P-ketoacyl synthase and the other on the ACP (reviewed by Hopwood and Sherman, 1990). Structural and functional studies, which became possible through the cloning and sequencing of genes encoding peptide synthetases, have led to a refinement of the thiol template mechanism of peptide formation (Schlumbohm et al., 1991; Marahiel, 1992; Stachelhaus and Marahiel, 1995a,b). Here we review the current state of research in this field with respect to the basic structural and functional molecular features of the building blocks of peptide synthesis systems. These molecular concepts will be illustrated in the course of the text by the description of selected examples of bacterial and fungal peptide synthesis systems. Finally, aspects of future developments of peptide synthesis systems such as the creation of recombinant enzymes through directed or combinatorial approaches will be discussed.

2. THE PEPTIDE SYNTHETASE DOMAIN

2.1. Peptide Synthetases The first peptide synthetase genes to be cloned and sequenced were those of GSI (Kratzschmar el al., 1989), tyrocidine synthetase I (TYI) (Weckermann et al., 1988), GSII (Krause et al., 1985; Turgay etal., 1992), and &(L-a-aminoadipyl)-Lcysteinyl-D-valine synthetase (ACVS) (Diez et al., 1990; Smith et al., 1990a,b; Gutierrez etal., 1991; MacCabe etal., 1991). The sequences ofthemulti-functional proteins activating more than one amino acid consist of repeating units, each encompassing a length of approximately 1000 amino acids corresponding to a molecular mass equivalent of 120 kDa. Each of these units show considerable sequence conservation to each other and the number of these units equals the number of the amino acids serving as substrates of the corresponding enzymes. Accordingly, single amino-acid activating enzymes such as TYI or GSI each consist

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THIOL TEMPLATE PEPTIDE SYNTHESIS SYSTEMS IN BACTERIA AND FUNGI

of only one unit. These findings lend compelling evidence for the view that each unit is responsible for the assembly of a single amino acid in the process of peptide formation. The sequence conservation in the units is particularly high in a 600 amino-acid-long central part (designated amino-acid activation domain) and is less, but still significantly, conserved in the interdomain sequences (spacers), which comprise some further 500 amino acids. Interpretation of the gene sequence data also revealed that the peptide synthetase proteins were much larger in size than had been estimated from previous mass determinations when no suitable MI markers were available. Thus, GSI and GSII were found to have molecular masses of 121 kDa and 510 kDa, respectively (Kratzschmar et al., 1989; Turgay et al., 1992).Inspection of the highly conserved region not only in the peptide synthetases mentioned above but also in a steadily increasing number of genes that have been cloned in the last 5 years has led to the identification of characteristic amino-acid sequence motifs (Marahiel, 1992; Stachelhaus and Marahiel, 1995a).Figure 2 shows that these motifs are spread over the entire length of the amino-acid activation domain and apparently play a role in functioning of the synthetase. Their sequence conservation is high enough to allow the construction of highly specific oligonucleotide primers for screening of new

peptide synthetase activation domain

I

I methyltransferase module -450 aa

thioester adenylylcarboxylate domain

racemuatiod transfer epimerization acy’

module

module

7 I I l l r n -l ---.LIT= C D E

- r - - v r - -I 200

400

I

u

I

600

I

I 800

I

I

1000

I

I

1200 aa

Figure 2 Assembly of the most highly conserved sequence motifs in peptide synthetase domains. Motifs A-E are highly conserved in adenylate forming enzymes. A(LKAGGAYVPID), B(YSGTTGXPKGV), C(GELCJGGXGXARGYL), D(YXTGD), E(VKIRGXRIELGEIE), F(DNFYXLGGHSL). Motif F represents the attachment site for the 4’-phosphopantetheine cofactor (thioestcr module). Motifs I-IV are conserved in peptide synthetase domains harbouring epimerase activity. I(AYXTEXND1LLTAXG). II(EGHGREXIIE), III(RTVGWFTSMYPXXLD), IV(FNYLGQFD). The acyltransfer module is characterized by the consensus HHXXXGD (“spacer or His motif’) which is found in acyltransferases. The methyltransferase module is present in peptide synthetase domains catalysing synthesis of N-methylated peptide bonds and has sequence motifs common with methyltransferases. The sequences of the various motifs are given in the one letter aa code.

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peptide synthetase genes in various organisms (Borchert et al., 1992; Turgay and Marahiel, 1994). Interestingly, five of the six strongest conserved motifs were also seen in a number of acyl CoA ligases such as coumarate ligase (Knobloch and Hahlbrock, 1975; Lozoya et al., 1988) and carboxyl-adenylate ligases, such as luciferase (de Wet et al., 1987), but not the aminoacyl-tFWAsynthetases indicating the existence of a distinct superfamily of adenylyl-carboxylate-formingenzymes (Turgay el al., 1992).

2.2. Motifs of the Carboxyl-adenylate-forming Domain Most of the work to elucidate the significance of the various conserved motifs in adenylate-forming domains (Fig. 2) has been done by studying peptide synthetase domains carrying site-directed mutations in various core motifs or by specific labelling of active site-located residues with ATP analogues or structurally related substrates. These analyses have been mostly done with TYI and to a limited extent with GSII. Experimental evidence was obtained that motif boxes B-E (Fig. 2) are involved in ATP binding and formation of carboxyl-adenylate. As suggested from its structuralresemblance with the Walker type A(phosphate-binding loop) (Walker etal., 1982), mutations in the glycine-rich sequence in motif B (GXXXXXGKT/S), such as replacement of the conserved lysine by unrelated amino acids, had drastic effects on the ability of the enzyme to activate D- or L-phenylalanine as adenylate. Similarly, mutations in the conserved aspartate in motif D resulted in loss of catalytic activity of the same reaction indicating a significant role of motifs B and D in ATP binding (Gocht and Marahiel, 1994). A specific role of motifs C and E in ATP binding and adenylation became clear through site-specific labelling of TYI with 8-azido-ATP and with fluorescein isothiocyanate, respectively (PavelaVrancic et al., 1994a,b). In other experiments it was shown that replacement of the conserved glycine in motif E (KIRCXRIEL) of the proline domain of GSII significantly affected the proline activation reaction (Tohika et al., 1993).

2.3. Modules in the Activation Domain

Analysis of peptide fragments carrying radioactively labelled substrate amino acids, such as leucine or valine (obtained after proteolytic cleavage of charged GSII), revealed that the motif F is that site of the amino-acid activating domain of peptide synthetases where the amino acid is attached to the enzyme in thioester linkage. Instead of the presence of a conserved cysteine, as was postulated in the previous model of the protein thiol template concept, the F consensus sequence GG IUD S WI shows similarity to the attachment site of the 4'-phosphopantetheine cofactor in numerous fatty acid and polyketide synthases (Hopwood and Sherman, 1990; Katz and Donadio, 1993). This sequence motif possesses a conserved senne

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91

leading to the view that each amino-acid activating domain of the peptide synthetase harbours a 4’-phosphopantetheine arm,which functions both in aminoacid attachment and peptidyl transfer (Schlumbohmetal., 1991; Stein etal., 1994). Replacements of the corresponding serine residues by alanine in various peptide synthetase domains, such as in surfactin synthetases (D’Souza et al., 1993) or in TYI (Gocht and Marahiel, 1994), abolished thioester formation activity while leaving the adenylation activity of the enzymes unaffected. These findings required a revision of the previous model of peptide synthetases in which each activating domain possesses a 4’-phosphopantetheine cofactor as carrier for both amino acid and peptidyl intermediates. This is in contrast to the situation in fatty acid or polyketide synthase (PKS) enzyme systems where 4’-phosphopantetheine is attached to ACP only as a single swinging arm (Hopwood and Sherman, 1990).

2.4. The NMethylation Module

Numerous peptides contain N-methylated amino acids in their peptide chains (Billich and Zocher, 1990). Biochemical evidence has indicated that, in the case of enniatin, cyclosporin and actinomycin biosynthesis, the N-methylation takes place after covalent binding of the amino acid on the surface of the corresponding peptide synthetases with S-adenosyl-L-methionine (AdoMet) as substrate (Zocher et al., 1982, 1986; Keller, 1987). Haese et al. (1993) eventually showed that the amino-acid activating domain of enniatin synthetase (Esyn), the enzyme catalysing cyclohexadepsipeptide synthesis from branched-chain amino acids and D-2hydroxyisovalerate, contains an additional stretch of 450 amino acids inserted into a position between the conserved sequences E and F, the thioester formation module (Fig. 2). The 450 amino-acid module contains characteristic motifs with similarity to conserved sequences of various methyltransferases (Haese et al., 1993).No such 450 amino-acid insertion is seen in the hydroxy-acid activating domain of Esyn. These findings clearly assign the N-methylation function to the 450 amino-acid insertion. The N-methylation module was later seen also in the sequence of the cyclosporin synthetase gene, where it is present in 7 of the 11 amino-acid activation domains (Weber et al., 1994). The cycloundecapeptide cyclosporin A contains seven N-methylnted amino acids. Their relative position in the cyclopeptide is consistent with the order of the N-methylation modules in the peptide synthetase sequence.

2.5. Acyltransfer and Epimerization Modules

The observation of conserved motifs in a 350-amino-acid stretch distal to the amino-acid activation domains of peptide synthetases with the potential to elongate and epimerize amino acid or peptidyl residues, has led to the tentative assignment

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of these motifs to acyl transfer and epimerization functions in these enzymes puma eta!., 1993; de CrCcy-Lagard et al., 1995; Stachelhaus and Marahiel, 1995a; Stein et al., 1995). A set of four highly conserved sequences is proposed to be involved in racemizatiodepimerization reactions. These motifs, designated I-IV (Fig. 2), were found only in peptide synthetase domains known to be involved in the incorporation of D-amino acids in the corresponding peptides. A HHXXXDG motif, previously named “spacer motif’ (Stachelhaus and Marahiel, 1995a) and which pervades all modules catalysing peptide transfer or epimerization, is also present in the Tn9 chloramphenicol acetyltransferase (CAT) family as well as in members of the dihydrolipoamide acyltransferase (E2p) family, which catalyse acyl transfers (Guest, 1987). The “spacer motif’, also called the “His motif’, is always located to the carboxyterminal side of activation domains in peptide synthetases at a constant distance ( 170-190 amino acids) from the 4’-phosphopantetheine attachment site (F), except in those rarer cases where it precedes an activation domain of peptide synthetases that receive acyl groups from another enzyme, such as in the case of surfactin synthetase E1A (see below). The second histidine in the “His motif’ has been suggested to have a catalytic role as a general base catalysed in the deprotonation events required for acyl transfer reactions as well as in the epimerization of covalently bound amino acids or peptides (de CrCcy-Lagard et al., 1995). The probable involvement of proton acceptor group was shown in epimerization reactions catalysed by peptidyl epimerase functions, such as of actinomycin synthetase I1 (see below). Stachelhaus and Marahiel (1995b) presented evidence that the 350-amino-acid stretch harbouring the putative four racemization motifs (I-IV, Fig. 2) is in fact responsible for amino-acid racemization. Deletion mutants of GSI lacking this region do not epimerize covalently bound phenylalanine, while other functions of the synthetase, such as adenylation and thioesterification of the substrate, were unaffected.

2.6. Thioesterase Modules in Peptide Synthetase Genes

Analysis of open reading frames (orfs) in several peptidc-synthetase gene clusters revealed the presence of thioesterase (TE) genes with similarity to predicted vertebrate fatty acyl thioesterase I1 enzymes from duck and rat, which can transfer and thus catalyse release of fatty acids from fatty acid synthase. Such orfs are olfl and orf2 of the bialaphos synthesis gene cluster of Srreptoinyces hygroscopicus and grsT, associated with gramicidin S synthetase genes grsA and grsB in Bacillus brevis (Kratzschmar et al., 1989; Raibaud et al., 1991). By analogy to fatty acid synthesis, such thioesterase genes probably catalyse, in peptide synthesis, release of the completed product from the synthetase. In the case of ACVS from various sources, a thioesterase module was found in the C-terminal region of the gene as an integral part of the protein (Smith etal., 1990a; MacCabe etal., 1991a,b).These data further support the prediction made by Lipmann (1971) that peptide and

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fatty-acid enzyme complexes share similarities with each other and may have common evolutionary origins. Meanwhile, cloning and sequencing of numerous polyketide synthase genes has revealed that polyketide antibiotics are also synthesized by multi-functional enzymes similar to FAS (Hutchinson and Fuji, 1995). All of these processes require cleavage of thioester bonds; therefore, thioesterase motifs are found either as an integral part of the polyketide synthases or as separate proteins. How product release (and cyclization) is accomplished in those cases of peptide synthetases where no thioesterase module is known, is unclear. It is also unclear in some cases whether thioesterase genes, which are associated with FAS genes, are really necessary for fatty acid synthesis (discussed by Hutchinson and Fuji, 1995).This point should be considered also in thiol template peptide synthesis.

2.7. Properties of Amino-acid Activating Domains

As shown in Fig. 1, the activation of amino acids proceeds in two steps. First, the amino acid is activated as adenylate, essentially as in the case of proteinogenic amino acids by the aminoacyl-tRNA synthetases. In the second step, the amino acid is covalently bound in thioester linkage by nucleophilic attack of the 4'-phosphopantetheine-SH to the acyladenylate. As shown by mutational analysis (Stachelhaus and Marahiel, 1995b) or by blocking the thioester formation module with sulfhydryl-directed agents (Zocher e l al., 1982), the adenylation reaction is independent of the presence of the thioester module. In this way the non-ribosomal system resembles the amino-acid activation by aminoacyl-tRNA synthetases, which activate their cognate amino acids as adenylates even in the absence of tRNA (Schimmel and Soll, 1979). In some cases inactivation or omission of the thioester formation module has led to significant reduction in adenylation reactions (Pfeifer etal., 1995), which might be due to structural constraints in the activation domain arising by deletion of such a long space-filling cofactor as 4'-phosphopantetheine. The 4'-phosphopantetheine also plays an important role in the epimerization reactions in peptide synthetases because the epimerized amino acids or peptides are attached to this cofactor in thioester linkage. The chiral centre inverted is the a-carbon adjacent to the thioestercarbonyl of the relevant residue. Interestingly, single arnino-acid activating enzymes, such as tyrocidine synthetase I or gramicidine synthetase 11, activate both L- and D-phenylalanine, and racemize these amino acids in their antipodal products (Gocht and Marahiel, 1994; Stein et al.. 1995). This is not true in the case of peptidyl epimerization. where the L-enantiomers are always activated and epimerization most probably takes place after peptide-bond formation (see the examples for actinomycin and surfactin below). Generally, one observes a rather strict stereospecificity in the activation reactions of the nonribosomal system, as in protein synthesis. By contrast, the specificity in terms of structural homology of substrates is lower than in protein synthesis, although in the mixtures of homologous series of peptides produced by microorganisms mostly

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conservative exchanges of amino acids occur. These are, of course, much more frequent than in protein synthesis, where misactivations are eliminated by proofreading mechanisms. How such proof-reading mechanisms in non-ribosomal peptide syntheses would work is not known.

3. ENZYME SYSTEMS

3.1. Organization of Activation Domains in Prokaryotes and Eukaryotes

Principally, peptide synthetase systems are defined as the arrangement of various amino-acid activation domains in the form of a multi-enzyme or a multi-enzyme complex. This has been shown by comparisons of various enzyme systems with their corresponding genes (Fig. 3). The order of the various activation domains is mirrored in the sequence of the peptide synthesized. In prokaryotes, one generally observes that the domains are distributed over more than one polypeptide chain with the exception of the single chain ACVS, which, however, is common to both the lower eukaryotes and prokaryotes (Aharonowitz et al., 1993). By contrast, all eukaryotic peptide synthetases, yet known, always consist of a single polypeptide chain encoded by an intronless gene. This single polypeptide chain harbours the various adenylate formation domains, thioester and additional modules necessary for the synthesis of a given product. This parallels observations concerning the modular organization of fatty acid synthases in prokaryotes and eukaryotes, which differ from each other in their structural and functional organization (Hopwood and Sherman, 1990).While in E. coli the FAS complex consists of ten different subunits each harbouring a distinct catalytic function, FAS from vertebrates is one polypeptide chain, and that of yeast consists of two polypeptide chains (Hopwood and Sherman, 1990).Similarly, in polyketide syntheses, which strongly resembles fatty acid syntheses, two types of polyketide syntheses have been described. While PKSI systems, as i n the case of erythromycin, consist of giant multi-enzymes containing the ACPdomain, P-ketoacyl synthase domain, sites for ketoreduction, dehydration, enoylreduction and thioesterase located on one polypeptide chain, PKSII systems consist of many separate largely monofunctional enzymes (Roberts et al., 1993). In contrast to the different FAS systems in prokaryotes and eukaryotes, no correlation of PKSI and PKSII systems with their occurrence in prokaryotes and eukaryotes is seen. The only exception is 6-methyl salicylic acid synthase of fenicilliuin patuluin, which is of PKSII-type but is represented by one single polypeptide chain (Hutchinson and Fuji, 1995). Besides determining the sequence of the peptide synthesized, the sequential order of activation domains and their accompanying modules is also important for

95

THIOL TEMPLATE PEPTIDE SYNTHESIS SYSTEMS IN BACTERIA AND FUNGI

Arrangement of Domains in Peptide Synthesis

ml I

ProkarVoteS

I I

1 I

Pro Val Orn Leu

Gramicidin S

Leu Asp D-Leu E

Glu Leu D-Leu E

Surlactin

Eukarvotes

m]

Enniatin B

I I I I

I I II I I I

I I

I

I

D-AlaI Leu Me1 Leu Me( Val Me] Bmt Me Abu Gly Me1 Leu Me1 Val Leu lMel Ala Cyclosporin A ACV

Figure 3 Organization of peptide synthetase domains in various multi-functional peptide synthesis systems. The order and organization of peptide synthetase domains in several thiol template peptide synthesis systems is shown as deduced from DNA sequences of the corresponding enzymes and from analysis of their enzymatic activities. ACV. 6-(L-a-aminoadipyl)-cysteinyl-D-valine synthetase. Symbols in the adenylate forming domains indicate the comesponding activated amino acids: Aad, 8-L-a-amino adipic acid; D-Hiv, D-2-hydroxyisovaleric acid; Abu, a-amino-L-butync acid; Bmt, 4-(E)-butenyl-4methyl-L-threonine. The symbols E and TE denote presence of epimerase modules and thioesterase module, respectively. Me, N-methyltransferase.

the class of product, i.e. linear or cyclic, homodetic or heterodetic, etc. In any case, acyl transfer modules such as the “spacer motif’ have to be present between each activation domain of a biosynthetic sequence to enable condensation of the amino-acid residues. In the case of prokaryotic peptide synthetases, which receive acyl or peptidyl intermediates from another multi-functional enzyme, the same motif has also to be present in front of the first amino-acid activation domain, in order to enable the acylation of the first amino acid. The structural principles of the peptide synthetase domains outlined above would allow one to make similar predictions for the synthesis of other peptide classes such as those containing N-methylated peptide bonds. Whether such predictions would really lead to successful synthesis is dependent on many factors, most important among which are programming and timing of events in the assembly of the peptide. Most important may also be the priming of the reaction, the reactivity of the peptidyl intermediates and eventually the size of the cavity in the multi-enzyme where condensation and cyclization reactions take place, which may play a significant role for the chain length of the product. In the case of polyketide synthesis, there

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are examples of the effect of exchanging domains from different biosynthesis systems with consequences for the nature and length of the polyketide chain formed by engineeredPKS complexes (Hutchinson and Fuji, 1995;Tsoi and Khosla, 1995). In peptide synthesis systems, factors governing the programming of chain growth are as yet largely unknown. Most of the information on the reactions of peptide synthetase domains can therefore be obtained only by combination of both genetic and biochemical investigations of the reactions with enzymes in their wild-type and mutated form. In the following, the principles presented above of the structures of peptide synthetase domains will be illustrated by description of some selected, important peptide synthesis systems, the enzymology of which is well developed. These systems include prokaryotic as well as eukaryotic peptide synthetases responsible for the synthesis of cyclic and linear peptides, as well as N-methylcyclodepsipeptides.

4. PEPTIDE SYNTHETASES FROM FUNGI 4.1. 6-(L-a-Aminoadipyl)-cysteinyl-D-valine

&(L-a-Aminoadipyl)-cysteinyl-D-valine(ACV) is the common precursor of the penicillins and cephalosporins (Fig. 4). The peptide is assembled by ACV synthase (ACVS), which has been isolated from a representative number of fungi and bacteria. Based on biochemical investigations in the cases of the enzymes from A. nidulans and S. clavuligerus (van Liempt etal., 1989;Jensen et al., 1990; MacCabe et al., 1991a; Schwecke et al., 1992) and also considering the sequences of a number of ACVS genes (see below), it is clear that this enzyme is composed of three peptide synthetdse domains lying on one polypeptide chain of 420 kDa (Aharonowitz e f al., 1993).

SH

Figure 4 Structure of 6-(L-a-aminoadipyl)-cysteinyl-D-valine. 6-(L-a-aminoadipyl)cysteinyl-D-valineis the common precursor of the penicillins and cephalosponns.

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4.1.1. ACVS Activates a- and 8-Carboxyl Groups ofAmino-acid Substrates Biochemical work has revealed that three independent sites (domains) are responsible for the activation of a-L-aminoadipic acid, L-cysteine and L-valine. While the first activation step of the latter two amino acids is clearly via the corresponding adenylates, conflicting results have been obtained regarding the a-L-aminoadipic acid. The enzyme of A. nidulans catalysed adenylation of a-aminoadipic acid as measured by aminoadipic acid-dependent ATP-pyrophosphateexchange, while the Streptomnyces clavuligerus ACVS did not (van Liempt et al., 1989; Schwecke et al., 1992). In the case of glutathione y-L-glutamyl-L-cysteinyl-glycine, the y-carboxylic group of glutamate is activated as phosphoryl-carboxylate from ATP with formation of ADP and the a-carboxyl group of y-glutamyl-L-cysteine is activated in the same way (reviewed by Meister, 1988). Thus, aminoadipic acid activation differs in its mechanism from the normal 0-carboxyl or peptide activation found in other systems of enzymatic peptide synthesis, such as of glutathione or muramyl peptide in bacterial cell-wall formation (Lipmann, 1980). However, covalent binding to ACVS through thioester linkage is observed with all of the three amino acids in all cases examined and 4'-phosphopantetheine has been shown to be present in several ACVSs tested (Baldwin etal., 1990, 1991). 4.1.2. Epirnerization in ACV Synthesis The fact that all of the valine attached to the enzyme in the activation step has the L-configuration indicates that epimerization of the L-valine is later during the events of peptide formation or in the peptide-bound state (van Liempt et al., 1989; Baldwin et al., 1991). Possibly, a thioesterase would be active in the release of the LLD peptide but not of the LLL intermediate, if the latter exists. Investigations of the reaction mechanism in terms of analysis of covalently bound peptidyl intermediates have not yet been reported. Recent findings, however, indicate that without L-aminoadipic acid, ACVS from Cephalosporiurn acrernoniurn forms and releases L-cysteinyl-D-valineas the product (Shiau et al., 1995b).Formation of the same product was enhanced in the additional presence of glutamate, which was activated by the enzyme but not incorporated in the product. Glutamate is a structural analogue of aminoadipate. The implications from these results are that the peptide bond between cysteine and valine is formed prior to epimerization, possibly in the peptide-bound state and also prior to the formation of the peptide bond between aminoadipate and cysteine. This mechanism would indicate a novel mechanism of peptide formation in thiol template peptide synthesis systems concerning the timing of peptide-bond formation events, which is not correlated with the order of the activation domains on the ACVS polypeptide chain. This mechanism would contrast with the previous model of ACV synthesis (Fig. 5). Interestingly, the cysteinyl-D-valine product was released from the enzyme

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I E I [TEl

Figure 5 Two possible schemes for events of peptide assembly catalysed by &@-aarninoadipy1)-cysteinyl-D-valine synthetase (ACVS). (a) Activation and peptide-bond formation steps in timely correlation with the order of peptide synthetasedomainson the ACVS polypeptide chain. (b) The start of the peptide-bond formation steps at the internal domain position with subsequent amino terminal peptide acylation. Note that in the threedimensional structure of ACVS, the Aad domain possibly could be vicinal in space to the Val domain. For details, see text. Aad, 6-L-a-aminoadipic acid; E, epimerase; TE, thioesterase.

indicating that the releasing enzyme activity in ACVS might recognize peptides shorter than ACV. Shiau et al. (1995a) have also shown that structural analogues of L-cys-D-Val such as 0-methyl-serinyl-D-valineare also efficiently formed and released from the synthetase together with a minor amount of 0-methyl-seryl-Lvaline. The latter finding would indicate that release of end product catalysed by the thioesterase module would not require strict stereospecificity at the a-carbon of valine in the peptidyl-thioester. 4.1.3.

ACVS Genes and Sequence Similarities of Peptide Synthetase Domains

Much of the knowledge on ACVS stems from sequence analyses of ACVS genes, which have been obtained from a variety of sources, such as actinomycetes, Gram-negative bacteria and fungi (Smith et al., 1990a,b; Coque et al., 1991; Diez et al., 1990, Tobin et al., 1990; Gutierrez et al., 1991; MacCabe et al., 1991b). In all cases, one intron-less orf has been found encoding a polypeptide of molecular mass around 420 kDa. This has facilitated determination of the conservation of fungal and prokaryotic peptide synthetase domains responsible for aminoadipic acid, cysteine and valine activation. Construction of phylogenetic trees on the basis

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of sequence similarities has clearly placed each of the three domains of ACVS in a separate branch regardless of their prokaryotic or eukaryotic origin (Turgay ef al., 1992). This might indicate horizontal gene transfer of ACVS domain genes between prokaryotes and eukaryotes, as has been discussed by Smith etal. (1990a). Similarly, genes encoding enzymes converting ACV to p-lactams, such as isopenicillin N-synthase. have led to similar conclusions of horizontal gene transfer of other p-lactam biosynthesis genes (Weigel et al., 1988). Interestingly, different fl-lactambiosynthesis genes involved in the formation of ACV and its modification have been found to be clustered in fungi and bacteria, which would support the horizontal gene transfer hypothesis (Smith ef al., 1990a; Gutierrez ef al., 1991; Martin and Gutierrez, 1995). Whether these hypotheseses will also lead to an evolutionary tree of peptide synthetase domains will be seen in the future.

4.2. Enniatins and Beauvericin

Enniatins are cyclohexadepsipeptides produced by various strains of the genus Fusariuin (Plattner et al., 1948). As shown in Fig. 6, enniatins consist of three residues of a branched chain N-methyl amino acid and D-2-hydroxyisovaleric acid (D-Hiv), arranged in an alternate fashion. Enniatins have antibiotic activity against various bacteria, exhibit immunomodulatory properties (Simon-Lavoine and Forgeot, 1979), and are potent inhibitors of mammalian cholesterol acyl transferase (Tomoda et al., 1992). Besides, enniatins are well known for their behaviour as ionophors with high specificity for potassium ions (Wipf et al., 1968). Enniatinproducing Fusaria are plant pathogens and enniatins were postulated to play a role as wilt toxin during infections of plants (Walton, 1990). Interestingly, enniatins like

Figure 6 Structures of enniatins and of beauvericin. Enniatin A: R1 = R 2 = R3 = sec-butyl; enniatin A l : R1 = iso-propyl, R2 = R3 = sec-butyl; enniatin B: R1 = R2 = R3 = iso-propyl; enniatin B1; R1 = R2 = iso-propyl, R3 = sec-butyl; beauvericin: R1 = R2 = R3 =

benzyl.

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the structurally related cyclodepsipeptides beauvericin and bassianolide also exhibit entornopathogenicproperties (Grove and Pople, 1980). 4.2.1. Enniatin Synthetase: Structure and Function Enniatins are synthesized by the multi-functional enzyme enniatin synthetase (Esyn), which was the first characterized N-methyl-cyclopeptide synthetase isolated from Fusarium scirpi (Zocher et al., 1982). Sequencingof the Esyn gene has revealed that the enzyme is one single polypeptide chain of 347 kDa (Haese et al., 1993). This is consistent with earlier biochemical investigations in which all catalytic functions of the enzyme were located on one polypeptide chain. Esyn synthesizes the enniatin molecule from its primary precursors D-Hiv and a branched-chain amino acid like L-valine or L-isoleucine in an ATP-dependent manner. AdoMet donates the methyl group in the N-methylated peptide bonds. As in other peptide synthetases 4'-phosphopantetheine is present as the prosthetic group. Dissecting the biosynthetic process in the individual steps catalysed by the individual domains of Esyn has revealed the picture of reaction steps shown in Fig. 7. Activation of D-Hiv and the branched-chain L-amino acids (e.g. L-valine) proceeds through adenylation and thioester formation. Characteristically, the thioesterified amino acid is methylated with AdoMet, and thus methylation takes place prior to peptide-bond formation and subsequent cyclization reactions. Omission of AdoMet in the in vitro system leads to formation of desmethyl enniatins (Zocher et al., 1982; Billich and Zocher, 1987a). Therefore, Esyn and other N-methylcyclopeptide synthetases can be considered as hybrid systems between peptide synthetases and N-methyltransferases. 4.2.2. Mechanism of N-Methylation of Amino A c i h in Thiol Template Peptide Synthesis A characteristic property of all N-methyltransferasesthat have been studied so far is their sensitivity to inhibition by S-adenosyl-L-homocysteine(AdoHCy), which is a reaction product of the methylation reaction derived from the methyl donor AdoMet. The antibiotic sinefungin, which is structurally related to AdoMet, is a potent inhibitor of a variety of methylases. Billich and Zocher (1987a) tested the effect of sinefungin and AdoHCy on product formation catalysed by Esyn, and found that sinefungin inhibits the methylation reaction of Esyn but allowed synthesis of desrnethyl enniatin even if present in excess amounts. Kinetic analysis showed a competitive inhibition pattern with respect to AdoMet, indicating direct competition of sinefungin with the AdoMet-binding site. By contrast, AdoHCy not only blocked formation of enniatin (methylated product) but also that of the unmethylated product. Kinetic analysis of the desmethyl enniatin synthesis revealed that, with respect to AdoMet, AdoHCy is a partial competitive inhibitor.

THIOL TEMPLATE PEPTIDE SYNTHESIS SYSTEMS IN BACTERIA AND FUNGI

Substrate activation 1

ATP + 0 - H i v

-

+E

( D - H ; ~ AMP)E

21 ATP + L-Val + E Cj ( L - V a l Thioester formation

3) (D-Hiv

-

AMPIE

4 ) (L-Val - A M P ) E

+

D-Hiv

+

L-Val

101

+ ppi

-

AMPIE

-

S-E + A M P

+ PPi

S-E + A M P

N-Methylation

5) L-Val

-

AdoMet

S-E-

L-MeVal

- S-E

Pep tide-bond forma tion

6) D-HIv

-S -

L-MeVal

1

S’

E-f

D-Hiv-L-MeVal

HS, S’

-

Ester-bond formation

7) 0-Hiv-L-MeVal

-

S-E

Cvcl i Z 8 t i O f l

cyclo-[ o - H i v - ~ - M e V a l ]+- ~E Enniatin B

Figure 7 Scheme of partial reactions leading to enniatin catalysed by enniatin synthetase.

These results indicate that Esyn must harbour a discrete binding site for the inhibitor AdoHCy but not for sinefungin. Thus, blocking the N-methyltransferase function of Esyn through AdoHCy results in inhibition of the peptide-bond formation ability of Esyn (R. Zocher, unpublished). These data suggest that the active sites for depsipeptide formation and for the methylation step in the Esyn system are not independent of each other as was confirmed later by the analysis of the sequence of Esyn (see below).

