Pseudomonas
Juan-Luis Ramos · Alain Filloux Editors
Pseudomonas Volume 6: Molecular Microbiology, Infection and Biodiversity
123
Editors Prof. Juan-Luis Ramos Consejo Superior de Investigaciones Cientificas c/Prof. Albareda, 1 18008 Granada Spain
[email protected]
Prof. Alain Filloux Imperial College, London Faculty of Natural Science Division of Cell & Molecular Biology London South Kensington Campus Flowers Bldg. United Kingdom SW7 2AZ
ISBN 978-90-481-3908-8 e-ISBN 978-90-481-3909-5 DOI 10.1007/978-90-481-3909-5 Springer Dordrecht Heidelberg London New York Library of Congress Control Number: 2010921600 © Springer Science+Business Media B.V. 2010 No part of this work may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording or otherwise, without written permission from the Publisher, with the exception of any material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)
Preface
Paris is a cosmopolitan city where roaring life, wonderful museums and excellent science can be found. It was during the XI IUMS conference held in this city that the Pseudomonas book series was first envisaged. On the first row of the auditorium sat a group of outstanding scientists in the field, who after devoting much of their valuable time, contributed in an exceptional manner to the first three volumes of the series, which saw the light simultaneously. The volumes were grouped under the generic titles of “Vol. I. Pseudomonas: Genomics, Life Style and Molecular Architecture”, Vol. II. Pseudomonas: Virulence and gene regulation; Vol. III. Pseudomonas: Biosynthesis of Macromolecules and Molecular Metabolism. Soon after the completion of the first three volumes, a rapid search for articles containing the word Pseudomonas in the title in the last 10 years produced over 6,000 articles! Consequently, not all possible topics relevant to this genus were covered in the three first volumes. Since then two other volumes were published: Pseudomonas volume IV edited by Roger Levesque and Juan L. Ramos that came to being with the intention of collecting some of the most relevant emerging new issues that had not been dealt with in the three previous volumes. This volume was arranged after the Pseudomonas meeting organized by Roger Levesque in Quebec (Canada). It dealt with various topics grouped under a common heading: “Pseudomonas: Molecular Biology of Emerging Issues”. Yet the “Pseudomonas story” was far from complete and a new volume edited by Juan L. Ramos and Alain Filloux was deemed necessary. The fifth volume was conceived with the underlying intention of collecting new information on the genomics of saprophytic soil Pseudomonas, as well as on the functions related to genomic islands and was published in 2006. At the request of a number of scientists and colleagues working in the field, we have collected a new set of chapters that are called on to provide further views on the biology of Pseudomonas. Chapters in Pseudomonas volume VI have been grouped under the following topics: Regulation and control of virulence, Life styles, Physiology and Metabolism. The chapters under the heading Life Styles constitute an in-depth analysis of the genome of Pseudomonas stutzeri, a denitrifier par excellence, and the behaviour and life style of P. aeruginosa in the human lung. The Physiology Metabolism and Markers section collects four chapters that deal with the v
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Preface
metabolism of acyclic terpenes by Pseudomonas, the biodiversity of siderophores, resistance to heavy metals and the role of relevant second messenger, a c-di-GMP, as a signalling molecule. Finally under Regulation and control of virulence we find several chapters dealing with sensing at the level of cell surfaces and quorum sensing, as well as the role of small RNAs in the control of gene expression. It would not be fair not to acknowledge that this sixth volume would never have seen the light if it were not for a group of outstanding scientists in the field who have produced enlightening chapters to try to complete the story that began with the five previous volumes of the series. It has been an honour for us to work with them and we truly thank them. The review process has also been of great importance to ensure the high standards of each chapter. Renowned scientists have participated in the review, correction and editing of the chapters. Their assistance is immensely appreciated. We would like to express our most sincere gratitude to: Bonnie Bassler Burkhard Tümmler Christophe Bordi Eduardo Díaz Eric Deziel Estrella Duque Hermann Heipieper Iñigo Lasa Isabelle Schalk
Norberto Palleroni Paul Visca Pierre Cornelis Regine Hennge-Aronis Simon Silver Soeren Molin Susanne Haussler Vittorio Venturi
We would also like to thank Carmen Lorente once again for her assistance and enthusiasm in the compilation of the chapters that constitute this sixth volume. Granada, Spain London, UK
Juan L. Ramos Alain Filloux
Contents
Part I
Regulation and Control of Virulence
1 Small RNAs of Pseudomonas spp. . . . . . . . . . . . . . . . . . . . Elisabeth Sonnleitner, Nicolas González and Dieter Haas
3
2 2-Alkyl-4(1H)-Quinolone Signalling in Pseudomonas aeruginosa . . Matthew P. Fletcher, Stephan Heeb, Siri Ram Chhabra, Stephen P. Diggle, Paul Williams, and Miguel Cámara
29
3 Cell-Surface Signalling in Pseudomonas . . . . . . . . . . . . . . . . María A. Llamas and Wilbert Bitter
59
4 Second Messenger c-di-GMP Signaling in Pseudomonas aeruginosa Massimo Merighi and Steve Lory
97
Part II
Life Styles
5 Emergence of Pseudomonas aeruginosa in Cystic Fibrosis Lung Infections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Joanna B. Goldberg 6 Insights into the Life Styles of Pseudomonas stutzeri . . . . . . . . . Elena García-Valdés, Magdalena Mulet, and Jorge Lalucat
141 177
Part III Physiology, Metabolism and Markers 7 Pyoverdine Siderophores as Taxonomic and Phylogenic Markers . . Jean-Marie Meyer
201
8 Metabolism of Acyclic Terpenes by Pseudomonas . . . . . . . . . . . Jesús Campos-García
235
9 Heavy Metal Resistance in Pseudomonads . . . . . . . . . . . . . . . Esther Aguilar-Barajas, Martha I. Ramírez-Díaz, Héctor Riveros-Rosas, and Carlos Cervantes
255
Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
283
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Contributors
Esther Aguilar-Barajas Instituto de Investigaciones Químico-Biológicas, Universidad Michoacana, UAM, Morelia, México Wilbert Bitter Department of Medical Microbiology, VU University Medical Centre, Amsterdam 1081 BT, The Netherlands Miguel Cámara School of Molecular Medical Sciences, Centre for Biomolecular Sciences, University Park, University of Nottingham, Nottingham NG7 2RD, UK,
[email protected] Jesús Campos-García Instituto de Investigaciones Químico-Biológicas, Universidad Michoacana de San Nicolás de Hidalgo, Edif. B-3, Ciudad Universitaria, CP 58030, Morelia, Michoacán, México,
[email protected] Carlos Cervantes Instituto de Investigaciones Químico-Biológicas, Universidad Michoacana, UAM, Morelia, México,
[email protected] Siri Ram Chhabra School of Molecular Medical Sciences, Centre for Biomolecular Sciences, University Park, University of Nottingham, Nottingham NG7 2RD, UK Stephen P. Diggle School of Molecular Medical Sciences, Centre for Biomolecular Sciences, University Park, University of Nottingham, Nottingham NG7 2RD, UK Matthew P. Fletcher School of Molecular Medical Sciences, Centre for Biomolecular Sciences, University Park, University of Nottingham, Nottingham NG7 2RD, UK Elena García-Valdés Microbiologia, Department de Biologia and IMEDEA (CSIC-UIB), Universitat de les Illes Balears, Palma de Mallorca, Spain Joanna B. Goldberg University of Virginia, Charlottesville, VA, USA,
[email protected] Nicolas González Institut de Microbiologie, Centre Hospitalier Universitaire Vaudois, CH-1011 Lausanne, Switzerland
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Contributors
Dieter Haas Département de Microbiologie Fondamentale, Université de Lausanne, CH-1015 Lausanne, Switzerland,
[email protected] Stephan Heeb School of Molecular Medical Sciences, Centre for Biomolecular Sciences, University Park, University of Nottingham, Nottingham NG7 2RD, UK Jorge Lalucat Microbiologia, Department de Biologia and IMEDEA (CSIC-UIB), Universitat de les Illes Balears, Palma de Mallorca, Spain,
[email protected] María A. Llamas Department of Medical Microbiology, VU University Medical Centre, Amsterdam 1081 BT, The Netherlands,
[email protected] Steve Lory Department of Microbiology and Molecular Genetics, Harvard Medical School, Boston MA 02115, USA Massimo Merighi Department of Microbiology and Molecular Genetics, Harvard Medical School, Boston MA 02115, USA,
[email protected] Jean-Marie Meyer Département Génétique moléculaire, Génomique et Microbiologie, UMR 7156 CNRS-Université de, Strasbourg, France,
[email protected] Magdalena Mulet Microbiologia, Department de Biologia and IMEDEA (CSIC-UIB), Universitat de les Illes Balears, Palma de Mallorca, Spain Martha I. Ramírez-Díaz Instituto de Investigaciones Químico-Biológicas, Universidad Michoacana, UAM, Morelia, México Héctor Riveros-Rosas Departamento de Bioquímica, Facultad de Medicina, UNAM, Mexico City, México Elisabeth Sonnleitner Département de Microbiologie Fondamentale, Université de Lausanne, CH-1015 Lausanne, Switzerland Paul Williams School of Molecular Medical Sciences, Centre for Biomolecular Sciences, University Park, University of Nottingham, Nottingham NG7 2RD, UK
Obituary
Yoshifumi Itoh, Professor of Microbial Biotechnology at Tohoku University (Sendai, Japan), suddenly passed away on October 4, 2008. He was well-known to the Pseudomonas community as a specialist of amino acid metabolism and plasmid replication. In this book series, he contributed two chapters, one on arginine and polyamine metabolism (in volume III) and one on histidine catabolism and catabolite repression (in volume V). He published almost 100 papers and book chapters during his career as a microbiologist. Yoshi, as he was known to his friends, was born in a small city near Sendai (Miyagi Prefecture) on August 5, 1951. Here he grew up with his elder sister and younger brother. After studies at Tohoku University, he obtained a Master of Agricultural Science in 1976 and a PhD of Agricultural Science in 1979. His PhD thesis was on the mode of action of a bacteriocin in Erwinia carotovora. In xi
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1980, he was hired as a Research Associate at Shinshu University (Matsumoto, Japan), where he worked with Yoshiro Terawaki and Hideki Matsumoto. He soon left to spend two years (1981–1983) as a postdoctoral fellow at the Department of Microbiology, ETH Zurich (Switzerland) where he worked in the group of Thomas Leisinger and Dieter Haas. He characterized the replication and partition functions of the Pseudomonas plasmid pVS1. Many years later, in 2000, he extended this work and, in collaboration with the group of Dieter Haas, made use of the minimal pVS1 replicon in the construction of a series of stable plasmid vectors, which are widely used and cited today. Upon return to Japan in 1983, he again joined the School of Medicine at Shinshu University. In 1988, he moved to the National Food Research Institute in Tsukuba where he eventually became the head of the Applied Bacteriology Laboratory. He developed an interest in metabolic functions of Pseudomonas aeruginosa and their regulation. Simultaneously, he also studied Bacillus subtilis as an organism used for natto (fermented soybean) production. In 2004, he became Director General of the Akita Research Institute of Food and Brewing. At that time he was somewhat reluctant to take up this position, as it did not allow him to spend much time on research. So he was very pleased that two years later he was appointed to his home University at Sendai as a Professor. Although he had substantial teaching commitments, he was finally able to return to his favourite research topics. Probably the most significant scientific contribution of Yoshifumi Itoh was his discovery of the CbrAB two-component system in P. aeruginosa. This system regulates the activity of sigma-54 RNA polymerase during the utilization of numerous carbon and nitrogen sources. Yoshifumi Itoh was a dedicated scientist and always enthusiastic about his research. He had a fine sense of humour. His untimely death came as a complete and very sad surprise to everyone. He is sorely missed by his wife Junko, his daughter, his son and his friends and colleagues. Dieter Haas
Part I
Regulation and Control of Virulence
Chapter 1
Small RNAs of Pseudomonas spp. Elisabeth Sonnleitner, Nicolas González and Dieter Haas
1.1 Introduction Small RNAs (sRNAs) of prokaryotes are 40–600 nucleotides in length and most have regulatory functions. As a rule, sRNAs do not encode polypeptides but important exceptions exist. Some bacterial sRNAs were discovered and characterized in the last two decades of the twentieth century, mostly with emphasis on sRNAs that regulate functions of plasmids, bacteriophages or transposable elements. Although it was already clear at that time that sRNAs could have diverse functions, their roles as cellular regulators were not fully appreciated [1]. This picture has changed since then with the discovery of many novel sRNAs, made possible by the introduction of new computational and genomic approaches. To date, close to 100 sRNAs have been reported in Escherichia coli and more than 150 sRNAs in prokaryotes altogether [2]. In this chapter, we will review the current status of sRNAs encoded by chromosomal genes in Pseudomonas species, with emphasis on P. aeruginosa where most genomic information is available. To date, > 40 sRNAs have been detected in this organism; however, for many of them, the functions have not yet been uncovered. Why has it been relatively difficult to find sRNAs and their genes in prokaryotes by classical genetical and biochemical approaches? There are several reasons for this. (i) Mutational inactivation of a particular sRNA gene often does not cause
D. Haas (B) Département de Microbiologie Fondamentale, Université de Lausanne, CH-1015 Lausanne, Switzerland e-mail:
[email protected] Note added in proof: After completion of this review in December 2008, Brencic et al. (Mol. Microbiol. 73:434–445, 2009) reported that the GacS/GacA two-component system achieves virtually all of its regulatory effects by positively controlling the expression of the rsmY and rsmZ genes in P. aeruginosa, implying that the GacA protein binds exclusively to the rsmY and rsmZ promoters. Moreover, the global H-NS-type regulator MvaT was recognised as a repressor of the rsmZ gene. Sonnleitner et al. (Proc. Natl. Acad. Sci. USA, doi: 10.1073/pnas. 0910308106) described a novel sRNA, CrcZ, mediating catabolite repression in P. aeruginosa. CrcZ is specified by the cbrB-pcnB intergenic region.