4.2.3. Substrate Specificity ofEsyn Owing to the relatively broad substrate specificity of Esyn for amino and hydroxy

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acids, a variety of different enniatins can be synthesized by Esyn if appropriate concentrations of substrates (depending on the various K , values) are used (R. Zocher, unpublished). Nevertheless, Esyns from different Fusarium strains differ in their amino-acid specificity, i.e. they can have different K , values for each of the various branched-chain amino acids (Pieper et al., 1992). For example, the enzyme from the enniatin A producer E sarnbucinum exhibits high affinity for the substrate amino acids L - ~ and U L-Ile. By contrast, Esyn from the enniatin B producer R Zateritiurn preferably accepts L-Val, the constituent amino acid of enniatin B and therefore strongly resembles the Esyn from R scirpi. The reasons for the altered substrate specificity among Esyns may lie in mutations in the amino-acid binding sites of the polypeptide chains. 4.2.4. Mulecular Structure of Esyn Monoclonal antibodies directed to the multi-enzyme Esyn were used to map the catalytic sites of the enzyme (Billich et al., 1987). The antibodies could be divided into three groups based on their influence on catalytic functions. Members of group one exclusively inhibited L-Val thioester formation, while members of group two interfered with D-Hiv thioester formation. Antibodies of group three inhibited both L-Val thioester and D-Hiv thioester formation as well as the N-methyltransferase. From these findings, it was concluded that the two domains of Esyn containing the two catalytically binding sites are situated very close to each other in the three-dimensional structure of the enzyme. The immunochemical data with group three antibodies indicate that this is also the case for the N-methyltransferase site, which is in the vicinity of the acyl and aminoacyl binding sites. Titration of Esyn with radioactive AdoMet revealed binding of one mole AdoMet per mole Esyn (Billich and Zocher, 1987a) indicating the presence of one methylase unit per enzyme molecule. Interestingly, the adenylation reactions for D-Hiv and L-Val were not affected by the monoclonal antibodies, indicating that these reactions have different sites on the multi-enzyme at some distance to the thiol sites confirming the modular structure of peptide synthetase domains. 4.2.5. Structure of the Esyn Gene Analysis of the sequence of the Esyn gene (esynl)of Fusarium scirpi revealed an open reading frame of 9393 bp encoding a 347 kDa polypeptide that contains two highly conserved peptide synthetase domains (designated EA and EB) (see Fig. 8) (Haese eta!., 1993).The domain EA lying on the amino-terminal side of the protein could be identified as the D-Hiv binding site and EB as the L-amino-acid binding site on the carboxytenninal side. In contrast to domain EA, domain EB is interrupted by insertion of a 434 amino-acid portion (M-segment) between motifs E and F (Fig. 2). The M-segment contains a sequence with similarity to a motif

THIOL TEMPLATE PEPTIDE SYNTHESIS SYSTEMS IN BACTERIA AND FUNGI

103

Figure 8 Structure of the enniatin synthetase as deduced from the gene sequence and biochemical characterizations.EA, D-Hiv-activationdomain; EB, L-Val-activation domain; M, methyltransferase; S, 4'-phosphopantetheine binding site.

apparently conserved within a number of methyl transferases (Haese et al., 1993). This portion of Esyn shows no homology to any region of known peptide synthetases (Haese et al., 1993). Tt was possible to express the M-segment in E. coli and to identify this protein as the methylase of Esyn by its binding properties for AdoMet. Three deletion mutants of this protein were shown to be inactive with respect to AdoMet binding (Haese et af., 1994). Similarly, a variety of other functional recombinant Esyn protein fragments could be expressed leading to the identification of the substrate activation sites for D-Hiv and L-Val (Pieper et al., 1995). A peculiar property of esynl is that it harbours two 4'-phosphopantetheine binding motifs in domain EB, one of which may represent a waiting position for peptidol units during chain growth.

4.2.6. Mechanism of Depsipeptide Formation Esyn consists of the two peptide synthetase domains EA and EB, but assembles three amino acids and three D-2-hydroxy acids in the final product enniatin. Studies on the mechanism of depsipeptide (enniatin B) formation revealed that the enniatin molecule is synthesized by three successive condensations of enzyme-bound (thioesterified) dipeptidols with each other (Zocher et al., 1983). This implies that, as in the fatty acid synthetases, Esyn contains a specific thiol group (waiting position) that picks up the intermediates of enniatin synthesis, i.e. the dipeptidol, tetrapeptidol and hexapeptidol to allow depsipeptide chain elongation (Fig. 9). After reaction of the thioester-bound N-methyl amino acid with the covalently bound D-Hiv (domain EA), the formed dipeptidol is transferred to the waiting position and attacked by the hydroxyl group of the newly formed dipeptidol yielding a tetrapeptidol. After transthiolation to the waiting position, the tetrapeptidol is attacked by the next new dipeptidol to fonn a hexapeptidol, which yields enniatin in the final condensation reaction. The presence of a second thioester formation module in the domain EB is consistent with this model and possibly this represents the waiting position. Apparently, the length of the growing depsipeptide chain is determined by the space provided by a putative cyclization cavity (Fig. 9).

104 (a) Cycle I

Pl-Hiv + P2-MeVal

-

( P2-MeVal-Hiv + P3-SH

RAINER ZOCHER AND ULLRICH KELLER

PI-SH + P2-MeVal-Hiv -+P2-SH + P3-MeVal-Hiv

P1-Hiv + P2-MeVal -* PI-SH + P2-MeVal-Hiv P2-MeVal-Hiv + P3-MeVal-Hiv-PP-(MeVal-Hiv)n + P3-SH PZ-(MeVal-Hiv)n + P3-SH --+ P3-(MeVal-Hiv)z + P2-MeVal-Hiv P1-Hiv + P2-MeVal --+Pl-SH P2-MeVal-Hiv + P3-(MeVal-Hiv)t --+ PZ-(Meval-Hiv)~+ P3-SH PZ-(MeVal-Hiv)s + P3-SH -~+ P2-SH + P3-(MeVal-Hiv)s + P3-SH + enniatin Cyclization P3-{MeVal-HIV)3 -

Figure 9 (a) The scheme of events of dipeptidol condensations on enniatin synthetase and the role of the additional 4'-phosphopantetheine containing thioester module as the waiting position. P1, P2, P3 = 4'-pliosphopantetheine groups. (b) Model of arrangement of catatytic sites of enniatin synthetase. Cy = cyclization cavity; EA, D-Hiv-activation domain; EB, L-Val-activation domain; M, methyltransferase.

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105

4.2.7. Biosynthesis of Beauvericin Beauvericin is a homologue of enniatins in which the branched-chain N-methyl amino-acid position always contains N-methyl-L-phenylalanine(see Fig. 6). Beauvericin synthetase catalysing beauvericin synthesis from L-Phe and D-Hiv under consumption of ATP and AdoMet has been isolated from the fungus Beauveria bassiana (Peeters et al., 1983). The enzyme strongly resembles Esyn with respect to its molecular size and the reaction mechanism. The main differences between both enzymes lie in the substrate specificity. Beauvericin synthetase exhibits a high specificity for aromatic substrate amino acids of the phenylalanine type, whereas Esyn is unable to incorporate such compounds.

4.3. Cyclosporin and Related Peptides

Cyclosporin is a cyclic undecapeptide with anti-inflammatory, immunosuppressive, antifungal, and antiparasitic properties (Borel, 1986). It is used world-wide in transplantation surgery and in the treatment of autoimmune diseases (Kahan, 1984; Schindler, 1985). The structure of cyclosporins is shown in Fig. 10. Besides the unusual amino acids a-aminobutyric acid (Abu), D-alanine and 4-(E)-butenyl4-methyl-~-threonine(Bmt), it also contains a number of N-methylated peptide bonds. Cyclosporins are produced by the fungus Beauveria nivea as a main component of 25 naturally occurring cyclosporins, mainly differing in positions 1, 2,4,5,7 and 11. Furthermore, unmethylated peptide bonds may occur in positions 1,4,6,9, 10 and 11 of the cyclosporin ring (Traber et al., 1987). 4.3.1. Biosynthesis of Cyclosporins Cyclosporins are synthesized via a thiol template mechanism, which, owing to the N-methylating steps, has strong resemblance to that of enniatin synthesis. This became evident from studies in vivo of the biosynthesis of cyclosporins (Zocher et al., 1984). In the initial cell-free studies on cyclosporin synthesis performed in the authors’ laboratory, a high molecular-weight-enzymefraction was obtained capable of activating all constituents of the undecapeptide and carrying out specific N-methylation reactions (Zocher ef al., 1986). Attempts to synthesize cyclosporin failed enzymatically. However, the enzyme synthesized the diketopiperazine c(D-Ala-N-MeLeu),representing a partial sequence of cyclosporin, which revealed the significance of the enzyme. Cell-free total synthesis of the cyclosporin molecule was finally established with an enzyme fraction obtained from a cyclosporin high producer strain (Billich and Zocher, 1987b). A number of different naturally occurring cyclosporins were synthesized by the multi-enzyme that has been designated cyclosporin synthetase (Cysyn). Further purification and characterization of

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Figure 10 Structure of cyclosporin A. Bmt, 4-(E)-butenyl-4-methyl-~-threonine; Abu, a-amino-L-butyric acid.

this multi-enzyme supported earlier findings that Cysyn strongly resembles the well-characterized Esyn system with respect to amino-acid activation and N-methylation reactions (Lawen and Zocher, 1990). Furthermore, 4'- phosphopantetheine was detected as a covalently bound cofactor in Cysyn. Like Esyn the multi-enzyme CySyn is unable to carry out racemization reactions. D-Ala is directly incorporated into the D-Ala position of cyclosporin. The D-Ala moiety is racemized from L-Ala by a specific alanine racemase, which plays a key role in cyclosporin biosynthesis (Hoffmann et al., 1994). 4.3.2. Mechanism of Cyclosparin Synthesis Studies on the mechanism of cyclosporin formation were reported by Lawen et al. (1994).The authors were able to isolate four enzyme-bound intermediate peptides of cyclosporin biosynthesis from a complex mixture of unidentified peptides. All four isolated peptides canied alanine as the N-terminal amino acid. One of the

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107

peptides represented the nonapeptide H-D-Ala-MeLeu-MeLeu-MeVal-MeBmtAbu-Sar-MeLeu-Val-OH.From these findings, the authors conclude that D-alanine provides the “starter amino acid” of cyclosporin synthesis leading to formation of a linear precursor undecapeptide with an N-terminal D-Ala. Cyclo-sporin is formed in a final cyclization step.

4.3.3. Substrate Specificity of Cysyn From the spectrum of naturally occumng cyclosporins, it seems obvious that some of the peptide synthetase domains of Cysyn have a rather broad substrate specificity and others, such as that responsible for Bmt, have a lower specificity, allowing incorporation of homologue substrates. Therefore, it is not surprising that a number of new immunosuppressive cyclosporins could be synthesized in vitro (Lawen et al., 1989). Results of studies on the substrate specificity of the different binding sites of Cysyn at the cell-free level (Lawen and Traber, 1993) agreed with findings from studies in vivo (Kobe1 and Traber, 1982; Traber et al., 1989). 4.3.4. Molecular Structure of Cysyn Cyclosporin synthetase is encoded by a giant 45.8 kb open reading frame. The predicted gene product is a polypeptide of about 1600 kDa containing 11 peptide synthetase domains, of which 7 are homologous to the EB domain of enniatin synthetase carrying the integrated N-methyltransferase module (Weber et al., 1994).From the arrangement of the domains and the fact that domains in peptide synthetases are colinear to the arrangement of amino acids in the peptides to be synthesized, the authors conclude that the 5’-terminal domain is responsible for D-Ala activation. The last domain at the 3‘-end would represent the L-Ala-activating pepride synthetase domain (see Fig. 3). This assumption was supported by the finding that a Cysyn fragment of 130 kDa could be isolated, which is capable of activating L-Ala. Edman degradation of this protein yielded a sequence with an N-terminus in the position of amino acid 13 601 of Cysyn (Weber et al., 1994). Like Esyn and other peptide synthetases, the peptide synthetase domains of Cysyn contain all highly conserved motifs and modules described as characteristic for this class of enzymes (see above).

4.3.5. Cyclosporiiz-related Peptolide SDZ 214-1 03 The peptolide SDZ 214-103 is a cyclosporin-related undecapeptide lactone produced by the fungus Cylindrotrichurn oligospennum (Dreyfuss et al., 1988). It is a peptidolactone analogue of the immunosuppressive undecapeptide cyclosporin, carrying a D-2-hydroxyisovaleratemoiety in position 8 of the ring system

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instead of D-Ala (see Fig. 10). This compound is synthesized by a multi-functional enzyme from its precursor amino acids and D-Hiv with consumption of ATP and AdoMet. The enzyme has been described to be a single polypeptide chain with a molecular mass of about 1400 kDa and strongly resembles Cysyn ( h w e n et al., 1991). Studies of the substrate specificities of peptolide synthetase revealed that most sites of this multi-enzyme appear to have narrower specificities than those of Cysyn. The D-2-hydroxy-acid position (corresponding to position 8 of the cyclosporin ring) can be occupied by a large range of substrates varying from D-lactic to D-2-hydroxyisocaproic acid.

4.4. Ergot peptide alkaloids

Ergot alkaloids of the peptide type are produced by the ergot fungus Claviceps purpurea. They consist of D-lysergic acid attached in amide-like fashion to a vipeptide arranged into a unique cyclol structure as in ergotamine (Fig. 11). The cyclol structure results from the modification of the amino acid adjacent to the lysergyl moiety into an a-hydroxy-a-amino acid. This.modification is believed to take place after the assembly of the D-lySergyhipeptide and most probably after the formation of the corresponding proline lactam (Fig. 11) (reviewed by Stadler, 1982; Kobel and Sanglier, 1986). Introduction of the hydroxyl group to the a-position of the amino acid is followed by spontaneous non-enzymatic closure of the cyclol ring (Hofmann et ul., 1963). The ergot cyclol peptide alkaloids-also called ergopeptines-constitute a family of compounds that differ from each other by substitutionsof amino acids in the first and second position of the peptide moiety adjacent to D-lysergic acid. Common to all naturally occurring ergopeptines is D-lysergic acid and proline, the latter located at the carboxy terminal position of the peptide chain. In the last two decades, members of an additional new class of D-lysergylpeptides have been isolated, which differ from the ergopeptines in that the cyclol structure is missing. They contain a proline lactam ring and therefore are called ergopeptams (Stadler, 1982; Kobel and Sanglier, 1986). In contrast to the ergopeptines, the proline here has the D-configuration. Most probably, these compounds arise from spontaneous isomerization of the corresponding L-prolinecontaining stereoisomer, e.g. D-lysergyl-L-alanyl-L-phenylalanyl-L-pro1in lactam (Fig. II), which can not be further converted to the corresponding ergopeptine. 4.4.1

I

D-Lysergylpeptide Assembly

Investigations of a putative ergopeptine synthetase indicate that the compound is synthesized by a non-ribosomal mechanism. However, this enzyme proved to be unable to incorporate free D-lySefgiC acid into the ergopeptine. Instead, it converts and incorporates biogenetic precursors of D-lysergic acid such as elymoclavin

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109

Figure 1 1 Biosynthetic relationship of structures of D-lysergyl-L-alanyl-L-phenylalanyl-L-proline lactani and ergotamine. Ergotanline (right) is a representative meniber of the ergopeptine family. The various groups of naturally occuming ergopeptines differ from each other by exchpnge of the amino acids in the two positions adjacent to the tetracyclic D-lysergic acid. Exchange is exclusively with branched-chain amino acids or phenylalanine. D-lysergyl-L-alanyl-L-phenylalanyl-~-proline lactam (left) is the immediate precursor of ergotarnine and is assembled by D-lySergyl peptide synthetase (see text).

(Maier et ul., 1983). This contrasts with the finding of a D-lysergic acid-activating enzyme that had been isolated from C. purpurea, suggesting that D-lysergic acid might be a free intermediate in ergopeptine synthesis. The enzyme is a single polypeptide chain of 62 kDa in its denatured form, which catalyses the ATPpyrophosphate exchange dependent on D-lysergic acid but, surprisingly, is unable to form a thioester with D-lySergiC acid or its structural analogue dihydrolysergic acid (Keller et al., 1984b, 1988). Therefore, it has been tentatively considered as a D-lysergic acid-AMP ligase acting in a similar fashion to actinomycin synthetase I and related proteins of prokaryotic origin (Keller, 1995). Keller et al. (1988) eventually reported a cell-free system of D-lysergyl peptide synthesis that catalysed formation of D-lysergyl-L-alanyl-L-phenylalanyl-L-pro1ine lactam from D-lysergic acid, alanine, phenylalanine and proline, which is the immediate precursor of ergotamine (see Fig. 11). The enzyme that catalyses formation of the D-lysergyl peptide was partially purified and shown to be a 500-550 kDa multi-enzyme complex under native conditions. Labelling the enzyme with radioactive substrate amino acids and dihydrolysergic acid enabled identification of the components of the enzyme complex by fluorography of SDS-PAGE gels. Radioactive dihydrolysergic acid (a reactive structural analogue of D-lySergiC acid) labelled a 160 kDa fragment, while radioactive phenylalanine labelled a 370 kDa fragment, that most probably also contains the binding sites for alanine and proline. Recent investigationswith antibodies raisedagainst the 62 kDa D-lysergic acid-activating enzyme previously isolated from Cluviceps purpurea revealed strong cross-reaction against various bands in the 100-200 kDa range, among them the 160 kDa fragment carrying the site of the D-lysergic acid thioester formation module (B. Riederer and U. Keller, manuscript submitted).The available data suggest that the D-lysergic acid-activating enzyme most probably arises through proteolytic degradation of the 160kDa fragment, which possesses an intact

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peptide synthetase domain. The 62 kDa is only able to activate D-lysergic acid as adenylate and not as thioester which indicates that the thioester formation is lost (Keller etal., 1988).In addition, degradation of the 370 kDa fragment was observed in partially purified protein fractions, where smaller fragments still activated individual amino acids as thioester, such as phenylalanine (M. Han and U. Keller, unpublished data). These data clearly indicate that proteolytic fragmentation of peptide synthetases can result in the formation of intact separate peptide synthetase domains still activating their cognate amino acids. From the data, therefore, it is most likely that the two fragments of the D-lysergylpeptide synthetase result from fragmentation of a 500-530 kDa single polypeptide as shown in Fig. 12. Similarly, results obtained in the case of HC-toxin synthesis indicate that HC-toxin synthetase from Cochliobolus carboneuin is a 550 kDa protein as estimated from the length of the 15.7 kb gene (Scott-Craig et al., 1992). However, purification and analysis of the enzyme revealed that it is always present as two fragments of about 300 kDa activating and epimerizing three of the four amino acids of HC-toxin (Scott-Craig et al., 1992).The previous isolation of a phage clone from a C. purpurea genomic

LSA

Ala

Phe

Pro

Fragmentation

160 kDa

t.’” I I I ....

S ‘LSA

62kDa

I-:::

I1 S

‘Ala

111 I S

‘Phe

D-Lysergyl peptide lactam

IV I

s‘Pt-O

D-Lysergic acid activating enzyme

LSA-AMP Figure 12 Scheme of domain assembly of D-lysergyl peptide synthetase from Cluviceps purpureu. The enzyme system catalysing D-lysergyl peptide synthesis (see Fig. 11) consists of two protein fragments of 370 kDa and 160kDa, which harbour sites for thioester formation of the amino acids and D-lysergic acid, respectively. The 62 kDa fragment represents the previously isolated D-lysergic acid activating enzyme of C. purpurea, which most probably is derived from the D-lysergic-acidactivating domain of D-lysergyl peptide synthetase.LSA,

D-lysergic acid.

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111

library containing four peptide synthetase domains in tandem raises the possibility that the ergot peptide synthetase also consists of one polypeptide chain with four activation domains for D-lysergic acid and the three amino acids of the cyclol nucleus (s. Riederer and U. Keller, unpublished). However, comparison of D-lysergylpeptide synthetase sequences with the DNA sequences are necessary to confirm the identity of that gene. The possible existence of one single orf for this enzyme would fit with the observation that eukaryotic peptide synthetases always consist of one polypeptide chain.

5. PROKARYOTIC PEPTIDE SYNTHETASE SYSTEMS

5.1. Acyl Peptide Lactones Much progress i n the enzymology of prokaryotic peptide synthetases has been achieved in the field of the acyl peptide lactone synthetases. Acyl peptide lactones consist of peptide lactone rings to which are attached aromatic or aliphatic side groups in an amide-like fashion. Acyl peptide lactones with aromatic side groups are mostly produced by streptomycetes whereas the various fatty acyl peptide lactones are mostly formed by Bacillus species (Vater, 1989). Both classes of compounds contain L- and D-amino acids in their chains. In addition, the streptomycete acyl peptide lactones contain N-methyl amino acids. No N-methyl peptides have been found in Bacillus yet. Examples of the acylpeptide lactones with aromatic side chains are the actinomycins (bicyclic pentapetide lactones), the quinoxaline antibiotic (monocylic octadepsipeptides), mikamycin BI (monocyclic hexapeptide lactones) and mikamycin BII (monocyclic heptapeptide lactones) antibiotics (structures reviewed by Okumura, 1983; Keller, 1995). Many of these compounds have therapeutic value as antibiotics or cytostatics. The enzymatic steps in the biosyntheses of the different aromatic acyl peptide lactones appear to be similar. 5.1.1. Actinomycin Biosynthesis as a Model of Aromatic Acyl Peptide Lactone Biosynthesis The large numbers of different actinomycins arise by substitutions in several positions of their pentapeptide lactone rings with homologous amino and imino acids (Katz, 1968; Meienhofer and Atherton, 1973). Actinomycins (e.g. actinomycin D) are bicyclic and originate from a monocyclic precursor, in which one pentapeptide lactone ring is attached to 4-methyl-3-hydroxyanthranificacid (4MHA). 4-MHApentapeptide lactone is the prototype of the whole class of aromatic acyl peptide lactones, although only short-lived in the cell because of dimerization

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MeVal

MeVal

MeVal

I

2 x

I

Thr

I

Thr I

&"*; CH3 OH

4-MHA pentapeptide lactone

Actinomycin D

Figure 13 Synthesis of actinomycin from its immediate precursor4-methyl-3-hydroxyanthranilic acid pentapeptide lactone. Oxidative condensation of 4-methyl-3-hydroxyanthranilic acid (4-MHA) pentapeptide lactone to actinomycin is enzymatic (see Keller, 1995). Sar, sarcosine (N-methyl-glycine); MeVal (N-methyl-L-valine).

through oxidative condensation of two such molecules immediately after its formation (Fig. 13). It is assembled from 4-MHA and the five amino acids constituting the peptide ring and S-adenosyl-L-methionine in an ATP-dependent manner. Synthesis is accomplished by a set of three peptide synthetases, which contain a total of six activation domains, two of which contain an N-methylation module (Fig. 14). 5.1.2. Actinornycin Synthelase I

Actinomycin synthetase I (ACMSI), the first enzyme involved in actinomycin synthesis, is a 4-MHA-AMP ligase, which catalyses the synthesis of adenylyl-4MHA from 4-MHA and ATP (Keller et al., 1984a; Keller and Schlumbohm, 1992). From its properties, it appears as merely consisting of an adenylate forming domain with a size of 45 kDa. The enzyme has broad substrate specificity with respect to various structurally related benzene carboxylic acids. However, heteroaromatic carboxylic acids, such as pyridine, quinoline or quinoxaline carboxylic acids are not activated (Glund et al., 1990; Schlumbohm and Keller, 1990; Keller and Schlumbohm, 1992). Structurally related benzene carboxylic acids with hydroxy groups in the 3-position or methyl group in the 4-position of the benzene nucleus serve as efficient substrates in peptide synthesis in vivo and in vitro. For example, strong evidence for the involvement of ACMSI in actinomycin biosynthesis came

THIOL TEMPLATE PEPTIDE SYNTHESIS SYSTEMS IN BACTERIA AND FUNGI

AdoMet

___)

Gly

Pro

Val Thr

AdoMet

-

Actinomycin Synthetase 111

113

MeVal

s

Sar

4

3

2

___)

Actinomycin Synthetose II Thr

1

NH

I

4-MHA

Actinomycin Synthetose I

Figure 14 Assembly of 4-methyl-3-hydroxyanthranilic acid pentapeptide lactone by actinomycin synthetases. Actinomycin synthetases activate, modify and polymerize the constituent amino acids and 4-methyl-3-hydroxyanthranilicacid (CMHA). Sar, sarcosine (N-methyl-glycine); MeVal (N-methyl-L-valine); AdoMet, S-adenosyl-L-methionine (methyl group donor).

from the finding that feeding structural analogues of 4-MHA, such as 4-methyl-3hydroxybenzoic acid, 3-hydroxybenzoic acid, p-toluic acid or p-aminobenzoic acid, to cells of actinomycin-producingS. chrysomallus and S.antibioticus resulted in the formation of new compounds instead of actinomycin (Keller, 1984). The new compounds were monocyclic acylpentapeptide lactones (actinomycin half molecules) containing the administered analogue instead of 4-MHA. 5.1.3. Actinomycin Synthetases II and III ACMSII and ACMSIII have been isolated from actinomycin-producingStreptomyces chrysomallus. ACMSII, a 280 kDa multi-enzyme, activates L-threonine and L-valine (occupying positions 1 and 2 in the ring) as thioesters via the corresponding adenylates (Keller, 1987; Stindl and Keller, 1993). ACMSIII activates proline,

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glycine and valine, which are in positions 3 , 4 and 5 of the pentapeptide lactone ring of actinomycin D, and has a molecular mass of 480 kDa (Keller, 1987; Stindl and Keller, to be published) (Fig. 14). In the presence of S-adenosyl-L-methionine, thioester-bound glycine and valine are N-methylated, which yields covalently bound sarcosine and N-methyl-L-valine,respectively. This indicates that ACMSIII has two N-methylation modules necessary for the N-methylation of covalently bound glycine and valine, respectively (Fig. 14). Several lines of evidence indicate that these modules are organized like the N-methylation module in the amino-acid domain of enniatin synthetase. First, the reaction catalysed by the two enzymes proceeds by the same mechanism. Second, monoclonal antibodies directed against the N-methyltransferdse domain of enniatin synthetase strongly cross-reacted with ACMSIII but not with ACMSII, confirming the presence of N-methyltransferase domain(s) in ACMSIII (A. Billich, R. Zocher and U. Keller, unpublished). Both ACMSII and ACMSIII contain 4'-phosphopantetheine as a covalently bound cofactor. 5.1.4.

Total Cell-free Synthesis of Acyl Pentapetide Lactone Biosynthesis

An enzymic total synthesis of acyl pentapeptide lactone analogues of 4-MHA pentapeptide lactone has been achieved as yet only in a system of permeabilized cells of S. chrysonmllus (A. Stindl and U. Keller, unpublished data). The various steps of peptide synthesis are summarized in Fig. 15. In this system, the incorporation into the complete product of all constituents could be demonstrated, which was not observed in cell-free fractions containing the three ACMSs at much higher concentration than in the permeabilized cell system. These findings indicate that functional activity of the synthetase complex strongly depends on the integrity of the cellular structure possibly through membrane association of the ACMS complex. Analysis of the individual synthetases with respect to their reactivity, such as in adenylate formation, thioester formation, N-methylation, peptide-bond formation and epimerization, indicate that they are functionally intact (Keller and Schlumbohm, 1992; Stindl and Keller, 1993,1994). Apparently, impairment of the correct positioning of the three synthetases in the complex andor of final steps in product release interfere with formation of the acyl pentapeptide lactone. 5.1.5. Priming of Reactions in initiation and Elongation of Peptide Synthesis on ACMSII ACMSII contains three distinct sites for covalent binding of 4-MHA, L-threonine and L-valine as thioesters (Keller, 1987;Stindl and Keller, 1993,1994). ACMSII is not able to activate 4-MHA as adenylate. Instead, ACMSI delivers adenylyl-4MHA to ACMSII, which binds 4-MHA as thioester (see Fig. 15). L-Threonine and L-valine are also bound as thioesters. However, prior to this, ACMSII activates

115

THIOL TEMPLATE PEPTIDE SYNTHESIS SYSTEMS IN BACTERIA A N D FUNGI

280 kDa

45 kDa 4MHA

4MHA-AMP7

L-2

Thr

+

Val

+

1

4 Val

GlY

4

AdoMet

AdoMet

4 Ro

480 kDa Figure 15 Organization of the actinomycin synthetase multi-enzyme complex and events of acylpentapeptide chain growth. For explanations of abbreviations, see Fig. 14.

these two amino acids as adenylates. This indicates the presence of two complete peptide synthetase domains on ACMSII for threonine and valine, and one additional thioester formation module for 4-MHA most probably separate from the Thr and Val domain. Functionally, the charging of ACMSII with 4-MHAcan be described as priming of the events taking place on the multi-functional enzyme. Two different initiation reactions can take place in the presence and absence of 4-MHA, respectively (Fig. 16). In the presence of 4-MHA (or its analogue p-toluic acid used in the experiments) peptide synthesis starts with formation of 4-MHA-threonine, the latter reacting further with L-valine to yield a mixture of 4-MHA-threonyl-~-valine

116

(a)

e I : -

RAINER ZOCHER AND ULLRICH KELLER

S -4MHA

4W-AMP Thr,Val, ATP

SH

(b) SH

Val - Thr - 4MHA LDVal- m-4m

es"

Thr ,Val, ATP

s~

S-Thr

sr Val

\D-Val

- Thr

-

Thr

Figure 16 Effect of priming the initiation reaction of acyl peptide lactone on actinomycin synthetase 11. For explanation of abbreviations, see Fig. 14. (a) Initiation in the presence of 4-MHA-adenylate (supplied by ACMSI). (b) Initiation in the absence of 4-MHAadenylate. For details see text.

and 4-MHA-L-threonyl-D-valine. If 4-MHA is absent in this reaction, ACMSII forms a mixture of threonyl-L-valine and threonyl-D-valine. However, the formation of these two dipeptides gives considerably lower yield than the yield of 4-MHA dipeptides, which are formed when 4-MHA is present. Thus, priming of the reaction by acylation of threonine with 4-MHA strongly influences the reactivity and the substrate specificity of the enzyme(s).

5.1.6. Epirnerizatioiz of Amino Acid in the Peptide-bound State ACMSII activates L-valine but not D-valine. Epimenzation of valine takes place after peptide-bond formation with threonine leading to 4-MHA-~-Thr-~-val (Stindl and Keller, 1994). During inversion at the a-carbon of L-valine in the covalently bound acyl dipeptide, a proton is abstracted that goes into solvent. This is strong evidence for the intermediacy of a carbanion structure at the chiral centre. Epimerizdtion proceeds independently of addition of cofactors, such as NAD, FAD or pyridoxalphosphate. Spectral analysis of ACMSII indicates the absence of any tightly bound cofactors in the enzyme. Thus, this mechanism of peptide epimerization is novel and may operate in the synthesis of a large number of nonribosomally as well as of ribosomally made peptides, which have D-amino acids at positions other than the amino-terminal end (Stindl and Keller, 1994). In the case of peptides with D-amino acids as amino-terminal residues of the growing peptide

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117

chain, the activating enzymes, such as gramicidin S synthetase I, racemize equally well the L- or D-isomer of the respective amino acid (Stein et al., 1995). Other systems, such as cyclosporin synthetase, do not possess racemase function and use D-amino acids as substrates that have been epimerized prior to activation. Stindl and Keller (1994) also propose an unknown proton acceptor group that is present on ACMSII, catalysing proton abstraction during inversion in a one- or two-base mechanism, which might be identical to the conserved His in the abovementioned “His motif’ of peptide synthetase domains. Meanwhile, a similar mechanism of epimerization of amino acid in the “peptide bound state” was also proposed for the epimerization of valine in 6-(L-a-aminoadipyl)-L-cysteinyl-D-vdinesynthesis catalysed by ACV synthetases (Shiau et al., 1995a,b). 5.1.7. Peptide-bond Formation Catalysed by ACMSIII Purified ACMSTII catalyses formation of prolylsarcosine and sarcosyl-N-methylL-valine. The latter two dipeptides cyclize spontaneously to the corresponding diketopiperazincs and are released from the enzyme. These partial reactions give relatively high yields. In the additional presence of ACMSI and 11, acylpeptide chain intermediates most probably representing acyltri-, tetra- and pentapeptide were identified as covalently bound intermediates. The whole sequence of events in acyl pentapeptide (lactone) synthesis based on these data is shown in Fig. 15 (U. Keller and A. Stindl, unpublished data). An enzyme fraction that catalyses the final acyl pentdpeptide lactone has not yet been isolated.

5.2. Surfactin Surfactin is a lipopeptide lactone composed of seven amino acids in the order Glu-Leu-D-Leu-Val-Asp-D-Leu-t-Leu in a heptapeptide ring to which is attached a P-hydroxy fatty acid (Fig. 17). The compound has been shown to be synthesized cell-free by a thiol template mechanism (Kluge et al., 1988; Ullrich et al., 1991; Menkhaus et al., 1993; Zuber et al., 1993). The multi-enzyme system responsible for the assembly of the compound has been resolved into its components and reconstituted from the latter ones into the functionally activecomplex by Menkhaus et al. (1993). These authors identified the enzyme fractions by their ability to catalyse the activation of the various amino-acid constituents of surfactin. Enzyme El consists of two multi-lunctional peptide chains of460 (Ela) and 435 kDa (Elb). Ela is responsible for adenylation and thioesterification of L-Glu, L-Leu (positions 1-3) and El b for adenylation and thioesterification of L-Val, L - A s ~and L - ~ U (positions 4-6). The 160 kDa enzyme E2 is responsible for activation of L - ~ U (position 7), while enzyme E3 has activity of an acyl transferase incorporating

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Leu I

- :0

D-Leu I ASP

Val I D-Leu I Leu I

Glu I

co I

CHZ I

CHL

2

Figure 17 Structure of surfactin.

L-hydroxy fatty acid into peptide from L-hydroxy fatty acyl CoA thioester. E3 has a mass of 40 kDa (Fig. 18).

5.2.1. Initiation of Surfactin Synthesis Menkhaus et al. (1993) have shown that the acyl transferase enzyme E3 of surfactin synthetase acylates L - G thioesterified ~ to El a with an L-P-hydroxy fatty acyl residue with consumption of L-P-hydroxy fatty acyl CoA thioester. The resultant product is L-P-hydroxy fatty acyl glutamate covalently bound to enzyme Ela. D-P-HY~~oxY fatty acyl CoA was a poor substrate under these conditions indicating specificity of the acyltransferase with respect to optical configuration. By contrast, the enzyme has broad specificity with respect to fatty acyl chain length. Whether the enzyme directly acylates Glu on enzyme Ela or transfers the fatty acyl chain prior to that step to a thiol group of Ela is not known. The whole mechanism of initiation differs from that of actinomycin synthesis in that ACMSI is not an acyl transferase (Stindl and Keller, 1993).

5.2.2. Elongation and Epiinerization Reactions in Amino-acid Positions of Surfactin Enzyme Ela forms thioesters with Glu and L - ~ inUa molar ratio of 1:2.Enzyme Elb binds L - A s ~L-Val , and L-Leu in a molar ratio of 1:l. Enzyme E2 binds L - ~ u . Determinations of the optical configurations of all the amino acids covalently bound to the enzymes revealed that all of the leucines were of the L-configuration. Thus, Menkhaus et al. (1993) argue that the enzymes are lacking racemization

THIOL TEMPLATE PEPTIDE SYNTHESIS SYSTEMS IN BACTERIA AND FUNGI

Glu

Leu

Leu

1

1

1

I

s\Glu

I

I

I

'\Leu

I

I El

119

EnzymeEla

'\D-Leu

Leu

+]

Enzyme E2

Figure 18 Enzyme organization of surfactin biosynthesis as deduced from gene sequences and biochemical data.

functions and that epimerization reactions take place in the course of the condensatiozdelongation reactions. Possibly these take place at the stage of peptidyl intermediates analogous to the situation encountered in the biosynthesis of actinomycin. Analysis of the 4'-phosphopantetheine contents in the various fractions indicated that this cofactor was present as a prosthetic group in enzymes Ela, Elb and E2. Details of the mechanisms of termination and release of the end product surfactin are not yet available.

5.2.3. Structure-ficnctiun Relationships in Surfactin Synthetases More information of the structure-function relationships of the surfactin synthetases became available by investigating surfactin synthetases carrying sitespecific mutations in the reaction centres of the various activation domains. The complete DNA sequence of the surfactin biosynthesis operon had been obtained previously by several groups who showed that part of it is important for competence establishment in Bacillus subtilis (Nakano et al., 1991;Tognoni el al., 1995) (see Fig. 3). This gene locus contains sequences coding for seven peptide synthetase

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domains distributed over a set of three large and one small transcriptionally coupled orfs designated s $ M , s$AB, s$AC and @AD, respectively (Cosmina et al., 1993; Fuma et al.,1993). Ser-to-Ala substitutions in the thioester formation motifs of the first four domains (i.e. all three motifs in s $ M and one in s$AB) of the cluster were introduced to see whether these had influence on surfactin production and competence development in vivo.Each substitution abolished surfactin production, while competence development remained unaffected (D’Souza et al., 1993). More detailed genetic analyses revealed that some unknown function of the Val-activating domain influences competence development. This will not be discussed here further. Further investigation of the products of mutated s.f genes showed that the corresponding enzymes were indeed lacking the expected functions such as thioester formation in Ser-to-Ala-substituted domains 2 and 3 of s $ M , which enabled assignment of EIa to the gene product of that or$ Further mutational analyses revealed that slfAB and srfAC encode enzymes Elb and E2, respectively (Vollenbroichet al., 1994). Interestingly, slfAD from its sequence apparently codes for a thioesterase like the gene product of grsT (Kratzschmar et al., 1989), which implicates the function of a thioesterase i n the terminatiodcyclization reactions of the biosynthetis of surfactin. Such an enzyme activity has not yet been characterized from surfactin-producingB. subtilis (Vollenbroich el al., 1994).

5.3. Bialaphos Bialaphos, a linear tripeptide consisting of two alanine (Ala) residues and an unusual amino acid called phosphinothricin (Pt) in the order Pt-Ala-Ala is synthesized by Streptornyces hygroscopicus and Streptornyces viridochromogenes (Fig. 19). Antibacterial activity of Pt is due to its release from the Pt-tripeptide through the action of intracellular peptidases after transport into bacterial cells (Diddens et al., 1976). The compound is active as an inhibitor of glutamine synthetases, which leads to cell death through glutamine starvation (Bayer et al., 1972). In plants, the same type of inhibition leads to accumulation of ammonium ions, which is toxic to the plant cell and therefore makes Pt a most effective herbicide. In addition, isolation of the bialaphos resistance gene from a bialaphosresistant Streptoniycrs strain has led to the construction of herbicide-resistant plants, removing the constraints i n using phosphinothricin tripeptide (Ptt) as a total herbicide. 5.3.1,

Isolation und Chamcterizatioiz of Peptide Synthetase Domain-related

orfs

Cloning and partial sequencing of the cluster of bialaphos biosynthesis genes has

THIOL TEMPLATE PEPTIDE SYNTHESIS SYSTEMS IN BACTERIA AND FUNGI CH3

CH3

O=b--OH

O=b-OH

121

1

CO-Ale-Ala Phosphinothricin (Pt)

Phosphlnothricyl-alanyl-alanine (Ptt, Bialaphos)

Figure 19 Structure of phosphinothricin and phosphinothricin-tripeptide.

identified many of the genes involved in the steps of Pt synthesis, and which are assembled in one large gene cluster in Streptomyces hygroscopicus and Streptomyces viridochrornogenes (Murakami ‘et al., 1986; Hara et al., 1991). By contrast, little is known about the process of non-ribosomal peptide-bond formation in bialaphos synthesis at both the biochemical and genetic level. Remarkably, genes that are involved in the steps of condensation of alanine with the immediate precursor of Pt, i.e. desmethyl-phosphinothricin (DMPT) or N-acetyl-DMPT, have been identified in disruption mutants and most of these mutations mapped to the bialaphos gene cluster (Hara et al., 1988). Sequencing of one particular clone mapping close to the bialaphos resistance gene bar revealed orfs ORFl and ORF2 with similarity to thioesterase genes in vertebrates (Raibaud et al., 1991). Wohlleben and co-workers (1992) isolated from the same region of the genome of S. viridochrornogenes an additional orf with similarity to various peptide synthetases. The deduced amino-acid sequence revealed a peptide synthetase domain encompassing all six highly conserved motifs necessary for adenylate and thioester formation. The gene termedphsA would encode a protein of 660 amino acids. From its size it would represent a single amino-acid activating domain. The presence of a single amino-acid activating enzyme and also single genes for thioesterases, indicates that Pt-tripeptide synthesis takes place on a multi-enzyme complex instead of one single polypeptide chain, as is the case of ACV synthesis (Raibaud et al., 1991). Consequently, one has to postulate additional orfs coding for the missing two activation domains because a total of three domains for the assembly of the Ptt-molecule has to be expected. These orfs could possibly be represented by the mutants isolated by Hara et al. (1 988). The cloning of these genes has not been reported yet.

5.3.2. PhsB is an Alanine-activating Peptide Synthetase Biochemical investigation on Ptt-peptide synthesis have shown the presence of an

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alanine-activating enzyme produced in coordinately elevated levels in mutants of

S. viridochromgenes, which had been previously selected for higher level Pt-tripeptide formation in a strain-improvement program (U. Keller and B. Riederer. manuscript submitted). The enzyme was purified to apparent homogeneity and identified by radioactive substrate labelling and autoradiography in SDS-PAGEgels. In the denatured form, it has an Mrof 147 OOO. Thus, it is clearly distinct from thephsA gene product, which would represent a polypeptide chain of Mr 80 OOO. The protein, which activates and binds alanine covalently as thioester, was designated PhsB. Like other peptide synthetases, it was shown to contain 4'-phosphopantetheine as cofactor. Attempts to detect a second alanine-activating enzyme from S. viridochmmogenes were unsuccessful. It cannot be ruled out that PhsB harbours two alanine-activating domains or is the degradation product of a still larger enzyme harbouring the complete set of two alanine-activatingdomains. Interestingly, PhsB also activates structurally related amino acids, such as leucine, valine and isoleucine, but not glutamate or Pt (U. Keller and B. Riederer, manuscript submitted). Leucine has been shown previously to be a component of a Pt-tripeptide-related protein called phosalacine from Kifasofosporiaphosalucime (Omura el al.. 1984). Phosalacine contains Pt, alanine and leucine in the order Pt-Ala-Leu. It cannot be excluded that Pt-Ala-Leu is synthesized by homologues of PhsA and PhsB under physiological conditions, where high leucine levels are present. Cloning of the gene encoding PhsB will enable us to see whether it contains one or two alanine-activating domains. The existence of PhsB as an alanine-activating enzyme also indicates that PhsA is most probably responsible for Pt activation. Meanwhile, biochemical work on Pt activation in S. viridochmmogenes has revealed the preFnce of an enzyme of molecular mass about 80 kDa, which activates acetyl-Pt and to a lesser extent Pt (U. Keller, unpublished). This enzyme is of the expected size of the phsA gene product. It will be interesting to test this enzyme with the substrate N-acetyl-DMPT to see whether this is an even better substrate than Pt or N-acetyl-Pt. These and the other data available on genes of Ptt synthetases and associated orfs characterize the Ptt biosynthesis system as typical for the organization of bacterial multi-enzyme peptide synthetases in which the various domains are spread over separate enzymes instead of one single enzyme polypeptide chain as in the eukaryotes.

6. FUTURE PROSPECTS OF PEPTIDE SYNTHETASE RESEARCH

Future developments in peptide synthesis research will aim at clarifying the mechanisms of initiation, elongation, epimerization and termination events on peptide synthetase. Parallel work will aim at the elucidation of the structures of peptide synthetase domains by biochemical techniques, such as cross-linking of

THIOL TEMPLATE PEPTIDE SYNTHESIS SYSTEMS IN BACTERIA AND FUNGI

123

motifs, mutational analysis of reaction sites and by elucidation of crystal structures of various domains by X-ray crystallography. This mechanistical and structural information will help to understand the factors determining substrate specificity and to circumvent possible constraints in combining peptide synthetase domains into novel enzymes by recombinant DNA techniques, which will aid in the design of new bioactive compounds. The success of such manipulations has been demonstrated i n several examples of polyketide synthase systems (Hutchinson and Fuji, 1995; Tsoi and Khosla, 1995).

6.1. Domain Exchange in Thiol Template Peptide Synthesis Systems

Construction of peptide synthetase-containing activation domains with altered substrate specificity or containing activation domains from other peptide synthesizing systems should allow us to synthesize new products in vivo and in vitm. Such an approach has been realized recently (Stachelhaus et al., 1995) when domaincoding regions of bacterial and fungal origin were combined with each other in hybrid genes that encoded peptide synthetases producing peptides with modified amino-acid sequences. In fact, replacement of the Leu-activation domain in the slfAC gene in B. subtilis by the cys domain of ACVS of Penicillium chrysogenum led to the expression of the engineered gene, as revealed by the formation of the corresponding protein as well as in the formation in vivo of a novel surfactin molecule containing cysteine instead of leucine in the 7-position of the peptide lactone ring. This approach is promising for the future development of new and valuable compounds.

6.2. Combinatorial Approaches in Future Peptide Synthesis Development The need for new compounds will require extensive screening programs using high numbers of samples of both natural and synthetic origin. Combinatorial approaches in synthetic chemistry allow the high throughput screening of millions of compounds. It is desirable to introduce such combinatorial approaches into biosynthesis systems by creating domain assemblies, which provide the best possibility to create libraries of compounds. Attempts to combine domains or modules of different PKSII synthases for combinatorial purposes with the aim of establishing libraries have been reported by Khosla, Hopwood and their co-workers (reviewed in Tsoi and Khosla, 1995). In the case of peptide syntheti;e systems, such approaches could be realized soon, provided that the number of different domains is large enough to create a pool of many different enzymes and detect a useful compound. Peptide synthetase systems lack the various modification steps met in PKS systems,

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such as ketoreduction, dehydration and enoylreduction, the programming of which are responsible for the great diversity in the product spectrum of PKS systems. Much more promising for future developments in this field, therefore, is the possibility of improving the production and structures of existing valuable peptides, such as the p-lactams, immunosuppressants and anti-tumor agents by domain exchange and directed mutagenesis affecting substrate specificity.

ACKNOWLEDGEMENTS The authors wish to thank A. Haese for valuable discussion. The help of M. Krause, F. Schauwecker and W. Weckwerth in drawing the figures as well as M. Glinski and B. Gorhardt for correcting the manuscript is gratefully acknowledged. Work done in the authors’ laboratory has been supported by the Deutsche Forschungsgemeinschaft, the Fonds der Chemischen Industrie, the European Community and the Erwin Riesch Foundation.

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(1987) Monoclonal antibodies to the multi-functional enniatin synthetase. Biol. Chem Hoppe-Seyler 368,521-529. Borel, J.F. (1986) Ciclosporin and its future. frog. Allergy 38.9-18. Borchert, S., Patil, S.S. and Marahiel, M.A. (1992) Identification of putative multifunctional peptide synthetase genes using highly conserved oligonucleotide sequences derived from known synthetases. FEMS Microbiol. Lett. 92,175-180. Coque, J.J.R., Martin, J.F., Calzada, J.G. and Liras, P. (1991) The cephamycin biosynthetic genes pcbAB, encoding a large multidomain peptide synthetase pcbC of Nocardia lactumduruns, are clustered together in an organization different from the same genes in Acremonium chrysogenum and Penicillium chrysogenum. Mol. Microbiol. 5,1125-1 133. Cosmina, P., Rodriguez, F., de Ferra, F., Perego, M., Venema, C. and van Sinderen, D. (1993) Sequence and analysis of the genetic locus responsible for surfactin synthesis in Bacillus subtilis. Mol. Microbiol. 8, 82 1-83 1. De Cdcy-Lagard, V., Marlikre, P. and Saurin, W. (1995) Multi-enzymatic non ribosomal peptide biosynthesis: identification of the functional domains catalysing peptide elongation and epinierization. C.R. Acad. Sci. furis, Sci. Ke/Lve Sci. 318,927-936. de Wet, J.R., Wood, K.V..DeLuca, M., Helinski, D.R. and Subramani, S. (1987) Firefly luciferase gene: structure and expression in mammalian cells. Mol. Cell. Biol. 7,725-737. Diddens, H., Zlhner, H., Kraas, E., Gohring, W. and Jung, C. (1976) On the transport of tripeptide antibiotic in bacteria. Eu,: J. Biochem. 66, 11-23. Diez, B., Gutierrez, S., Barredo, J.L., van Solingen, P., van der Voort, L.H.M. and Martin, J.F. (1990) The cluster of penicillin biosynthetic genes. Identification and characterisation of the pcbAB gene encoding the a-aminoadipyl-cysteinyl-valinesynthetase and linkage to thepchC and penDE genes. J. Biol. Chem. 265,16358-16365. Dreyfuss, M.M., Schreyer, M., Tscherter, H. and Wenger, R. (1988) European Pat. Appl. 0296 123 A2. D’Souza, C., Nakano, M.M., Coibell, N. and Zuber, P. (1993) Aminoacylation site mutations in amino-acid-activating domains of surfactin synthetase: effects on surfactin production and competence development in Bacillus sublilis. J. Bacteriol. 175,3502-35 10. Fuma, S., Fujishima, Y., Corbell, N., D’Souza, C., Nakano, M.M., Zuber, P. and Yamane, K. (1993) Nucleotide sequence of S’-portion of srfA that contains the region required for competence establishment in Bacillus subtilis. Nucl. Acids Res. 21,93-97. Gevers, W., Kleinkauf, H. and Lipmann, F, (1969) Peptidyl transfers in gramicidin S biosynthesis from enzyme-bound thioester intermediates. f roc. Nut1 Acad. Sci. USA 63, 1335-1 345. Glund, K., Schlumbohni, W., Bapat, M. and Keller, U. (1990) Biosynthesis of quinoxaline antibiotics: Purification and characterisation of the quinoxaline-2-carboxylic acid activating enzyme from Sfrepfomycestriosfinicus. Biochemistry 29,3522-3527. Gocht, M. and Marahiel, M.A. (1994) Analysis of core sequences in the D-Phe activating domain of the multi-functiona1peplide synthetase TycA by site-directed mutagenesis. J. Bacteriol. 176, 2654-2662. Grove, J.F. and Pople, M. (1980) The insecticidal activity of beauvericin and the enniatin complex. Mycopafhologiu 70, 103-105. Guest, J.R. (1987) Functional implications of structural homologies between chloramphenicol acetyltransferases and dihydrolipoamide acetyltransferases. FEMS Microbiol. Len. 44,417-422. Gutierrez, S., Diez, B., Montenegro, E. and Martin, J.F. (1991) Characterisation of the Cephulosporiurn acreinortiurn pcbAB gene encoding a a-aminoadipyl-cysteinyl-valine synthetase, a large niultidomain peptide synthetase: linkage to thepcbC gene as a cluster of early cephalosporin biosynthetic genes and evidence of multiple functional domains. J. Bacteriol. 173,2354-2365.

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Haese, A., Schubert, M., Hemnann, M. and Zocher, R. (1993) Molecular characterisation of the enniatin synthetase gene encoding a multi-functional enzyme catalysing N-methyldepsipetide formation in Fusariurn scirpi. Mol. Microbiol. 7 , 905-914. Haese, A., Pieper, R., von Ostrowski, T. and Zocher, R. (1994) Bacterial expression of catalytical active fragments of the multi-functional enzyme enniatin synthetase. J. Mol. Biol. 243, 116-122. Hara, O.,Anzai, H., Imai, S., Kumada, Y.,Murakami, T., Itoh, R., Takano, E., Satoh, A. and Nagaoka, K. (1 988) The bialaphos biosynthetic genes of Streplomyces hygroscopicus: cloning and analysis of genes involved in the alanylation step. J. Anribiotics41,538-547. Hara, O., Murakami, T., Imai, S., Anzai. H., Itoh, R., Kumada, Y., Takano, E., Satoh, E., Atsuyuki, S., Nagaka, K. and Thompson, C.J. (1991) The bialaphos biosynthetic genes of Streptomyces viridochrornogenes: cloning, heterospecific expression and comparison with genes of Streptomyces hygroscopicus. J. Gen. Microbiol. 137,35 1-359. Hoffmann, K., Schneider-Schener, E., Kleinkauf, H. and Zocher, R. (1994)Purification and characterisation of eukaryotic alanine racemase acting as a key enzyme in cyclosporin biosynthesis. J. Riol. Chem. 269, 12710-12714. Hofmann, A., Ott, H., Griot, R., Stadler, P.A. and Frey, A.J. (1963) Die Synthese und Stereochemie des Ergotamins. Hel. Chim. Acta 46,2306-2328. Hopwood, D.A. and Sherman, D.H. (1990) Molecular genetics of polyketides and its comparison to fatty acid biosynthesis. Ann. Rev. Genet. 2 4 , 3 7 4 6 . Hutchinson, C.R. and Fuji, I. (1995) Polyketide synthase gene manipulation: structurefunction approach in engineering novel antibiotics. Annu. Rev. Microbiol. 49: 201-238. Jensen, S.E., Wong, A,, Rollins, M.J. and Westlake, D.W.S. (1990) Purification and partial synthetase from Strepcharacterisation of 6-(L-or-aminoadipyl)-L-cysteinyl-D-vahe tomyces clavuligerus. J. Bacterial. 172,7269-727 1 . Kahan, B.D. (ed.) (1 984) Cyclosporin: Biological Activity and Clinical Applications. Grune and Straton Inc., Orlando, FL. Katz, E. (1968) Actinomycin. In: Antibiotics I1 (D. Gottlieb and PD. Shaw, eds), pp, 276-341. Springer-Verlag, New York. Katz, L. and Donadio, S . (1993) Polyketide synthesis: prospects for hybrid antibiotics. Annu. Rev. Microbiol. 47,875-912. Keller, U. (1984) Acyl pentapeptide lactone synthesis in actinomycin-producing streptomycetes by feeding with structural analogues of 4-methyl-3-hydroxyanthranilicacid (4-MHA) J. Biol. Chem. 259,8226-823 1. Keller, U. (1987) Actinomycin synthetases: multi-functional enzymes responsible for the synthesis of the peptide chains of actinomycin. J . B i d . Chem. 262,5852-5856. Keller, U. (1995) Peptidolactones. In: Generics and Biochemisfry ofAntibioric Production (L.C. Vining, ed.), pp. 71-94, Heinemann-Butterworths, Toronto, New York. Keller, U. and Schlumbohm, W. (1992) Purification and characterisation of actinomycin synthetase I, a 4-methyl-3-hydroxyanthranilic acid: AMP ligase from Strepromyces chrysomallus. J. B i d . Chem. 267, 11745-1 1752. Keller, U., Kleinkauf, H. and Zocher, R. (1 984a) 4-Methyl-3-hydroxyanthranilic acid (4-MHA) activating enzyme from actinomycin producing Streptomyces chrysomallus. Biochemistry 23, 1479-1484. Keller, u., Zocher, R., Krengel, U. and Kleinkauf, H. (1984b) D-Lysergic acid activating enzyme from the ergot fungus Claviceps purpurea. Biochem. J. 218,857-862. Keller, U., Han, M. and Stoffler-Meilicke, M. (1988)D-Lysergic acid activation and cell-free synthesis of D-lysergyl peptides in enzyme fractions from the ergot fungus Claviceps purpurea. Biochemistry 27, 61644170. Kleinkauf, H. and von Dohren, H. (1987) Biosynthesis of peptide antibiotics. Ann. Rev. Microbiol. 41,259-289.

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Microbial Dehalogenation of Halogenated Alkanoic Acids, Alcohols and Alkanes J. Howard Slater', Alan T. Bull' and David J. Hardman3 1

Molecular Ecology Research Unit, School of Pure and Applied Biology, University of Wales, PO Box 915,Cardiff CFI 3T4 UK 2 Research School of Biosciences, University of Kent at Canterbury, Canterbury, Kent CT2 7NJ, UK 3 Cadbury Herne Lrd, Research and Development Centre, Canterbury, Kent CT2 7PD, UK

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Dehalogenation of halogenated alkanoic acids . . . . . . . . 2.1. 2-Haloalkanoic acid hydrolytic dehalogenases . . . . . . 2.2. Regulation of dehalogenase synthesis . . . . . . . . . . 2.3. Genetic organization of 2 H M hydrolytic dehalogenases 3. Dehalogenation of halogenated alcohols . . . . . . . . . . . 3.1. A pathway involving the formation of H M s . . . . . . . 3.2. Haloalcohol hydrolytic dehalogenases . . . . . . . . . . 3.3. Other systems for the transformation of haloalcohols . . 4. Dehalogenation of halogenated alkanes . , . . . . . . . . . . 4.1. Haloalkane hydrolytic dehalogenases . . . . . . . . . . 4.2. Oxygenase-type haloalkane dehalogenases . . . . . . . 4.3. Cofactor-dependent dehalogenases . . . . . . . . . . . 5. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . References , . , . , . . . . . . . . . . . . . . . . . . . . .

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Abbreviations: 1BP, 1-brornopropanol; CA, chloroacetone; ZCE, 2-chloroethanol; lCP, 1 -chloropropanol; 2CPD, 2-chloro-l,3-propandiol,2-chloropropan1,301; 3CPD, 3-chloro-1,2-propandiol,3-chloropropan-l,2-01; 13DBP, 1,3-dibromopropanol; DCA, dichloroacetic acid; 13DCA, 1,3-dichIoroacetone;DCM, dichlorornethane; 13DCP, 1,3-dichloro-2-propanol,1,3- dichloropropan-2-01; 23DCP, 2,3-dichloro-I-propanol, 2,3-dichloropropan-1-01; 22DCPA, 2,2- dichloropropionic acid; DMSO, diinethylsulfoxide; 24DNP, 2,4-dinitrophenol; ECH, epichlorohydrin; GDL, glycidol; GSH, glutathione; HAA, halogenated alkanoic acid; 2HAA. 2-halogenated alkanoic acid; MBA, monobromoacetic acid; 2MBPA, ADVANCES IN MICROBIALPHYSIOLOGY VOL 38 ISBN 0-12-027738-7

Copyright Q 1997 Academic Press Limited All rights of reproduction in any form reserved

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2-monobromopropionic acid; MCA, monochloroacetic acid: 2MCPA, 2-monochloropropionic acid; MFA, monofluoroacetic acid; NEM, N-ethylmaleimide; PAGE, polyacrylamide gel electrophoresis; PMS, p-mercunbenzoate sulfonate; SDS, sodium dodecyl sulfate; TOL, toluene.

1. INTRODUCTION

About one hundred years ago, investigations began into the effect of halogenated compounds on the physiology and biochemistry of microbes. W. J. Penfold found that the growth of Bacterium coli (Escherich) and Bacterium lactis aerogenes was inhibited by chlorinated and brominated compounds, such as monochloroacetate (MCA), 2-monobromopropionic acid (2MBPA) and monochlorohydrin, and showed that the pattern of sugar fermentations was altered. Growth in the presence of these compounds led to the selection of mutants resistant to the toxic effects of halogenated compounds (Penfold, 1913). It was established that halogenated analogues of intermediary metabolites were toxic because they inhibited key reactions of central metabolism. For example, fluoroacetate (MFA) inhibited the tricarboxylic acid cycle (TCA) because the lethal synthesis of fluorocitrate generated by the activity of the citrate-condensing enzyme inhibited the next TCA cycle enzyme, aconitase (Peters, 1952). The removal of halogens, particulzrly fluorine and chlorine, was identified as one mechanism to relieve the inhibitory effects of these compounds and, moreover, provide novel carbon and energy sources for growth. Den Dooren de Jong (1926) showed that bromopropionate and bromosuccinate were used as the sole growth substrates by some bacteria. Subsequently many halogenated compounds have been shown to be degraded by a variety of microorganisms (Hardman, 1991; Janssen et al., 1994; Slater, 1994). However, not all halogenated organic compounds can be degraded and there are many that are resistant to microbial attack. In general the greater the number of halogens per molecule, the more difficult it is to isolate microbes that dehalogenate and grow on the compound (Commandeur and Parsons, 1994). This fact is of considerable environmental importance since many man-made (xenobiotic) compounds released into the biosphere during the last 50 years are multi-halogenated: indeed, their usefulness as pesticides, dielectrics, flame retardants, preservatives or whatever, depends on their chemical and biochemical inertness. More recently, the application of dehalogenating enzymes as catalysts for the resolution of racemic mixtures and the production of chiral intermediates in chemical syntheses has achieved significance (Taylor, 1985,1988; Hasan etal., 1991; Kasai etal., 1992). How biological systems, in particular microbes, handle halogenated compounds is an important and intriguing topic. Knowledge of the mechanisms of dehalogenation is not only intrinsically important for basic principles of biochemistry, but is

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crucial in delineating what products and processes are environmentally and industrially acceptable. This review is concerned with the catabolism of halogenated aliphatic compounds in the series: alkanoic acids, alcohols and alkanes. Microorganismsremove halogens from halogenated aliphatic compounds by the activity of enzymes generally called dehalogenases. Microbes that produce dehalogenases are widely distributed in nature, apparently having evolved to degrade naturally-occurring halogenated compounds in order either to exploit them as carbon sources for growth, or as a means of protection against the toxicity of these compounds (Fowden, 1968; Slater, 1994).

2. DEHALOGENATION OF HALOGENATED ALKANOIC ACIDS The discovery of dehalogenases was the result of the pioneering work of Jensen who isolated bacteria and fungi that grew on halogenated alkanoic acids (HAAs) and who was responsible for introducing the term dehalogenase (Jensen, 1951, 1957, 1959, 1960, 1963). He was the first to assay dehalogenases in cell-free systems and it was this work that promoted almost all subsequent studies, which have concentrated on HAAs substituted in the C2 position (2HAAs). Some microbes can grow on 3-halogenated alkanoic acids, but the biochemistry and enzymology is far from clear (Bollag and Alexander, 1971; Hughes, 1988). Interest in 2HAA biodegradation was stimulated by the introduction of the herbicide Dalapon (2,2-dichloropropionic acid; 22DCPA) and the isolation of many soil microorganisms that grew on 22DCPA as the sole carbon and energy source, thus explaining the rapid removal of Dalapon from soil (Magee andcolmer, 1959; Hirsch and Alexander, 1960; MacGregor, 1963; Kearney et al., 1964,1965; Kearney, 1966; Burge, 1969; Foy, 1975; Senior et al., 1976). Degradation of the highly toxic MFA by soil pseudomonads provided another stimulus for the characterization of new dehalogenases (Goldman, 1965; Kelly, 1965; Tonomura et al., 1965; Goldman andMilne, 1966; Goldman etal., 1968;Lien etal., 1979). Most research has focused on aerobic microbes. 2HAA dehalogenases calalyse the removal of halogens from organic compounds thereby forming simple compounds which are either intermediates of central metabolism or can be converted to intermediary metabolites. The general hydrolytic mechanism for mono-substituted alkanoic acids is as follows, where X represents the halogen atom: R - (CHX) - COOH + H20 + R - (CHOH) - COOH + H+ + Xand for di-substituted alkanoic acids: R - (CX2) - COOH + H20 R - (CO) - COOH + 2H+ + 2XGenerally 2HAA dehalogenases are inducible enzymes with low affinities for

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substrates (K, values in the millimolar range) and pH optima around pH 9.0. Typically these enzymes have broad substrate specificities and do not dehalogenate acids substituted other than at the C2 position. The stability of the carbon-halogen bond increases with increasing halogen electronegativity and this probably explains the greater ease of isolating microbes growing on chlorinated compounds compared with fluorinated acids (Hoffman, 1950). Principally studies have concentrated on halogenated acetic, propionic and butyric acids, and much less is known about the characteristics of enzymes that attack C5 or haloalkanoic acids of longer chain lengths. Enzymes that dehalogenate both D- and L-, or D- or L-stereoisomers are known. Frequently more than one dehalogenase is expressed by any given microbe (Hardman and Slater, 1981a,b). Some of the general properties of dehalogenases have been reviewed recently by Slater (19941, Janssen et al. (1994) and Slater et al. (1995) and will not be repeated here.

2.1. 2-Haloalkanoic Acid Hydrolytic Dehalogenases

Historically dehalogenases were grouped on the basis of substrate affinities, reaction kinetics, proposed or known catalytic mechanisms, and sensitivities to inhibitory compounds (Little and Williams, 1971; Hardman, 1991; Slater, 1994). We have recently updated a scheme for classifying dehalogenases (Table 1)(Slater et al., 1995) in order to accommodate the current range of known enzymes and their properties (cf. the earlier classifications of Weightman et al., 1982, and Hardman, 1991). Protein and nucleotide sequence data are defining significant similarities and differences between dehalogenases, and, in due course, accurate descriptions at these levels will determine evolutionary relationships (Leisinger and Bader 1993; Janssen et al., 1994; Slater, 1994). Presently the classification and naming of dehalogenases are based primarily on catalytic properties, with subgroups based on other factors, such as substrate specificities, and, where available, nucleotide or amino-acid sequence information.On this basis four classes of 2HAA hydrolytic dehalogenases are recognized (Table 1).

2.1.1. Class IL 2HAA Hydrolytic Dehalogenases These dehalogenases remove halides from L-2-haloalkanoic acids, inverting the product configuration with respect to the substrate, and react to sulfhydryl-blocking reagents to varying degrees. Goldman et al. (1968) postulated two mechanisms to explain the inversion of substrate configuration: an electron-donating group within the enzyme's active site yields a hydroxyl which displaces the halogen by a nucleophilic substitution; or 0 a carboxyl group within the active site acts as the nucleophile displacing the halogen with subsequent hydrolysis of the ester which is formed.