J.L. Ramos, A. Filloux (eds.), Pseudomonas, DOI 10.1007/978-90-481-3909-5_1, C Springer Science+Business Media B.V. 2010
3
4
E. Sonnleitner et al.
an easily observable phenotype. (ii) Numerous sRNAs are functionally redundant and therefore loss of one element does not have important consequences. (iii) In transposon mutagenesis, sRNA genes may be spared because they are small. (iv) Point mutations are much better tolerated in sRNA genes than in coding genes where missense, nonsense or frameshift mutations usually lead to loss of function. (v) Overexpression of sRNAs, while providing a useful tool for probing sRNA function, is a technically demanding method in that it often requires a strong external promoter to be fused precisely to the transcription start site of an sRNA gene. There are several strategies that have proved useful in the elucidation of prokaryotic sRNAs. These approaches have been explored mostly in E. coli and several have also been applied to Pseudomonas spp. (i) A few sRNAs are sufficiently abundant so that they can be isolated from cells in pure form and sequenced. Historically, this has been the case for 4.5S RNA (the product of the f f s gene, the RNA component of the signal recognition particle), 6S RNA (the ssrS gene product, a regulator of RNA polymerase), 10Sa RNA (the ssrA gene product, termed tmRNA today) and 10Sb RNA (the rnpB product, the RNA component of RNase P) in E. coli [1]. However, most sRNAs are present in low amounts that preclude this approach. (ii) Computational prediction of sRNAs, followed by experimental verification using Northern blotting, has proved a broadly applicable strategy. Various algorithms have been developed to this end. They usually assume that sRNA genes are located in intergenic regions (IgRs) rather than within open reading frames and that putative promoter and rho-independent terminator sequences must be within a short distance of each other (< 500 nucleotides). Furthermore, computational predictions often include the criterion that sequences as well as secondary structures of sRNAs should be conserved in closely related bacterial species [3]. The principal limitation of this approach is that it is difficult to predict bacterial promoters except for those that have highly conserved RpoD, RpoN or RpoS recognition elements. Another limitation is that in some cases no typical rho-independent terminator can be identified. (iii) sRNAs that have a high affinity for an RNA-binding protein can often be copurified with the protein. Typically, the protein is isolated by immunoprecipitation or affinity chromatography and cDNAs are prepared from the sRNAs attached to the protein [4]. This strategy has been exploited successfully with proteins of the Hfq and CsrA families. Here, an inconvenient feature is the fact that a large proportion of non-specifically bound rRNAs and mRNA fragments are also enriched in the purified protein fraction. At any rate, the roles of sRNAs recognized in (ii) or (iii) have to be verified by genetic tests. (iv) Genetic screens – often involving multicopy expression of certain phenotypes – can be a fruitful strategy, especially when post-transcriptional regulation of gene expression is suspected, as for example is the case of the rpoS gene and genes encoding outer membrane proteins in E. coli [5–7]. Here, a caveat is that some overexpression effects might be missed because they are slight and transient or toxic to the cell [8, 9]. (v) Transcripts emanating from IgRs can be detected by microarrays when these cover the IgRs entirely; tiling microarrays are particularly useful in this respect. However, this approach can only be a
1
Small RNAs of Pseudomonas spp.
5
first step towards the elucidation of sRNAs and needs to be combined with other methods [10]. (vi) Direct shotgun cloning of cDNAs that are derived from sizefractionated RNAs can also be a useful first step, but this approach yields a high background of clones that are isolated due to RNA degradation products [11]. (vii) Finally, once an sRNA gene is well established in one bacterium, homologous sRNA genes may be found in related bacteria by BLAST searches. Such an approach may be facilitated when recognition sites for activators or repressors are conserved in promoter regions of sRNA genes or when neighbouring coding genes are conserved [12, 13]. It has to be pointed out, however, that sRNA genes diverge more strongly than do coding genes [2]. For this reason, for example, only few sRNA genes of Pseudomonas spp. could be deduced from sRNA genes of E. coli by homology searches.
1.2 Overview of Observed sRNAs in P. aeruginosa To date, three genomic surveys of sRNAs in P. aeruginosa strain PAO1 have been published. The first study by Livny et al. [14] used a computational tool termed sRNAPredict2, which attempts to identify sRNA genes in IgRs for which conservation of sRNA sequence and secondary structure is found among multiple species. Furthermore, appropriately located rho-independent terminators are taken into consideration [3, 14]. From 38 predicted candidate genes, about half were found to specify transcripts that were detectable in Northern blots, in addition to four genes (rsmY, rsmZ, prrF1 and prrF2) that had previously been identified (Table 1.1). A second survey by González et al. [15] took a similar approach, but used the QRNA program [23] instead of sRNAPredict2. From a preliminary list of 162 candidate IgRs, 14 were found to specify short transcripts and of these, eight were new. The latter number probably represents an underestimate because not all candidate IgRs were subjected to Northern blot analysis [15]. In a third study by Sonnleitner et al. [16], total RNA was extracted from P. aeruginosa, size-fractionated in the range of 50–500 nucleotides, and incubated with Hfq protein. After addition of anti-Hfq antibodies, the immunoprecipitate was recovered and RNA bound to Hfq was converted to cDNA, which was cloned and sequenced. After elimination of clones derived from mRNA fragments, tRNAs and rRNAs, eight candidate genes were recovered and sRNAs were verified by Northern blotting or RT-PCR. In addition, two sRNAs were predicted by RNAz, a bioinformatics tool based on the conservation of RNA structures, and verified by RT-PCR. In total, five new sRNAs were found. The consolidated data of the three studies [14–16] and an early publication describing 6S RNA [20] are summarized in Table 1.1, which shows observed transcript sizes, the orientation of the sRNA genes, map coordinates according to the genomic sequence of strain PAO1 [24], and flanking genes. Clearly, the total of 41 sRNA genes or candidate sRNA genes observed (Table 1.1) represents a preliminary estimate and this number is likely to grow as more studies will become available.
334456334733 586664587016 785174785969 901047901933 912780913085 971625972166 11173911118157 11408601141267 12047821205770 14363971436663 16688331669085 19286271928893
Coordinates of intergenic regions
12050311205330 14364911436618 16689111669081
586867586990 785498785570 901520901872
P11
P9
888–4.5S RNA 26/–o
25/102o
300
645
171 (113f ) 100
128
200, 300
622
<<<
>><
>>>
><>
>>>
>>>
>>>
< >>
<<<
353n 90
>>>
<><
<< >
Orientationc
73
124
300
Size (nt)b
130
ffsf
ssrA
rsmYd
Gene
Locationa
P8
–/96o
tmRNA
PhrD
RsmY
Sonnleitner et al. (2008) [16]
140
491
González et al. (2008) [15]
P7 (?)e
P5
RsmY
P1
Livny et al. (2006) [14]
Designation according to
sRNAs
Table 1.1 Experimentally found sRNAs of Pseudomonas aeruginosa PAO1
PA1781nirB
PA1530
PA1324
PA1112
PA1052
PA0887acsA PA1030
PA0836
PA0826
PA0527dnr PA0714
PA0296
Name
5 gene
PA1782
PA1531
PA1325
PA1113
PA1053
PA0837 - slyD PA0888aotJ PA1031
PA0827
PA0715
PA0297spuA PA0528
Name
3 gene
Flanking genes
6 E. Sonnleitner et al.
19968071997508 29182122918965 31067523107002 31121513112876 32062533206915 32989223299492 33186573318881 33606543360873 37029513703166 37051613705888 37780003778265 40574834057910
Coordinates of intergenic regions
amiL
37053093705521 37780343778133 40575434057658
RsmZ
amiE leader
rsmZi
116
100
213
PhrS
P20
1887
100
200
P18
1714
122
300
250, 300
76
40/129o
33187473318868
31069193106994
150, 200, 300 300
Size (nt)b
P16 rgsA
Gene
Locationa
180
–/118o
Sonnleitner et al. (2008) [16]
–/132o
1698
1559
1466e
1059
González et al. (2008) [15]
P15
P14 (?)h
P13 (?)g
Livny et al. (2006) [14]
Designation according to
sRNAs
Table 1.1 (continued)
> <<
<<>
<<<
<<<
>> <
< >>
< >>
>?>
<<<
>?<
Orientationc
PA3366amiE PA3621fdxA
PA3305
PA3304
PA3001
PA2958
PA2942
PA2852
PA2744thrS PA2750
PA1838cysI PA2581
Name
5 gene
PA3622rpoS
PA3367
PA3306
PA3002mvf PA3305
PA2959
PA2853oprL PA2943
PA2751
PA2745
PA2582
PA1839
Name
3 gene
Flanking genes
1 Small RNAs of Pseudomonas spp. 7
53084255309326 53449045345085
44445974444977 45364934536919 47806184780838 47817864781985 49391884939299 49560294956732 49857314985846 51968335197184 52839065284368
Coordinates of intergenic regions
180 80
–/29o –/27o
P30 (?)k
P32
113
prrF2j
92/32o
2667
PrrF2
62
115
PrrF1
52839955284110 52842075284319
49857824985843
350
84
prrF1j
rnpB
49391944939277
90/31o
PhrW
72/101l,o
250
54/72o 90
180
50/88o
Size (nt)b 300
Gene
Locationa
–/94o
Sonnleitner et al. (2008) [16]
200
2626
2510
2315
González et al. (2008) [15]
–/34
P35
P28
P27 (?)e
P26
P24
Livny et al. (2006) [14]
Designation according to
sRNAs
Table 1.1 (continued)
<<<
><>
>><
>><
>>>
<<<
<<<
<><
<><
<<<
<<<
><>
Orientationc
PA4726cbrB PA4758carA
PA4704
PA4628lysP PA4704
PA4451
PA4055ribC PA4270rpoB PA4272rplJ PA4406lpxC PA4421
PA3964
Name
5 gene
PA4727pcnB PA4759dapB
PA4705
PA4705
PA4629
PA4452
PA4056ribD PA4271rplL PA4273rplA PA4407ftsZ PA4422
PA3965
Name
3 gene
Flanking genes
8 E. Sonnleitner et al.
P34
Livny et al. (2006) [14]
González et al. (2008) [15]
Designation according to
102/16l,o
101/26o
PhrYl
PhrXl
Sonnleitner et al. (2008) [16]
ssrSm
Gene
58364295836579 58594805859674 58843205884502 59861205986170
Locationa
51
183
195
151
150
Size (nt)b
<>>
>>>
<>>
><>
><>
Orientationc
PA5316rpmB
PA5204argA PA5227
PA5183
PA5181
Name
5 gene
PA5228ygfA PA5317
PA5205
PA5184
PA5182
Name
3 gene
Flanking genes
a
A location is given when the 5 and 3 ends can be deduced with reasonable certainty from available data or when the predicted and observed lengths are in agreement [14, 15]. b Experimental values found in Northern blots. c The middle arrow indicates the orientation of the sRNA, while the flanking arrows indicate the orientation of the adjacent genes. Unknown orientation of sRNAs is indicated by a question mark. d Identified by Kay et al. [17]. e Signal intensities observed for these transcripts were particularly weak and the possibility of non-specific detection remains [14, 15]. f Identified by Toschka et al. [18]. The mature form is reported to consist of 113 nt [18]. g Transcript size and intensity observed differed using two distinct probes [14]. h Due to its size, the P14 transcript could correspond to the mRNA of the upstream PA2853 gene. i Identified by Heurlier et al. [19]. j sRNAs encoded in tandem, identified by Wilderman et al. [13]. k In the authors’ laboratory, P30 has not been detected but there is evidence for sRNAs including CrcZ encoded by the opposite strand. l Detection by RT-PCR. m 6S RNA gene identified by Vogel et al. [20]. n Mature tmRNA according to sequence homologies [21, 22]. o Predicted by a bioinformatic approach (NcDNAlign/multiz) in Sonnleitner et al. [16].