137

MICROBIAL DEHALOGENATION

1

Figure 1 Two proposed dehalogenation mechanisms (from Slater, 1994). A third mechanism, proposed by Little and Williams (1971), was a general base catalysis involving a histidine residue as the nucleophile (Fig. 1). Whatever the exact mechanism may be, presently there are 12detailed examples of this class of enzyme (Table I). The nucleotide sequences of nine of the 2HAA hydrolytic dehalogenases have been reported, clearly demonstrating nucleotide sequence similarities and defining this class as a coherent group of relatedenzymes (Table 2). Class 1L dehalogenases probably include the halidohydrolases I and I1 of the dichloroacetate (DCA)-degrading pseudomonad described by Goldman et al. (1968) and the dehalogenase of fseudumonas dehalogenans NCIMB 9061 described by Little and Williams (1971), although this cannot be asserted without the

Table 1 Classification o f 2-haloalkanoic acid hydrolytic dehydrogenases.

Class

1L

Organism

Enzyme

Gene

Number of amino acids Native Subunit Number from Calculated mol. mass mol. mass of nucleotide subunit mol. (kDa) (kDa) subunits sequence mass(kDa)

Reference

Pseudomom species CBS3

DehCI

dehCI

42.0

28.0

2

227

25.4

Pseudomoms species CBS3

DehCII

dehCii

64.0

29.0

2

229

25.7

Pseudomoms cepacia MBA4 Pseudomoms species 109 Pseudomoms putida AJl Xanthobacter autotmphicus a10 Pseudomoms species Y L Moraxella species B Rhizobium species

HdlIVa

hdlNa

45.0

23.0

2

23 1

25.9

DehHlO9

dehHI09

34.0

25 .O

2

224

25.2

Klages et al., 1983; Morsberger et al.. 1991; Schneider et al., 1991, 1993 Klages ef al., 1983; Morsberger etal., 1991; Schneider et al., 1991, 1993 Tsang et al., 1988; Murdiyatmo et al., 1992 Kawasaki et al.. 1994

HadL

hadL

79.0

26.0

2 (3?)

227

25.7

Jones et al., 1992

DhlB

dhlB

36.0

28.0

1

253

27.4

van der Ploeg et a[.,199I

L-DEXYL

LdexYL

54.0

27.0

2

232

26.2

Nardie-Dei et al., 1994

DehH-2

dehH-2

225

25.3

Kawasaki et al., 1992

280

30.9

Cairns, 1994; Cairns and Cooper, personal communication

DehI/HadL

26.0 60.0

2

Table 1 continued.

Class 1D

2R

21

Organism

Enzyme

Gene

Number of amino acids Native Subunit Number from Calculated mol. mass mol. mass of nucleotide subunit mol. (kDa) (kDa) subunits sequence mass(kDa)

hadD

Pseudomoms putidaAJ1 Rhizobium species

HadD

Pseudomoms putida PP3 A lcaligenes xylosooxidans Unidentified isolate K37 Pseudomonas put& PP3 Pseudomoms 113 Rhizobium species

DehI

dehl

DhlC

dehlC

DehI

dehl

DehII

dehll

DehIIVHadD

58.0 46.0

300

33.6

2

266

29.4

2

296

32.7

2

296

32.7

Barth et al., 1992; Smith et al., 1990 Cairns, 1994; Cairns and Cooper, personal communication Thomas, 1990; Topping, 1992 Brokamp and Schmidt, 1991 Murdiyatmo, 1991

52.0 68.0

DehII

33.0

4

Reference

Topping, 1992 35.0

2

Motosugi etal., 1982a,b Leigh et a [ . , 1988

Table 2 Amino-acid identity between Class 1L 2HAA hydrolytic dehalogenases (after Leisinger and Bader, 1993).

Organism Pseudomonas species CBS3 Pseudomonas species CBS3 I! putida All Xanrhobacter aurotrophicus GJlO I? putida 109 I? cepacia MBA4 Mornxella species B Rhizobium species Pseudomonas species Moraxella species P. putida AJ 1 Rhizobium species

Enzyme Class

Gene

dehCI dehCII hadL dhlB deh109 MlWa dehH2 hadL(R) LdexYL dehHI hadD hadD(R)

1L

dehCI

100

1L

dehCII

36

100

1L 1L

hadL dhlB

38 42

51 42

100 43

67 35

37 49

40 44

38

53

36 49 20 56

12 19

16 13

16 19

15 14

1L 1L

1L

100

1L 1L

1D 1D

100 100 33

100

39

51

100

19 21

19 17

12 19

100

44

57

100 16

100 23

100

MICROBIAL DEHALOGENATION

141

benefit of nucleotide sequence data. Furthermore, the I? dehalogenuns enzyme may be identical to halidohydrolase I. All three enzymes dechlorinated L-2-monochloropropionic acid (2MCPA) yielding D-lactate as the product but they did not dechlorinate D-2MCPA. Halidohydrolase I1 had a limited ability to use D,L2MBPA, but the product configurations are unknown and it seems unlikely that the D-isomer was dechlorinated. All three enzymes were unaffected by sulfhydrylblocking reagents, such as p-chloromercuribenmate and N-ethylmaleimide. Halidohydrolase I differed from halidohydrolase 11in that MCA was the main substrate for the former, and DCA was the principal substrate for the latter. The dehalogenase I of a Rhizobium species isolated on Dalapon was considered to be similar to these three enzymes (Berry et al., 1976, 1979; Allison el al., 1983; Leigh er al., 1988). Pseudomonas species CBS3 was isolated on 4-chlorobenzoate but synthesized two 2HAA dehalogenases, DehCI and DehCII, which dehalogenated L-2MCPAbut not D-2MCPA (Klages et al., 1983; Morsberger et al., 1991). Protein analyses showed that both of these enzymes were dimeric proteins with overall molecular masses of 41 and 64 kDa, and subunit molecular masses of about 28 and 29 kDa, respectively (Klages et al., 1983; Morsberger efal., 1991). Schneider et al. (1991) reported that the dehC1 gene (located on a 1.1 kb SinaI - SstI fragment) yielded a protein of 227 amino acids with a molecular mass of 25.4 kDa. The first 23 amino acids of the proposed sequence corresponded exactly with the N-terminal amino acid sequence from DehCl. The gene was preceded by a promotor of the -lo/-35 consensus sequence type and appeared to be negatively regulated by the enzyme’s substrates. The dehCII gene was similar and coded for a protein of 229 amino acids (molecular mass of 25.7 kDa) with the first eight amino acids of the N-terminus corresponding exactly with the sequence predicted from the nucleotide sequence. There was 45% nucleotide sequence homology, which corresponded to 37.5% amino acid sequence identity, and over 70% amino acid similarity (Table 2 ) . In this case there appeared to be a close evolutionary relationship between the dehCI and dehCIIgenes, which suggested a common origin from an ancestral gene. The observed pattern may have resulted from a gene duplication in the ancestral Pseudoinonasspecies CBS3 followed by separate but parallel evolutionary events. Alternatively, the genes may have evolved in parallel in another host organism and been transfei~edto Pseudoinonas species CBS3. Either way there was a close, and maintained, structural relationship between these two genes. In addition, there was some sequence homology between the N-terminal amino acid sequences of DehCI and DehCII, including a common AsplO, and a region containing the Aspl 24 residue in the haloalkane dehalogenase of Xanthobacter autotrophicus GJ 10, which was involved in the nucleophilic substitution catalysed by this enzyme (Section4.1). Schneider etal. (1993) used site-directedmutagenesis to replace the Aspl 0 with alanine, completely eliminating dehalogenase activity and concluded that AsplO was H nucleophiiic residue involved in the active site. The inference was that there was a defining relationship between these two major classes of dehalogenases i n the crucial region of the active site, suggesting a deep

142

J. HOWARD SLATER

eta/.

and relatively conserved evolutionary link within this region of the proteins. However, Janssen et al. (1994) questioned the direct role of Asp10 in the active site since “the proposed alignment neglects the fact that the sequence is determined largely by the position of the Asp124 in the nucleophile elbow between the p strand 5 and strand 3 and by the presence of Trp125 involved in the leaving-group stabilisation (halide binding)” and suggested that other Asp residues, Asp15 and Asp175, were involved in the active site. Tsang et al. (I 988) isolated Pseudomunus cepucia MBA4 from a batch culture growing on inonobromoaceticacid (MBA) and showed that under these conditions the organism synthesized two dehalogenases, 111 and IVa. The former was not studied in any detail, but the latter was found to be a Class 1Lenzyme (Murdiyatmo, 1991; Asmara, 1991; Murdiyatmo et al., 1992; Asmara et al., 1993). The sequence of the gene, hdllVa, for DehIVa (located on a 1.6 kb genornic fragment) predicted a gene product of 231 amino acids and a molecular mass of 25.9 kDa, corresponding to proteins observed by SDS-PAGE of about 23 kDa and by gel filtration of about 45 kDa. The sequence contained two regions which indicated -101-35 promotors, and it was suggested that the gene was under positive regulation. Unquestionably, this dehalogenase was closely related to others in the class: for DehIVa and DehCI there was 67% amino acid identity and 81% similarity, and for DehIVa and DehCII the corresponding values were 37% and 56%, respectively. Other relationships between Class 1L enzymes, based on amino acid identities, are shown in Table 2. Asmara et al. (1993), on the basis of chemical modification, random and site-directed mutagenesis, suggested that two amino acid residues, His20 and Arg42, were the key active site residues, with Asp18 also implicated in water activation by His20, again providing a possible link with the haloalkane dehalogenases (Janssen et al., 1989; Franken et al., 1991). Dehalogenase I (HadL) of a Hhizobiuin species has, for the moment, been included in Class 1 L on the basis of its overall catalytic properties. However, the protein showed minimal identity with Ha& of Pseudoinonasputida AJ1 (Table 2) and its subunit size appeared to be significantly different from the mean value of 231 for the eight dehalogenases that were unequivocally members of this class (Table 1). Similarly, DehH-2 of Moraxella species B has been included in Class 1Lon the basis of amino acid sequence information (Table 2) (Kawasaki et al., 1981a,b,c, 1992). This organism was isolated with MFA as the growth substrate and the two dehalogenases it contained dehalogenated only haloacetic acids; its response to stereo-specific compounds is not known. This example serves to reinforce the difficulty of using substrate specificities as the method of classification. The sequence data strongly suggested that this enzyme should be in Class lL, but it has evolved substrate specificities that are more limited and exclude C3 alkanoic acids. Basic information on the other microbial dehalogenases in Class 1L will not be discussed in detail, but may be found in the earlier reviews of Leisinger and Bader (1993), Slater (1994), Janssen et al. (1994) and Slater etal. (1995). The information

MICROBIAL DEHALOGENATION

143

presently available shows that this is an important and coherent class of 2HAA dehalogenases. The similarity of subunit sizes in terms of amino acid residues is striking (Table 1),although there may be interesting differences with respect to the number of subunits in the functional proteins. All the biochemical information strongly indicates a uniform class of proteins at the functional and mechanistic levels. In addition, there are suggestions of relationships with haloalkane dehalogenases, which should be of considerable interest in the future as evolutionary relationships are resolved (Section 4.1). 2.1.2. Class I D 2HAA Hydrolytic Dehalogenases The key feature of this class of 2HAA hydrolytic dehalogenases is their ability to dehalogenate selectively D-isomeric substrates, such as D-2MCPA, with inversion of product configuration. Many groups have tried, with little success, to isolate microbes that produce this class of dehalogenase; the most detailed work by the Zeneca plc (formerly ICI plc) group, involved Pseudomonas pufida AJl (Taylor, 1985, 1988; Barth, 1988; Smith e f al., 1989a,b, 1990; Barth et al., 1992). This industrial research group was interested in exploiting D-specific dehalogenase properties in a commercial process using L-2MCPA as the starting point for the synthesis of a novel herbicide. With a racemic mixture of D,L-~MCPA(manufactured cheaply by simple chemical routes) as the starting material, 50% of the final product did not have the required biological activity, and so half the synthesis was a waste of manufacturing time and resources. An initial treatment of the inexpensive racemic mixture to remove the unwanted D-2MCPA selectively ensured the efficient manufacture of a product with full biological activity, whilst the added manufacturing costs of the dehalogenase treatment step were offset by the improved unit costs of the L-specific process. Pseudornonasputida AJ 1, isolated from soil pre-enriched with racemic 2MCPA, contained two dehalogenases. Both enzymes were highly stereospecific: HadL was active against L-2MCPA (Section 2.1,l), whilst HadD dehalogenated D-2MCPA only (Table 1). Both native enzymes were probably tetramers with HadD having a molecular mass of 135 kDa (Smith et al., 1990), and HadLa mass of 79 kDa (Jones et al., 1992). The two genes, ha& and ha&, were closely linked on the genome. The hadD gene encoded a polypeptide of 33.6 kDa, and the first 21 amino acids deduced from the nucleotide sequence corresponded to the N-terminal analysis of the protein. Apart from the dehalogenase 111 from the Rhizobiurn species (see below), this enzyme had no significant homology with any other dehalogenase represented i n nucleotide or protein databases (Table 2). There was no -lo/-35 promotor preceding the structural gene. Recently, Cairns and Cooper at the University of Leicester have examined the Dalapon-degrading Khizobiuriz species (Section 3.1) containing dehalogenases I and 111, and have suggested that these enzymes corresponded to HadL and HadD

144

J. HOWARD SLATER eta/.

of r! putida AJI, respectively (Cairns, 1994; R.A. Cooper, personal cornmunication). The equivalent enzyme to HadD in Rhizobium, dehalogenase 111,had a native molecular mass of 58 kDa and a subunit molecular mass of 28 kDa, suggesting a dimeric structure. Nucleotide sequence data indicated a polypeptide of 29.4 kDa and 266 amino-acid residues. Sequence analysis suggested 23% identity with the amino-acid sequence of HadD, but no other relationships were found (Table 2). This was a rather low level of identity and there were other significant differences, such as its dimeric structure compared with the tetramer of HadD. Furthermore, the Rhizobiunz D-specific dehalogenase I11 subunit size was smaller than the H a m subunit size of P puridu AJ1 (Table 1). It is possible that these two D-Specific enzymes were related but it seems more likely that this was only at the functional level. 2.1.3. Class 21 2 H A A Hydrolyric Dehalogenases Enzymes of this class differ from Class 1 enzymes by their ability to dehalogenate both L- and D-isomers by a mechanism that inverts substrate-product configurations. This in turn distinguishes them from Class 2R enzymes (Section 2.1.4). Class 21 dehalogenases are unaffected by sulfhydryl-blocking reagents unlike Class 2R enzymes (Table 1). Motosugi et al. (1982a,b) isolated Pseudornonus species 113 which grew on both D- and L-2MCPA and synthesized a single dehalogenase which dehalogenated the L-isomer more rapidly than the D-isomer (D-isomer at 72% of the L-isomer rate). The dehalogenase had a molecular mass of 68 kDa and subunits of 35 kDa suggesting that the protein was a dimer. In a series of interesting experiments, Hasan et al. (1991) demonstrated that lyophilized preparations of the purified dehalogenase, when dissolved in organic solvents, such as anhydrous dimethyl sulfoxide (DMSO) or toluene, catalysed the dehalogenation of long-chain haloalkanoic acids and aromatic substituted haloalkanoic acids. For example, 2-bromohexadecanoic acid, which was not dehalogenated in aqueous solution, was dechlorinated in these solvents at 2.7 times the rate of 2MCPA. In solvents, L-2MCPA was only dehalogenated at 6% of the rate in aqueous solutions, leading to the suggestion that activity against higher chain length acids was due to their greater solubility in organic solvents. Pseudornonns putidu PP3 evolved in a stable microbial community growing continuously on the herbicide Dalapon and was found to synthesize two 2HAA dehalogenases (Slater et al., 1976, 1979; Senior et al., 1976; Senior, 1977). Dehalogenase 11 (DehII) (previously termed the fraction I1 dehalogenase) was active against a similar range of HAAs as the other enzyme, dehalogenase I (DehI) (Section 2. I .4), but differed by being unaffected by sulfhydryl-blocking reagents (Weightman el al., 1979, 1982; Weightman, 1981). The native molecular mass of DehII was 52 kDa (Topping, 1992). The differences were such that it seemed very

MICROBIAL DEHALOGENATION

145

unlikely that the two enzymes were related mechanistically and, therefore, originated from different ancestral genes, a situation closer to the unrelated HadL and HadD of l? putida AJI, rather than the two related enzymes of Pseudomoms species CBS3. So far DehII has not been sequenced and so comparisons with other dehalogenases remain tentative. However, it may be closely related to dehalogenase I1 of the Rhizobium species discussed in Sections 2.1.1 and 2.1.2, although the Rhizobium enzyme showed some sensitivity towards thiol reagents. 2.1.4.

Class 2R 2HAA Hydrolytic Dehalogenases

Enzymes of this class differ from Class 1 enzymes in their ability to dehalogenate both L- and D-isomers by a mechanism that retains substrate-product configurations, thus separating them from Class 21 enzymes (Section 3.3). These dehalogenases are strongly inhibited by sulfhydryl-blocking reagents unlike Class 21 dehalogenases (Table 1). Dehalogenase I (DehI) (formerly fraction I dehalogenase) (Slater et al., 1979; Weightman et al., 1979) was estimated to have a molecular mass of 46 kDa as the native protein. However, the nucleotide sequence for d e N predicted that the polypeptide had a molecular mass of 32.7 kDa which corresponded to SDS gel estimates of 33 kDa, suggesting possibly that the DehI protein was a dimer in the active state (Thomas, 1990; Topping, 1992; S.J. Hope and J.H. Slater, unpublished observations). The N-terminal sequence of the purified DehI has not been determined to date, but Murdiyatmo (199 1) purified an enzyme called HdlV from an unidentified isolate, strain K37, and determined the first 13 N-terminal amino acids. The sequence corresponded exactly with that of a protein encoded by the putative dehl open reading frame beginning at the second methionine residue. In the nucleotide sequence between codons specifying the first and second methionine residues (the DehI protein start), there was a strong Shine-Dalgarno sequence region separated by eight bases from the initiation codon, a distance considered to be optimal for transcription (Gold, 1988). Another Shine-Dalgarno sequence ten bases upstream from the codon specifying the first methionine was also located, and so the precise polypeptide sequence remains to be resolved. Recently we have shown that the amino acid sequence for DehI is identical to the sequence for DhlC fromAZcaligeizesxylosooxidnns (J.H. Slater, S.J. Hope, M.R. Lewis, A.W. Thomas, A.W. Topping, S.D. Greenaway and D.J. Hardman, unpublished observations). Unlike the genetic organization for all other dehalogenases described to date, a -12/-24 type promotor motif was identified upstream from dehl, These promotors are recognized by the 054family of sigma factors of RNA polymerases and have been associated with the expression of many metabolic functions in Gram-negative and Gram-positive bacteria (Dixon, 1986; Thony and Hennecke, 1989; Merrick, 1993). This confirmed earlier observations that expression of dehl was dependent

146

J. HOWARD SLATER eta/.

on the presence of a functional d4polymerase, since it was not expressed in rpoN mutants of t.i puticla suggesting that dehl was under positive regulatory control (Thomas et al., 1992b).

2.2. Regulation of Dehalogenase Synthesis

Very little is known about dehalogenase regulatory mechanisms at the molecular level, although the picture is becoming clearer with respect to the genetic organization of the regulation system for DehI. We observed, in gene transfer experiments (Section 2.3.1) that, when hybrid plasmids carrying dehl were transferred to other strains oft? putidu, l? aeruginosa or E. coli, dehl was regulated in a manner that was identical to the parental strain (Beeching et al., 1983; Slater et al., 1985; Thomas, 1990; Thomas et al., 1992a). Subsequently, mapping of an 11.6 kb EcoRI-G fragment from a derivative TOL plasmid containing the dehl gene, by transposon mutagenesis and complementation analyses, identified a region that was crucial to the expression of dehl. Topping ef al. (1995) demonstrated that dehl was adjacent to its regulatory gene, dehRI (Fig. 2). Analysis of dehRI showed that the gene had an open reading frame, which encoded a 64 kDa protein containing 571 amino acid residues. About 60 bases upstream from the putative translation start of dehR, was a highly conserved -35/-10-type promotor motif. A DNAbinding helix-turn-helix motif was identified at the C-terminus (Dodd and Egan, 1990). Comparison of the dehRI amino acid sequence with other d4-dependent activators showed that there was significant amino acid conservation within a central domain of approximately 230 amino acids, whereas there was little amino-acid sequence homology in the carboxyl- and amino-terminal domains. The conserved central region, termed the C-domain by Drummond et al. (1986), was responsible for interactions with RNA polymerase and bound ATP (Thony and Hennecke, 1989; Huala and Ausubel, 1989; Kustu et al., 1991; North et al., 1993) and may be expected to show significant similarities between different d4dependent activators. A cladograin showing the relationships between a number of d4-dependent activators is shown in Fig. 3 . This family of activators includes various nitrogenase regulators (NifA-like) and other nitrogen regulators (NtrC-like). This group contained only two activators for genes associated with biodegradative pathways, namely, DmpR, which regulated dimethylphenol catabolism (Shingler et al., 1993; Fernandez el al., 1994),and XylR, which regulated methylbenzoatecatabolism (de Lorenzo et ul., 1991). However, in the C-domain region of 229 amino acids, DehRI showed only 25.6% similarity with XylR and 26.1% similarity with DmpR. DehR1 did not cluster with either the NifA or NtrC groups; instead it had greatest C-domain similarity (48%) with the putative product of the hyuR sequence from a Pseudornonas species, a plasmid-borne regulatory gene involved in L-amino acid biosynthesis (Watabe et a/., 1992). Despite the similarity of DehRl to NifA-like and

0

I

2.00

4.00

6.W

8.00

10.00

12.00

I

I

I

I

I

I

H M l

PSI

3.80

4.41

4.69

4.91

6.69

6.91

Figure2 Physical map showing the arrangement of the DEHtransposon containing the structural gene dehl and its regulator gene dehRI, showing the key endonuclease restriction sites (after Topping et al., 1995).

148

J. HOWARD SLATER

I I

eta/.

NifA- like

NtrC-like

Figure 3 Cladogram illustratin the relationships between the derived DehRI aminoacid sequence and those of other J4-dependent activators. The cladogram was produced after an alignment of the conserved central regions (see text) of the selected activators using CLUSTALV (Higgins el a/., 1992) and manual editing. Sequence similarities were determined by the Dayhoff PAM matrix method and Neighbor-Joining using the PHYLIP package (Felsenstein, 1989). The following sequences were obtained from the GenBank database unless noted otherwise: AcoR-A.e; acetoin catabolic regulator from Alculigenes eutrophus (P28614-SWISSPROT database); AcoR-C.m, acetoin catabolic regulator from Clostridiurn magnu/n (L3 1844); DctD-R.I, C4-dicarboxylate transport regulator from Rhizobiirm /egu/ninosunrm (X06253); DehRI-ttp, dehalogenase regulator from Pseudomonus pufida (U237 16); DmpR-Psp, dimethylphenol catabolic regulator from Pseudomonas sp. CF600 (X68033); FixD-R.m, nitrogenase regulator from Rhizobium meliloti (X03065); GlnBXc, glutamine metabolic regulator from Escherichiu coli (S67014); HoxA-A.e, hydrogenase regulator from A. eurrophus (M64593);HydG-Ec, hydrogenase regulator from E. coli (M28369); HydG-S.t, hydrogenase regulator from Salmonella typhimurium (M64988); “HyuR’-Rsp, hydantoin catabolic regulator from Pseudomoms

MICROBIAL DEHALOGENATION

149

NtrC-like activators, the dehl gene was not activated in putatively trans-acting plasmid constructions with these nitrogen metabolism activators (A.W. Topping, unpublished observations), suggesting that the significant sequence differences at the termini, particularly the N-terminus, resulted in effector specificity and defined the regulatory capability of DehRI. The G + C content of dehRI was much lower than that expected for a Pseudumunas species (51.7% compared with about 63%). Murdiyatmo et al. (1992) and Jones el al. (1992) also reported lower G + C ratios for dehalogenase structural genes, suggesting that these genes originated in other species and were transferred to Pseudoinonas species later in their evolution, perhaps by plasmid transfer or as a component of an appropriate transposon (Section 2.3.2).

2.3. Genetic Organization of 2HAA Hydrolytic Dehalogenases

2.3.1. Plasrnids Frequently, dehalogenase genes are located on plasmids (Slater and Bull, 1982; Reanney et al., 1983). Kawasaki and his colleagues showed that the MFA- and MCA-utilizing capability in Moruxella species B was associated with a plasmid (Kawasaki el al., 1981b,c,d, 1982, 1983a,b, 1984, 1985, 1992). Strain B contained a single large plasmid, p U 0 1, with a size of 66 kb and encoded the genes for both Deh-H1 and Deli-H2. In some mutants, for example, strain 86, in which Deh-H2 was lost, there was an accompanying decrease in the size of the remaining plasmid, whilst in other strains, for example strain 123,complete loss of the plasmid resulted in the loss of both dehalogenases. Brokamp and Schmidt (1991) demonstrated the transfer of plasmid pFL40, which carried the Class 2R 2HAA dchalogenase in species NS671 (QOl265-SWISSPROT); LuxO-Vh,luminescence regulator from vibrio harveyi(L26221);NifA-Azb.v, nitrogenaseregulatorfrom Azofobacter vineZandii(Y00.554); NifA-Azxc, nitrogenase regulator form Azorhizobium caulinodans (X08014); NifA-K.p, nitrogenase regulator from Klebsiella pneumoniae (X02616);NifA-R.E. nitrogenase regulator from R. legiiminosarum (LI 1084); NifA-Rhd.c, nitrogenase regulator from Rhodabacter capsulatus (X07567); NtrC-Agt, nitrogen regulator from Agrobacterium tumefaciens (J03678); NtlC-Azs.b, nitrogen regulator from Azospirillum brasiiiense (X67684); NtrCB K ~nitrogen , regulator from Bradyrhizobium parasponia (M14227); NtrC-E.c, nitrogen regulator form E. coli (X05173); NtrC-K.p, nitrogen regulator from K. pneumoniae (X02617); NtK-R.m, nitrogen regulator from R. meliloti (M15810); NtrC-TJ nitrogen regulator from Thiobacillrrsferrooxiduns(L18975); PilR-Pa, fimbriae expression regulator from Pseudomonos aei-uginosa (Q00934-SWISSPROT); RocR-B.s, ornithine aminotransferase regulator from Bacillus subtilis(L22006);vrR,E.c, tryptophan biosynthetic regulator from E. coli (M12114); VnfA-Azb.v, nitrogenase regulator from A. vinelandii (M26752); XylR-Pp., methylbenzoate catabolic regulator from P. putida (P06519-SWISSPROT) (from Topping et al., 1995).

150

J. HOWARD SLATER eta/.

Alcaligenes xylosooxidans ABlV to Pseudoinonas fluorexems and other soil bacteria in sterile soil microcosm experiments. Hardman (1982) showed that almost all novel, freshly isolated 2HAA-utilizing strains of soil bacteria contained large plasmids, varying in size from about 150 kb to 300 kb (Hardman etal., 1986).Spontaneous loss or curing of the plasmidresulted in loss of the ability to grow on 2HAAs. Beeching et al. (1983) showed that it was possible to mobilize the dehl gene on to suitable target plasmids such as RP4 and RP45, and we have come to realize that this property was associated with the transposon structure carrying the dehl gene (Section 2.3.2). 2.3.2. The DEH Transposon Many 2HAAs are toxic and their uptake into cells causes cessation of growth and cell death if exposure is prolonged. Active synthesis of dehalogenases removes the inhibitory problem and growth resumes (Slater et al., 1979). In selection experiments originally designed to produce mutants with impaired uptake mechanisms, it was found that mutants of Pseudoinonas putida PP3 resistant to concentrations of MCA or DCA as high as 100 m~ were obtained at very high frequencies. For example, at 42 m~ MCA, resistant mutants appeared at a frequency of 3.2 x lo-' and, at 42 mM DCA, the frequency was 5.3 x (Slater etal., 1985). The mutants were stable and on analysis five classes were defined (Table 3), all of which had reduced capabilities for the uptake of 2HAAs (Slater et al., 1985). On the basis of these observations, we proposed that the mutants were the result of the loss of one or more transposons carrying DchI and DehII, and their associated uptake systems, which impaired the uptake of 2HAAs leading to the resistant phenotypes. Subsequently the transposon, DEH, carrying dehl and dehRI, was characterized (Thomas, 1990; Thomas et al., 1992a,b,c; Topping, 1992). DEH was an unusual transposon, since it transferred to target DNAs as an element of varying size, ranging from 6 to 13 kb. To date there is no evidence to confirm that dehll was also located on a transposon, and it is possible that mutations involving the loss of this gene were associated with the movement of DEH and topological interactions between DEH and the dehll gene which resulted in changes of expression of both genes. Furthermore, changes in the growth environment, in particular the presence of toxic 2HAAs, resulted in high-frequency mutations which switched off one or both genes, and other conditions resulted in the expression of dehllbeing switched back on (Thomas et al., 1992a; Hope and Slater, 1995; Slater and Hope, 1995). R putida PP3 and its associated mutants (Table 3) routinely formed papillae in colonies grown under appropriate conditions as cryptic dehalogenase genes were switched on (Thomas et al., 1992a; Hope and Slater, 1995; Slater and Hope, 1995). Movement of DEH under these conditions led to major rearrangements of the DNA as shown by altered restriction enzyme digest patterns (Thomas et al., 1992a). In other systems, loss of Deh-H2 was associated with a discrete piece of DNA

151

MICROBIAL DEHALOGENATION

Table 3 Characteristicsof various W A Rmutants of Pseudornonusputida PP3 isolated by growth in the presence of 42 mM DCA and succinate.

Strain or mutant class

Specific growth rate' (h-')

PP3 PP40 PP411 PP4 12 PP4 120 PP42

0.3 1

0.00 0.14 0.23 0.04

0.26

Response Dehalogenase complement to DCA Permease (85 mM) DehI DehII activityb S R R R R R

+ +

-

-

+

+

-

+ + +

8000 100 1900 2100 100 657

%Total mutants -

16 54 7 3

20

a Average

values based on three independent determinations,except for the growth rates of PP412 mutants (four detenninations) and PP4120 mutants (six determinations). Determined from the initial rate of uptake of [ 14C]MCAwith rates expressed as counts per minute of radioactivity uptake per minute of incubation per 1 mg of cell protein (from Slater, 1994).

with a constant size of 5.4 kb and it was suggested that this gene might be located on a transposon (Kawasaki et al., 1983a,b, 1984, 1985). Kawasaki et al. (1992) showed that the dehH2 coding region was sandwiched between two repeated sequences of about 1.8 kb, and suggested that these homologous sequences might be implicated in the excision of the gene and/or its transposition. Topping (1992) also found that there were short repeated, homologous regions within the DEH transposon. Janssen et al. (1994) suggested that the overexpression of DhlB in mutants of Xmthobacter autotrophicus was due to the insertion of a small DNA fragment upstream of the dhlts gene. Clearly a great deal of further information about the mobility of dehalogenase genes is needed to shed light on their expression and control, and the interaction between different dehalogenase genes.

3. DEHALOGENATION OF HALOGENATED ALCOHOLS Dehalogenation of haloalcohols has been known since Castro and Bartnicki (1965) showed that 3-bromopropanol was biodegradable. These workers isolated a Flavobacteriurn species which grew on 2,3-dibromopropan-l-ol and found that it converted halohydrins to epoxides (Castro and Bartnicki, 1968; Bartnicki and Castro, 1969). At present it appears that inicroorganisms have evolved two main pathways for the mineralization of haloalcohols.

152

J. HOWARD SIATER eta/.

3.1. A Pathway Involving the Formation of HAAs

The first pathway is mediated by enzymes that accept halosubstituted molecules as their substrates, with dehalogenation taking place only after the haloalcohols have been oxidized to the corresponding haloalkanoic acids. This mechanism was described for the degradation of 2-chloroallylalcohol(van der Waarde et al., 1993), and was found in the Gram-negative bacterium CElr (Stucki and Leisinger, 1983) and Pseudornorzas putida U2 (Strotmann er al., 1990). In Gram-negative bacteria, the haloalcohol pathway was part of the core pathway described for the degradation of 1,2-dichloroethaneby Xanfhobacter autotrophicus GJlO (Janssen ef al., 1985) (Section 4.1). Here catabolism of the disubstituted haloalkane resulted in the formation of 2-chloroethanol and, thereafter, enzyme recruitment resulted in the conversion of the halogenated alcohol to MCA, which in turn was dehalogenated to glycolate by a 2HAA hydrolytic dehalogenase. For example, Stucki and Leisinger (1983) showed that strain CElr synthesized alcohol and aldehyde dehydrogenases, which specifically converted 2-chloroethanol to 2-chloroaldehyde and thence MCA. The 2-chloroethanol-degrading strains expressed the same catalytic pathway as Xarzthobacter autufrophicus GJ10, without the first hydrolytic haloalkane dehalogenase, and constitutively expressed an MCAspecific hydrolytic dehalogenase. A comparison of the enzymes from the pseudomonads and the Xanfhobacter strain at a molecular level would be very interesting in order to determine the exact relationships between these pathways. The possibility that they are closely related and that acquisitive evolution has led to the construction of this pathway was supported by the observation that the structural genes for the haloalkane dehalogenase and the aldehyde dehydrogenase for the 1,2-dichloro-ethane pathway were encoded on a 200 kb plasmid (Tardif et al., 1991). In a related pathway, Rhodotorula glutinis catalysed the asymmetric reduction of prochiral chloroacetone to chiral l-chloro-2-propanol using an NAD(P)H-dependent alcohol dehydrogenase (Weijers ef al., 1992), which suggested that these enzyme systems are related and widely distributed in nature.