58350715835481 58364025836909 58594575859792 58842865884508 59861205986474
Coordinates of intergenic regions
sRNAs
Table 1.1 (continued) 1 Small RNAs of Pseudomonas spp. 9
10
E. Sonnleitner et al.
Not all transcripts listed in Table 1.1 are necessarily sRNAs sensu stricto. Some may be processed untranslated 5 or 3 leader sequences and two such examples (2315 sRNA, amiL) will be discussed in Section 1.3.5. It is interesting to note that three sRNAs that had been known for many years were sighted in at least one of the three screens: tmRNA (ssrA), 4.5S RNA (f f s) and the RNA component of RNase P (rnpB), whereas a forth traditional RNA, 6S RNA (ssrS), was not revealed although it is sufficiently abundant to be isolated directly and can be predicted by bioinformatics [16, 20]. One strongly expressed sRNA (P20/1887/PhrS) was found in all three genomic surveys and its properties as a quorum sensing regulator will be described in Section 1.3.9. Another distinctly expressed sRNA (P16/1698), which was termed RgsA [15], was seen in two studies and was investigated in some detail (see Section 1.3.7). A further sRNA (P5/491), which was also revealed in two studies, has not yet been characterized with respect to its function.
1.3 Examples of sRNAs in Pseudomonas spp. An overview of several Pseudomonas sRNAs with known functions and the methods leading to their identification are presented in Table 1.2. In the following, we will discuss the major properties of these characterized sRNAs. Clearly, this should be seen as an initial appraisal of a rapidly growing area of research.
1.3.1 6S RNA 6S RNA has been mainly studied in E. coli. It has about 10,000 copies at the end of exponential growth, is about 180 nucleotides in length and adopts a large hairpin-like secondary structure with several bulges. 6S RNA specifically binds to the σ70 RNA polymerase holoenzyme whereby the central major bulge of about 15 nucleotides mimics the open complex structure of promoter DNA. Thus, by interacting with σ70 RNA polymerase, 6S RNA inhibits the transcriptional expression of certain σ70 promoters in stationary phase, whereas transcription from σs (σ38 ) promoters appears to be favoured in vivo (Fig. 1.1). Mutants devoid of 6S RNA (ssrS) do not have any pronounced growth defect, but are affected in survival during late stationary phase. When E. coli cells are transferred from stationary phase cultures to fresh medium, 6S RNA is used as a template for transcription of a short RNA (< 20 nucleotides), which frees the enzyme of 6S RNA and restores RNA polymerase activity at σ70 promoters [33, 34]. 6S RNA occurs in many bacterial species. Owing to its conserved sequences bordering the central bubble and those in the closing stem formed by the 5 and 3 terminal segments, 6S RNA genes can be annotated with relative ease in a variety of bacteria including Pseudomonas spp. Furthermore, the conserved presence of a downstream gene, ygfA, facilitates the annotation of the ssrS gene in α- and γ-proteobacteria [12, 31]. In P. aeruginosa, 6S RNA was experimentally demonstrated by Vogel et al. [20]. These authors purified the RNA, which was associated with 70S ribosomes, labelled it at the 3 end and
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Small RNAs of Pseudomonas spp.
11
Table 1.2 Approaches used to detect and characterize sRNAs in Pseudomonas spp Experimental approach
Examples of sRNAs
Direct isolation of abundant sRNA
6S RNA: regulator of RNA polymerase 4.5 RNA: RNA in signal recognition particle 10Sa RNA: tmRNA 10Sb RNA: RNaseP RNA PrrF1 and PrrF2: functional RyhB homologs RsmY: GacA-dependent regulator of exoproducts RgsA: GacA-dependent regulator of oxidative stress response RsmX, RsmZ: GacA-dependent regulators, binding to RsmA protein PhrS: regulator of PQS biosynthesis, binding to Hfg protein TRR: RsmY homolog, multi-copy suppressor of temperature control of toxin synthesis PrrB: RsmZ homolog, multi-copy suppressor of gacA
Similarity with known sRNA of E. coli
Computational prediction of sRNA
Co-purification of sRNA with a protein
Genetic screen
a b
Speciesa
Gene
References
Pa
ssrS
[20]
Pa
ffs
[18]
Pa
ssrA
[22]
Pf
rnpB
[25]
Pa
prrF1, prrF2
[13]
Pf
rsmY
[26]
Pf
rgsA
[15]
Pf
rsmX, rsmZ
[27, 28]
Pa
PhrS
[16]6
Psp
rsmY
[7, 29]
Pf
rsmZ
[30]
Pa, P. aeruginosa; Pf, P. fluorescens; Psp, P. syringae pv. phaseolicola E. Souuleitner et al., unpublished data
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6S RNA EXPONENTIAL GROWTH
σ70
DNA open complex
RNA polymerase holoenzyme
σ38
σ70
3’ 5’
6S RNA STATIONARY PHASE
Fig. 1.1 During exponential growth (shown in green) bacterial cells strongly express housekeeping genes. These are usually under the control of σ70 promoters, which are recognized by σ70 RNA polymerase holoenzyme. At the beginning of the stationary phase (shown in blue) the expression of 6S RNA is increased. 6S RNA binds specifically to σ70 RNA polymerase holoenzyme and mimics the open complex structure of a σ70 promoter. This interaction inhibits the transcription from certain σ70 promoters and favors the transcription from σ38 promoters
used it as a probe to clone the chromosomal ssrS gene, which was then sequenced. The predicted secondary structure of 6S RNA from P. aeruginosa is very similar to that of 6S RNA from E. coli and there is 60% sequence identity between the 6S RNAs of both bacterial species [20], suggesting that these RNAs have similar functions in both organisms. The ssrS gene of P. aeruginosa appeared in the bioinformatic screen of Sonnleitner et al. [16], whereas neither Livny et al. [14] nor González et al. [15] detected this gene (Table 1.1). This is probably due to the fact that the latter screens rely on the presence of rho-independent terminators. However, 6S RNA does not have a rho-independent terminator and is formed by processing from a longer precursor, at least in E. coli [12].
1.3.2 4.5S RNA The bacterial signal recognition particle (SRP), which consists of a protein (the f f h gene product) and 4.5S RNA (the f f s gene product), serves essentially to target proteins to the cytoplasmic membrane, immediately following translation. This is
1
Small RNAs of Pseudomonas spp.
4.5S RNA
13
Ribosome mRNA
SRP Peptide signal sequence
4.5S RNA Ffh protein
SRP receptor FtsY
Cytoplasmic membrane SecYED
Sec translocon
Fig. 1.2 4.5S RNA and the Ffh protein form together the signal recognition particle (SRP). This complex is essential for targeting membrane proteins to the cytoplasmic membrane. The SRP recognizes a hydrophobic signal sequence in the nascent protein and binds to it. The ribosome-SRP complex moves to the cytoplasmic membrane where the SRP interacts with the SRP receptor, FtsY. The peptide signal sequence is transferred to the Sec translocon and the membrane protein is co-translationally inserted into the cytoplasmic membrane. At the end of this process, the SRP dissociates from its receptor and can be recovered
achieved by an interaction between the SRP and the hydrophobic signal sequence of the nascent membrane protein, which is thus prevented from folding (or misfolding) in the cytoplasm (Fig. 1.2). SRP is essential for viability [35]. The bacterial 4.5S RNA adopts a long hairpin structure with several small bubbles; they are smaller than the open complex-like bulge of 6S RNA. The loop region of 4.5S RNA contains a strongly conserved motif of about 20 nucleotides. This motif is useful to identify the f f s gene in bacteria [36] and has served to clone the P. aeruginosa f f s gene [18]. In this organism, 4.5S RNA (113 nucleotides) is formed from a larger precursor by processing of the 5 region, possibly by RNase P, as in E. coli. The 3 end of 4.5S RNA might be generated by a poorly conserved rho-independent terminator or by processing of a transcript having 34 extra nucleotides at the 3 end [18]. Although the function of the f f s gene has not been verified experimentally in P. aeruginosa, there can be little doubt that it has the same function as in E. coli, given the 75% sequence identity of the f f s genes between both organisms. The fact that the P. aeruginosa ffs gene was not revealed by the screen of Livny et al. [14] but was found in those of González et al. [15] and Sonnleitner et al. [16] (Table 1.1), probably reflects a less stringent terminator definition in the latter studies.
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tmRNA SmpB E P
EFTu A
A
EFTu E P
A
A
tmRNA
mRNA
stalled ribosome
E
PA A
A E P N A
E P
N
A
D N
A NDDNYALAA
C
proteolysis
Fig. 1.3 Under certain conditions (e.g., premature termination of transcription or truncation of mRNA) a translating ribosome can be stalled. The release of the unfinished polypeptide from the ribosome is accomplished with the help of tmRNA, which has features of both tRNA and mRNA. The tRNA part of tmRNA is charged with alanine and enters the ribosome A (acceptor) site in a complex with the SmpB and EF-Tu proteins. The nascent polypeptide is transferred to the alanyltmRNA. After transfer of the peptidyl-tmRNA-SmpB complex to the P (peptidyl) site, a small open reading frame is provided by the mRNA part of tmRNA. This allows further elongation and termination of translation. The resulting peptide is tagged, which provides a signal for proteolysis
1.3.3 tmRNA The function of tmRNA (for transfer-messenger RNA, the product of the ssrA gene, formerly termed 10Sa RNA) has been elucidated primarily in E. coli [37]. When translating ribosomes stall because of premature termination of transcription or truncation of mRNA, a special trans-translation mechanism elongates and releases the unfinished polypeptides from the ribosomes (Fig. 1.3). In this process, the tRNAlike part of tmRNA is charged with alanine. Alanyl-tmRNA enters the ribosome A (acceptor) site as a complex with the SmpB and EF-Tu proteins. The stalled nascent polypeptides are transferred to alanyl-tmRNA; upon translocation of the peptidyltmRNA-SmpB complex to the P (peptidyl) site, the mRNA part of tmRNA provides a small open reading frame that allows continuation and termination of translation. This results in the addition of a C-terminal peptide tag (ANDENYALAA) to the stalled nascent polypeptides. Subsequent termination of translation releases the ribosomes, and tagged proteins are targeted for proteolytic degradation (Fig. 1.3). The
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Small RNAs of Pseudomonas spp.
15
presence of tmRNA appears to be universal in bacteria. Null mutations in ssrA are tolerated but result in reduced fitness of the organisms [37]. The 5 and 3 termini of tmRNA are conserved. This enabled Williams and Bartel [22] to clone the ssrA gene (353 bp) from P. aeruginosa and other proteobacteria. The deduced tag sequence of P. aeruginosa tmRNA (ANDDNYALAA) is very similar to that of E. coli. A tmRNA fragment was found to be bound to Hfq in the screen of Sonnleitner et al. [16] (Table 1.1). However, it is not clear whether Hfq binding to tmRNA occurs in vivo.
1.3.4 RNase P RNA RNase P is an essential enzyme required for processing tRNA precursor molecules at the 5 end. It contains a catalytic RNA (the rnpB product consisting of 276 to ∼500 nucleotides, depending on the organism) and has been found in bacteria, archaea and eukaryotes. In E. coli and other bacteria, a small protein encoded by the rnpA gene is needed for in vivo activity of RNase P. The RNA of RNase P has several highly conserved nucleotides in its core sequence of about 200 nucleotides. This information can be used to identify the rnpB gene in various microorganisms [38, 39]. The first Pseudomonas species from which the gene for RNase P RNA was isolated, was P. fluorescens [25]. More recently, RNase P RNA was detected in P. aeruginosa in three independent screens [14–16] (Table 1.1).
1.3.5 Processed Leader Transcripts Some sRNAs do not have regulatory functions per se, but are products of regulatory transcription termination processes. For example, the expression of the P. aeruginosa amiLEBCRS operon is regulated by aliphatic amides via an antitermination mechanism, as discussed and illustrated in volume 2 [40]. In this operon, the amiE gene encodes aliphatic amidase and the amiB and amiS genes are thought to function in amide transport [40]. The AmiR protein, a positive regulator, acts as an antiterminator by allowing transcription to proceed beyond the amiL (5 untranslated leader) sequence. Mutation in amiR leads to premature termination of transcription at the rho-independent terminator downstream of amiL, producing the 100-nucleotide AmiL leader RNA [40]. It is assumed that the AmiR protein binds to the leader sequence and thereby interferes with the formation of the stem–loop structure of the amiL terminator. The AmiC protein acts as a negative regulator and senses aliphatic amides by binding them. Accordingly, amiC disruption leads to constitutive amidase synthesis. In the absence of inducing aliphatic amides, AmiC forms a complex with AmiR, which occludes the antitermination activity of AmiR. In the presence of an inducer, the AmiR-AmiC complex is dissociated, which allows antitermination activity of AmiR [40, 41]. AmiL RNA was recovered from non-induced P. aeruginosa cells as an sRNA bound to Hfq [16]. It is possible that Hfq binds to the AU-rich sequence upstream of the amiL terminator; however, the biological significance of the observed Hfq-AmiL interaction is not clear. Artificial overexpression of amiL
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did not reveal significant effects on the transcriptome and proteome in P. aeruginosa (E. Sonnleitner, unpublished data), suggesting that AmiL does not have global regulatory functions. The 2315 sRNA of P. aeruginosa (Table 1.1) is a homologue of SroG of E. coli [11, 15]. These sRNAs are part of a riboswitch mechanism, in which premature termination of transcription in a 5 untranslated leader mRNA results in the formation of the sRNAs. The particular riboswitch producing SroG is characterized by a conserved RNA structure element termed RFN [42]. Flavins such as FMN repress the expression of riboflavin biosynthetic (rib) genes by binding to the RFN element and thereby causing premature termination of transcription in the 5 untranslated rib leader mRNAs. In P. aeruginosa, the 2315 sRNA (∼180 nucleotides) is generated from the ribC leader sequence [15].