3.2. Haloalcohol Hydrolytic Dehalogenases The alternative pathway involves specific haloalcohol dehalogenases. When enrichment cultures were established to select for halohydrin-degrading bacteria, the competent isolates normally catalysed the direct dehalogenation of vicinal haloalcohols via a process that is basically hydrolytic, but in which the actual dehalogenation step involved an internal rearrangement of the substrate and a simultaneous elimination of a proton, with the halide resulting in the formation of an epoxide (Fig. 4). This mechanism, first describedby Castroand Bartnicki (1965) in the 2,3-dibromo-1-propanol-degrading Flavobacteriurn species was catalysed

153

MICROBIAL DEHALOGENATION

CH,CI-CHOH-CH,CI 1,3-dichloro-2-pmpnol CH,OH-CHOH-CH,CI 3-c hl oro-l,Zpropandiol

- /"\ + H,O

-HCI

(a)

-HCI .

(C)

CH,CI epicNomhydrin

(b)

0

+HO 4C%OH 2 (d) glycidol

CH,OH-CHOH-CH,OH gly c e d Figure 4 Pathway for the catabolism of 1,3-dichloro-2-propanol(DCP). (a) and (c), haloalcohol dehalogenases; (b) and (d), epoxide hydrolases.

by enzymes termed halohydrin epoxidases (Castro and Bartnicki, 1968). Subsequently, these enzymes have been variously described as haloalcohol dehalogenases and haloalcohol halogen-halide lyases (van den Wijngaard et al., 1989, 1991) and halohydrin halogen-halide lyases (Nagasawa et al., 1992). In the complete pathway, the epoxide resulting from the dehalogenation reaction is hydrolysed by an epoxide hydrolase to the corresponding alcohol (Fig. 4). Haloalcohol dehalogenases showed a restricted range of activities being active only when the halogen is vicinal to a hydroxyl group or to a keto group, such as in chloroacetone (van den Wijngaard et al., 1991). The most commonly used substrates for the isolation of haloalcohol-competent strains were 1,3-dichloro-2propanol (13DCP) (Nakamura et al., 1992), 3-chloro-l,2-propandiol (3CPD) (Suzuki and Kasai, 1991; Suzuki eral.. 1992) and epichlorohydrin (ECH) (van den Wijngaard el a/., 1989).The dehalogenation reaction was reversible, which allowed the interconversion ofvic-haloalcohols and epoxides. This reversibility also means that the enzymes catalyse the trans-halogenation of haloalcohols. The pathway for the biodegradation of 13DCP mimics the chemical degradation of the substrate when exposed to solutions of high pH (Fig. 4).In all of the systems reported to date, two different epoxide hydrolases were required to open the ECH and glycidol ring structures resulting from the dehalogenation of 13DCP and 3CPD, respectively. However, when a strain lacked the epoxide hydrolases, for example, Arthrobacter species AD2 (van der Wijngaard et al., 1989), chemical hydrolysis of the dehalogenation products still enabled it to utilize the metabolites for growth, albeit at a much reduced rate. In common with other aliphatic biodehalogenation systems, multiple dehalogenases are frequently observed in isolates. The most common profile is two dehalogenases, although isolates with only a single enzyme are known (Fig. 5).

154

J. HOWARD SIATER

1

2

eta/.

3

Dehalogenase D Dehalogenase C

Dehalogenase B Dehalogenase A

i

Dehalogenase A with different subunit

combinations

Figure 5 Zymogram of the common haloalcohol dehalogenase forms as separated by activity PAGE. Examples: 1, Agrobacterium tumefaciens HK7 (Bull et al., 1992), Pseudomonas species AD1 (van den Wijngaardet al., 1989);2, Arthrobacterhistidinolovorans (Bull e l al., 1992). Arthrobacter AD2 (van den Wijngaard el al.. 1989);3 “Arthrobacter erithii” (Assis et al., 1996). Corynebacteriurnspecies N-1074 (Nagasawa et al., 1992).

Five alcohol dehalogenases have been described in detail from P s e u d o m m s species AD1 and Arthrobacter species AD2 (van den Wijngaard et al., 1989), Colynebacterium species N-1074 (Nagasawa et ul., 1992) and another pseudomonad capable of growth on 2,3-dichloro-l-propanol(23DCP)(Kasai et al., 1990, 1992).In our laboratory, we have identified six electrophoreticallydistinct enzymes from species of Pseudomonas, Agrobacteriurn, Arthrobacter and Rhodococcus (Bull etal., 1992; Assis et al., 1996; A.J. Cotton, M. Huxley, D.J. Scherr, A.T. Bull, J.H. Slater and D.J. Hardman, unpublished observations). DNA profiling studies indicated that Pseudoinonas species AD1 may in fact be a strain ofAgrobacterium tumefaciens (A.J. Cotton, S.D. Greenaway, A.T. Bull, J.H. Slater and D.J. Hardman, unpublished observations). In comparison with the other classes of dehalogenases, the biochemical and molecular data are limited (Table 4). The current information suggests that there is limited diversity among these enzymes isolated from organisms across a wide geographical distribution. The

MICROBIAL DEHALOGENATION

155

available data suggest that there are only two classes of haloalcohol dehalogenases, defined ds Class 4 s and Class 4C (Slater et al., 1995) (Table 4). Class 45 enzymes are structurally simpler than Class 4C enzymes, being simple dimeric proteins formed from a single polypeptide subunit; Class 4C enzymes are composed of at least two different polypeptides. Class 4C enzymes exhibit a narrower substrate profile that Class 4 s enzymes, but they also have chiral selective properties. In the same way that the 2HAA dehalogenases are grouped according to their relative activities towards different substrates, so can the haloalcohol enzymes. The Class 4C enzymes, as represented by haloalcohol dehalogenase Ib from Corynebacteriurnspecies N-1074 (Nakamura et al., 1992) and DehA from “Arthrobacter erithii” (Assis et al., 1996), showed little activity towards 3CPD, but high activity towards 13DCP. In contrast, other enzymes, such as the enzyme from Alcaligenes species DS-S-7G (Suzuki et aE., 1992), had no activity towards 13DCP and greatest activity towards 3CPD. Such differences are analogous to the different specificities of the 2HAA dehalogenases towards mono- and di-halogenated acetic acids (Hardman and Slater, 1981a). Activity towards 23DCP also served to differentiate these enzymes (Kasai et al., 1990, 1992; A.J. Cotton, A.T. Bull, D.J. Hardman and J.H. Slater, unpublished observations), whereas Pseudomonas species AD 1 and Arthrobacter species AD2 (van den Wijngaard et al., 1989), Corynebacteriurn species N-1074 (Nagasawa et al., 1992) and Flavobacteriurn species (Castro and Bartnicki, 1968) had little or no activity towards 23DCP. Dehalogenation of 2CPD has not been reported. Detailed biochemical information is available for only four of the haloalcohol dehalogenases. The enzyme from Arthrobacter species AD2 was purified and characterized (van der Wijngaard et at., 1991). This Class 4 s dehalogenase (molecular mass 65-69 kDa) was a dimer consisting of two identical subunits of mass 29 kDa. The enzyme was iinmunologically distinct from that of Pseudoinonas species AD1 and also demonstrated very different substrate profiles. The haloalcohol dehalogenase I, from Corynebacteriurn species N-1074 (Nagasawa et al., 1992)was catalytically similar to the AD2 enzyme and, despite the fact that enzyme I, was a hoinotetrainer (molecular mass 105 kDa), its subunits showed a similar molecular mass (28 kDa) and a very Similar N-terminal amino acid structure to the AD2 dehalogenase. In contrast to the similarities between the dehalogenase 1, and AD2 enzymes, the second enzyme, Ib, in Corynebacteriurnspecies N-1074 was markedy different from the I, dehalogenase. Again the presence of genetically, immunologically and mechanistically different dehalogenases within the same organism has been described previously for 2HAA dehalogenases (Weightman et al., 1982). Dehalogenase Ib, a representative of Class 4C, was a tetrameric protein (molecular mass 115 kDa) forined from two different polypeptide subunits (32 and 35 m a ) . On PAGE, a ladder of five dehalogenase-active bands was created as a consequence of the formation of homo- and hetero-tetramers from different combinations of the

Table 4 Properties of selected haloalcohol dehalogenases.

Organism Pseudomonas species AD1 Dehalogenase Enzyme class Molecular mass

4s

orb>

Subunits Subunit molecular mass (ma) Relative substrate activity 13DCP 100.00 23DCP 0.00 3CPD 19.00 1CP 13DCA CA 13DBP 134 OOO.00 1BP 2CE 13DCPaffinity constant

Alcaligenes

Arthrobacter

Corynebacterium species

species AD2

N- 1074 Ia

Ib

4s 65-69

4s 105

4c 115-118

DehA 4c 200

2 29

4 28

4 32 and 35

6 3 1.5 and 34

100.00

100.00 0.30 37.50 22.70

100.00 0.09 0.92 18.50

12 500.00

161.00

60.00

8.50

0.26 2.44

0.13 1.03

1.20 0.11

9.00

3.13

148.00

236.00

10.00 11.00 356.00 10.00 31 000.00 3780.00

N-1074

species OS-K-29

100.00 10.00 33.00

species DS-K-S-38

100.00 47.00 106.00

“Arthrobacter erithii H l Oa”

100.00 0.00 0.10 27.00 72.00

(Km)(m)

v m (Fr;nOl

min- mg-’) Stereopsecific pH optimum Temperature optimum (“C)

No

No 8.5 50

No 8.0-9.0 55

Yes 45

Yes

Yes

Yes 8.5-9.5 50

Table 4 continued.

Organism Pseudomonas species AD1

Arthrobacter species AD2

Inhibitors (% max. activity) Mercuric Slight chloride NEM None MCA (ki) (mM) 0.05 Reference van der van der Wijngaard et Wijngaard et al. (1989) af. (1991)

Corynebactenum species N- 1074

N-1074

26.0

80.4

Pseudomonas species OS-K-29

A lcaligenes species DS-K-S-38

“Arthrobacter erirhii H 1Oa”

100.0

None Nagasawa et al. Nakamun el 01. (1992) (1992,1994)

Kasai et nl. ( 1992)

Suzuki et al. ( 1992)

Assis ef nl. (1 996)

13DCP, 1,3-dichloro-2-propanol;23DCP, 2,3-dichlorc+I -propanol; 3CPD. 3-chloro-1.2-propandiol; 1CP 1-chloropropanol; 13DCA, 1.3-dichloroacetone; CA, chloroacetone; 13DBP. 1,3-dibromopropanol; I BP, I-bromopropanol;2CE, 2-chloroethanol; NEM. N-ethylmaleimide; MCA, rnonochloroacetic acid.

158

J. HOWARD SLATER eta/.

two subunits (4:O; 3:l; 2:2; 1:3; 0:4) (Nakamura et al., 1992). A second example of a Class 4C haloalcohol dehalogenase, DehA, was purified and characterized by Assis et al. (1996) from “Arfhrubacfererithii” HlOa. This enzyme also was composed of two different subunits (molecular masses 31.5 and 34 kDa) and had a molecular weight of 200 kDa, suggesting that it was a hexameric protein. The Corynebacteriurn I b and the Arthrobacter DehA enzymes were the first dehalogenases reported to be composed of two different polypeptides. The Class 4C haloalcohol dehalogenases appeared to have a greater affinity for 13DCP than the Class 4 s enzymes. However, Fauzi et al. (1996) isolated oligotrophic bacteria growing on very low concentrations of 13DCP.These bacteria possessed haloalcohol dehalogenases that belonged to Class 4s (in terms of electrophoretic mobilities) but demonstrated K, values in the order of 0.1 mM, more akin to the values for Class 4C enzymes. The main functional characteristic of Class 4C haloalcohol dehalogenases was their enantioselective catalytic activity which may be related to their relative complexity. The similarity witli the complex structure of the D-specific 2HAA dehalogenase which was a tetrameric protein and was structurally significantly different to the dimeric L- and D,L-2-haloalkanoic acid dehalogenases, may be significant (Section 2). The enantioselective dehalogenation of 13DCP, leading to the formation of R-ECH, has commercial interest for the synthesis of chiral pharmaceuticals, such as P-adrenergic blockers, platelet-activating factor, vitamins, pheromones, agrochemicals and ferro-electric crystals. Most of the reported methods for the production of chiral epoxides using dehalogenating bacteria were based on the enantioselective degradation of racemic mixtures. The process patented by Zeneca plc for the production of L-2-~nonochloropropionate(Section 2.1.2), utilized a D-specific dehalogenase to remove selectively the D-enantiomer from the racemic mixture. In the case of the chiral epoxides, either 23DCP (Kasai et al., 1990,1992) or 2-chloro-1,3-propandiol (2CPD) (Suzuki and Kasai, 1991; Suzuki et al., 1992) have been used. The unassimilated haloalcohol accumulated in the growth medium and was subsequently converted chemically to the epoxide by addition of hydroxide. Using this approach, both isomers of ECH and glycidol (GDL) were obtained in high enantioineric excess (>99%). The disadvantage of enantiomer resolution based on enantioselective biodegradation was that the yield of the desired product was less than 35%. From an industrial point of view, the production of optically active compounds by enantioselective microbial transformation of prochiral starting materials is more attractive, since a quantitative yield of the desired enantiomer can be obtained. Nakamura et al. (1992, 1994) studied such a transformation of 13DCP to R-2CPD by Corynebacterium species N-1074. Using this method, they obtained a molar conversion yield of 97.3% but the enantiomeric excess of the R-CPD produced was too low (87.3%) to be of commercial interest. The characterization of the haloalcohol dehalogenases is not as advanced as the other groups of aliphatic dehalogenases. It is apparent, however, that there is

MICROBIAL DEHALOGENATION

159

diversity of structure and function, which is comparable to that of haloalkane dehalogenases (Section 4).

3.3. Other Systems for the Transformation of Haloalcohols

A novel enzymatic mechanism responsible for the catalysis of the NAD-dependent oxidative dechlorination of K-3CPD to acetic and formic acids was recently described in Alcaligenes species DS-S-7G (Suzuki ef al., 1994). The enzyme (Enzyme 1) that demonstrated enantiospecificity was characterized and shown to be a dimeric flavoprotein (molecular mass 70 kDa) composed of two different polypeptides (58 and 16 kDa). This protein was associated with a second protein (Enzyme 2), a dimer (86 kDa) comprised of two subunits (33 and 53 kDa). which, whilst unable to catalyse the dehalogenation reaction itself, promoted a 4-5-fold increase in the haloalcohol dehalogenation activity when combined with Enzyme 1. Although Enzyme 1 was capable of the direct conversion of 3CPD, this was the least efficient pathway. The Enzyme 1 and Enzyme 2 complex, the more effective “dehalogenase”, converted the 3CPD to acetic and formic acids via hydroxyacetone and formaldehyde intermediates at rates that were 4-5 times more rapid than Enzyme 1 alone. The Km of the “combined” enzyme for 3CPD was 322 pM with a Vmx of 3.34 pmol min-’ mg-’.

4. DEHALOGENATION OF HALOGENATED ALKANES

Haloalkanes are significant environmental compounds, occurring as both natural products (Gschwend etal., 1985)and as xenobiotic compounds (Keith andTelliard, 1979). Their biodegradability was uncertain until Omori and Alexander (1978a,b) reported that about 1% of soil microorganisms were able to utilize 1,9-dichlorononane under aerobic conditions. Brunner et a/. (1980) showed that dichloromethane was fully mineralized by microbial populations, and Wilson and Wilson (1985) showed that trichloroethylene was completely mineralized by soil populations when also provided with methane under aerobic conditions. Murphy and Perry (1983, 1984) isolated fungi with an ability to use 1-chlorooctadecane and I-chlorohexadecane aerobically, and also showed that the fatty acids of these organisms were heavily halogcnated. Under anaerobic conditions, methanogenic bacteria can degrade a wide range of halogenated alkanes, such as chloroform, carbon tetrachloride and dichloroethylene (Bouwer et al., 1981; Bouwer and McCarty, 1983a,b). Polychlorinated compounds were reductively dechlorinated to less heavily chlorinated products. Belay and Daniels (1987) showed that halogenated compounds were toxic to methanogenic bacteria and that brominated compounds were more toxic than

160

J. HOWARD SLATER

eta/.

chlorinated compounds. Vogel and McCarty (1985) proposed a series of reductive dehalogenations to mineralize tetrachloroethylene completely via trichloroethylene, dichloroethylene and vinyl chloride, a route that was subsequently supported by others (Fathepure et al., 1987; Mikesell and Boyd, 1990). Fathepure and Boyd (1988) suggested that electrons released during the reduction of carbon dioxide to methane were transferred instead to the halogenated alkanes leading to the elimination of the halogen; certainly methanogenically active consortia degraded tetrachloroethylene more rapidly (Fathepure er al., 1987). The microbiological evidence is for a wide diversity of haloalkane dehalogenation mechanisms under aerobic conditions, and there seems to be a much wider diversity of systems and mechanisms involved in haloalkane dehalogenation than for either halogenated alkanoic acids or alcohols. Haloalkane degradation involving NADH-linked reactions (Omori and Alexander, 1978a,b), oxygenases (Hartmans et al., 1985, 1986; Yokota et al., 1986), glutathione (GSH)-dependent reactions (Kohler-Staub and Leisinger, 1985) (Section 4.3), as well as hydrolytic mechanisms (Janssen et al., 1985; Yokota et al., 1986) have been proposed and demonstrated. In general terms the pH activity profiles are broad with an optimum that is slightly lower than that seen for the dehalogenation of haloalkanoic acids, and the enzymes show broad substrate specificities. The most detailed knowledge is available for the hydrolytic mechanisms found in aerobic microbes. These mechanisms have been discovered and studied in Xanthobacter autotrophicus GJlO (Janssen et al., 1985, 1989; Keuning el al., 1985), various Corynebacteriwn species (Yokota et al., 1986), Arthrobacter species (Scholtz et al., 1988a), Rhodococcus species (Sallis et al., 1990) and Ancylobacter aquaticus (van den Wijngaard et al., 1992).

4.1 Haloalkane Hydrolytic Dehalogenases On the basis of substrate specificity, these enzymes presently can be divided into two classes (Slater et al., 1995). 4.1.1. Class 3R Haloalkane Hydrolytic Dehalogenases This class of enzymes found in Gram-negative bacteria shows a restricted range of substrate specificities and has been most extensively studied in Xanthobacrer autotrophicus GJlO (Janssen et al., 1989; Keuning et al., 1985). This organism, which was isolated from enrichment cultures on dichloroethane, also synthesized a Class 1L 2HAA hydrolytic dehalogenase (Section 2.1.1). The hydrolytic haloalkane dehalogenase was heat-labile and acted principally on C 1 4 4 substituted alkanes, such as dichloroethane, bromoethane, 1-chloropropane, l-chlorobutane, 1,3-dichloropropane and 3-chloropropene, yielding the corresponding alcohols,

161

MICROBIAL DEHALOGENATION

CI-$CI-CH,CI 1,2-&chloroet haw

+ HO ,

CH$I-CHO + NADH lchloroacctaldehyde

+

CH,OH-CH,CI l-chlwoethallol

+

+ NADH CH,CI-COOH rnonochloroacetic acid

H+

+

HCI

-

NAD+

+ HO , +N AP

H+

+ HO ,

:

CH,OH-COOH glycolate

+

HCI

Figure 6 Pathway of haloalkane catabolism.

such as ethanol, I-propanol and 1-butanol, as reaction products. The enzyme was expressed constitutively and the enzyme accounted for 2-3% of the total cellular protein content. Janssen et nl. (1985) proposed a simple pathway for haloalkane metabolism (Fig. 6 ) . The purified enzyme had a molecular mass of 36 kDa and functioned without any cofactors (Keuning et al., 1985). dehalogenating l-chloroalkanes up to C4 and l-bromoalkanes up to C10. The substrate affinities of the enzyme were in the millimolar range; for example, the K,,, for dichloroethane was 1.1 m ~and , the pH optimum was 8.2. Janssen et al. (1988) proposed that the enzyme catalysed a nucleophilic substitution with water, and it was also observed that the reaction was strongly inhibited by thiol-blocking reagents, possibly implicating a cysteine residue at the active site (Keuning et al., 1985). Superficially there appeared to be some relationship with 2HAA hydrolytic dehalogenases, but no immunological cross-reactions were demonstrated (Keuning ef al., 1985). Janssen et al. (1989) determined the nucleotide sequence, finding a gene that was predicted to encode for a polypeptide of 310 amino acid residues with a molecular mass of 35.1 kDa. This suggested that the native enzyme was composed of one polypeptide subunit. The three-dimensional structure of the enzyme was determined and shown to be composed of two domains (Rozeboom et al., 1988; Franken et al., 1991; Verschueren ef al., 1993a,b,c). The main domain was composed of a central eight-stranded P-sheet surrounded by a-helices, a structure which is common to many hydrolytic proteins and leads to the concept of the general a / p hydrolase. Surmounting the main domain was a second, termed the cap domain, which was composed of five a-helices linked by loops. The active site was located between the two domains and formed an internal hydrophobic cavity. X-ray crystallography at low temperatures (Verschueren et al., 1993~)revealed three key amino acid residues at the active site: Asp124, which functioned as the nucleophile; His289,

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which was involved in the hydrolysis of the enzyme-substrate covalent intermediate; and Asp260, which stabilized the positive charge, which developed on His289 (Fig. 7). The key role of Asp1 24 was demonstrated by site-directed mutagenesis, since replacement of this residue with alanine, glycine or glutamic acid inactivated the enzyme (Pries ef al., 1994).The halide binding was achievedvia two tryptophan residues, namely, Trpl25 located in the main domain, and Trp175 located in the cap domain. Clearly the enryme is a dehalogenase and not a general broadspectrum hydrolase, which functions as a hydrolytic dehalogenase in a fortuitous manner. Van den Wijngaard et al. (1992) isolated a number of facultative methylotrophs, including various Ancylobacter aquaticus strains and Xanthobacter autotrophicus GJ11, growing on dichloroethane by virtue of a constitutive haloalkane dehalogenase. Sequence data (N-terminal amino acid analyses and polymerase chain reaction (PCR)-amplified sequences of the putative dhlA gene from these strains) showed that the enzymes were identical to the dichloroethane dehalogenase from X.aututropliicus GJlO (Janssen et al., 1985). Tardif et al. (1991) demonstrated that this dehalogenase was plasmid-encoded and therefore the evidence for horizontal gene transmission was strong (Section 2.3.1). Janssen et al. (1994) recently suggested that the 2HAA dehalogenase DehHl from Moraxellu species B, despite limited sequence similarity, may be closely related to the haloalkane dehalogenase of X. aufotrophicus GJ10. The evidence relates to the sequence similarity around the nucleophilic aspartate within the active site, and the overall structure (i.e. main and cap domains) and catalytic mechanisms. However, similar relationships can also be defined between the Xunthornoms enzyme and other enzymes, so that the relationship may reflect common ancestry with a general dfi hydrolase. These enzymes are: tetrachlorocyclohexadiene hydrolase (LinB) from I? paucirnobilis UT26; 2-hydroxy-muconic semialdehyde hydrolase (DmpD) from P srudornonas species CF6OO (Norlund and Shingler, 1990); 2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoate hydrolase (BphD) from I? putida KKS 102 (Kimbara et al., 1989); 4-methyl-2-hydroxymuconic semialdehyde hydrolase (TodF) from f? putidu (Menn et al., 1991); dienelactone hydrolase (TcbE) from Pseudomonus species P51 (van der Meer et af., 1991); dienelactone hydrolase (TdfE) from A. eutrophicus (Perkins et al., 1990); and Figure 7 Catalytic mechanism of haloalkane dehalogenase. (a) The carboxylate group of Asp124 acts as a nucleophile, which displaces the halide from the substrate. The leaving group is stabilized by two hydrogens bound to the ring nitrogens of Trpl25 and Trp175. The covalent alkyl-enzyme intermediate is hydrolysed by a water molecule that is activated by His289, which acts as a base catalyst. (b) Attack by water at the carbonyl function of the ester results in hydrolysis of the intermediate, leading to incorporation of oxygen from water in the enzyme. Asp260, the third residue of the catalytic triad, may stabilize the temporary positive charge on His289. (c) After hydrolysis of the covalent intermediate, the alcohol, halide ion, and proton leave the active site. (Reproduced, with permission, from Janssen et al. Annual Review of Microbiology, Volume 48,O 1994, by Annual Reviews Inc.)

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(a)

80%decrease in germanium accumulation, inferring an active transport process. EDS revealed that germanium was associated with the cells. In contrast, Ps. stutzeri AG259 accumulate germanium by an energy-independent mechanism (Van Dyke et al., 1990).

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7.6.4. Lead Although toxicity of lead compounds in higher animals, including humans, has been widely studied, relatively little information is available on lead resistance in bacteria. Lead resistance has been found in different bacterial isolates such as S. aureus, Micrococcus luteus, Azotobacter spp., and K.aerogenes (Novick and Roth, 1968; Tornabene and Edwards, 1972; Aiking et al., 198s). Both methylation and immobilization of Pb2+have been suggested as resistance mechanisms. Detoxification of Pb2+by K. aerogenes NCTC 418 has been studied in some detail and the formation of PbS was suggested to be the resistance mechanism (Aiking et al., 1985). AAS has been used to study Pb2+uptake by bacteria. In the study of Tornabene and Edwards (1972), cellular distribution of lead was analysed. After separation and digestion in concentrated nitric acid, different cellular fractions were analysed by AAS. Approximately 99% of the cellular Pb was bound to the cell envelope. Using electron microscopy, Aiking et al. (1985) showed that PbS precipitate was deposited in the cell envelope of K.aerogenes NCTC 418. This method avoids the complication of separating different cell fractions.

7.6.5.

Mercury

Mercury compounds have been used as catalysts in industry as well as disinfectants for clinical purposes. Both mercuric ion (Hg2') and organomercurial compounds are highly toxic to biological systems owing to their strong affinity for thiol groups in proteins. Hence, accumulation of Hg contaminants in nature causes serious environmental and public health problems (Belliveau and Trevors, 1989a; Misra, 1992). Mercury resistance involving enzymatic reduction of Hg2+to volatile elemental mercury and/or decomposition of organomercurialcompounds have been observed in Gram-negative and Gram-positive bacteria (Misra, 1992). Mercury-resistance phenotypes are plasmid mediated and can be divided into two groups. First, broad-spectrum resistance determinants detoxify organomercurial compounds by a two-step process that includes cleavage of the C-Hg bond by an intracellular organomercurial lyase enzyme (MerB) followed by reduction of Hg2+to Hgo by an FAD-containing, NADPH-dependent mercuric reductase (MerA). The second system is a narrow-spectrum pathway, which confers resistance only to inorganic mercury ions via MerA. nierP and merT encode mercuric ion transport proteins. MerP is a Hg2+-bindingprotein located in the periplasmic space. It functions as a shuttle to transfer Hg2+ to the membrane-bound MerT protein, which internalizes the Hg2+into the cytoplasm. The Hg-resistance operon is controlled by the MerR regulatory protein. In addition to studies of the genes required for mercury resistance and their

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individual roles, the MerR protein and its reaction with Hg2+have been intensively studied in vitm, but this work is outside the scope of the present chapter (see O'Halloran et al., 1989; Wright et al., 1990). Similar1 203Hg2+(Sahlman and Jonsson, 1992) has been used to show that MerP is a Hg +-bindingprotein, which binds one Hg2+per MerP monomer. Resistanceto mercuric chloride and phenyl mercuric acetate in a Bacillus cereus strain was encoded on a self-transmissible plasmid pGB130 (Belliveau and Trevors, 1990). A mating system for a Gram-positive bacterium like Bacillus allowed study of the transfer frequency (lo4 to and levels of mercuric reductase activity in the wild-type and transformants. Electrotransformation was also used to introduce this plasmid from B. cereus to B. thuringiensis (Belliveau and Trevors, 1989b).

F

'

7.6.6. Selenium Although selenium is an essential trace element for animals (Gadd, 1993),elevated concentrations of selenium oxyanions are highly toxic to living systems (Painter, 1941). As analogues to sulfur compounds, selenite and selenate inhibit sulfite reductase and sulfate transport systems in bacteria, respectively (Brown and Shrift, 1980; Harrison etal., 1980).Several fungal and bacterial spp., such as a Fusarium sp., Wolinellasuccinogenes, some Salmonella strains and Desulfovibrio desulfuricans, can reduce selenate and/or selenite to elemental selenium and hence detoxify the selenium oxyanions (Gharieb et al., 1995;Tomei et aE., 1992, 1995). Quantitative measurements of selenium can be done by usin either radioactive "Se or fluorometric procedures. Using Na?%e03 and Na27FSe04, Brown and Shrift (1980) showed that selenate ion was transported and assimilated by the same process as was sulfite in S.+phimurim. However, selenite was not transported by the sulfate carrier. Selenium reductase of Desulfovibrio desulfuricans was studied by Tomei et al. (1995). Cell samples were digested with perchloric and nitric acids. After treating with 2,3-diaminonaphthalene and extracting by cyclohexane, fluorescence of samples was recorded (excitation and emission at 369 and 525 nm, respectively). D. desulfuricans was found to reduce 95 and 97% of 1 pM selenate and 100 p~ selenite, respectively. Using electron microscopy and EDS, selenium granules were shown to be deposited in the cytoplasmic fraction of the bacteria. 7.6.7. Silver Soluble silver ions are probably toxic to all microorganisms. Indeed, silver compounds have long been used as anti-microbial agents for treating burns and eye infections of newborn infants (Slawson et al., 1992). A study by Modak and Fox ( I 973) showed that '"Ag was detected intracellularly in f! aeruginosa cells exposed to this isotope, where the majority of the silver

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was complexed to DNA. Binding of '"Ag to RNA and cell envelope was about 1/40 and 1/10, respectively, of the amount bound to DNA. Bacteria that exhibit tolerance to Ag+ have been isolated. For example, Ag"-resistant E. coli and €? stutzeri strains have been isolated from a burn patient and a silver mine, respectively (Gadd et al., 1989; Goddard and Bull, 1989; Starodub and Trevors, 1989). Uptake and accumulation of Ag' have often been associated with silver resistance and detoxification, inferring exclusion or immobilization of Ag' as a mechanism to alleviate Ag' toxicity in cells. There is experimental evidence to suggest a plasmid-encoded Ag' resistance mechanism in an E. coli strain (Starodub and Trevors, 1989). TEM and EDS showed that the Ag+-resistant strain did not accumulate silver, whereas the sensitive strain contained dense silver deposits. Quantitative analysis by AAS showed that the Ag+-sensitiveE.coli strain accumulated about five-fold more Ag than the resistant strain (Starodub and Trevors, 1990). In addition, the Ag+-resistantstrain produced one third more H2S and intracellular acid-labile sulfide than the sensitive strain. Similar Ag' resistance phenomena have been observed in €? stutzeri (Slawson et al., 1992). There is no evidence that Ag' is transformed to Ag(0) by a reductase enzyme.