1.3.6 sRNAs Sequestering Proteins of the RsmA Family RsmY and RsmZ are two sRNAs of P. aeruginosa whose roles have been studied extensively in the context of quorum sensing regulation and virulence factor expression. The sequences and predicted secondary structures of RsmY and RsmZ of strain PAO1 have been given in volume 2 [43] and aspects of their expression via the GacS/GacA two-component system have been discussed in volume 5 [44, 45]. In the following, we will briefly review the salient features of the regulatory network involved. In P. aeruginosa, quorum sensing involves at least three signal molecules, i.e. N-(3-oxododecanoyl)-homoserine lactone (the reaction product of the LasI enzyme and principal activator of the LasR transcription factor), Nbutanoyl-homoserine lactone (the reaction product of the RhlI enzyme and principal activator of the RhlR transcription factor), and the Pseudomonas quinolone signal (PQS), which activates the PqsR transcription factor [44, 46]. Quorum sensing positively regulates biofilm polysaccharide biosynthesis and the expression of a range of extracellular virulence factors such as exotoxin A, lytic enzymes, and toxic secondary metabolites. In addition, quorum sensing negatively controls the type III secretion system (TTSS) and pilus formation. Several global regulators including the GacS/GacA two-component system influence the expression of quorum sensing [44, 47]. Mutations in either gacS (sensor kinase) or gacA (response regulator) strongly attenuate virulence of P. aeruginosa for different host organisms including burnt mice, nematodes, and insects [48, 49]. The activity of the GacS/GacA system is modulated by two sensor kinases, RetS and LadS [50, 51, 52]. RetS acts as an antagonist of GacS and evidence for the inhibitory activity of RetS has been obtained both in vivo and in vitro [50, 51]. A retS null mutant exhibits a small colony phenotype due to strong cell-cell aggregation, overproduction of extracellular polysaccharides, enhanced biofilm formation, lack of the TTSS and reduced twitching motility due to downregulation of type IV pili [50, 53, 54]. By contrast, when LadS is mutationally inactivated, biofilm formation is reduced and TTSS expression is enhanced [52]. Evidence for cooperation between LadS and GacS comes from studies in a related bacterium, P. fluorescens CHA0 [55, 56]. These
1
Small RNAs of Pseudomonas spp.
17
GacS
RetS
LadS
Cytoplasmic membrane
Phosphorelay GacA Transcriptional activation
RsmY
RsmZ
Inhibition Translational activation TTSS Mobility
RsmA
Translational repression Biofilm QS
Fig. 1.4 In P. aeruginosa, RsmA, a translational regulator, acts as an activator of the type III secretion system (TTSS), pilus formation and mobility and as a repressor of biofilm formation and quorum sensing (QS). Under conditions of high cell population densities and slow growth, the expression of two sRNAs, RsmZ and RsmY, is elevated. These two sRNAs bind to the RsmA protein and inhibit its function. Both sRNAs are under the control of the GacS-GacA two-component system, where GacS is the sensor protein and GacA the response regulator. Phosphorylation of GacS is regulated by two membrane-bound sensors, RetS and LadS, which physically interact with GacS. RetS blocks GacS autophosphorylation, whereas LadS appears to stimulate this activity
findings can be rationalized by a model (Fig. 1.4) which stipulates that RetS and LadS interact directly with GacS and that the combined input from the three sensors determines the level of phosphorylation of the response regulator GacA. The signals and environmental cues that activate the three sensors are largely unknown [45, 57]. In P. aeruginosa, RsmY and RsmZ are expressed under strict positive control by GacA, which is an activator when it is in the phosphorylated state [51]. An rsmY rsmZ double mutant has the same phenotype as a gacA-negative mutant. Both RsmY and RsmZ antagonize the action of the RsmA protein, by avidly binding to it [17, 19]. RsmA (acronym for regulator of secondary metabolism) is a small dimeric RNA-binding protein that represses translation of target mRNAs, many of which are involved in virulence factor expression [58, 59]. RsmA is a member of a large protein family found in > 150 bacterial species, including E. coli where the homologue is called CsrA (acronym for carbon storage regulator [60]). In P. aeruginosa, RsmA also indirectly activates the promoters of the rsmY and rsmZ genes via an unknown mechanism [17, 19]. Hfq binds to RsmY and stabilizes it [61, 62].
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A proteomic analysis suggests that the RsmA/RsmY/RsmZ triad mediates virtually all important functions of the GacS/GacA two-component system [17]. Loss of virulence in gacS/gacA mutants correlates with markedly diminished biofilm formation and with reduced expression of extracellular virulence factors that are positively regulated by quorum sensing [17, 63]. On the one hand, the GacS/GacA system positively influences the expression of the quorum sensing machinery, in particular that of the rhlI gene, which encodes the enzyme for the biosynthesis of the quorum sensing signal N-butanoyl-homoserine lactone [17, 59, 63]. On the other hand, the GacS/GacA system regulates the expression of certain virulence factors directly at the level of translation [64], as has been shown for the Gac/Rsm signal transduction pathway in other γ-proteobacteria [65]. The first observation of RsmY sRNA was not made in P. aeruginosa, but in P. syringae pv. phaseolicola, a producer of phaseolotoxin and pathogen of bean (Table 1.2). Some phaseolotoxin-negative mutants, which were not mapped but presumably carried mutations in gacS or gacA, were functionally restored by multicopy suppression with a 0.4-kb locus termed TRR (for thermoregulatory region) [29]. Although the TRR product was not identified at the time of publication (1993), in retrospect it is evident that TRR codes for an RsmY-like sRNA [26]. Overexpression of TRR not only restores phaseolotoxin production in the mutant background but also overrides temperature control of phaseolotoxin production. The wild type synthesizes the toxin at 20◦ C, but not at 28◦ C, whereas the suppressed mutants produce the toxin at both temperatures [29]. Similarly, initial evidence for RsmZ sRNA came from multi-copy suppression of gacA and gacS mutations (Table 1.2) in P. fluorescens F113, a soil bacterium that colonizes the roots of different crop plants and protects these from fungal pathogens [30]. The suppressor locus (termed prrB) specified an sRNA closely related to RsmZ. Overexpression of PrrB restored the production of typical antifungal GacA-dependent metabolites, i.e., 2,4-diacetylphloroglucinol (DAPG) and hydrogen cyanide (HCN), to gacS and gacA mutants of strain F113. However, mutation of the prrB gene had relatively minor effects on DAPG and HCN production, suggesting that PrrB may not be the only GacA-dependent regulatory sRNA in strain F113 [30]. A more complete picture of GacA-dependent sRNAs has emerged from studies of P. fluorescens CHA0. Like strain F113, strain CHA0 has biocontrol properties; they depend on a blend of antifungal secondary metabolites (including DAPG, HCN, pyoluteorin and pyrrolnitrin), extracellular lytic enzymes and poorly characterized elicitors of induced systemic resistance, i.e., a mechanism that renders the host plant less susceptible to pathogens [57, 66]. Biocontrol activity of strain CHA0 is lost in gacS or gacA null mutants, because most secondary metabolites and extracellular enzymes are produced at very low levels in such mutants, by comparison with the wild type [67–69]. In strain CHA0, the GacS/GacA twocomponent system is required for the expression of RsmY (118 nt) and RsmZ (127 nt) as well as that of a third sRNA termed RsmX (119 nt). A conserved upstream sequence element (consensus TGTAAGN6 CTTACA), which might be a GacA binding site, is found in the rsmX, rsmY and rsmZ promoters as well as in promoters of
1
Small RNAs of Pseudomonas spp.
19
GacA-controlled sRNA genes in other γ-proteobacteria [26–28, 43]. The three sRNAs have a high affinity for the RNA-binding protein RsmA and for its paralogue RsmE in strain CHA0 [70]. Both proteins act as translational repressors of target mRNAs carrying, e.g., hcnA (for an HCN synthase subunit), aprA (for an extracellular metalloprotease) or phlA (for a subunit of the DAPG biosynthetic enzyme complex) [70, 71]. Together, RsmX, RsmY and RsmZ sequester the regulatory proteins RsmA and RsmE. This allows target mRNAs to be translated and hence biocontrol factors to be synthesized. Deletion of rsmX, rsmY and rsmZ mimics the gacS or gacA mutant phenotype, whereas mutation of only one or two sRNA genes has smaller effects [28]. Thus, the three sRNAs can functionally replace one another. Nevertheless, their expression patterns are different during growth in batch culture and therefore their physiological functions might be distinct [28]. In addition to regulating exoproduct formation, the GacS/GacA two-component system positively controls the expression of the stress sigma factor σS (RpoS) and, as a consequence, the resistance of strain CHA0 to oxidative stress in stationary phase [72]. The expression of rsmX, rsmY and rsmZ is temperature-sensitive; the temperature-responsive element has been identified as RetS [55]. It appears that the inhibitory effect of RetS on GacS is stronger at 35◦ C than at 30◦ C, resulting in decreased GacA activity and, therefore, in diminished biocontrol factor expression at the higher temperature [55]. In the plant-pathogen P. syringae pv. tomato DC3000 there is computational evidence for three GacA-controlled sRNAs (RsmX, RsmY, RsmZ), two of which have been demonstrated experimentally, and for four RsmA-like RNA-binding proteins [32, 58, 73]. In this organism, the GacS/GacA two-component system is essential for the production of the phytotoxin coronatine [73]. RsmX, RsmY and RsmZ sRNAs of Pseudomonas spp. all have multiple GGA motifs in unpaired regions; about half of these motifs are present in an extended form, ANGGA (where N is any nucleotide). The GGA motifs are essential for function, as demonstrated by mutational analysis of RsmY of P. fluorescens [74], the construction of artificial sRNAs containing multiple GGA motifs [75] and binding studies with RsmY of P. aeruginosa [62]. A fully conserved ACANGGANGU motif was found in synthetic RNAs that were enriched for high-affinity binding to E. coli CsrA protein [76]. This motif can adopt a structure consisting of an ANGGAN loop placed on a 2-bp stem and mutational analysis has shown that such a hairpin is important for CsrA binding [76]. Similar motifs can be found in numerous target mRNAs whose translation is regulated by the Gac/Rsm regulatory pathway in γ-proteobacteria [60, 65, 77]. For instance, the major RsmA/RsmE binding site in the 5 leader region of hcnA mRNA of P. fluorescens contains a similar sequence, UCACGGAUGA, which encompasses the hcnA ribosome binding site. The solution structure of this target mRNA segment, in a complex with the RsmE homodimer, shows that RsmE makes important contacts with the conserved nucleotides in the ANGGAN loop and helps the flanking nucleotides to form a short stem [78]. This mRNA configuration blocks ribosome access. By contrast, when sRNAs of the RsmX/RsmY/RsmZ-type are abundant in the cell, they sequester the RsmA and RsmE proteins and the unfolded target mRNA becomes
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accessible to ribosomes. This enables translation of the target mRNA to occur [65, 77, 78]. In many γ-proteobacteria, the GacS/GacA two-component system and RsmA/CsrA-type proteins are conserved and can be identified by BLAST searches [58, 67]. Furthermore, the mechanistic principle of GacA-controlled sRNAs titrating the RsmA/CsrA-type proteins also appears to be conserved [60, 65]. However, in this case BLAST-driven homology searches have a limited scope: they will usually identify RsmX, RsmY and RsmZ homologues – if present – in Pseudomonas spp., but not the functionally equivalent sRNAs of other bacterial species. Similarly, the CsrB and CsrC sRNAs of E. coli, which sequester the CsrA protein, have homologues in other enteric bacteria, but it is not clear whether these sRNAs actually share common ancestors with the RsmX, RsmY and RsmZ sRNAs of pseudomonads. Therefore, to predict GacA-dependent sRNAs in γ-proteobacteria, Kulkarni et al. [32] have used an algorithm that combines the putative GacA-binding site (which is conserved in the upstream promoter region) with multiple ANGGA or AGGA motifs present in the downstream transcribed region. In this way, GacAdependent sRNAs could be identified or predicted in 23 species of γ-proteobacteria [32].
1.3.7 RgsA sRNA Involved in Oxidative Stress Response The genomic search for sRNAs in P. aeruginosa conducted by González et al. [15] reveals an sRNA (1698) whose expression depends strongly on the stress sigma factor RpoS and partially on the response regulator GacA. This sRNA, which also appears in the lists of Livny et al. [14] and Sonnleitner et al. [16], has been termed RgsA (for regulation by GacA and stress). A homologous sRNA occurs in P. fluorescens CHA0 and its regulation by RpoS and GacA is similar. RgsA of strain CHA0 has a single unpaired ANGGA motif and appears unable to sequester RsmA and RsmE. Deletion of the rgsA gene in strain CHA0 results in enhanced sensitivity to hydrogen peroxide, via an unknown mechanism [15]. RgsA of P. aeruginosa binds Hfq (E. Sonnleitner, unpublished results) and might engage in base-pairing interactions with mRNAs. However, such target mRNAs still need to be identified.