7.6.8. Tellurium

The metalloid tellurium is used in batteries, alloys and rubber and as a colouring agent in glass. Tellurium compounds are toxic to many microorganisms, particularly Gram-negative bacteria. However, some strains of Corynebucreriurndiphtheriae, Streptococcusfaecalis, S. aureus, A. faecalis and A. denitri'cans, and some members of the family Enterobacteriaceae are resistant to tellurite (Walter and Taylor, 1992). Plasmids from three incompatibility groups have been shown to control tellurium resistance in bacteria (Walter and Taylor, 1992). Unfortunately, the mechanisms of plasmid-mediated tellurite resistance are not known, but increased efflux or reduced uptake have been ruled out as possible mechanisms. Although a tellurite-reducing protein of approximately 53 kDa was purified from 7: thermophilus, it is not related to any of the known tellurite-resistance determinants (Chiong el al., 1988). Investigations of tellurite resistance have been hampered by the lack of a convenient assay to measure the concentration of tellurite and tellurium. Quantitative measurement of the black metallic tellurium was used by Chiong et al. (1988) to estimate tellurite reduction in different cellular fractions of Thennus thermophilus. Electron spectroscopic imaging and X-ray diffraction have also been used to locate metallic tellurium deposition in a teilurite-resistant E. coli mutant (Taylor et al., 1988). Metallic tellurium was shown to be accumulated just inside the inner membrane of the bacteria. An AAS procedure has been developed to detect

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23 1

tellurium in animal tissues (Cooper, 1971). The detection limit of this method is 2 pg Te g-' tissue. 7.6.9. En Tin compounds are used for protective coating of metals or as anti-fouling agents in marine paints (McDonald and Trevors, 1988). Although many studies on tin toxicity have focused on organotin compounds, inorganic hydrated tin chloride can also be toxic to microbial populations at concentrations above 10 mg 1-' (Hallas and Cooney, 1981). The toxicity of tin compounds is caused in part by inhibition of oxidative phosphorylation and energy metabolism. The form in which tin is present is critical to its toxicity. Trialkyl tin compounds are the most toxic, followed by dialkyl tin. Monoalkyl tin compounds are non-toxic, and tetraalkyl tin is only toxic when converted to trialkyl tin. Bacterial degradation of triorganotin compounds can be considered as a detoxification mechanism, since the conversion of bis(tributy1tin) oxide (TBTO) to monobutyltin derivatives results in lower toxicities (Barug, 1981). Gas chromatography-mass spectrometry (GCMS) was used by Barug (1981) to study the degradation of TBTO. In this study, none of the bacterial and fungal species was able to utilize TBTO as sole carbon source. However, Ps. aeruginosa and some fungal strains degraded TBTO and converted the biocide to a less toxic monobutyltin compound. Using HPLC coupled with a graphite furnace atomic absorption spectrophotometerand gas chromatography coupled with a tin-specific flame photometric detector, Blair et al. (1982) showed that some marine bacterial isolates were able to degrade TBTO and immobilize 3.7-7.7 mg tin g-' dry weight of cells. Radioactive '13Sn has been used to study uptake and accumulation of inorganic tin compounds (Wong et al., 1984). By studying the accumulation of "'Sn in different cellular fractions of an Ankistrodesrnus falcatus strain, these researchers suggested that the uptake of tin was a physicochemical surface adsorption process; 85% of the Il3Sn recovered was in the polysaccharide fraction. Using GCMS analysis, Hallas et al. (1982) showed that inorganic Sn(1V) was rnethylated in sediments. Although evidence has implicated methyltin species as being responsible for increased formation of methylmercury in sediments, the significance of this reaction is presently not clear.

8. CONCLUSIONS AND OUTLOOK Achieving the ultimate objective of a comprehensive understanding of the interactions between metals and microorganisms requires taking a rnulti-

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disciplinary approach. This involves microbiology, particularly microbial physiology, genetics and molecular biology, bioinorganic chemistry, analytical chemistry and the application of instrumental techniques. Our aim in this contribution has been not to review this field but to survey these approaches. We firmly hold the view that the exploration of the effects of any metal ion on microbial physiology is hazardous unless due attention is paid to the underlying chemistry. There are examples in the literature of conclusions that are erroneous or suspect because the speciation and bioavailability of the metal ion under study have not been rigorously considered. Equally important is the ability to control intracellular concentrationsof metals. The last point is illustrated well by the recent use of photolabile compounds for binding calcium. Undoubtedly, such new techniques will find valuable applications in the hands of ingenious investigators. Genetics and molecular biology hold great promise for advancing studies of metal-microbe interactions,particularly those aspects that are currently perceived as especially significant. These include gene regulation by metals (metalcontaining transcription factors and zinc fingers), control of specificity at metalbinding sites and genetic modification of microorganisms for application in biohydrometallurgy.The availability of extensive sequence data-in an increasing number of cases, of the whole genome-will allow hunting for genes and their proteins that display clear binding motifs for metal ions or prosthetic groups. It is clear, however, that microbial physiology coupled with bioinorganic chemistry will remain as key contributors in this field.

Danielle Fortin and Suzanne Schultze-Lam of TJB's laboratory supplied the TEM images and this TEM research was supported by the Natural Science and Engineering Research Council of Canada. RKP and MNH acknowledge support from the Natural Environment Research Council (UK). Research by JTT is supported by the Natural Sciences and Engineering Research Council of Canada in the form of an operating grant.

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Author Index Numbers in bold refer to pages on which references are listed at the end of each chapter Abdel-Wahab, N.H., 53,57,59,60,67,83 A M , A., 223,233 A h , T., 2M), 242 Abraham, E.P., 49,81 Abul-Haj, Y.J., 6, 8, 18,31,41 Adams, M.D.. 211,236 Adams, M.W.W., 180,232 Adams, S.R., 188,189,232 Adams, T.H., 27.36 Adams, T.J., 217,233 Adler, J,, 189, 242 Adlington, R.M.,49,81 Agosin, E., 13,40 Aharonowitz, Y., 94,%, 97,124,129 Ahmad, A., 186,233 Ahmed, E.D., 165,166 Aiking, H., 227.228,233 Ainsworth, A.M., 2,40 Aitken, A., 60.81 Alatalo, E., 5, 8,40 Alexander, M., 135, 159,160,167,169,172, 173 Alijah, R.,121,131 Alison, J.E., 88, 92,98,99,129 Allison, J.D., 186,233 Allison, N., 141,166,167 Altier, D.J., 55, 74,76 An, D.,50,55,58,76 Andersen, N.H., 8.37 Anders, M.W., 165,166 Anderson, G.L., 225,233 Andrew, S.C., 216,233 Anzai, H., 121,126,128 Apel, K., 9.37 Aramaki, H., 57, SO Archibald, F.S., 188, 190,233 Arends, J., 17,37 Armes, L.G., 226,238

Armfield, S.J., 160, 164,165,166,173 Am. N., 58.40

Aronson, A.I., 11,37 Arreguin, B., 8,37 Asante-Owusu, R.N., 23,24,37 AsgeirsdMr, S.A., 3, 4. 5 6 , 10. 12, 13, 14, 19, 20,21,41,44 Askendal, A., 16.38 Asmara, W., 138,142,149,166,167,172 Assis, H.M.S., 154, 158, 167 Asther, M., 10,38 Atherton, E.. 111.128 Atsuyuki, S., 121,126 Atto-Asato-Adjei, E., 68,69,70.71,72 Augier, H., 197,233 Aumercier, M., 224,241 Ausubel, EM.,146,169 Avery, S.V., 183.233 Avi-Dw, Y.,209,237 Axcell, B.C., 50,53,58,76 Axelrod, B.,73,81 h a m , F., 227,233 Babich, H., 182, 183, 185,233 Bachman, B., 48,W Bader,R.. 136,137,140,142,171 Bailey, G.W., 183.240 Baines, A.J.. 138, 142,149,167,172 Bairoch, A., SO, 55,61,66,73,81 Bak, F., 197,233 Baker, D.P., 56,82 Baker. P.B., 165,166 Baldwin, J.E., 49,81,97,98, 117, 124,129 Ballou, D.P., SO, 51,53,55,56,57,60,61,64,

65,66,67,69,72,75,76,77,78,79, 80,= Banham, A.H., 23,24,37 Bapat, M., 112,125 Barra, G.E., 230,235

246 Barra, R., 230,235 Bmedo, J.L., 88,98,125 Barrett, J., 194,226, 233 Bamault, D., SO, 80 Barth, P.T., 138, 139, 142, 143, 149, 165,167. 170 Bartilson, M., 146,173 Bartnicki,E.W., 151,152,153,155,167 Barton, B., 88,92,98,99,129 Barton, L.L., 229,242,243 Barug, D., 231,233 Basham, LP., 183,239 Bashford, C.L., 219,233 Bates, R., 166,169 Batie, C.J., 50.51, 53,5556, 57,60, 61, 64.65, 66. 67,69, 72, 76, 77, 79, 80,83 Batley, G.E., 185, 233 Bauerfeind, P., 224, 240 Bayer, E., 120,124 Bazett-Jones, D.P., 230,242 Bazylinski, D.A., 223,233 Beall, M.L., 135,171 Beard, S.J., 214, 222,234 Beardwood, P., 76 Beck, J.N., 196,240 Beckett, A., 29,38 Beckmann, J.D., 68,76 Beeching, J.R., 146,150,167 Beever. R.E., 5, 8,9, 11, 12, 17,28,33,37 Beg, F., 60,81 Belay, N., 159, 167 Belliveau, B.H., 228,229, 234 Bell-Pedersen, D.. 5 , 11, 12, 28,37 Benitez, T., 5 , 39 Benkovic, S.J., 49, 81 Berg, T.L.. 86, 124 Bergeron H., 50,83 Bergman, B., 215,240 Bergmeyer, J., 94,96,124 Bemhard, M.,185, 187,234 Bemhardt, FAI., 49, 75.77, 83,84 Benistein, LA., 226, 239 Berry, E.K.M., 141,167 Bertrand, P.,65,76,77 Bessen, R., 227,239 Best, D., 165.169 Beveridge. T.J., 178, 183, 201, 202,203,234, 236,237,239,240,242 Bezkorovainy, A., 191,216,238 Bidcchka,M.J., 11, 12, 13.37 Bill, E., 75.77, 209,216, 239 Billich, A., 100, 102. 105, 114, 124

AUTHOR INDEX Bird, J.W., 97,124 Bird, N.P., 185,234 Birkenkamp, K.U., 3,5,24,25,26,44 Bitton, G., 227,234 Blair, W.R., 231,234 Blake, C.K., 49,82 Bleasby, A.J., 148, 169 Blum, ED., 6, 8, 18,31,41 Blum, H.,72,82,220,241 Blumenthal, H.J., 10.38 Bohlmann, H., 9.37 Bohlool, B.B., 215,238 Bolin, J.T.,73,81 Bollag, J.M., 135, 167 Boller, A., 99, 128 Bolton, L., 139, 142, 167 Bolyard, M.G., 9, 18, 33,37,42 Booij, H., 9,41 Borchert, S.,90,125 Borel, J.F., 105,125 Borst-Pauels, G.W.F.H., 180, 234 Bomvka, L., 16.41 Bosscher, J., 5.6, 12, 26,39 Boucias, D.G., 29,37 Bouwer, E.J., 159, 165,167 Bowden, C.G., 5, 18, 19, 31,37 Boyd, S.A., 160,168,172 Boylan, M.T., 28,36 Brackman, J., 161, 170 Bradley, EC., 73,77 Bradshaw, R.M., 11,44 Brandt, U., 68, 72.79 Brand, J.M., 48.49, 53,77,79, 80 Brasier, C.M., 18, 31,37 Bratt, P.J., 67, 82 Braun, D.G., 102,124 Braun, V., 189,237 Bremer, P.J., 207, 234 Bnnckman,F.E., 185, 187, 231, 234 Bnscoe, P.A., 230,236 Britt, R.D., 67,69, 72,77 Broda, E., 224,234 Brodexick, J.B., 73, 75,77 Broekaen, W.F., 8,9,40,42 Broer, A., 226,234 Brokamp, A., 139,149,167 Bronchart, R., 10.37 Brown, D.S., 186,233 Brown, G.A., 17,38 Brown, N.L., 214,234 Brown, T.A., 229,234 Brown, T.R., 209,241

AUTHOR INDEX Browne, J., 88. 92,98, 99, 129 Bruins, A.P., 162,173 Brunner, W., 165,167 Bruschi, M., 59, 62.77 Bucheder, F., 224,234 Bull, A.T., 135, 136, 138, 142, 144,145, 149, 150,154, 155, 160,164,165,166, 167,168,169,172,173,174,175, 176,222,235,237 Bull, J.H., 88,92,98,99, 129 Bunz, P.V., 50,53, 56,58,59,74,77 Burgess, T., 5, 33, 39 Burgett, S.C., 99,130 Burge, W.D., 135,167 Burnham,M.K.R., 88,92,98,99,129 Busscher, H.J., 16. 17, 29, 30,37, 40,43, 44 Butler, C.S., 59, 83, 219,242 Butt, T.M., 29.38 Byford, M.E, 97,98, 117, 129 Byrom, D., 138, 143, 149,170 Cabanthik, Z.I., 217,239 Cabral, J..P.S., 196, 234 Caceres, O., 162.173 Cairns, S.S., 138, 139,167 Calzada, J.G., 98, 125 Cammack, R., 50, 53,57,58,59,60,61,63,65, 67, 68,71,72, 73,75,77,79,80,81, 82,83,208,219,235,242 Cammue, B.P.A., 9,42 Cantoral, J.M., 94,%, 124 Cao, B., 8, 37 Cao, J., 99, 130 Carey, P.R., 5,6, 9, 18,44 Carpenter, C.E., 5,6, 8, 9, 13, 18, 19,32,38 Cam, M.T., 223,235 Casselton, L.A., 5, 6, 23, 24,37, 38,39 Casserly, D.M., I%, 240 Casteras-Simon,M.,221,239 Castonguay, Y.,9.38 Castro, C.E., 151, 152, 153, 155,167 Cavalier-Smith, T., 22,38 Cervantes, C., 213,214,225,226,234,240 Cha, J.-S., 221, 234 Chakrabarty, A.M., 164,168 Chakravarti,B., 49,81 Chalatnish, S.. 209,237 Chambers, J.G., 185,234 Chance, B., 219,233,234,241 Chang, Y.C., 5,12,27,33,38,40 Charlent, O., 197,233 Chau, YK.,198,231,234,235,243 Cheetham, A.K., 209,239

247 Chen, C., 226,238 Chen, R.F., 50.84 Chen, S.S., 219,235 Chen, V.J., 49, 73,77,81, 99, 130 Cheong, C.-M., 60,61,83 Cheung, T.T., 200,242 Chiong, M., 230,235 Chmielowski, J., 227,235,238 Chu, L., 226,240 Chumleym, F.G., 29,30,38 Ciccognani, D.T.,189,222,234,235 Ciriolo, M.R., 223,235 Civitareale, P., 223,235 Claverie-Martin, E, 11,38 Clayton, R.A., 211,236 Clement, J.A., 29.38 Cline, J.F., 65, 66, 77 Clutterbuck, A.J., 27.38 Ccdd, G.A.. 183,233 Cohen, G., 94,%, 124 Colby, J., 139, 143, 165, 168, 174 Cole, A.L.J., 135, 171 Cole, G.T.,3, 10, 11.38 Cole, R.M., 201,202,203, 234 Collins, Y.E., 185, 186, 187,235 Colmer, A.R., 135,172 Columbo, E., 119,130 Commandeur, L.C.M., 134,168 Connolly, T.N., 186, 237 Cook, A.M., 50,53,55,56,57,58,59,60,64, 77,80,82, 160, 164,169,173 Cooksey, D.A., 214,221,234,235 Cooney, J.J., 231, 237 Cooper, C.E.. 59,83, 219,242 Cooper,R.A., 139, 141, 144, 166, 167,171 Cooper, W.C., 230,235 Ccque, J.J.R., 98, 125 Corbell, N., 91, 92, 120,125 Corbisier, P., 212,235 Correll, C.C., 55,56,77, 78 Correll, C.J., 51, 55,76 Cosmina, P., 119,120, 125,130 Costerton, J.W., 11.44 Cotoras, M., 13,40 Cotton, A.J., 154, 155 Cowart, R.E., 217,233 Crawford, B.F., 179,221,239 Crawford, R.L., 49,81 Crestani, B.. 5, 6, 8, 12,33,42 Cruden, D.L., 49.53,77,79,82 Crutcher, S.E., 53,64,77 Cui, X.Y., 208,235

248

AUTHOR INDEX

Cummins, D., 185,234 Curry, J.. 119,128 Daldal, F., 67.68.69, 70,71, 72, 77 Dalton. H., 48,77, 165,168,174 Daniels, L., 159,167 Davidson, E.. 67,68,69,70,71,72,77 Davies, M.M.. 165, 169 Day, E.P., 63,65,68,72,78 de Boer, P., 17.37 de Bont, J.A.M., 152, 160,169,176 de Cavalho, D., 5,33,39 De Cky-LagWd, Y.,92,125 de Ferra, F., 120, 125 de G m t , P.W.J., 5 , 3 8 de Jong, H.P., 17,37 de la Cruz, I.,5,39 de Lorenzo, V., 146,168 De Martino. A., 223,235 De Rome, L., 222,235 de Vos, W.M., 162,175 devries, O.M.H., 3,4,5,6,9, 10, 11, 12.13, 14, 17, 19,28,30,34,38,39,42,43,

44 de Vries, S.C., 19,38,41 de Wet, J.R., 90,125 de Wit, P.J.G.M., 9, 43 Dean, A.C.R., 209,227,239 Dean,R.A.,4,5, 8, 11, 12, 13, 27,28,42 Debrunner, P.G., 65,81 Deising, H., 29.40 DeLuca, M., 90,125 Dernain. A.L., 94.96.124 Demoulin, V., 10.37 Dempsey, G., 11, 12, 17,37 Den Dooren de Jong, L.E., 134,168 den Hollander, J.A., 209,240 Dengis, P.B., 17,41 Denome, S.A., 67.77 dev~e, r.w., 188,233 Diaz-Torres, M.R., 11,38 Dibrov, P., 193,200,237 Dickson, D.P.E., 65,78 Diddens, H., 120,125 Diez, B., 88,98,99,125 Dijkhuizen,L., 152, 160,161, 162,170 Dijkstra, B.W., 142, 161,168, 173,175 Dilworlh, M.J., 182,215,236 Dittmann, J., 106,127 Dixon, R.A., 145,168 Dodd, LB., 146,168 Doi, S., 50,W Dolenw, E.K., 218,239

Donamo, S., 90,126 Dons, J.J.M., 19,38 Doolittle, R.F., 6,7,39 Dorendorf. J., 121,131 Douglas, C., 90,127 Dowell. R.M., 91,129 Doyle, R.J., 178, 234 Drenth, J.H.H., 4, 8,9, 10, 12. 13, 14,20,21, 3% 44 Dreyfuss, M.M., 107,125 Drummond, M., 146,168 DSwza, C., 91,92,120,125 DuBow, M.S., 224,237 Dudding. T.. 48, 80 Dufrene, Y.F., 17,41 Dunham, W.R., 63,65,68,72,78 Dunlap, J.C., 5, 11, 12,28,37 Dunlap, J.R., 215,241 Dunn, G.M.,222,235 Ebbole, D.J., 5.8.28, 29, 30.42 Ebdon, L., 198,235 Ebersold, H.-R., 165, 174 Ebersptkher, J., 50, 53, 56.57.59, 64,82 Eckart, K., 117,127 Edwards, H.W.. 228,243 Egan, J.B., 146,168 Eggen. R.I.L., 162, 175 Eggermont, K., 9.42 Eggink, G., 57.78 Eilatn, Y,200,236 Eizember, L.E., 218,239 Ellis-Davies, G.C.R., 188,235 Elwing, H., 16,38 Emery, T., 217,235 Engel, H., 57, 78 Ensley, B.D., 50,53,55, 64,67,78, 82 Epstein, L., 29, 30, 31,40 Erge, D., 109,128 Erickson, B.D., 50,53,60,61,67,73,78 Esaki, N., 134, 138, 139,144,169,172 Ewart, D.K., 194,226,233 Farrell, R.E., 186,235 Fathepure, B.Z., 160,168 Fauzi, A.M., 158,168 Fee, J.A., 59,63,65,66,67,68, 69, 72,77, 78, 79, 80 Feig, A.L., 49,72,73,78 4, 9, 11. 13, 28, 34,38 Fekkes, M.P., Felix, A., 165,173 Felsenstein, J., 148, 168 Femandez, S., 146,168 Ferrari, M.A., 30.39

AUTHOR INDEX Fems, EG., 183,235,240 Feteke, F.A., 217,235 Fetzner, S.,48,49, 50,53,56, 57.60, 64, 78, 79.82 Field, R.A., 97,124 Finding, K.L., 63,65,68,72,78 Finette, B.A., 53, 84 Fink,N.H., 229,243 Fink, U., 94,96,124 Firtel, M., 202,203,236 Fisher, D.J., 17,38 Fitz-James, I?, 11,37 Fleischmann, R.D., 211,236 Flemming, C.A., 183,240 Flis, S.E.,182,215,236 Florence, T.M., 195,236 Forge, A., 11.44 Forgeot, M., 99,129 Foster, T.J., 226, 236 Fowden, L., 135, 168 Fox, B.G., 55, 73,74,76,81 Fox, C.L., 229,240 Foy, C.L., 135, 168 Franchi, E., 119,130 Frank, R., 138, 141,173 Franke, P.,91,93, 117,129 Frankel, R.B., 223,233 Franken, S.M., 142,161,168,175 Frantz, B., 164, 168, 229, 240 Fraser, C.M., 211,236 Frederick, C.A., 73.82 Freihofer, V.,227,234 Frey, A.J., 108, 126 Fridovich, I., 188, 189,241 Frimmel, F.H., 187, 236 Friihner, C., 50, 53,56, 57.59, 64,82 Fr(dholm,L.O., 86,124 Frolik, C.A., 49,77,99,130 Fu, C., 224, 236 Fuchs. J.A., 226,238 Fuchs, R., 148, 169 FiihrbaB, R., 88,130 Fuji, I., 93, 94,96, 123, 126 Fujisawa, H., 50,53,56,57,60,64,84 Fujishima, Y.,92, 120,125 Fukuda, M., 57.80,162,171 Fukuyama, K., 62.78 Fuma, S., 92, 120,125 Furukawa, K., 53,60,61,62,73,78,83 Fusamoto, H., 219,235 Fuse, H., 160,164,176 Futsi, F., 135, 175

249 Fyfe, W.S., 183,235 Gabellini, N., 63,67,78 Gadd, G.M., 178, 180,182, 183, 187,221,222, 226,229,230,233,235,236,238, 240.243 Galiazzo, F., 223,235 Galli, R., 165, 168, 171,174 Gangola, P., 191,236 Garber, E.A.E., 226,238 Garner, R.M., 224,240 Garnon, J.. 50,83 Garratt-Reed, A.J., 223,233 Gassner, G.T., 55,75,78 Gatti, D.L., 5 5 7 8 Gay, N.J., 90, 130 Gayda, J.-P., 65, 76, 77 Geary, P.J., 50,53,56,57,58,59,60,64,65,75, 77,78,79,81 Geddie, J.L., 197, 236 Geesey, G.G.. 207,234 Genet, M.J., 17, 41 Geoghegan, M.G., 11,38 Gerba, C.P.,192, 221,239,243 Gerin, P.A., 10,17,38,41 Gennida, J.J., 186,235 Gemtse, J., 161, 170 Gersonde, K.. 75,83 Gevers, W., 87,125, 129 Geyl, D., 107, 108,127 Geywitz, J., 187,236 Gharieb, M.M., 229,236 Ghiorse, W.C., 183,236 Ghislain, M., 200,236 Gibson, D.T., 47,48,49,50,53,55,56.57,58, 59,60,64.66,77,78,79,80,82,83, 84,162,172 Gibson, J.F., 62, 63,76,79 Gierlich,A., 9,41 Gilboa, H., 209,237 Glenn, A.R., 182,215,236 Glund, K., 112,125 Gocayne, J.D., 211,236 Gocht, M., 90,91,93,125 Goddard, P.A., 230, 237 Goffeau, A., 200,236 Giihring, W., 120,125 Gold, L., 145, 168 Goldman, P., 135, 136, 137,168,169 Goncalves, M.L.S., 195,237 Good, N.E., 186,237 Gorby, Y.A., 183,239 Gordon, A.S., 214,223,237

250 Gordon, M.P., 162,173 Gtittgens, B., 23,24,37 Conschalk, G., 200,237 Covers, H., 227,228,233 Gowland, EC.,149, 150,169,173 Goyal, A.K., 50,79 Graham,L.A., 68,69,70,71,72,79 Graham,L.L., 202,203,237 Grandi, G., 119,130 Gravina. S.A., 200,242 Greenaway, S.D.. 145,154,169 Greene, A.C., 207,237 Greenwood, D.R., 5,8,11,42 Creer, C.W., 152,162,174 Griffith, G.S., 2,40 Griffiths, A.J., 183,187,226,236 Griot, R., 108,126 Croger, D.. 109, 128 Grossmann, A.D., 119,128 Grove, J.F., 100, 125 Grynkiewicz, G., 188,232 Gschwend, P.M., 159,169 Gucken, J.B., 207,240 Gueffroy,D.E., 186,237 Guerinot, M.L., 218.240 Guerlesquin, F., 59, 62,77 Guest, J.R., 92, 125,216,233 Gugel, K.H., 120,124 Guida, L., 187, 199,237 Guigliarelli, B., 76 Gull, K.. 10, 11,39 Gunsalus, LC., 65, 81 Gupla, A,, 226,237 Curbiel, K.J.,65, 66, 67, 69, 72, 79 Curies, R.P., 18, 31,37 Gutfinger, T., 68,80 Gutierrez, S.. 88,98,99, 125,128 Gutierrer-Corona. E, 214,234 Gutteridge, J.M.C., 49,80 Guzzo. A,, 224,237 Haak, B.. 50, 79 Haddock, J.D., 50,53,60,79 Haese, A,. 91, 100, 102, 103,126,128 Hagete, K., 120, 124 Hagenmaier, H., 120, 124 Hahlbrock, K., 90,127 Hahn, M., 9.41 Haigler, B.E., 49,53,56,58,59,60,78,79 Halachmi, D., 200,236 Hall. B.G., 146, 150. 174 Hall, D.M., 29, 43 Hall. D.O., 62, 63, 79

AUTHOR INDEX Hallas. L.E., 231,237 Halliwell, B., 49, SO Hamer, J.E., 5 , 8 , 28, 29.30, 38,42 Hammond, R.C., 165,169 Hansen. R.E., 63,82 Hanukoglu, I., 68,SO Han, M., 109, 110,126 Hara, O., 121,126 Harada, N., 219,235 Harashima, S., 67.81 Harayama, S., 49,50,55,60,61,66,73,80,81 Harcourt, A., 67,82 Hardman, D.J., 134, 136, 138, 142, 145, 149, 150, 154, 155, 158, 160, 164, 165, 166,167,168,169,172,173,174. 175 Harel-Bronstein, M., 193, 200,237 Harley, J.L., 3,38 Harmsen, M.C., 24,41 Harpel, M.R., 49,73,77,81 Harris, A., 60,81 Harris, C.I., 135, 171 Harrison, G.I., 229,237 Harrison, K., 139, 143,174 Harrison, EM., 216,233 Hartmann, A., 189,237 Hartmans. S., 160,165, 169,171 Hartnett, C., 50,55,61,66,73,81 Hanvood, V.J., 214,237 Hanvood-Sears, V., 223,237 Hasan,A.K.M.Q., 134, 144,169 Hase, T., 62.78 Hashim, R., 207,232 Hashimoto, T., 10, 38, 162, 171 Hassett, R., 221, 237 Hausinger, R.P., SO, 59,82 Hauska, G., 63,82 Hawkins, C . .216,233 48, 49, 80 Hayaishi, 0.. Hayashi, S., 149, 170 Hayashida, S., 53,60,61,73,78,83 Hazen, K.C., 32,39 Hazlett, R.D., 17, 39 Hearshen, D.O., 63, 65,68,72,78 Hederstedt, L., 218,242 Hegner, J., 24, 43 Heise,R., 200,237 Helinski, D.R., 90,125 Helm, D., 207,238 Hennecke, H., 145, 146,175 Henson, J.M., 207,240 Heppel, L.A., 165,169

AUTHOR INDEX

Hermann, H., 9,41 Hemero, M., 146,168 Hemnann,M., 91, 100, 102,103,126 Hess, W.M. 10,39,41 Higgins, D.G., 539, 148, 165,169 Hill, S.J., 198, 235 Hille, R., 65,68,72,78, 225,233 Hillemann, D., 121, 131 Hinata, M., 57,W Hinotozowa, K., 122,128 Hintz, W.E., 5, 18, 19,37 Hiratsuka, Y., 13, 19,42 Hirose, J.. 53.60, 61, 73,78.83 Hirsch, P., 135,169 Hobnt, J.A., 10, 11, 39 Hoch,H.C., 3,30,38,42 Hochkeppel, H.K., 102,124 Hodgson, J.E., 88,92,98,99,129 Hoffman, B.M., 65,66,67,69,72,77,79 Hoffman, EW., 136,169 Hoffmann, K., 106,126 Hofmann, A.. 108,126 Hofmann, H., 90,105, 107,127 Hohnadel, D., 189,239 Hol, W.G.J., 57,84 Holan, Z.R., 224,238 Holcombe, J.A., 193, 225.239 Hollander, J.A.. 209, 240 Holloway, P.J., 17, 29,38, 39 Holt. S.C., 11.39 Holt, T.G., 92, 121,128 Honneger, R., 3,10,33,34,39 Hooker, P.J., 183,239 Hope, S.J., 145, 150,169,174 Hopwood,D.A., 88,M, 91,94,126 Horgen, P.A., 5, 18, 19,31,37 Hori, K., 90,130 Horiuchi, T., 57,80 Home, R.W., 11,44 Hoskins, J.A., 98,130 Hou, C.T.. 165,173 Howard, R.J., 29,30,38,39 Huala, E., 146,169 Huang, P.M., 186,235 Huang, X.-H., 99,130 Hubbard, J.A.M., 189,216,222,238 Hubbell, J.A., 35,39 Hubbes, M., 5, 13, 18, 19,31.37,42 Hudlicky, T., 48,M Huet, J.-C., 9, 40 Hughes, M.N., 178, 180, 185, 186, 187, 188, 189,190, 191,194,195, 196, 198.