1.3.8 Fur-Regulated PrrF sRNAs Iron is an essential nutrient for P. aeruginosa, like for most bacteria. Abundance of iron in the environment leads to repression of iron acquisition functions (e.g., via siderophores) and to derepressed synthesis of the iron storage protein bacterioferritin (bfrB) and iron-containing enzymes such as superoxide dismutase (sodB) and succinate dehydrogenase (sdhABCD) in P. aeruginosa. The Fur repressor mediates these effects [79]. Together with iron, Fur represses the transcription of key iron acquisition genes as well as that of the tandem prrF1 and prrF2 genes. The
1
Small RNAs of Pseudomonas spp.
21
HIGH IRON sodB mRNA 5’
RBS Start codon CCGCGCCAGUCCUGACCUGAGGAAGAAUAGGAGAGACACCAUG 3’
Translation of sodB
Fe2+
Fe2+
Fur
Fur
-35 -10
prrF1
-35 -10
Ribosome
Transcription prrF2
-35 -10
prrF1
-35 -10
prrF2
Blocked transcription
PrrF core 3’ 5’
GGCACUAAUCGGACU--ACUCCUCUAUUAGACUUCUC 5’ ||| | | ||||| |||||| | ||| |||||| CCGCGCCAGUCCUGACCUGAGGA-AGAAU-AGGAGAGACACCAUG
sodB mRNA
Blocked translation/degradation
RBS
3’
Start codon
LOW IRON
Fig. 1.5 Under iron-replete conditions (shown in red), Fe2+ binds to and activates the Fur repressor. The activated Fur-Fe2+ protein recognizes a specific sequence (Fur box) within the promoter regions of the prrF1 and prrF2 genes and blocks transcription of these two sRNA genes. It is controversial whether repression is brought about by one or two Fur-Fe2+ dimers interacting with a Fur box [80]. During iron depletion (shown in blue), Fur is not active and the prrF1 and prrF2 genes are expressed. PrrF1 and PrrF2 sRNAs both bind to sodB mRNA in the vicinity of the ribosome binding site (RBS). This base-pairing event prevents ribosome binding and subsequent translation of sodB [13]. Such conditions might lead to degradation of the sodB transcript
products of the prrF1 and prrF2 genes are two highly similar sRNAs of about 110 nucleotides each [13]. By a base-paring mechanism, these sRNAs occlude the ribosome binding site of sodB mRNA and hence inhibit translation initiation (Fig. 1.5). As a result, when iron is replete, the two sRNAs are not expressed and sodB translation can proceed. When iron is limiting, the sRNAs are produced and sodB mRNA translation is blocked [13]. The PrrF1 and PrrF2 sRNAs are thought to act similarly on a range of other mRNAs and are reminiscent of the Fur-repressible sRNAs RyhB of E. coli and FsrA of Bacillus subtilis, which have analogous functions in their organisms, however, without showing sequence homology with PrrF1 and PrrF2 of P. aeruginosa [81, 82]. Recently, evidence for a PrrF2 homologue has been obtained in P. syringae [83]. The extensive regulatory effects of PrrF1 and PrrF2 also provide a link between iron and quorum sensing regulation in P. aeruginosa. This occurs when these sRNAs cause translational repression of the antABC genes, which code for enzymes degrading anthranilate to catechol. Thus, under iron-limiting conditions, the degradation of anthranilate is hindered in the wild type and cells can use anthranilate as aprecursor
22
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for the synthesis of PQS, a late quorum sensing signal. In a prrF1,2 mutant, PQS synthesis is decreased and can be restored by the addition of anthranilate [84]. Following these observations, one would expect that, under iron-replete conditions, PQS levels should decrease in the wild type as well – however, the opposite result was found (PQS levels increased), suggesting that an alternative pathway may supply anthranilate in high iron conditions. Preliminary results suggest that the PrrF1 and PrrF2 sRNAs also regulate the expression of two aconitases [84] and that of other enzymes of the citric acid and glyoxylic acid cycles [79]. These observations could explain why the prrF1,2 mutant has problems growing on succinate, which is an excellent carbon source for the P. aeruginosa wild type [79].
1.3.9 PhrS sRNA, an Anaerobic Regulator of PQS Synthesis P. aeruginosa preferentially respires with oxygen as the terminal electron acceptor, but when oxygen concentrations are low and nitrate is present, the organism uses nitrate as the electron acceptor in a process called denitrification [85]. Aerobic respiration and denitrification are not mutually exclusive; rather they overlap when cells experience low oxygen concentrations [86]. P. aeruginosa is particularly well adapted to such hypoxic conditions [87, 88]. The oxygen-sensing transcription factor ANR plays a key role in the adaptation to hypoxic and anaerobic conditions [89]. ANR is essential for all steps of denitrification and also activates the expression of the cbb3-2 oxidase, which is instrumental under hypoxic conditions and appears to have a high affinity for oxygen [87, 90–92]. ANR recognizes a conserved binding site (consensus TTGATN4 ATCAA), which is typically located at -41 bp from the transcription start site in ANR-dependent promoters [93]. The PhrS sRNA (alternatively termed P20 or 1887) [14–16] modulates PQS synthesis during the transition from aerobic to hypoxic conditions in batch culture of P. aeruginosa. The phrS gene is strongly induced from an ANR-dependent promoter when oxygen becomes limiting (E. Sonnleitner et al., unpublished results). This regulation may stabilize PQS production in environments of fluctuating oxygen concentrations. Anaerobic conditions abrogate PQS formation; an important reason for this may be the fact that the last step of PQS biosynthesis, the hydroxylation of 2-heptyl-4(1H)-quinolone (HHQ), requires oxygen. Under limiting oxygen conditions, PQS synthesis is low [46, 94]. This tendency appears to be countered by enhanced phrS expression. PhrS contains an open reading frame of 37 codons, which is translated [16]. The resulting small protein is predicted to be anchored in the cytoplasmic membrane by a single transmembrane helix. The function of this protein is unknown. Interestingly, PhrS is the first example of a coding sRNA of Pseudomonas spp. Recent results indicate that small proteins of 16–50 amino acids are common in E. coli and that more than half of them are membrane-bound. However, their functions are completely unknown [95].
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1.4 Concluding Remarks In this overview, we have presented the proverbial tip of the iceberg: the number of sRNAs in Pseudomonas spp. is certainly larger than that reported in the current summary of Table 1.1. P. aeruginosa has a larger genome than E. coli; by simple extrapolation, we can assume that P. aeruginosa should have as many sRNAs as E. coli, if not more. The tools to find new sRNAs in Pseudomonas spp. exist and will undoubtedly be put to application. The example of the three seemingly redundant sRNAs (RsmX, RsmY, RsmZ) in the Gac/Rsm signal transduction pathway of P. fluorescens illustrates one of the common problems, namely that several sRNA genes need to be inactivated simultaneously for phenotypic effects to become apparent. Not always are functionally related sRNA genes clustered and can be knocked out by a single deletion, as in the case of the P. aeruginosa prrF1,2 genes. Often, sRNA genes are scattered over the bacterial chromosome(s) and, unless they have a common property such as a conserved promoter element, they are hard to find. A major challenge that lies ahead is to unravel the functions of new sRNAs and to obtain a coherent picture of the impact that sRNAs have on cell physiology. Transcriptomic and proteomic approaches, in combination with mutant or overexpression strategies, are clearly very useful in this respect. Considering the metabolic versatility of pseudomonads, we predict that sRNAs may well have important roles in fine-tuning metabolic functions in these bacteria. Acknowledgments This work was supported by the Swiss National Foundation, the ErwinSchrödinger Research fellowship (to E.S.) and a genomics project of the University of Lausanne. We thank Cornelia Reimmann for critically reading the manuscript.
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Chapter 2
2-Alkyl-4(1H)-Quinolone Signalling in Pseudomonas aeruginosa Matthew P. Fletcher, Stephan Heeb, Siri Ram Chhabra, Stephen P. Diggle, Paul Williams, and Miguel Cámara
2.1 Introduction to 2-Alkyl-4(1H )-quinolone Quorum Sensing in Pseudomonas aeruginosa Quorum sensing (QS) is the process whereby bacteria co-ordinate gene expression at the population level. This process is usually mediated by the production of a signal molecule, or “autoinducer”, of which the concentration in the extracellular environment is related to the population density of the bacteria producing the signal. The QS system of Pseudomonas aeruginosa consists of two linked sets of genes, coding for proteins that generate and utilise N-acyl-homoserine lactones (AHLs) as signal molecules. The first of these sets is termed las and consists of the N-(3-oxododecanoyl)-L-homoserine lactone (3-oxo-C12 -HSL) synthase LasI [1, 2], and the transcriptional regulator LasR [3]. The second of these systems is termed rhl and comprises the N-butanoyl-L-homoserine lactone (C4 -HSL) synthase RhlI [4, 5], and the transcriptional activator RhlR [6]. 3-Oxo-C12 -HSL accumulates within the growing bacterial population until molecule it reaches a threshold concentration resulting in the activation of LasR which in turn activates numerous virulence factors such as elastase [1], pyoverdin [7], lasI (creating a positive feedback loop) [8] and the rhl QS system [9]. The rhl system with the aid of C4 -HSL activates transcription of rhamnolipid biosurfactants [10], cytotoxic lectins, pyocyanin and elastase, amongst other virulence factors [11]. The QS control of gene expression is not always system specific, with many genes being activated by both AHLs to differing extents [12]. The recognition of QS target promoters by LasR-3-oxo-C12 -HSL and RhlR-C4 -HSL usually depends on the recognition of conserved palindromic sequences, termed “las-rhl boxes” which function as binding sites for either or both regulators [13, 14]. The above QS hierarchy is also further modulated by the existence of other regulators, for reviews see Williams and Cámara [15] and von Bodman et al. [16]. M. Cámara (B) School of Molecular Medical Sciences, Centre for Biomolecular Sciences, University Park, University of Nottingham, Nottingham NG7 2RD, UK e-mail:
[email protected]
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In addition to the above QS systems, P. aeruginosa employs a non-AHL QS signalling system that is an integral part of the QS hierarchy [17]. This system utilises 2-alkyl-4(1H)-quinolone (AQ) molecules, of which the primary signals are 2-heptyl-3-hydroxy-4(1H)-quinolone (named the Pseudomonas quinolone signalPQS) [17] and 2-heptyl-4-hydroxyquinoline (HHQ) [18]. Other major molecules belonging to this family are 2-nonyl-3-hydroxy-4-quinolone (C9-PQS), 2-nonyl4-hydroxyquinoline (NHQ) and 2-undecyl-4-hydroxyquinoline (UHQ). 2-alkyl-4hydroxyquinoline N-oxide derivatives of AQs (AQNOs) have also been discovered, the main molecules being 2-heptyl-4-hydroxyquinoline N-oxide (HQNO), 2-nonyl4-hydroxyquinoline N-oxide (NQNO) and 2-undecyl-4-hydroxyquinoline N-oxide (UQNO) [18–20]. A related molecule, 2,4-dihydroxyquinoline (DHQ), has also been discovered [21, 22] although this is not technically an AQ as it contains no 2-alkyl chain. Figure 2.1 shows the structures of these molecules. In addition, P. aeruginosa produces over 50 different AQ congeners, many of which are produced only in small quantities and whose physiological function, if any, is unclear. AQs were first recognised as antibiotics and isolated (although not completely characterised) from P. aeruginosa by Hays et al. [23], who built on over half a century of previous work stretching back to the days of Louis Pasteur in the 1870s. At that time it had been demonstrated that if a highly virulent strain of a certain bacterium that caused severe or fatal disease in a test animal was co-inoculated with selected cultures of other bacterial species, then the test animal failed to develop signs of disease. This was subsequently confirmed at the turn of the century by Emmerich and Löw [24] who found that cell-free culture supernatants from P. aeruginosa, concentrated to a tenth of their original volume prevented Bacillus anthracis from causing disease in rabbits. They named this concentrated extract pyocyanase due to its supposed enzymatic properties. It later became apparent that the antibiotic activity of pyocyanase was not due to enzymes, but to lipid-soluble molecules present in the concentrated supernatant as the antibacterial activity of the concentrate was found to partition almost entirely into organic solvents. Despite this initial promise for clinical applicability, the use of pyocyanase was abandoned due to problems with commercial manufacture. However the compounds responsible for antibiotic activity were extracted, fractionated, isolated, crystallised and renamed as “Pyo” compounds [23] (also called “Pseudanes”). Pyo I-IV were found to have a high antibacterial activity against Gram-positive organisms but only a slight activity against Gram-negative bacteria. Also, while these compounds had been partially chemically and physically characterised and shown to be structurally related, their identities were partially solved by Wells et al. [25, 26]. Using ozonolysis and UV absorption spectra in comparison with synthetic standards, they identified Pyo Ib as HHQ, Pyo Ic as NHQ and Pyo III as a mono-unsaturated alkyl side chain variant of NHQ. The identity of Pyo II was revealed from concurrent studies of P. aeruginosa, which showed that the organism produced a substance that inhibited the action of the antibiotics streptomycin and dihydrostreptomycin [27]. This substance was isolated [28] and comparison of the UV spectrum of its reduction product with synthetic standards suggested that this inhibitor was a mixture of quinoline N-oxides, HQNO and NQNO. Subsequent structural analysis confirmed the inhibitor was indeed a
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AQs O
N H
OH
R
N
R
R= n-C5H11 ; 2-pentyl-4-hydroxyquinoline (PHQ) 2-pentyl-4-quinolinol (IUPAC) R= n-C7H15 ; 2-heptyl-4-hydroxyquinoline (HHQ) 2-heptyl-4-quinolinol (IUPAC) R= n-C9H19 ; 2-nonyl-4-hydroxyquinoline (NHQ) 2-nonyl-4-quinolinol (IUPAC) R= n-C11H23 ; 2-undecyl-4-hydroxyquinoline (UHQ) 2-undecyl-4-quinolinol (IUPAC)
R= n-C5H11 ; 2-pentyl-4(1H)-quinolone R= n-C7H15 ; 2-heptyl-4(1H)-quinolone R= n-C9H19 ; 2-nonyl-4(1H)-quinolone R= n-C11H23 ; 2-undecyl-4(1H)-quinolone
PQS and its analogues O
OH OH
N H
OH
R
N
R= CH3 ; 3-hydroxy-2-methyl-4(1H)-quinolone (C1-PQS)
R
R= CH3 ; 2-methyl-3,4-dihydroxyquinoline 2-methyl-3,4-quinolinediol (IUPAC) R= n-C7H15 ; 2-heptyl-3,4-dihydroxyquinoline 2-heptyl-3,4-quinolinediol (IUPAC) R= n-C9H19 ; 2-nonyl-3,4-dihydroxyquinoline 2-nonyl-3,4-quinolinediol (IUPAC)
R= n-C7H15 ; 2-heptyl-3-hydroxy-4(1H)-quinolone (PQS) R= n-C9H19 ; 3-hydroxy-2-nonyl-4(1H)-quinolone (C9-PQS)
AQNOs OH
N O
O
R
N
R
OH
R= n-C7H15 ; 2-heptyl-4-hydroxyquinoline N-oxide (HQNO)
R= n-C7H15 ; 2-heptyl-1-hydroxy-4(1H)-quinolone
R= n-C9H19 ; 2-nonyl-4-hydroxyquinoline N-oxide (NQNO) 4-hydroxy-2-nonylquinoline N-oxide (IUPAC) R= n-C11H23 ; 2-undecyl-4-hydroxyquinoline N-oxide (UQNO) 4-hydroxy-2-undecylquinoline N-oxide (IUPAC)
R= n-C9H19 ; 2-nonyl-1-hydroxy-4(1H)-quinolone R= n-C11H23 ; 2-undecyl-1-hydroxy-4(1H)-quinolone
DHQ O
N H
OH
OH
OH
2-hydroxy-4(1H)-quinolone
N H
O
4-hydroxy-2(1H)-quinolone
N
OH
2,4-dihydroxyquinoline (DHQ) 2,4-quinolinediol (IUPAC)
Fig. 2.1 Structure, IUPAC names and abbreviations of AQ molecules synthesised by P. aeruginosa. Both the tautomeric lactam and phenolic forms of each molecule are shown. Arrows indicate the equilibrium of these molecules as would exist under physiological conditions and pH. Where more than one name exists for a molecule, the correct IUPAC designation is indicated. The IUPAC designation is not necessarily the name that will be adopted in this review (see section on nomenclature)
mixture of three N-oxides- HQNO, NQNO produced in the approximate ratio of 2:1 respectively with a trace of 2-undecyl-4-hydroxyquinoline N-oxide (UQNO) [29]. These compounds were also capable of inhibiting the cytochrome systems of heart
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muscle and the growth of bacteria [30] such as Staphylococcus and Bacillus spp. It was around this time that PQS was first isolated, identified and chemically characterised, bearing the name “Substance B” [31]. A tentative, method of synthesis of these compounds was suggested from condensation of anthranilate with β-keto fatty acids, releasing CO2 and water [29]. Later studies using feeding of radiolabelled isotopes of these compounds to P. aeruginosa and subsequent analysis of their degradation products confirmed that these molecules were formed by this method [32, 33]. Notwithstanding sporadic observation at various intervals by different researchers, usually for their properties as antibiotics or their presence in P. aeruginosa infection, the true significance of AQs to the organism was not yet appreciated. PQS was first described as a QS signal molecule by Pesci et al. [17], who found that addition of spent culture media extract from a P. aeruginosa wild type (PAO1) induced expression of lasB (elastase) in a PAO1 lasR mutant more than by adding 3-oxo-C12 -HSL or C4 -HSL. The mutant was AHL deficient and this therefore strongly suggested the presence of a third signal in this organism. Solvent extraction, HPLC separation and mass spectrometry (MS) identified the molecule as PQS. This AQ thus represented a new class of signal molecules, whose mechanism of regulation and action was linked to AHL-based QS. AQs were initially thought to be produced primarily in stationary phase. Using lasB expression as an indirect way of monitoring PQS production, negligible amounts were detected in late log phase cultures and maximal production of PQS was detected in late stationary phase after 30–42 h [34]. However, two other independent studies on the timing of PQS production, using more sensitive and direct detection methods, revealed that substantial levels of PQS are produced in logarithmic/early stationary phase cultures. Using a stable isotope dilution method coupled to mass spectrometry (MS) in P. aeruginosa strain PA14, PQS and HQNO were detected near the end of the log phase and at maximal levels at the onset of stationary phase, reaching a final concentration of 16 μM [35]. A second study using thin layer chromatography (TLC) in conjunction with a synthetic PQS standard in P. aeruginosa strain PAO1 estimated that at the onset of stationary phase, PQS concentrations were approximately 5–10 μM, increasing to 25 μM in late stationary phase [36]. It would appear therefore that PQS is produced maximally at late log/early stationary phase, and accumulate to higher concentrations later in growth. In contrast, in PA14 HHQ levels rise initially until around 6 h, but then plateau and begin to drop, presumably as this HHQ is converted into PQS [18].
2.2 Nomenclature and Abbreviations The structures, IUPAC based nomenclature and abbreviations of all the major AQs produced by Pseudomonas aeruginosa are summarised in Fig. 2.1. To help the reader, some of the non-IUPAC names that have been used to describe these same molecules in the scientific literature have also been included. AQs,
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2-alkyl-4(1H)-quinolones (lactam form) are tautomeric with 2-alkyl-4hydroxyquinolines (phenolic form), of which the predominance of one form over the other is pH dependent [37–39]. For example, it has been demonstrated using pKa values for 2-methyl-3-hydroxy-4(1H)-quinolone (C1-PQS) that over physiological pH ranges, the neutral 4-quinolone form is the predominant tautomer [40]. These tautomeric forms are shown in Fig. 2.1, with their relative ratios indicated by the arrows. Ideally, for consistency and structural accuracy, nomenclature and abbreviations based on only one tautomeric form should be uniformly adopted. However, this poses difficulties for a review article, since in the available scientific literature, individual laboratories have subjectively referred to these molecules in either one form or the other. For example, even the two arguably “lead” molecules of the class in this field- PQS and HHQ, are inconsistent with each other with reference to naming and abbreviation; Pseudomonas quinolone signal (PQS), 2-heptyl-4-hydroxyquinoline (HHQ). The nomenclature and abbreviations used in this review therefore amount to a compromise between what is technically correct, taking into account IUPAC designations and structural predominance due to physiological pH, and also what is the prevalent terminology utilised in the scientific literature for each molecule. Hence, the abbreviation PQS (Pseudomonas quinolone signal; 2-heptyl-3-hydroxy4(1H)-quinolone) has been maintained and the alkyl side chain variants of this molecule abbreviated by the number of carbon atoms in the side chain e.g. C1-PQS, C9-PQS. The designation HHQ for 2-heptyl-4-hydroxyquinoline has also been maintained, and the other alkyl side-chain derivatives have been abbreviated accordingly (e.g. PHQ- 2-pentyl-4-hydroxyquinoline etc). It should be noted that AQNO series of compounds (e.g. HQNO, NQNO; 2-heptyl-4-hydroxyquinoline N-oxide, 2-nonyl-4-hydroxyquinoline N-oxide) can adopt the 2-alkyl-1-hydroxy-4(1H)-quinolone form (Fig. 2.1) but not at physiological pH. 2,4-Dihydroxyquinoline (DHQ) can exist in both 4-hydroxy-2(1H)quinolone (predominant at physiological pH) and 2-hydroxy-4(1H)-quinolone tautomeric forms (Fig. 2.1) but to avoid confusion and to conform to the literature citations, DHQ is used to denote this molecule in this chapter. To further help the reader, Table 2.1 shows a list of the proposed nomenclature of AQs that are used in this chapter, along with abbreviations. Included in this table are the associated synonyms that have been used to describe these same molecules in the scientific literature.
2.3 AQ Spectrum Produced by P. aeruginosa The AQ biosynthetic machinery of P. aeruginosa enables this organism to produce a diverse range of AQs. An early study used gas chromatography and electron capture mass spectrometry (MS) to identify over twenty different AQs [20]. HHQ was the most prevalent, then NHQ. Also present were homologues of these compounds
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Table 2.1 Nomenclature and abbreviations of AQs used in this review. Intended for quick reference; synonyms of these molecules used elsewhere in the scientific literature are also shown Proposed nomenclature
Synonyms
2-alkyl-4(1H)-quinolone (AQ)
2-alkyl-4-hydroxyquinoline (AHQ) 4-hydroxy-2-alkylquinoline (HAQ) 2-alkyl-4-hydroxyquinoline N-oxide (AQNO) 4-hydroxy-2-alkylquinoline N-oxide 2-alkyl-1-hydroxy-4(1H)-quinolone 2-heptyl-3-hydroxy-4(1H)-quinolone 2-heptyl-3,4-dihydroxyquinoline Pseudomonas quinolone signal (PQS) 2-heptyl-3,4-quinolinediol 3-hydroxy-2-nonyl-4(1H)-quinolone 3,4-dihydroxy-2-nonylquinoline (C9-PQS) 2-nonyl-3,4-quinolinediol 2-pentyl-4-hydroxyquinoline (PHQ) 2-pentyl-4(1H)-quinolone 4-hydroxy-2-pentylquinoline 2-pentyl-4-quinolinol 2-heptyl-4-hydroxyquinoline (HHQ) 2-heptyl-4(1H)-quinolone 4-hydroxy-2-heptylquinoline 2-heptyl-4-quinolinol 2-nonyl-4-hydroxyquinoline (NHQ) 2-nonyl-4(1H)-quinolone 4-hydroxy-2-nonylquinoline 2-nonyl-4-quinolinol 2-undecyl-4-hydroxyquinoline (UHQ) 2-undecyl-4(1H)-quinolone 4-hydroxy-2-undecylquinoline 2-undecyl-4-quinolinol 2-heptyl-4-hydroxyquinoline N-oxide 4-hydroxy-2-heptylquinoline N-oxide (HQNO) 2-heptyl-1-hydroxy-4(1H)-quinolone 2-nonyl-4-hydroxyquinoline N-oxide (NQNO) 4-hydroxy-2-nonylquinoline N-oxide 2-nonyl-1-hydroxy-4(1H)-quinolone 2-undecyl-4-hydroxyquinoline N-oxide 4-hydroxy-2-undecylquinoline N-oxide (UQNO) 2-undecyl-1-hydroxy-4(1H)-quinolone 2,4-dihydroxyquinoline (DHQ) 4-hydroxy-2(1H)-quinolone 2-hydroxy-4(1H)-quinolone 2,4-quinolinediol
containing saturated and mono-unsaturated alkyl chains varying from one to thirteen carbons in length, and the N-oxides HQNO and NQNO. Two more recent studies used electrospray ionisation and liquid chromatography MS to obtain the mass spectra of over 50 different AQs. These mainly consisted of PQS, HHQ, HQNO and NHQ, with other saturated and mono-unsaturated alkyl side chain length homologues [35, 19]. Many of these latter compounds are produced at seemingly biologically insignificant levels however, perhaps due to a lack of fidelity of the AQ biosynthetic proteins for the different lengths of β-keto fatty acids present during AQ manufacture, rather than for any specific biological function. In addition, a new metabolite, identified as 2,4-dihydroxyquinoline (DHQ), was found in cultures of both Pseudomonas aeruginosa and Burkholderia thailandensis [21, 22]. DHQ is neither a degradation product nor a precursor of AQs and the precise function
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of this molecule is unknown. However it was found that DHQ was able to inhibit the growth and cell viability of mouse lung epithelial MLE-12 cells [22], therefore DHQ may have a role in pathogenicity in lung infection.