251 199,206,207,208,213,214,216, 222,224,224,233,234,235,237, 238,241 Hughes, S., 135,169 Huh, C., 17.39 Hurtubise, Y.. 50.80 Hutchinson, C.R., 93,94,%, 123,126 Huxley, M., 154, 166,169 Hyslop, E.K., 183.239 Igarashi, K., 200,240 Imai, S., 121,126,128 Imai, T., 165,170 Imamura, N., 122,128 Ingolia. T.D., 99,130 Ipsen, J.D., 6.8, 18, 31,41 Irie, S., 50, So Ishiama, T., 31.44 Ishikawa, T., 146, 176 Itoh, R.. 121,126 Iverson, W.P., 231,234 Iwasaki, H., 192,238 Izawa, S., 186,237 Jager, D., 161, 170 Jakeman, R.J.B., 209,239 Jansen, K.H., 197,233 Janssen, D.B., 134, 136, 138, 142,151, 152, 153, 154, 155, 160, 161, 162,170, 171,173,174,175 Jardim, W.F., 186,238 Jarrell, K.F., 186, 238 Jayatilake, G.S.,49.81 Jeng, R.,5, 13, 18, 19, 37.42 Jennings, D.H., 2.39 Jensen, H.L., 135,170 Jensen, S.E., 96,126 Jerina, D.M., 48,84 Jeronirnus-Stratingh, C.M., 162,173 Jessipow, S., 120.124 Ji, G., 212, 225, 226,234,235,238 Joannou, C.L., 53,56,57,58,59,60,64.67,79, 82,83,208,219,235,242 Johnson, A.C., 215,238 Johnson, D.B., 220,238 Johnson, K.A., 49.81 Jones, C.W., 218,238 Jones, D.H.A., 138, 143, 149,170 Jones, E.B.C., 29.39 Jones, R.P., 180,238 Jonssnn, B.H., 229,241 Joosten, M.H.A.J., 9, 43 Joshi, L., 11, 12, 13,37 Jung. G., 120,125

252 Kabuto, K., 48, 84 Kahan, B.D., 105,126 Kaida, N., 50.83 Kaldenhoff, R., 28,39 Kalk, C., 170 Kalk, K.H., 142, 161,168,175 KaIkkinen, N., 5, 8,40 Kallio, R.E., 50.79 Kamada, T.,219,235 Kamakura, T.,31,44 Kamp, R.M., 117, 118, 120,128,130 Kanda, M.,90,130 Kao, J.RY., 188, 189,232 Kaplan, J.H., 188,235,238 Kasai,N., 134, 153,154,155,158,170,174 Kasai, R.L., 10, 11,38, 153,155, 158, 159,174 Katsube, Y., 62.78 Katz, E., 111, 126 Katz, L.. 90,126 Kaufman, D.D., 135.171 Kawano, S., 219,235 Kawasaki, H., 138,142, 149,151,170,171 Kazemier, B., 142, 160,161, 162,170,175 Kazmierczak, P., 5,6,8,9, 13, 18, 19,32,38,45 Kearney, P.C., 135,171 Keister, D.B., 135, 136, 137, 169 Keith, L.H., 159,171 Keller, U., 91,93, 100, 103, 105, 109, 111, 112, 113, 114, 118, 122,129,130,131 Kelly, M., 135, 171 Kent, T.A., 63, 65, 68,72,78 Kershaw, M., 30.42 Kester, A., 9.42 Keuning, S., 160, 161,171 Khosla, C., 96. 123,130 Kikucbi, Y.,57.80 Kimbara, K., 57,80, 162,171 Kinghorn, J.R., 88,92,94,%, 124,127 Kingma, J., 161, 162,173,175 Kingsley, M.T., 215, 238 Kingsnorth, C.S., 23,24,37 215,238 Kinraide. T.B., Kirk, S.A., 31,37 Kirkness, E.F., 211,236 Kiyohara, H., 50.83 Kjelleberg, S., 4,41 Klages, U., 138, 141,171 Klapatch, T., 218,240 KIapcinska, B., 227,235,238 Klein, M.P.,67,69, 72.77 Kleinkauf, H., 86, 87,88,90,91,93,94,%, 97, 100,102, 103, 105,106,107, 109,

AUTHOR INDEX 112,124,126,127,128,129,130, 131 Klose, K.E., 146,172 Klube. K.D., 138,141,172 Kluge, B., 93, 117,118,127,128,129,130 Knaff, D.B., 67,69,72,77 Knobloch, K.-H., 90,127 Knogge, W., 9,41 Koana, T.,162,171 Kobal, V.M., 48,84 Kobayashi, H.. 200,240 Kobel, H., 107,108, 127,130 Koch, J.R., SO, 79 Kodama, T., 164. 165,170,176 Koga, H., 57.80 Kogut, M., 209,237 Kohata, K., 197,243 Kohler-Staub, D., 160, 165,171 Koiso, A., 149, 151,171 Kok, M.,49,80 Kok, R., 152,175 Kong, S.E., 188, 189,241 Konig, W.A., 120,124 Konings, W.N., 200,242 Kosaka, H., 219,235 Koshikawa, H., 138,172 Kosman, D.J., 179,221,222,237,239 Kot, E., 191,216,238 Kothe, E.M.,24,26,41,43 Kovacevic, S., 98,130 Kovaleva, V., 9.42 Kraas, E., 120,125 Kraepelin, G., 105.128 Kramar,0.. 231,243 Kratzschar, I., 88,89,92, 120,127 Krause, M.,88, 89,90,92, 120,127,129,130 Krauss, S., 138, 141, 171 Kreger, D.R.. 10,43 Krekel, D., 50,53.61.64, 80 Krengel, U., 109,126 Krianciunas. A., 67,69,72,73,77,81 Krook, J.H., 4, 10, 12, 13, 14, 20, 21, 44 Krouse, H.R., 229,237 Kruft, V., 88,91,129 Kues, U., 23,38,39 Kuila, D., 59,65,66,69,77,78,80 Kumada. Y., 121,126 Kunoh, H., 30.40 Kunst, L., 215,240 Kunugi, M.,197, 243 Kurahashi, K., 86.127 Kurane, R., 50.83

AUTHOR INDEX

Kurihara, T., 138,172 Kurkela, S.. 59.80 Kurotsu, T., 90,130 Kushner, D.J., 185, 241 Kustu, S.. 146, 171, 172 Kutz, S.M., 221,243 Kwart, L.D., 49,83 Kwon-Chug, K.J., 5, 12. 33,40 Kyte, J., 6, 7,39 La Roche, S.D., 165,171 Labbe,D.,50,83,152,162,174 Labbe, P., 221,239 Laberge, S., 9, 38 Labischinski, H., 207.238 LaBorde, A.L., 53.55.78 Laddaga, R.A., 180,224,226,227,238,239, 242 Laddison, K.J., 24, 43 LaHaie, E., 50,53,55,57,61,64,65,66,76,77 Laishley, E.J., 229,237 Laland, S., 86,124 Landa, E.R., 183,239 Landeen, L.K., 192,221,239,243 Landrum, G.A., 75,78 Langley, M.P., 196, 240 Laskin, A.J., 165,173 Latgb. J.-P., 5, 6, 8, 12, 33,42 Lau, P.C.K., 50, 83, 152, 162,174 Laurence, O.S., 230,236 Laurent. P., 5. 33.39 Lauter. E-R., 5, 11, 28, 29, 39,41 Lavanchy, D., 102,124 Lawen, A., 106,107, 108,127 Leadbetter, E.R., 11.39 Leadlay, I?, 94,128 Lee, B.T.O., 214,234 Lee, H., 227,229,230,239,242,243 Lee,H.-I., 8,40 Lee, K., 49,79 Lee, L., 48,49,80 Lee, R.S., 48.80 Leech, R.W., 185,234 Lehvaslaiho, H., 59,80 Leigh, J.A.. 139, 141, 171 82, Leisinger, T., 50, 53, 55,56, 57, 60,64,80, 136,137, 140, 142, 152, 160,164, 165,167,168,169,171,173,174 Leitner, E., 91, 106,130 Lernanski, C.L., 229,242,243 Lengeler, K.B., 24, 43 Lesuisse, E., 221, 239 Lewandowska, K.B., 189,216,222,238

2 53 Lewington, J., 150, 175 Lewis, M.R., 145 Ley, A., 186, 238 Ley, S.V.. 48,80 Librnan, J., 217,239 h e n , B.C., 135,171 Lin, C.-M., 179,221,222,239 Lindstedt, S., 73,77 Lingens, F., 48,49,50,53,55,56,57,59,60, 61,64,79,80,82, 138, 141, 171, 172, 173 Lion, R., 197,233 Lipmann, F., 86, 87, 92, 97,125,127,129 Lippard, S.J., 49,72,73,78,82 Lipscomb, J.D., 49, 73, 76,77,81 Liras, P., 98, 125 Litjens, M.J.J., 152, 176 Little, M., 136, 171 Liu, J.Q., 138, 172 Liu, T.-N., 50,53, 55,56,58,59,82, 83 Ljungdahl,P.O., a,76 Llobell, A., 5 3 9 Lloyd, D., 219,233 Locher, H.H., 50.53,56, 57,60,64,80 Loosli, H.R., 105,130 Lopez, J.L., 68,76 Lora, J.M., 5 3 9 Loros. J.J., 5, 11, 12, 28,37 Lovatt, D., 144, 145, 150, 174 Lovley, D.R., 183, 239 Low, B.J., 8,9, 38 Lomya, E., 90,127 Lubben, M., 218,242 Ludwig, M.L., 55.56, 77,78 Lugones, L.G., 3, $6, 12, 24, 25, 26,39, 44 Lundstrom, L., 16.38 Lurquin, P.F., 162, 173 Luyben, K.C.A.M., 160,169 Lynen, F., 88, 127 Lytton. S.D., 217,239 Macaskie, L.E., 209,227,239 MacCabe, A.P., 88,92,96,98,127 MacFarlane, J.K., 159,169 MacGregor, A.N., 135,172 Madduri, K., 98,130 Madgwick. J.C., 207,237 Madry, N., 91. 105, 128,131 Maeda, T., 138,171 Magee, L.A., 135,172 Magistrelli, C., 119, 130 Magnuson, R., 119,128 Maguire, R.J., 231, 243

254 Mahan, C.A., 193,225,239 Maier, R.J., 224, 236 Maier, W., 109, 128 Majidi, V., 193,225,239 Malkin, R., 67,69, 72,77 Mandel, M., 48, 80 Marahiel, M.A., 88, 89, 90,9492, 93, 117, 120, 123,125,127,129,130,131 Marchant, R., 10, 26,43 Markus, A., 50,53,55,56,57,60,61,64,80,82 Marliere, P., 92. 125 Martin, F., 5, 33,39,42 Martin, J.F., 88,98,99, 125 Maseles, EC., 50,79 Mason, J.R., 49,50,53.56,57,58,59,60,63, 64,67,68,71,72,73,80,81,82,83, 84,219,242 Mason, S.G., 17,39 Mather, M.W., 59,78 Matsubara, T., 192,238 Matsumoto, S., 62.78 Matsunaga, K., 57, SO Matsnshita, I., 138, 142, 149, 151,171 Matzanke, B.F., 209,216,239 McCabe, A., 94,96,124 McCarty,P.L., 159, 160,165,167,175 McCombie, W.R., 53,84 McCray, J.A., 188,239 McDonald, L., 230,239 McGinness, s., 220,238 McKee, J.A., 218,239 Mclean, R.J.C., 183,239 Means, J.C., 231,237 Mehta, N., 120, 130 Meienhofer, J., 111,128 Meister, A,, 97, 128 Mellor, E.J.C., 23, 24,37 Mendgen, K.,29.40 Menkhaus,M., 117,118,128 Menn, E-M., 162,172 Mergeay, M., 212,235 Merola, J.S., 48.80 Menick, M.J., 145,172 Messi, F., 160, 173 Messina, M.C., 221,243 Mester, B., 217,239 Metze, M., 146,168 Meyer, H.E., 68.81 Meyer, J.-M., 189,239 Mikesell, M.D., 160,172 Miller, C.G., 200, 242 Miller, J.R., 98, 130

AUTHOR INDEX Miller, M.J., 218, 239 Miller, W.G., 6, 8, 18, 31.41 Milne, G.W.A., 135, 136, 137,169 Mildowski, A.E., 183,239 Mms, W.B., 65.66.77 Minami, F.,149,170 Minamiura,N., 153, 155,158, 159,174 Minnick, A.A., 218.239 Mincda, Y., 160, 164, 165,170,176 Minor, W., 73.81 Minta, A,, 188,232 Miranda, R., 213,240 Misra, K.T., 228,239 Misra, T.K., 180,226,242 Mitra, R.S., 226,239 Miyoshi, K., 30,40, 142. 149,170 Mobley, H.L.T., 224,240 Modak, S.M., 229,240 Mondello, EJ., 50, 53, 60.61, 67,73,78, 81 Money, N.P., 30.39 Monod, M., 5,6,8, 12, 33,42 Montenegro, E., 88,99,125 Moodie, F.D.L., 53.81 Moore, R.T., 2,40 Moore, T., 146, 173 Morby, A.P., 226,237 Moritani, T., 50, 83 Morrice, N., 60,81 Morns, H.R., 91,129 Morsberger, E-M., 138, 141,172 Motosugi, K., 139, 144,172 Moukha, S.M., 2.4, 15.20.44 Mowll, J.L., 221, 236 Mozes, N., 17,41 Mueller, C.S., 196,240 Mueller, R.J., 5, 6, 8,9, 13, 18, 19, 32,38 Muisers, J.M., 9, 43 Mukohara, Y.,146.176 Mulder, G.H., 3,5, 14, 19,20.24,25,26,40,43 Mullen, M.D., 183,240 Muller, A., 63,82 Muller, G.I., 209, 216, 239 Muller, R., 50,56, 57, 64,78, 138, 141, 172, 173 Muller, V., 200,237 Muller-Rober, B., 28, 29, 41 Munck, E., 49,65,12,73,76,77,78,81 Muiioz. G.A., 13,40 Murakami, T., 121,126,128 Murata, M., 122, 128 Murdiyatmo, U., 138, 139, 142, 145, 149,167, 172 Murphy, G.L., 159,172

AUTHOR INDEX

Murphy, P., 5, 33 Myers, A., 119,128 Nadeau, P.,9.38 Nadim, L.M., 50,79 Nagaka, K., 121,126 Nagao, R., 99,130 Nagaoka, K., 121,126,128 Nagasawa, T.,153, 154, 155. 158,172 Nagata, Y.,57,M Nager, U., 99,128 Nakagawa, M., 67.81 Nakagawa, S., 165,170 Nakamura, H., 146,176 Nakamura, K., 213,240 Nakamura, T., 153, 154, 155, 172 Nakano, M.M., 91, 92, 117, 119, 120, 125, 128, 131 Nakari-Seala, T., 5,8,40 Nakatsu, C.H., 50, 53, 55.81 Nardi-Die, V., 138,172 Narro, M., 50,53,55,56,82 Nasse, B., 5, 33,42 Naumann. D., 207,238 Nehls, U., 5, 33,39 Neidle, EL., 49,50,55,61,66,73,81 Neilands, J.B., 181, 217, 240,241 Nelson, W.H., 207,240 Nengu, J.P., 160, 168 Nespoulous, C., 9,40 Netting, A.G., 29.40 Neumann, A.W.. 16,41 Newby, P.H., 187,240 Newman, K.A., 159,169 Nicholls, P.D., 207,240 Nicholson, R.. 29. 30, 31, 40 Nies, A., 226.240 Nies, D.H., 226, 240 Nihira, T.,91, 105,131 Nilssorn, U., 16, 38 Nishida, H., 99, 130 Nishino, H., 138,172 Nitschke, W., 63,82 Niven, D.E., 207,240 N0bar.A.M.. 186,198, 199,206,233,241 Nomura, Y., 67,81 Noordmans, J., 16.40 Nordin, J.H., 18,37 Nordlund, P., 73,82 Norlund, I., 162, 172 North, A.K., 146,171,172 Novick, R.P., 228, 240 Novo-Gradac, K.J.,186,233

255 Nozawa, Y,10, 11,38 Nussbaumer, B., 121,131 Nuyts, G., 212, 235 Nyandoroh, H.. 164 O’Callaghan, N.M., 97,124 Oelrichs, P.B., 105,128 Ogawa, N., 67,81 Ogawa, S., 209,241 Ogino, T., 209,240 O’Halloran, T.V.,73,75,77,229,240,243 Ohlendorf, D.H., 73,81 Ohnishi. T.,68.72.77, 82 Ohshima, A., 31,44 Ohyama, T., 200,240 Oishi, M.,162, 171 Okamura, O., 138.172 Okuda, S., 99,130 Okumura, Y,111,128 Olami, Y., 193,200,237 Olson, E X , 67,77 Olson, G.J., 231, 234 Omon, T., 159,160,164,172.173,176 Omura, S., 99, 122,128,130 Omston, L.N., 50, 55,61,66,73,81 Orville, A.M., 49,73,77, 81 Osbom, R.W., 9,42 Oshima, Y., 67,81 Osslund, T.D., 67.82 0 t h H., 108.126 Otto, A., 93, 117,129 Otto, M.K., 138, 141, 172 Otwinowslu, Z., 73.81 Ozaki, H., 138,172 Pacheo, S.V., 213,240 Padan, E., 193,200,237 Painter, E.P., 229,240 Palacz, Z., 117, 130 Palissa, H., 88,92, 96.97, 124, 127, 129 Palva, E.T., 59,M Panaccione, D.G., 110,129 Pang, C.-P., 49.81 Panico, M., 91, 129 Paris, S., 5,6, 8, 12, 33, 42 Parker, W., 227,243 Parsons, J.R., 134,168 Parta, M., 5,8, 12,33,40 Patel, R.N., 165,173 Patil, D., 53,59,75,79 Patil, S.S., 90,125 Paul, E., 91, 105,131 Pavela-Vrancic, M., 90,93,128 Pearce, J.M., 183,239

256 Pearson, H.W., 186.238 Peters, H., 91, 105, 128,131 Pelzer, S., 121, 131 Pember, S.O., 49.81 Pendland, J.C., 29,37 Penfold, W.J.. 134.173 Penner-Hahn, J.E., 66,67,83,229,242 Penlenga. M., 152, 162,173, 174 Penttila, M., 5, 8,40 Perego, M., 120,125 Perkins, E.J., 162, 173 Pernollet, J.-C., 9, 40 Perry, J.J., 159, 172 Peter, M., 34.39 Peters, J.A., 135,171 Peters, R.A., 134,173 Pettersson, A., 215,240 Pfefferkom, B., 68.81 Pfeifer, E., 88, 90, 92, 93,94,%, 124,127, 128 Pfister, R.M., 11,41 Pfleger, K., 49,84 Phillips, E.J.P., 183, 239 Phillips, S.E., 225, 226,240 Phung, L.T., 180,242 Pieper, R., 102, 103,126,128 Pinner, E., 193, 200, 237 Pinto-DaSilva, P., 5, 12, 33,40 Pintor-Toro, J.A., 5,39 Pioda, L.A.R., 99,131 Pirt, S.J., 179, 188, 189,222, 224, 240 Plathler, P.A., 99, 128 Plessner, O., 218, 240 PKard, J.-A., 110, 129 Ponelle, M., 105, 130 Poole. R.K., 178, 180, 185, 186, 187, 188, 189, 190, 191, 195, 196, 198,199,206, 207, 209,213,214,216,218,219, 220,222,224,226,233,234,235, 237,238,241 Poolnian, B., 200,242 Popkin, T.J., 201,202,203, 234 Pople, M.,100, 125 Porter, R., 29.38 Porterfield, V.A., 165,169 Poulos, T.L., 49, 81 Powlowski, J., 50,80 Pries, F., 134, 136, 142, 151, 160, 161, 162,170, 173,175 Privalle, C.T., 188, 189, 241 F’usztai-Carey, M., 5, 6, 9, 18, 44 Que, I,., 73. 77 Queener, S.W., 99,130

AUTHOR INDEX Que, L.Jr., 49,77,81 Raag, R., 49,81 Raibaud, A., 92, 121,128 Raikhel, N.V., 8, 9,40, 42 Ralston, D.M., 229.240 Ramamoorthy, S., 185,241 Ramelow, G.J., 196,240 Ramirez, J.L., 225,226, 234 Raper, C.A., 22,24,40,43 Raper, J.R., 2, 19,22,40 Raudaskoski, M., 14,24,26,40,44 Rayner, A.D.M., 2,40 Rayner, M.H., 209,241 Read, D.J., 3,40 Reanney, D.C., 149,173 Redgewell, R.J., 11, 17,37 Rees, S.B., 9,42 Regensburg, B.A., 10,43 Regev, R., 209,237 Rekik, M., 50.55,60,61,46,73,80,81 Remsen, C.C., 10,39,41 Resnick, S.M., 49.79 Reutlinger, M., 195, 237 Reuvekamp, P.T.W., 153, 155,175 Riach, M.B.R.. 88,92,96,98,127 Ricci, M., 28, 29,41 Richards, W.C., 5,6, 8,9, 18, 31,37,40,42,44 Richardson. J.S., 8.9.38 Riedel, A., 63,82 Riederer, B., 111,122 Rieske, J.S., 63,82 Rikkerink, E.H.A., 8, 9.42 Rittman, B E , 159, 167 Roach, D.H., 30.39 Roberts, D.W., 5.8, 11, 12, 13, 30,37,41 Roberts, G.A., 94,128 Robinson, N.J., 226,237 Robson, C.D., 2,43 Rodriguez, E, 120, 125 Rodriguez-Romero,A., 8,37 Roe, A.L., 73,77 Rohe, M., 9,41 Rolland, C.. 5, 6, 8, 12, 33,42 Rollins, M.J., 96,126 Romanov, V., SO, 59.82 Ronneau, C., 197,233 Roomans, G.M., 215,240 Rosahl, S., 9 , 4 1 Rosche, B., 49,53,60,82 Rosen, B.P., 180, 183, 191,236,241,242 Rosenberg, G., 50,53,56,57, 59,&1,82 Rosenberg, H., 217,241

AUTHOR INDEX

Rosenberg, M., 4.41 Rosenzweig, A.C., 73,82 Roskoski, Jr., R., 87,129 Rosner, J.L., 224,241 Rotenberg, Y., 16,41 Roth, C.. 228, 240 Roth, L.E., 215,241 Rotilio, G., 223,235 Rouch, D.A., 214,234 Roucoux, P., 197,233 Rouden, J., 48,W Rouxhet, P.G., 10, 17,38,41 Roy, C.. 5, 6, 9, 18.44 Rozeboom, H.J., 142, 161,173,175 Ruardy, T.G., 16,29,44 Ruf, H.H., 53,55,56,57,60,82 Ruiters, M.H.J., 14, 20,24,25, 26,41,44 Rulong, S., 5, 12,33,40 Rundgren, M., 73.77 Runswick, M.J., 90, 130 Russell, N.J., 209, 237 Russo, P.S., 6, 18, 31,41 Russo, V.E.A., 5 , s . 11, 28, 29,39,41 Rutherford, A.W., 63,82 Saboowalla, F., 53, 59,75,79 Sadler, P.J., 185, 187,208,209, 234 Sahhan, L., 229,241 Said, 2, 187, 191,199,237 Saigo, T., 192,238 Saito, Y, 90,130 Salerno, J.C., 72,82 Sallis, P.J., 138, 142, 154, 158, 160, 164, 165, 166,167,173,175 Salnikow, J., 117,127 San Martin, K.,13,40 Sanglier, J.-J., 108, 127 Saraste, M., 90. 130 Sargent, J.S., 68,72,79 Sariaslani, F.S., 165,169 Sassen, M.M.A., 10,39,41 Satoh, A., 121,126, 128 Satoh, E., 121, 126 Satoh, S., 50, 83 Sauher, K., 50,53, 56,57,59,64,82 Sauer, K., 67,69,72,77 Saulnier, M., 186,238 Saunders, R., 67,82 92,125 Saurin, W., Savvaidis, I., 195, 196, 198, 199,206, 222,241 Sawada. T., 50,83 Schaap, P.J., 5,38 Schafer, H.J., 90,128

257 Schallehn, G., 207,238 Scheer, J.H.J., 24,26,41 Schellekens, G.A., 9,41 Scheper, A., 152,154,160, 161,162,170 Scherr, D.J., 154 Schimmel, P., 93, 129 Schnder, R., 105,129 SchISfli, H.R., 50, 55,56, 64.82 Schlegel, H.G.. 225,226,241,242 Schlumbohm, W., 88,91, 112,114.125,126, 129 Schmidhauser, T.J.,28,29,41 Schmidt, B.,9,41 Schmidt, F.R.J., 139, 149, 167 Schmidt, T., 226,241 Schmuckle, A., 160,169 Schneider, A., 121,129 Schneider, B., 138, 141,173 Schneider-Scherzer, E., 106,126,130 Schocken, M.J., 49, 79 Schofield, C.J., 97. 98, 117, 124, 129 Scholtmeyer, K., 5, 6, 12.26.39 Scholtz, R., 160, 164,173 Schoonover. J.R.,66.69.80 Schorgendorfer, K., 91,106,130 Schreyer, M., 107,125 Schrljder, W., 90, 103,128 Schubert. M.. 91, 100, 102, 103,126 Schuhmann, A.. 197,233 Schuldiner, S., 193,200,237 Schulze, R., 221,243 Schuren, F.H.J., 3,4, 5, 12, 13, 14, 15, 16, 19, 20, 22,24,30. 32, 35, 41, 43, 44 Schuurs, T.A., 3,5, 20.24, 25,26,43,44 Schwecke, T., 94,96,97,124,129 Schweizer, D., 53, 55.56, 57,60,82 Schwyn, B.. 217,241 Scott, R.L, 219, 220, 241 Scott-Craig, J.S., 110, 129 Sebald, W., 63, 67.78 see^, M.. 53, 55, 56, 57,60, 82 Seiyama, A., 219,235 Sekiya, M., 10, 11,38 Selitrennikoff, C.P., 12, 28,41 SeljQ, E 161, 175 Senior, E., 135, 144, 145,150,173,174 Serdar, C.M., 50,53,58,59,83 Service, R.F., 34, 41 Sewall, T.C. 4,5,8, 11, 12, 13,27, 28.42 Shanxr, A., 217,239 Sharp, P.M., 5.39 Sheets, T.J., 135, 171

258

AUTHOR INDEX

Sheldrake, G.N., 48.82 Shen, G.-J., 165, 170 Sherburne, R., 230,242 Shergill, J.K., 67, 82 Sherman, D.H., 88,90,91,94,126 Shiaris, M.P., 50. 84 Shiau, C.Y. 97,98, 117,129 Shiga, T., 219,235 Shin, M.K., 229,240 Shingler, V., 146, 162,168, 172, 173 Shotyk, W., 183,235 Shrift, A,, 229,234 Shulman, R.G., 209,240,241 Shuttleworth, K.L., 187, 194,241.242 Sietsma, J.H., 2, 3,4, 9, 15, 20.24, 25, 41,43,

44 Sigg, L., 195, 237 Sillerud, L.O., 229, 243 Silver, S., 178, 179, 180,183,210,212, 213, 224,225,226,227,234,235,238, 239,240,241,242,243 Simon, M.J., 67.82 Simon, W., 99,131 Simon-Lavoine, N., 99,129 Singh, R.M.M., 186,237 Sivaraja,M., 65,66,67,69, 72,79 Skamoulis, A.J., 209,239 Skatrud, P.L., 98,99,130 Skinner, A.J., 139, 141, 166,167, 171 Slater, J.A., 18,41 Slater, J.H., 134, 135, 136, 142, 144, 145, 146, 147,149, 150, 151, 154, 155,160, 166,167,169,173,174,175,176 Slavin, W., 193,242 Slawson, R.M., 227,229,230,242 Sleytr, U.B.. 10, 38 Smalley, E., 18.31.37 Smith, D.J., 88, 92, 98, 99, 129 Smith, J.M., 139, 143,174 Smith, J.M.A., 216, 233 Smth, S.E., 3,38 Smucker, R.A., 11,41 Snavely. M.D., 200,242 Sobey, W.J., 98, 117. 129 Soda, K., 134, 138. 139, 144,169,172 Sokolovsky, V.Y, 28,29,41 SB11,D., 93,129 Somlyo, A.P., 188,238 Sonnenberg, A.S.M., 5.38 Southam, G., 183,202, 203,236 Spain, J.C., 49, 50, 55, 58. 76, 82 Specht, C.A., 24,43

Speelmans, G., 200,242 Spence, J.T., 217, 235 Sperry, J.F., 207, 240 Springer, J., 3, 13, 14, 19, 20, 24,26,38,41, 43,44 StLeger, R.J., 5 . 8 , 11, 12, 13,30,37,41 Stacey. G.. 215.241 Stachelhaus, T., 88, 92,93, 123, 129 Stadler, P.A., 108, 126, 129 Stahl, li., 9, 44 Stanley, D.C., 67,77 Staples, R.C., 5, 8, 30,41 Starcdub, M.E., 230,242 Staub, D., 165,167 Staunton, J., 94, 128 Steczkn, J., 73.81 Stedman, K.M., 146,172 Stefanac, Z., 99,131 Stein, T., 88,91, 93, 117, 129 Sterk,P.,9,41 Sternfeld, F., 48,80 Stevenson, K.J., 5, 6, 9, 18, 41.44 Stewart, A., 221,236 Sticklen, M.B., 9, 18,33,37,42 Stindl, A,, 113, 114, 116, 117, 118,129,130 Stirling, D.I., 165, 168, 174 Stijffler-Meilicke,M., 109, 110, 126 Stookey, L.L., 190,221,242 Stoppel, R.-D., 225,242 Stotzky, G., 182, 183, 185,186, 187,233,235 Stratton, G.W., 226, 243 Straus, N.A., 50,81 Stringer, M.A., 4, 5,6, 8, 11, 12, 13, 27,28,42 Strotmann, U.J., 152, 174 Stubbs, B.M., 154,167 Stucki, G.R., 152, 165, 174 Stumm, W., 195,237 Stuttard, C., 98, 130 Subramani, S., 90,125 Subramanian, V., 50,53,55,56,58,59,64,82,

83 Suemori, A., 50, 83 Suen, W.-C., 53, 60,64.67, 82,83 SUggS, S., 67.82 Summers, A.O., 179,242 sun,Y.,5,45 Surerus, K.K., 49.77 Surewicz. W.K., 5,6, 9, 18,44 Suler, F.,164, 165,171,173 Sutherland, I.W., 197,236 Suyama, A., 53,60,61,73,78 Suzuki,T., 134,153,154,155,158,159,170,174

AUTHOR INDEX

Svensson, B., 218,242 Svircev, A.M., 13, 19.42 Sylvestre, M., 50,W Tagu, D., 5,53,39,42 Taira,K., 53.83 Takada, H., 134,144,169 Takagi, M., SO, 57, 80, 162, 171 Takai, S., 5,6,8, 9, 13, 18, 19,37,41,42,44 Takamatsu. T., 197,243 Takano, E., 121, 126 Takao, M., 149, 151,171 Takigawa, H., 165,170 Takuziwa, N., SO, 83 Talbot, N.J.. 5, 8, 28, 29, 30,42 Tan, H.-M., 53, 57,58, 59, 60,61, 67,83, 219, 242 Tanabe, O., 135,175 Tanaka, H., 99,130 Tang, H.-Y., 53, 57, 59,60, 67,83 Tardif, G., 152, 162,174 Tam, G.E., 65,68,72,78 Taylor, D.E., 230,242, 243 Taylor, F., 165,169 225, 226,240 Taylor, M.L., Taylor, S., 48,W Taylor, S.C.. 134, 143,174 Teen, T.H., 5 9 , 8 Tegli, S., 31,37 Telliard, W.A., 159,171 Temple, R., 18, 31,37 Templeton, M.D., 5, 8.9, 11, 33, 42 Te-Ning, L., 50,64,82 Terashima, Y , 138, 172 Terhune, B.T., 30,42 Terpstra, P., 57,78,84, 142, 160, 161,170 Terns, F.R.G., 9, 42 Thau, N., 5 , 6 , 8 , 12,33,42 Thomas, A.W., 139, 145, 146, 147, 149, 150, 174,175 Thomas, C.M., 138, 143, 149,170 Thomas, D., 36,42 Thompson, C.J., 92, 121,126,128 Thompson, R.L., 196,240 Thompson, S.A.J., 23,24,37 Thomson, J.C., 139, 142,167 Thony, B., 145,146,175 Thornley, J.H.M., 62, 63, 79 Timberlake, W.E., 4,5, 6, 8, 11, 12, 13, 27, 28, 36,38,42 Timmis, K.N., 60,80, 146, 168 Ting, H.-H., 49, 81 Tisa. L.S., 180, 189,242

259 Tizard, R., 92, 121, 128 Tobin, M.B., 98,130 Tognoni, A., 119,130 Tohika, K., 90,130 Tomei, EA., 229,242, 243 Tomizuka, N., 50,83 Tomoda, H., 99,130 Tone, H., 142,149,170 Tonomura, K., 135, 138, 142, 149, 151,170, 171,175 Topping, A.W., 139, 144, 145,146, 147, 149, 150, 151, 174 Torigce, S., 50,83 Tomabene, T.G.. 228,243 Torok, D.S., 49.79 Torrekens, S.,9,42 Toyama, T.. 138, 171 Traber, R., 105, 107, 108, 127,130 Tramper, J., 160, 169 Trautwein, A.-X., 75,77,209,216,239 Trenlham, D.R., 188,239 Trevors, J.T., 226,227,228,229,230,234,236, 239,242. 243 Trinci, A.P.J., 2. 43 Tronchin, G., 5, 6, 8, 12, 33, 42 Troughton, J.H., 29,43 True, A.E., 65,66,67,69,72,79 Trumpower, B.L., 63,68,69,70,71,72,76,79, 83 Tryhorn, S.E., 165,169 Tsang, H.-Y., 66, 67, 83, 229, 243 Tsang, J.S.H., 138,142, 149,172,175 Tscherter, H., 107,125 Tshisuaka, B., 49, 53, 60, 82 Tsibns, J.C.M., 65,81 Tsien, R.Y., 188, 189, 232 Tsoi, C.J., 96, 123, 130 Tsuda, K., 138, 142, 149, 151,171 Tsujimura, K., 134. 154, 155, 158,170 Tsukihara, T., 62.78 Turgay, K., 88,89,90,130 Turner. G., 88.92, 98,99,129 nuner, J.S., 226,237 Twilfer, H., 75,83 Ugurbil, K., 209,241 Ulbrich, N., 9,44 Ullrich, C., 88,91, 117, 118,128,129,130 Unkles, S.E., 88.92,96,127 Unoum, K., 134, 154, 155, 158,170 Unz, R.F., 187, 194, 241,242 Vaillancourt, L.J., 24, 43 Valent, B., 29, 30, 38