2.4 Properties of AQs AQs are lipophilic molecules with a low aqueous solubility. PQS has a solubility of around 1 mg/L (∼5 μM) in water [35] and due to this hydrophobic nature a high proportion is associated with the outer membrane and membrane vesicles (MVs) of the bacterium [41]. It has been shown in vitro that rhamnolipid biosurfactants can increase the solubility of PQS [42] and that this increased solubility correlated with the ability of PQS to induce lasB expression (although too much rhamnolipid is apparently counter-productive, as PQS is possibly sequestered into micelles) [42]. However, whether rhamnolipids are utilised for this purpose in vivo is unknown. PQS initiates the formation of MVs and around 80% of total PQS produced by P. aeruginosa PA14 was reported to be contained within vesicles, in contrast with less than 1% of both 3-oxo-C12 -HSL and C4 -HSL [43]. The PQS contained within these vesicles was biologically active and could restore the production of pyocyanin in a PQS-deficient mutant, independent of the vesicles themselves. Mutants deficient in PQS production, but not other AQs were highly defective for vesicle formation, but vesicle formation did not rely on active growth or protein synthesis and would seem to be a spontaneous process in the presence of PQS. A mechanism of vesicle formation is suggested through PQS interaction with the acyl chain and 4 -phosphate of purified bacterial lipopolysaccharide [41]. This activity seems dependent on the 3-hydroxy group (since HHQ induces much less vesicle formation) and is optimal with C7 -alkyl chain moieties of PQS, although C5 , C3 alkyl side chain variants also induce MV formation [44, 41]. Interestingly, MVs can still be induced by PQS analogues lacking an acyl side chain moiety, suggesting that this group is dispensable. It was suggested packaging into MVs might serve to concentrate and protect PQS from catabolism by surrounding cells, although it is unclear at present if PQS is broken down to any significant extent.
2.5 Biosynthesis of AQs The genes responsible for AQ biosynthesis and response were identified by screening a transposon library for mutants displaying reduced pyocyanin production and termed pqsABCDE, pqsR, pqsH and pqsL [45, 46, 19]. In P. aeruginosa PAO1, these chromosomal genes are organised in an operon comprising pqsABCDE (PA0996-PA1000) which is adjacent to phnAB (PA1001-PA1002) and pqsR (PA1003) (see Fig. 2.3). The pqsH (PA2587) and pqsL (PA4190) genes are located elsewhere on the chromosome.
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The pqsR (also termed mvfR) gene encodes the transcriptional regulator of both the pqsABCDE operon and the phnAB genes. PqsR is a member of the LysR-type family of transcriptional regulators as it has a helix-turn-helix motif at the N-terminus with the first 280 amino acids sharing high similarity (62– 71%) to other LysR-type regulators [47, 48]. A mutation in this regulator in P. aeruginosa PA14 resulted in reduced pyocyanin, elastase, exoprotein and 3oxo-C12 -HSL production, abolition of phnAB, pqsABCDE expression and AQ biosynthesis together with the inability to cause disease in plants and animals [47, 18]. The pqsABCD genes are responsible for the production of the PQS precursor HHQ and structurally related compounds such as NHQ and UHQ, plus the N-oxide series of AQs such as HQNO and NQNO [18]. PqsA is an anthranilate coenzyme A ligase involved in the activation of anthranilate for AQ synthesis [49]. The pqsB and pqsC genes are predicted to encode proteins with functional similarity to β-ketoacyl-ACP (acyl-carrier protein) synthases. PqsD shares some sequence similarity to the E. coli initiation condensing enzyme FabH and its Cys-His-Asn active site moiety [50, 32, 33, 22]. Mutants in pqsA, pqsB and pqsD are devoid of AQs [36, 22]. PqsE has structural similarities to proteins from the zinc metallo-β-hydrolase superfamily. This family contains numerous hydrolytic enzymes that can perform various functions, such as β-lactamases, glyoxalases, AHL-lactonases and arylsulfatases. These proteins, and also PqsE, contain the conserved amino acid motif “HXHXDH”, responsible for binding to zinc. PqsE is involved in the cellular response to PQS since pqsE mutation or overexpression has effects on several virulence factors such as pyocyanin, lectin and HCN but does not affect AQ production [51, 36, 52, 46]. The PqsE substrate is not yet known (PqsE will be covered in further detail later in this section). The pqsH gene codes for a predicted FAD-dependent mono-oxygenase required for the conversion of the immediate PQS-precursor, HHQ, into PQS [18]. Mutants in pqsH do not produce 3-hydroxy-4(1H)-quinolones, but do produce other AQs [18]. The pqsL gene also encodes a putative mono-oxygenase, required for the synthesis of HQNO and associated N-oxides [19]. Addition of deuterated HHQ to a culture of P. aeruginosa wild type revealed overproduction of deuterated PQS, but not HQNO [18]. This suggests that HHQ is a precursor of PQS, but not of HQNO. Mutants in pqsL over-produce PQS compared to the wild type [45] suggesting that the AQ biosynthetic intermediates are diverted in these mutants towards the production of this molecule. Figure 2.2 presents a simplified scheme for the biosynthesis of AQs. A tentative pathway was first suggested, based on the condensation of anthranilate with β-keto fatty acids, releasing CO2 and water [29]. Subsequent radiolabelling studies also indicated that this was indeed the case [32, 33] and this was further confirmed by Bredenbruch et al. [50]. Using feeding experiments with isotope-labelled precursors and analysis of extracted AQs by MS and NMR, a second possible route of AQ biosynthesis via kynurenic acid and involving the reaction of anthranilate and orotic acid, was excluded [50]. DHQ biosynthesis requires only the PqsA and PqsD proteins [22]. Anthranilate is activated by PqsA to form anthraniloyl-coA [22].
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2-Alkyl-4(1H)-Quinolone Signalling in Pseudomonas aeruginosa O
O Glycolysis
37
OH
OH O
Pyruvic acid
N H
CO2H
AroCK HO
N H
O
OH OH
OH
PqsH
O
CO2H
CO2H
PQS
HHQ
Chorismic acid
Shikimic acid PhnAB O
PqsBCD? O
O
CO2H NH2
TrpEG
KynU
PqsA
OH NH2 Anthranilic acid
NH2 Kynurenine
S
CoA
NH2 Activated anthranilate
+
O
O
ACP S Acivated b-ketodecanoate TrpGDF
KynB
PqsBCD? PqsL
O
CO2H CO2H NH2
CHO N H N-Formylkynurenine
OH
NH2
KynA N H Tryptophan
N O
HQNO
Fig. 2.2 Proposed mechanism of synthesis of PQS, HHQ and HQNO in P. aeruginosa. AQs are derived from a condensation reaction between anthranilate and β-keto fatty acids. Anthranilate is derived from either the Trp/Phn or Kyn metabolic pathways using either chorismate or tryptophan as the substrate respectively. Anthranilate is firstly activated with Co-enzyme A (CoA) by PqsA. Anthranilate-CoA and an activated β-ketodecanoate are then condensed, possibly via the PqsBCD enzymes to HHQ, releasing CO2 and water. The mono-oxygenase PqsH then converts HHQ to PQS. HQNO is derived from the same starting products as HHQ but utilises the additional mono-oxygenase enzyme PqsL. HHQ is not a precursor for HQNO. ACP = Acyl Carrier Protein
This molecule is then transferred to the cysteine active site of PqsD (unactivated anthranilate does not transfer) and reacts with either malonyl-CoA or malonyl-ACP to form the intermediate compound 3-(2-aminophenyl)-3-oxopropanoyl CoA. This compound is short lived and undergoes an internal rearrangement to form DHQ. It is possible that some variation of this mechanism, using longer chain β-keto-fatty
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acids instead of malonyl-CoA or malonyl-ACP, is responsible for synthesis of the more prominent AQ molecules such as HHQ, although the functions of the PqsB and PqsC proteins in this process are still unclear. The substrates required for AQ biosynthesis have also been investigated. Two pairs of genes responsible for anthranilate biosynthesis had previously been described: phnAB [53], located adjacent to the pqs operon [46], and trpEG, which code for the genes involved in tryptophan biosynthesis [54]. TrpEG direct the synthesis of anthranilate from chorismate, which can then be utilised for tryptophan or AQ biosynthesis. A third source of anthranilate investigated by Farrow et al. [55] requires the homologues of the tryptophan-2,3-dioxygenase- KynA, KynB and KynU [56] to obtain anthranilate from tryptophan. They proposed that in rich media containing the aromatic amino acid tryptophan, the kynurenine pathway is the main source of anthranilate for PQS production whereas the phnAB genes supply anthranilate in minimal media in the absence of exogenous tryptophan. Interestingly, Palmer et al. [57] noted that raised PQS levels in certain human CF clinical isolates of P. aeruginosa correlated with the presence of aromatic amino acids in the growth medium. The expression of pqsA was raised in presence of tryptophan, phenylalanine and tyrosine. Serine (a non-aromatic amino acid) had little effect on pqsA expression. This may therefore be the pathway of anthranilate generation in a clinical context. By growing both a PQS-producing (PAO1) strain and a mutant unable to make PQS in the presence of 14 C heteroaromatic ring labelled anthranilate, Calfee et al. [58] showed that the majority of radioactivity was found in the AQ extract for those strains that could make PQS (PAO1-parent) (around 50%), compared to around 3–4% in those strains that could not. This suggested that the strains unable to make PQS could not convert anthranilate into AQs which was confirmed using TLC and autoradiography. PAO1 was then grown with increasing amounts of methyl anthranilate (a competitor of anthranilic acid as a precursor for PQS synthesis) and the supernatant was extracted for AQs and analysed on TLC [58]. PQS levels were reduced as methyl anthranilate concentration increased independent of any growth effects. Methyl anthranilate also inhibited elastase production by PAO1 in a concentration-dependent manner, with 1.5 mM of methyl anthranilate practically abolishing its production. This showed the necessity of anthranilate for PQS production in this growth medium, since an anthranilate competitor was able to decrease this production, with subsequent effects on virulence factor production. Bredenbruch et al. [50] also performed feeding experiments with isotope-labelled AQ intermediates such as N15 -anthranilate coupled with gas chromatography and MS to determine the exact chemical mechanism of AQ synthesis. The resulting AQs generated in these experiments contained around 66% of N15 , further demonstrating that anthranilate serves as a common precursor for AQs and that the heteroaromatic nitrogen in the quinolone ring originates from this molecule. Feeding unlabelled anthranilate and labelled 13 C-acetate to PAO1 Bredenbruch et al. [50] also showed that the heteroaromatic ring moiety of quinolone was formed from acetate. The resulting fragmentation pattern, confirmed with nuclear magnetic
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resonance spectroscopy indicated that the mechanism of this reaction was via a direct head-to-head reaction involving anthranilate and β-keto fatty acids derived from acetate. β-keto fatty acids are therefore essential precursors of AQ biosynthesis. It was suggested that there was a link between AQ production and rhamnolipid biosynthesis, which was interesting since rhamnolipids increase PQS solubility and are likely to be utilised in vivo partially for this purpose [42]. Rhamnolipids are composed of rhamnose and fatty acids of the same chain length as those involved in AQ synthesis. It was initially thought that the gene rhlG, encoding a potential β-ketoacyl-ACP reductase directed the incorporation of these fatty acids during rhamnolipid production [59, 60, 61] and could have participated in the provision of fatty acids utilised as a substrate for AQ biosynthesis [50]. However, recent studies have contradicted this, since a rhlG mutant was found to be unaltered in rhamnolipid production compared to the corresponding wild type [62] and the crystal structure was found to be inconsistent with the proposed fatty acid biosynthetic pathway [63]. Therefore, it would appear that RhlG is not involved in rhamnolipid and AQ biosynthesis. It appears that PQS is either the end product of the AQ synthetic pathway or is not substantially converted to another compound, as when labelled PQS was added to wild type P. aeruginosa cultures, no extra compounds were identified [18].