260 van Alfen, N.K., 5,6, 8,9, 13, 18, 19, 3 2 , s . 45 van den Ackerveken, G.F.J.M., 9’43 van den Mjngaard, A.J., 153, 154, 155, 160, 162,175 van der Karnp, K.W.H.J., 160, 162,175 van der Lende, T.R., 19,24,41 van der Meer, J.R., 162,175 van der Mei, H.C., 16, 17, 29,30,43, 44 van der Plceg, J., 134, 136, 138, 142, 151, 160, 161,162,170,175 van der Valk, P., 26,43 van der Vegt, W., 16, 17,43 van der Vmn, L.H.M., 88,98,125 van der Waarde, J.J., 152, 175 Van Dyke, M.I., 227, 229,230,239, 242,243 van Griensven, L.J.L.D., 5.38 van Hall, G., 138, 175 Van Kammen, A,, 9,41 Van Kan, J.A.L., 9.43 Van Leuven, F., 9,42 van Liempt, H., 88,!30,92,94, %,97,124,127, 128,129,130 Van Loon, J.C., 198,243 van Pelt, A.W.J., 17,37 van Pouderoyen, G., 162,173 van Sinderen, D., 120,125 van Solingen, P., 88, 98, 125 van Wetter, M.-A., 3, 5, 19, 20, 23, 24,25,26, 37,43,44 Vanderleyden, J., 9,42 Vano, K., 162,171 VanRiet, J., 227, 228,233 Vartivarian, S., 217,233 Vasquez, C., 230,235 Vater, J., 88. 91, 111, 117, 118, 120, 128, 129, 130 Venema, G., 120,125 Verbakel, H.M., 9,43 Verschueren, K.H.G., 160, 161,175 Vbzina, L.-P., 9, 38 Viitanen, H., 26,40 Villafranca, J.J., 49,81 Villalon, D.K., 5,6, 8, 9, 13, 18, 19, 32,38,45 Mning, L.C., 98, 130 Visser, I., 5, 38 Vogel, T.M.. 135,175 Volcani, B.E., 227,233 Volesky, B., 224,238 Vollenbroich, D., 117, 118, 120, 128,130 Volpe, D., 13,40 von Dohren, H., 86,88,90,93,94,%, 97,124, 126,127,128,129,130

AUTHOR INDEX von Ostrowsla, T.,103,126 von Wartburg, A., 105,130 von Wettstein-Knowles, P., 29, 40 Vnend, G., 57,78 Wackett, L.P., 49,50,53,58,59,79,83 Wakley, G.E.. 30.42 Walderhaug, M., 178,210,242 Walker, J.E., 90.130 Walker, J.R.L., 135,136 Walter, E.G., 230,242 Walton, A.P., 198, 235 Walton, J.D., 99, 110, 129, 130 Wang, Y., 50,83 Ward, J.M., 88, 92, 98, 99, 129 Ward, R.W., 198,235 Waring, A.J., 219,241 Watabe. K., 146,176 Watanabe, I., 153.1.54, 155, 158,172 Watanabe, M.,197,219,235,243 Watterson, J.R., 183,243 Weber, G., 91, 106.130 Weber, P.C., 73, 81 Weckermann, R., 88,130 Wehrli, E., 11, 44 Weigel, B.J., 99, 130 Weightman, A.J., 136, 144, 145, 146, 147, 149, 150, 155,167,174,175, 176 Weijers, C.A.G.M., 152, 176 Weiss, D.S., 146, 171 Weiss, M.A., 50,55, 64,82 Welin, S., 16.38 Welz, B., 193,243 Wende, P., 49,84 Wendland, J., 24.43 Wenger, R.M., 106, 107,125,127 Wessels, J.G.H., 2, 3.4, 5,6,7, 9, 10. 11, 12, 13, 14, 15, 19, 20, 21, 22, 23,24,25, 26, 28, 29, 30, 32, 33, 34, 3.5,37,38, 39,41,42,43,44 West, J.M., 183,239 Westlake. D.W.S., 96,126 Whatley, F.R., 62,63,79 White, C., 180,221,236 White, D.C., 207,240 White, O., 211,236 White, R.L., 49,81 Whittaker, J.W., 75, 78 Whitton, B.A., 226,237 Witty, P., 146, 168 Widom, J., 49,81 Wiebe, M.G., 2,43 Wierenga, R.K., 57,84

AUTHOR INDEX

Wildermuth, H., 11,44 Wilkinson, S.C., 229,236 Williams, J.M., 216,225,233 Williams, J.W., 225, 243 Williams, J., 225, 233 Williams, P.A., 136,171 Williams, S.J., 11, 44 Willis, A.C., 97,124 Wilson, B.H., 159,176 Wilson, J.T., 159, 176 Winget, G.D., 186,237 Winter, W., 186,237 Wipf, H.K., 99,131 Witholt, B., 57.78, 142, 152, 153, 154, 155, 160,161,162,170,171 Withann-Liebold,B., 88,91,93, 117,129 Wnendt, S., 9.44 Wohlleben, W., 121,131 Wolf, D.C., 183,240 Wolfe, S., 94,%, 124 Wong, A., 96,126 Wong, P.T.S., 198,231, 235, 243 Wood, K.V., 90,125 Wood, M., 215,238 Woodland, M.I?, 53,81 Woodruff, W.H., 66.69.80 Wootton,J., 146, 168 Wosten, H.A.B., 2, 3,4, 5, 6, 9, 10, 11, 12, 13. 14, 15, 16,24, 25,26,28, 29.30.32. 34,35,38,43,44 Wright, C.S., 8, 9,38 Wright, J.G., 229,240,243 Wu-Yuan, C.D., 10.38 Wyndham, R.C., 50,53,55,81 Xiao, J.-2.. 31,44 Yaguchi, M., 5,6,9, 18,44 Yahara, H., 149,151,170,171 Yahya, M.T.,192, 221,239,243 Yamada, H., 153,154, 155. 158,172 Yamaguchi, E., 57,80

261 Yamaguchi, I., 31,44 Yamaguchi, M., 50,53,56,57,60,64,84 Yamamoto, R., 153, 155, 158,159,174 Yamane, K., 92, 120,125 Yamaoka, T, 135,175 Yanase, N., 149, 151,170 Yang, Y., 50,84 Yano, K., 50,57,80 Yanovsky, C., 5, 11,28,39 Yeh, W.-K., 50,53,55,56,58,59,64,82,83 Yli-Mattila, T., 14,26,44 Yokota, T., 160, 164,176 Yokoyama, T., 10,11,38 Yorifuji, T.. 50.80 Yoshida, T., 59,63,65,68,72,78 Yoshihara, H., 219, 235 Yost, K.J., 48.80 Young, J.C., 196,240 Young, K.D., 67,77 Yu, C.A., 67,69,72,77 Yu, F., 153, 154, 155, 158,172 Yu, L., 67, 69, 72,77 Zahner, H., 120,124,125 Za~ki. T.. 53,60,61,73,78 Zalacain, M., 91, 92,93,128 Zamanian, M., 53,60,64,84 Zaugg, W.S., 63.82 Zehnder, A.J.B., 162,175 Zentmeyer, G.A., 18,44 Zhang, J., 94, %, 124 Zhang, L., 5.6.8.9, 13. 18,19,32,38,45 Ziffer, H., 48.84 Zocco, T.G., 229, 242,243 Zocher, R., 91, 100, 101,102.103,105,106, 107,109, 112,114,121,124,126, 127,128,131 Zuber, P., 91,92,117. 119,120,125,12%,130, 131 Zylstra. G.J., 49, 50, 53,57, 60,66,77,79, 82, 84, 162,172

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Subject Index Figure and table references are shown in italic

abaA A. nidulam gene, in conid~ogenesis,27 ABHl hydrophobin, 4 , 5 , 6 in fruit bodies, 26-7 actinomycin synthesis, 91, 111-14 synthetase I, 112-13 synthetase II, 113-14 in amino acid epimerization, 116-17 reaction priming on, 114-16 synthetase 111, 113-14 in peptide bond formation, 117 synthetases in cell-free synthesis, 114, 115 ACVS see S-(L-aIpha-aminoadipyl)cysteinyl-D-valine synthetase (ACVS) acyl peptide lactones, 111 synthesis, actinomycin as model, 111-12 see also actinomycin synthesis acyl transfer, by peptide synthetases, 92 S-adenosyl-L-homocysteine, and enniatin synthetase, 100-1 S-adenosyl-L-methionine, and enniatin synthetase, 100 AES see atomic emission spectroscopy Agancus bisporus. hydrophobins in fruit bodies, 26-7 see also ABHl hydrophobin agglutinins, and hydrophobins, 8-9 Agrobacteriurn turnefaciens, haloalcohol dehalogenases, 154 Alcaligenes, haloalcohol dehalogenases, 156-7 Alcaligenes eutmphus. cadnuum resistance, 226 alcohol dehalogenation, 151-9 dehalogenases, 152-9 classes, 154-5, 158 pathway, 152-3 profiles, 153-4 properties, 156-7 enzymatic oxidative dechlorination, 159

halogenated alkanoic acid formation, 152 algae, lichen symbiosis, hydrophobins in, 3 3 4 alkane dehalogenation, 1 5 9 4 5 class 3B dehalogenases, 164 catabolism pathway, 161 evolutionary relationships, 162, 164 from Xanthobacfer aurotrophicus, 162,163 class 3R dehalogenases, 1W cofactor-dependent dehalogenases, 165 discovery, 159 diversity of mechanisms, 1M) methanogenic bacteria in, 1 5 9 6 0 oxygenase-type dehalogenases, 164-5 alkanoic acid dehalogenation, 135-51 dehalogenases characteristics, 1 3 5 6 class ID, 139. 140, 1 4 3 4 class lL, 136-7, 138,140, 141-3 class 21, 139, 144-5 class 2R, 139, 1 4 5 4 classification, 136,138-9 genetic organization, 146,147, 149-51 synthesis regulation, 146-9 hydrolytic mechanism, 135 aluminium, toxicity, 182, 214-16 &(L-a-aminoadipyl)-cysteinyI-D-valine(ACV), structure, 96 6-(I.-a-aminoadipyl)-cysteiIiyl-D-valine synthetase (ACVS), 96-9 activation sites, 97 epimerization in synthetic action, 97-8 genes, 98-9 Anabaena qlindrico, aluminium accumulation, 215 Ancylobacter aqwficus, dehalogenase, 162 Ankistmdesmwfalcatur, tin accumulation, 23 I anodic stripping voltammetry (ASV), 194-5 antimony, 182 arsenic, 182

264 arsenic (continued) bacterial resistance, 225-6 Arthrobacter dehalogenases haloalcohol, 154, 155, 156-7 haloalkane, 164 ascomycetes, hyphal structure, 2 Aspergillus hydrophobins from, 13 rodlet layer, 11, 12, 13 Aspergillus nidulans ACV synthase from, 96,97 conidiogenesis, molecular genetics, 27-8 hydrophobin gene, 4 atomic absorption spectroscopy, 193 with chromatography, 198 atomic emission spectroscopy, 193 atomic fluorescence spectrometry, 194 axisymmetric drop shape analysis, for hydrophobins, 16 Azotobacter, lead resistance, 228 Bacillus germanium accumulation, 227 mercury resistance, 229 blotting analysis, 213 bacteriofemtin, 216 basi&omycetes fruit bodies, 3 hyphal structure, 2

Beauveria, 105 beauvericin, 105 benzene dioxygenase, 61 ferredoxin, 5 8 , 5 9 4 non-haem iron, 75 reductase, 57 spectroscopic analysis, 65 benzoate dioxygenases, 55,56,57 iron site, 74-5 beta-lactam antibiotics, hydrolysis by metals, 222 bialaphos, 120-2 biocompatibility. and hydrophobins, 35 biosensors, 21 1-12 hydrophobins in, 35 biphenyl dioxygenases. substrate specificity, 61 BLAST, 21 1 blotting hybridization techniques, 212-13 Bradyrhizobium japonicum, nickel metabolism, 224 brlA A. nidulans gene, in conidiogenesis, 27 cadrmum

INDEX bacterial resistance, 2 2 6 7 fungal toxicity, and pH, 187 precipitation, 209 uptake, zinc competition, 224 caesium, radioactive, from Chernobyl, 183 calcium analysis, 191 cellular concentration, and photoreactive ligands, 187-8 as essential metal, 180 Cephalosporium acremonium, ACVS from, 97 cerato-ulmin, 4,6, 13 disulphide bridges, 9 function, 19 hydropathy pattern, 6.7 phytotoxic mechanism, 3 1-2 sequence determination, 18 surface activity, 18 Chelex- 100 for copper metabolism study, 222 for metal removal from m d u m , 189 Chernobyl fall-out, 183 R-3-chloro- 1,2-propandiol, enzymatic dechlonnation, 159 chlorocatechol dioxygenase, 73 iron site, 75 I-chlorohexane dehalogenase, 164 4-chlorophenyl acetate benzoate dioxygenase, 55,56,57 chromatography, of metals, 198 ion chromatography, 197-8 chrome azurol (CAS), in siderophore assay, 217-18 Chviceps purpurea, 108 Cochliobolus carboneum, HC-toxin synthetase from, 110 CoHl hydrophobin, 4.5 in aerial hypha formation, 21-2 conidia, hydrophobins in formation, 27-9 copper binding proteins, 222-3 cellular uptake, 221-2 microbial corrosion. 207-8 in oxygenase catalysis, 49 resistant bacteria, 214 speciation, and cell growth, 186 transport, 181 copper-zinc superoxide dismutase, 223 Corynebacterium dehalogenases haloalcohol, 154, 155, 156-7, 158 haloalkane, 164 cryofrxation, 202-3

265

INDEX cryparin, 4,6, 13 function, 19 hydropathy pattern, 6 , 7 lectin-like activity, 9 cyclosporins, 105 structure, cyclosporin A, 106 synthesis, 91, 105, 106-7 synthetase, 105-6 molecular structure, 107 substrate specificity, 107 see also SDZ 214-103 Cylindrotrichumoligospennum. 107 Dalapon, 135 Pseudomonasputida growing on, 144 defensins, cyskine residues, 9 dehalogenation, microbial of alcohols see alcohol &halogenation of alkanes see alkane dehalogenation of alkanoic acid see alkanoic acid dehalogenation depsipeptide formation, enniatin synthetase in, 103,104 Desuljovibria desulfuricans,selenium resistance, 229 dewA A. nidulans gene, in conidiogenesis,27-8 dibenzofuran dioxygenase ferredoxin, 59 reductases, 57-8 1,3-dichloro-2-propanol, enantioselective dehalogenation. 158 1,2-dichloroethane,Xanthobacter autotrophicus degradation, 152 dichloromehane, microbial dechlorination, 165 2,2-dichloropropionicacid (22DCPA), 135 diffuse reflectance IR spectra (DRJlT), 207 3.4-dihydroxyphenylacetate2,3-dioxygenase,49 dioxygen, chemistry,48 dioxygenases,ring-hydroxylating,49 biochemical organization,50,Sl catalytic non-haem Fe centre, 72-5 catalytic terminal oxygenase component, 60-1,62 classification, 51, 52-3,754 ferredoxin component, 5 8 - 4 9 ferredoxins sequence analysis, 58-9 specificity,58 iron-sulphur clusters, 61-72 amino acid sequence comparisons,67-8,

71 classes, 61-3

ligand analysis, 63 site-dmcted mutagenesis, 68-72 S ~ ~ C ~ ~ O63,657.66 SCO~Y, reductase component,S 1 , 5 5 4 subunit composition, 50,54 disulphide bridges, in hydrophobins, 9 DM-nitrophen, 188 DRIFT spectra, 207 Dutch elm disease, cerato-ulmin in, 18, 31-2 dye displacementmetal analysis, 198-9

crassa gene, in coni&ogenesis.2.8-9 ectomycollhiza, hydrophobin genes in formation, 33 electrochemicalmetal analysis, voltammetry, 194-5 electron microscopy see transmission electron microscopy electron spin resonance (ESR) spectroscopy, 208 enantioselection,microbial, 158 enniatin synthetase, 1 0 0 4 depsipeptide formation, 103,104 gene structure, 102-3 molecular structure, 102 N-methylationmechanism, 1 W 1 structureffunction,100,101 substrate specificity, 101-2 enniatins, 99-100 synthesis, 91 R-epichlorohydrin, 158 epimerization, by peptide synthetases, 92 ergot peptide alkaloids, 108-11 ergotamine, 108,109 Escherichia coli arsenic resistance, 226 chemotaxis, calcium ions in, 188 copper metabolism study, 222 genes in inorganic ion physiology, 210 iron in, 216 Mossbauer spectroscopy,209 iron-limited growth, 189 metal-tolerand-sensitive,isolation, 214 silver nistance, 230 Eucalyptus globulus, Pisolifhus tinctorus association, 33 e m N.

fatty acid synthase, 88 activation domain organization, 94 FBF, in fruit body formation, 24 ferredoxins, of dioxygenases, 58-60 femtin, 216 ferrozine, for iron determination, 190-1

266 flavoproteins in droxygenase system, 50,51 iron-sulphur, of dioxygenase reductases, 55-7 Fourier transform IR spectroscopy, 206-7 fruit bodies of basidiomycetes, 3 formation, 22-7 fungi biology, 1-3 peptide synthases 6-(L-a-aminoadipyl)-cysteinyl-o-valine. 96-9 beauvericin, 105 cyclosporin, 105-7 enniatins, 99-104 ergot peptide alkaloids, 108-11 SDZ 214-103,107-8 fura-2, in calcium analysis, 191 Fusaria, enniatin-producing, 99, 102 gas chromatography, 198 gene probing, 212-13 gene sequence libraries, 21 1 germanium, 182 microbial accumulation, 227 glutathione, in dichloromethane dechlorination, 165 gramicidin S, synthesis, 86-7 gramicidin synthetases, 87 molecular masses, 89 haem proteins analysis in vivo, 218-19 Mbssbauer spectroscopy, 209 halocompound dehalogenation see alcohol dehalogenation; alkane dehalogenation; alkanoic acid dehalogenation halohydrin epoxidases, 153 haploid fruiting alleles (hfa), in monokaryons, 24 HC-toxin, 110 HD genes, in fruit body formation, 2 3 4 herbicides Dalapon, 135 phosphinothricin as, 120 Hefernsigma akashiwo, phosphorus metabolism, 197 His motif, in peptide synthetases, 92 hybridization techniques, 212-13 hydrophobins, 3-45

INDEX in aerial hypha formation, 19-22 cerato-ulmin. sequence determination, 18 in conidiogenesis, 27-9 Aspergillus nidulanr genes, 27-8 Neumspora crassa genes, 28-9 discovery, 3-4 in fruit body formation ABHl gene expression, 26-7 functions, 26-7 SC gene expression, 24-6 identity, 4-10 agglutinin relationships, 8-9 assembly, 9-10 characteristics, 6, 8 cysteine residues, 9 hydropathy patterns, 6, 7 sequence diversity, 5 . 5 4 in pathogenesis, 29-33 Dutch elm disease, 31-2 fungal host adhesion, 29-31.32 human infections, 32-3 as plant defence response elicitors, 33 purification, enhancement, 34 in rodlet formation, genetic experiments, 11-12

sc3 purification, SC3, 14 surface activity experiments, 14-18 surface activities, 13-19 cerato-ulmin, 18-19 in symbiosis, 33-4 in technology, 34-6 applications, 35-6 and hydrophobin properties, 34-5 4-hydroxyphenylpymvate droxygenase, 73 hyphae aerial formation, SC3 hydrophobin in, 19-22 growth, 1-2 transport processes, 2 Hyphomicrnbim, dehalogenase, 165 inductively coupled plasma AES, 193 inductively coupled plasma MS, 194 ion chromatography, 197-8 ion-exchange resins, for metal removal from medium, 189 ion-selective electrodes, 1 9 5 4 cadmium, 226-7 copper, 222 iron, 216-21 aluminium interference, 215 analysis, 190-1

INDEX assays, 216 forms of,216 growth limitation, 189-90 oxidation/reduction, 220-1 in oxygenase catalysis, 49 protein analysis in vivo, 218-20 siderophores, 181,217-18 lransport, 181,217 iron-sulphur clusters of dioxygenases. 61-72 amino acid sequence comparisons, 67-8, 71 classes, 61-3 Iigand analysis, 63 sitedirected mutagenesis, 68-72 Spectro~c~py, 63,65-7.66 iron-sulphur proteins, analysis in vivo, 219-20 isopenicillin N synthase, 49, 73 isotope transport assays, 199-200 Klebsiella aemgenes cadmium resistance, 227 lead resistance. 228 Lactobacillus planrarum, iron requirement, I 9 0 lead, microbial resistance, 213,228 lectin-like activity of cryparin, 9 lectins, cysteine residues, 9 libraries, gene sequence, 211 lichens, 3 algal symbiosis, hydrophobins in, 3 3 4 Listeria monocytogenes, iron transport assay, 217 D-lysergylpeptide assembly, 108-1 1 Magnaporrhe grisea, hydrophobic host adhesion, 29-30 manganese oxidation, microbial, 207 mass spectrometry, inductively coupled plasma, 194 MAT genes in fruit body formation, 22-5 in SC3 regulation, 19-20 meiospores, fungal, 3 membrane proteins, in metal transport, 181 mercury biosensor, 212 microbial resistance, 228-9 Bacillus, 213,229 metallothioneins, copper, 221 metaldmtalloids, 177-241 aluminium, 182,214-16 analysis, 190-205 cell treatment for, 191-2

267 chromatography, 198 colorimetry, 190-1 contamination avoidance, 192 dye displacement, 198-9 inductively coupled plasma-mass spectrometry, 194 ion chromatography, 197-8 ion-selective electrodes, 1 9 5 4 isotope transport assays, 199-200 neutron activation, 196-7 proton displacement, 199 sample treatment, 192 spectroscopy, 1 9 3 4 transmission electron microscopy, 201-5 voltammetry, 1 9 4 5 antimony, 182 arsenic, 1 8 2 , 2 2 5 4 biosensors, 21 1-12 cadmium see cadmium copper see copper detoxification processes, 183 essential, 180-2 gene probing, 212-13 germanium, 182,227 “indifferent”, 183 ion complexation in media, 185-90 and bioavailability, 185, 186-7 calcium concentration control, 187-8 and growth limitation, 188-90 and speciation, 185-7 iron see iron lead, 213,228 mercury see mercury molecular genetics, 210-1 I , 232 molybdenum, 181 nickel, 224-5 potassium see potassium properties, 183-5 d block, 184 p block, 184-5 s block. 183-4 resistanfhlerant hacteria isolation, 213-14 selenium, 182, 229 silver, 229-30 sodium, 181 S p e C t r ~ ~ ~205-9 ~py, analytical techniques, 1 9 3 4 electron spin resonance, 208 electronic, 205-6 metal binding sites, 205 MBssbauer, 209 nuclear magnetic resonance, 208-9

268 metaldmetalloids, spectroscopy (continued) vibrational. 206-8 tellurium, 230-1 terminology, 179-80 tin, 231 toxicity and resistance, 182-3, 225 zinc, 223-4 Metarhizium anisopliae, hydrophobic host adhesion, 30 methane monooxygenases, and haloalkane metabolism, 164-5 4-methoxybenzoate monooxygenase, 49 iron site, spectroscopy, 75 4-methyl-3-hydmxyanthranilic (4-MHA) pentapeptide lactone, 111-12,112 Micrococcus luteus, lead resistance, 228 molybdenum, transport, 181 L-2-monochloropropionic acid, commercial production, 143, IS8 monooxygenases, 49 methane, and haloalkane metabolism, 164-5 4-methoxybenzoate, 49 iron site, spectroscopy, 75 Moraxella plasmids, 149 species B, dehalogenases, 138, 141, 142, 162 Mossbauer spectroscopy, 209 of iron-sulphur clusters in dioxygenases, 64. 65 mycorrhiza see ectornycorrhiza naphthalene dioxygenase, 49 ferredoxin, 58, 5 9 , m reductases, 58 Neumspora cmssa conidiogenesis genetics, 28 spores, rodlet layer, 11 neutron activation analysis, 196-7 nickel, microbial interactions, 224-5 Nitr-5, 188 nitrilotriacetic acid, 189 7'-nitrobenz-2-oxa-l,3-diazde (NBD). in iron analysis, 217 Nitachiu a h , germanium uptake, 227 nuclear magnetic resonance (NMR) spectroscopy, 2 0 W

oils, water dispersion, hydrophobins in, 35 oxygen see dioxygen oxygenase-type dehalogenases, 164-5 oxygenases, bacterial, 48-9

INDEX as biocatalysts, 48 monooxygenases/dioxygenases, 49 see also dioxygenases, ring-hydroxylating; monooxygenases

Paracoccus denitn'ficans. iron-limited growth, 189 peptide synthesis systems, backridfungi, 85-131 activation domain organization, 9 4 4 . 9 5 fungal, 96-1 11 delta-(L-alpha-aminoadipy1)cysteinyl-wvaline, 96-9 beauvericin, 105 cyclosporin, 105-7 enniatins, 99-104 ergot peptide alkaloids, 108-1 1 SDZ 214-103,107-8 peptide synthetase domain, 88-94 acyltransfer/epimerization modules, 91-2 amino acid activation, 9 3 4 modules, in activation domain, 90-1 motifs in carboxyl-adenylate-forming domain, 90 N-methylation module, 91 peptide synthetases, 88-90 thioesterase modules in genes, 92-3 prokaryotic, 111-22 acyl peptide lactone synthetases, 112-17 bialaphos, 120-2 surfactin, 117-20 research prospects, 1 2 2 4 tho1 template model, 86-8 periodic table, 183, I84 pbenylalanine bydroxylase, 49 phosphinothricin, 120, 120-2, I21 PhsB peptide synthetase, 121-2 phthalate dioxygenase, 55,56,57 ferrous active site, 75 spectroscopic analysis, 65-7.66 Pisolithus tinciorus, Eucalyptus globulus association, 33 plastics, hydrophobin adsorption, 15-17 polarography, 194 polyketide synthetases, 93 activation domain organization, 9 4 , 9 5 4 polymerase chain reaction, 212 Posidonia oceanica, trace element analysis, 197 potassium as essential metal, 180 transport, 181 rubidium as analogue, 200

INDEX protocatechuate 3,4-dioxygenase, 73 proton displacement metal analysis, 199 Pseudomonas, 47 dioxygenases, 50 haloalcohol dehalogenases, 154, 1 5 6 7 species 113, dehalogenase, 139,144 species CBS 3, 138, 141 Pseudomonas aeruginosa, silver accumulation, 229-30 Pseudomonas cepacia dehalogenases, 138, 141, 142 phthalate hoxygenase analysis, 65-7,66 Pseudomnas dehalogennns, dehalogenase, 137, 141 Pseudomnas putida benzene dioxygenase analysis, 65 dehalogenases, 139, 140, 143, 144-5 germanium uptake, 227 iron-sulphur cluster analysis, 219-20 Pseudomnas srurzeri, germanium accumulation, 227 Pseudomonas syringae, copper-resistant, 214 putidaredoxin, Mossbauer parameters, 65 pyridine, in haem protein analysis, 218 Pyrococcusfuriosus, tungsten in, 180 Rumaline stenospora, metal analysis, 196-7 RETRIEVE, 21 1 Rhizobium aluminium toxicity, 215 dehalogenases, 138, 139, 141, 142, 143-4 Rhodobacter capsulatus, Rieske proteins amino acid sequence, 67.7 1 sibdirected mulagenesis, 69, 71-2 Rhodococcus rrythmpolis, dehalogenases, 164 oxygenase type, 165 Rhodopseudomoms sphaemides, Rieske proteins, 67 Rieske proteins, 63 amino acid sequences, 67-72 site-directed mutagenesis, 68-72 Sacchammyces cerevisiae, 6&9,70 specm 64 spectroscopic analysis, 65-7 dA. nidulam gene, in conidiogenesis, 27 rodlet layer, 4, 8 fornation, 20,21 and hydrophobin wettability, 17-18 in lichedalgal synibiosis, 34 in pathogenicity, 33 rodlets, 10, 10-13

269 bacterial, 11 genetic experiments, 11-12 hydrophobins in formation, 11-13 isolation from fungal spores, 10-1 1 rubidium in potassium transport studies, 200 S-glucan, and rodlet location, 10 Sacchammyces cerevisiae copper uptake, 221 genetics of conidiogenesis, 27 iron metabolism, 22 1 Rieske proteins amino acid sequence, 68 site-directed mutagenesis, 68-9, 70 SC3 hydrophobin, 5 . 6 in aerial hypha formation, 19-22 discovery, 3-4 hydropathy pattern, 6 , 7 purification, 14 rodlet layer formation, 4 surface activity experiments, 14-18 SC4 hydrophobin, 5 . 6 discovery, 3-4 Schizophyllum commune fruit body formation, 2 2 4 gene expression, 2 3 , 2 3 4 hydrophobin function, 26 hydrophobin genes, 3 hydrophobins from, 14 dendrogram, 5 hydropathy patterns, 6, 7 in rodlet formation, 12-13 hyphal adhesion, 30-1 rodlet layer, 4 rodlets, 10, 12-13 SDZ 214-103, 107-8 selenium, microbial toxicity, 182, 229 siderophores, 181,217-18 silver, microbial toxicity, 229-30 sinefungin, as methylase inhibitor, 100-1 snake toxins, cysteine residues, 9 s d u m , transport, 181 Southern blotting, 212 spacer motif, in peptide synthetases. 92 spectrometry atomic fluorescence, 194 inductively coupled plasma-MS, 194 spectrophotometry, in haem protein analysis, 218-19 spectroscopy atomic absorption, 193 atomic emission, 193

270

INDEX

spectroscopy (continued) energy-dispersive X-ray, for TEM detection, 203-4 of iron-sulphur clusters in dioxygenases, 63, 657,154 for metal-microbe interactions, 205-9 electron spin resonance, 208 electronic, 205-6 metal binding sites, 205 Mossbauer, 209 nuclear magnetic resonance, 208-9 vibrational, 2Ch5-43 spinach ferredoxin, 61 Staphylococcus aureus arsenic resistance, 226 lead resistance, 228 Streptomyces clirysornallus, 113 Streptomyces clavuligerus, ACV synthase from, %,97 Streptomyces hygroscopicus, phosphinothricin from, 120 Streptomyces viridochromogenes, phosphinothricin from, 120 superoxide dismutase, Cu-Zn. 223 surfactin, 117-20 enzymes in assembly, 117-18,119 reactions in amino acid positions, 118-19 structure, 118 synthesis initiation, 118 synthetases, structdfunction, 119-20 symbiosis, fungal, 3 3 4 Syncephulastrum racemsum spores, rodlet layer, 11 Teflon, hydrophobin adsorption, 15-17.35 tellurium, microbial resistance, 230-1 tetrahionate, ion chromatography, 197-8 Thermm thermophilus iron isotope studies, 209 ironsulphur clusters, spectroscopy, 65, 66

Kieske proteins, amino acid sequence, 68 tellurium resistance, 230 Thiobncillusfermoxidans, 220 thioesterase genes, in peptide synthesis, 92 thiol template peptide synthesis model, 86-8 see also peptide synthesis systems, bacterialfungi thionins, cysteine residues, 9 thiosulphate, ion chromatography, 197-8 THN gene, in fruit body formation, 24 thri mutation, and SC3 expression, 19

tin, microbial toxicity, 231

tissue engineering, hydrophobins in, 35 toluene dioxygenase, 49 ferredoxin, 5 8 , 5 9 4 reductase, 57 toxin-agglutinin fold proteins, hydrophobin relationship, 8-9 trace elements, 180 deficiencies, and growth, 188-90 see also metaldmetalloids transition metals in oxygenase catalysis, 49 see also metaldmetalloids transmission electron microscopy for metals, 201-5 energy-dispersive X-ray spectroscopy detection, 2 0 3 4 selected-area diffraction with, 204-5 thin section preparation, 201-3 whole mounts, 201 transport in hyphae, 2 metals, 180-2 iron, 181. 217 isotope assays, 199-200.217 bis(tributy1tin)oxide (TBTO), bacterial degradation, 231 Trichodema hanionum, conidiospores, 13 Mchophyton menmgrophytes, microconidial rodlet layer, 10-11 tungsten, as essential metal, 180 Uromyces appendicularus, hydrophobic adhesion, 30 Vibrio alginolyticus copper-binding proteins, 223 copper-resistant/-sensitive, 214 voltammetry, 194-5

water, disinfection, copper/silver, 221 Western blotting, 213 wetA A. nidulans gene, in conidiogenesis, 27 X-ray absorption spectroscopy, phthalate dioxygenase, 6 7 X-ray photoelectron spectroscopy ( X P S ) , for hydrophobins, 17 Xanthobacfer aurotrophicus dehalogenases alkane, 16&2,163 alkanoic acid, 138. 141 DhlB overexpression, 151

271

INDEX 1,2-dichloroethane degradation, 152 zinc, microbial interactions, 223-4

zinc-sensitive E. cofi,214 zygomycetes, hyphal structure, 2

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