2.6 Regulation of AQ Production A simplified model of AQ regulation in P. aeruginosa is shown in Fig. 2.3.
2.6.1 AQs and the las/rhl QS Systems of P. aeruginosa Initial studies revealed that AQ production and regulation were linked to the las and rhl QS systems of P. aeruginosa because the las system was involved in AQ production and the rhl system in its biological effects [34, 17]. Further work elaborated upon this and revealed that the las system acts to increase production of AQs by raising the expression of both the pqsR and pqsA genes, whilst the rhl system negatively regulates AQ signalling [64, 65, 66]. There is a las/rhl box sharing 80% identity to that of the QS- regulated rsaL gene at -513 base pairs from the pqsR translational start site [64, 66]. Removal of this box greatly reduced pqsR expression, suggesting that this may constitute the LasR binding site. In line with this, a lasR mutant showed a reduction in pqsR expression to approximately 25% compared to the wild type [65]. Also, addition of 3-oxoC12 -HSL to E. coli expressing recombinant LasR and a pqsR::lacZ transcriptional fusion showed a great increase in pqsR expression [65], indicating that LasR-3oxo-C12 -HSL serves as the activator. A large reduction in pqsA expression in a lasI mutant has also been reported [64]. LasR also controls the expression of the pqsH gene [18, 46, 67]. A lasR mutant accumulates the HHQ series of compounds but
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3-oxo-C12-HSL
LasI
lasR
lasl
LasR
C4-HSL
RhlI
rhlI rhlR
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pqsA pqsB pqsC pqsD pqsE
PqsABCD
phnA phnB pqsR
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pqsH PqsH
O
O OH
N H
HHQ
N H
PQS
O
N H
PqsR
PqsR O OH N H
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1 kb
Fig. 2.3 Regulation of AQ production in P. aeruginosa. The las QS system positively regulates the expression of PqsR, which up-regulates the expression of pqsABCDE. PqsABCD synthesize HHQ which is converted to PQS by PqsH. pqsH expression is also positively controlled by las. Autoinduction occurs when either HHQ or PQS bind to PqsR and enhance the expression of the pqs operon. The rhl system represses the AQ system, although it is itself regulated by AQs. Arrowed lines represent positive regulation, blunt-ended lines represent negative regulation
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produces very little PQS early in growth, suggesting that it is lasR that controls this conversion [18], probably via the mono-oxygenase PqsH which is required for conversion of HHQ to PQS [46]. The requirement for las to up-regulate AQ biosynthesis is not absolute, since a lasR mutant is capable of producing PQS in late stationary phase, possibly via a lasR-independent mechanism [36]. This finding has been supported by other studies which also showed that in a lasR mutant, expression of pqsR was delayed [66]. The pqsH gene is also expressed in a lasR mutant after both 8 and 24 h, indicating that at higher cell densities, the genetic machinery for making PQS is transcriptionally active in this genetic background. At present, the exact mechanism activating AQ production in a lasR mutant is unknown, however it may involve RhlR [68]. In PA14, it was found that provision of constitutively expressed RhlR on a plasmid was able to partially overcome the delay in PQS production in a lasR mutant [68]. Thus, it would seem that RhlR is able to assume some of the functions of LasR with respect to pqsABCD and pqsH expression to generate PQS. This is somewhat paradoxical, since RhlR is generally considered to be a repressor of AQ production (see below) and indicates that the traditional LasR-RhlR-AQ QS hierarchy in P. aeruginosa may be more complex than was originally thought. While the las system positively regulates PQS and AQ production, the rhl system works as a negative regulator. In a rhlR mutant, a 50% increase in pqsR expression was observed, suggesting RhlR is a repressor of pqsR expression [65]. Additionally, expression of pqsA in a rhlI mutant was greatly enhanced and C4 -HSL inhibited the activation of pqsA by 3-oxo-C12 -HSL in a dose-dependent manner, with a corresponding reduction in PQS levels [64]. The pqsA promoter contains two putative las/rhl boxes at -311 and -151 base pairs before the pqsA transcriptional start site [66]. Deletion of the las/rhl box at -311 causes increased pqsA expression and therefore it is likely that a repressor binds at this site. Deletion of both boxes does not further alter pqsA expression. In a rhlR mutant, expression of pqsA was unaffected with or without the presence of the box at -311, suggesting that RhlR binds to this box to repress pqsA transcription as a deletion of this box in a wild type strain resulted in increased pqsA expression. A previous study investigated the binding of C4 -HSL and RhlR (from E. coli lysates) to the pqsA promoter using DNA mobility shift assays and found they did not bind to the pqsA promoter [65]. However, the promoter region of pqsA studied was only 253 base pairs in length and it is likely this region did not include the las/rhl box situated at -311 base pairs. Interestingly, this negative regulation of the AQ system is itself is driven by the production of AQs and so acts as a negative feedback loop. PQS added to a culture of PAO1 produced enhanced levels of RhlR and C4 -HSL after 8 h [36]. Exogenous provision of C4 HSL and PQS was also found to reverse the loss of rhlI expression in a lasR mutant, causing expression to increase to that observed in the wild type [34]. Individual addition of these compounds did not cause notable increases in rhlI expression, indicating a possible need for synergy between the two compounds. Thus, in P. aeruginosa, AQs are positively up-regulated by the las QS system and are negatively down-regulated by the rhl QS system. AQs indirectly negatively regulate their own production via positive regulatory effects on the rhl QS system.
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2.7 Autoregulation of AQs PqsR is the transcriptional regulator for the pqsABCDE biosynthetic operon [46] and in a pqsR mutant, expression of phnAB, pqsABCDE and AQ production is abolished [18, 64]. The pqsR promoter was mapped by primer extension analysis and this showed that two primary transcriptional start sites exist for pqsR at –190 and –278 base pairs upstream from the pqsR start codon. The –278 base pair start site matched 5 out of 6 bases of the consensus sequence for both the –35 and –10 regions of a σ70 type promoter, suggesting pqsR is basally expressed [65]. Using recombinant PqsR from E. coli lysates in DNA mobility shift assays, PqsR was found to bind to the pqsA promoter in the absence of PQS, but when a PAO1 extract was added in combination with this molecule, significantly enhanced binding to the pqsA promoter was observed [65]. Therefore PQS is a co-inducer for PqsR. Interestingly, two bands were observed in these gel shift assays suggesting that PqsR and PqsR-PQS bind to two different locations in the pqsA promoter region [65]. Primer extension analysis indicated that the pqsA transcriptional start site is -71 base pairs upstream of the pqsA start codon [64] and the pqsA promoter contains a putative LysR-type box with dyad symmetry at –45 base pairs before the pqsA transcription initiation site [66]. A point mutation of this LysR box results in a major reduction in pqsA expression and if the dyad symmetry is destroyed, pqsA expression is abolished. This LysR box would therefore appear to be a central pqsABCDE regulatory element. PqsR binding to this box was severely reduced in the LysR box mutants; it is likely therefore that this is the element to which PqsR-PQS binds to up-regulate the pqsABCDE operon. PqsR may be involved in the regulation of anthranilate production since overexpression of PqsR in PAO1 practically abolished the expression of antA, a gene involved in anthranilate degradation [69]. Presumably, this helps to ensure an adequate supply of anthranilate is available for AQ biosynthesis. Interestingly, PQS is not the only AQ capable of up-regulating AQ expression via PqsR. The pqsABCDE and pqsR genes were cloned into E. coli and this system produced both HHQ and NHQ but not PQS as the pqsH gene was absent [70]. Additionally, in P. aeruginosa, pqsA expression and AQ production were comparable in the wild type versus a pqsH mutant [70]. Consequently, it was speculated that HHQ is also capable of inducing a conformational change in the PqsR protein, and enhance PqsR binding to the pqsA promoter in vitro, although not as effectively as that seen with PQS. In a PA14 pqsAH mutant, PQS was 100 times more potent at activating the expression of pqsABCDE than HHQ. This finding was corroborated by a study using a novel PAO1 AQ bioreporter to show activation of pqsA by HHQ in a pqsAH mutant [40]. Another study compared the whole gene expression profiles of a pqsH mutant in PA14 with and without the exogenous addition of synthetic PQS compared to that for the pqsR mutant [70]. This showed that there was less than 2fold change of all genes regulated by pqsR in the pqsH mutant and that this was not increased when PQS was added. This therefore implies that HHQ, already known to be capable of binding to pqsR [70], is also capable of activating the expression of many of the genes that are down-regulated in a pqsR mutant (except phzA1 as PQS is critical for pyocyanin expression) [70]. These studies indicate that HHQ can
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induce its own biosynthesis independently of PQS. Other AQs, such as NHQ, can also activate PqsR, although at lower levels than PQS [71, 70].
2.8 Other Regulators of AQs Other regulators of AQs have also been identified. PhoB is part of a phosphate regulon responsible for sensing phosphate levels in the environment. AQ production is raised under low phosphate conditions but this induction is abolished in a phoB mutant [72]. There is a putative PHO box preceding pqsR to which this induction may be linked [72]. The gene PA0964, designated pmpR has recently been shown to affect AQ regulation [73]. PmpR belongs to the YebC family and binds to the pqsR promoter negatively affecting PqsR expression and consequently AQ production. This may have numerous effects on other phenotypes since a pmpR mutant showed increased pqsH and pqsA expression, increased expression of phzA1 and pyocyanin production, swarming motility and biofilm formation. PtxR is a LysR-type positive transcriptional regulator of exotoxin A and is regulated by PvdS and Vfr. PtxR negatively regulates the pqsABCDE operon and consequently reduces expression of phz1 [74]. However, PtxR was also found to positively regulate the las QS system, while negatively regulating the rhl QS system. Since this would be expected to give an increase in the expression in pqsABCDE, rather than a decrease it is likely there may be other intermediate regulators also working on this system. The gene PA2663 (ppyR), is a putative NO-mediated novel membrane sensor [75]. In a ppyR mutant, whole genome expression analysis showed that numerous genes, including genes involved AQ biosynthesis (pqsABCDEH) and anthranilate degradation (antABC) were repressed compared to the wild type. Correspondingly, there was no detectable PQS is this mutant, as viewed using TLC. The mechanism of regulation by ppyR on the pqs operon is not known, but also repressed were genes directing biofilm formation (psl), pyoverdin biosynthesis (pvd) and elastase production (lasB). In PAO1, the psl operon is responsible for the biosynthesis of a mannose/galactose rich exopolysaccharide thought to be responsible for biofilm formation [76, 77] Since PQS is thought to be responsible for the release of some of the DNA that is a component of the biofilm extracellular matrix [78, 45, 79, 80], ppyR may form a regulatory link between these two operons.
2.9 Effects of AQs on Expression and Production of Virulence Factors It had been previously thought that AQs were necessary for the activation of several different virulence phenotypes and that this activation was effected via the β-metallohydrolase-like protein, PqsE. In pqsE and pqsR mutants, pyocyanin
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production [47, 36, 46], phzA1 expression [51], lectin production [36], elastase production and rhamnolipids levels were considerably reduced compared to the wild type [47, 36]. PQS [51, 36], HHQ or HQNO [51] addition could not restore these phenotypes in these mutants [46]. Whole genome expression studies have examined the subset of genes regulated by the AQ system [51]. A pqsR mutant displayed 22 genes whose expression had been repressed by pqsR, with another 121 genes whose expression was enhanced by pqsR compared to the wild type. Expression of the pqs and phn operons was abolished and that of pqsR reduced. The phz operon, hcnABC, chiC (chitinase), mexGHI-opmD (broad substrate efflux pump) and lecA transcription were also reduced. The role of AQs in the production of virulence factors was also demonstrated by addition of methyl-anthranilate, an inhibitor of PQS biosynthesis, to PAO1 as this resulted in reduced elastase production [58], lecA expression and pyocyanin production in a dose-dependent manner [36]. A concentration of 500 μM methylanthranilate completely inhibited pyocyanin and lecA without affecting growth, which could partially be restored by the provision of exogenous PQS. Addition of PQS to PAO1 at concentrations greater than 100 μM caused an extended lag phase and reduced stationary phase optical densities [36], however, the onset of expression of lecA was observed at lower cell densities and therefore maximal lecA expression was seen at the onset of stationary phase. These effects were not seen with HHQ or formyl-HHQ [36]. PQS was however, able to both advance and enhance elastase and pyocyanin production into the logarithmic phase. PQS failed to restore lecA expression in a rhlR or rpoS mutant as these two regulators are essential for lecA expression. However, PQS was able to overcome the repression of lecA by the HNS type protein MvaT and the post-transcriptional regulator RsmA and it increased lecA expression in mutants overexpressing these repressors [36]. A recent paper [52] has shed new light on these findings and revealed that PqsE alone could be the main driver for the expression of these virulence phenotypes, through the rhl QS system. The expression of PqsE in pqsA or pqsR mutants (both AQ negative) caused increases in pyocyanin, rhamnolipid and elastase production in the absence of AQs. AQs are therefore not required for production of these phenotypes. This restoration of exoproduct production was not observed in a rhlR mutant, indicating that PqsE may exert its effect through the rhl system. These findings raise the question as to the role of AQs in P. aeruginosa. As only PqsE is necessary to activate these, and possibly other virulence phenotypes, it may be that a primary regulatory function of PQS and HHQ in P. aeruginosa is to bind to PqsR, up-regulate the pqsABCDE operon and increase the production of PqsE. Another intriguing question raised by the data is to the function of PQS itself. There are conflicting reports of the necessity of PQS for virulence in different P. aeruginosa wild type strains (although different hosts were used). PQS has been shown to be necessary for virulence in PAO1 (nematodes) [46], but unnecessary in PA14 (burned mice) [70]. There are also conflicting reports as to the efficacy of PQS at upregulating pqsA via PqsR. PQS is less effective at up-regulating pqsA than HHQ in PAO1 [71], although another study using a different reporter in PA14 found that PQS was
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more effective [70]. PQS may therefore not be as important to the QS system of P. aeruginosa as first envisioned, and may have evolved as a fortuitous by-product with other functions [81, 40]. This may explain its absence in other AQ producing bacteria [82, 83]. The role of PqsR is also cast in a new light. PqsR is possibly important for the virulence of P. aeruginosa in that it is responsible for the expression of pqsE via the production of AQs and corresponding auto-up-regulation of the pqsABCDE operon, at least as far as the production of pyocyanin and expression of lecA are concerned. It is therefore likely that the loss of virulence noted in pqsR mutants [47, 51] is primarily due to the corresponding loss of PqsE, not the loss of PqsR alone, seen in the fact that mouse mortality in PA14 was much decreased from the wild type and was equivalent in both pqsA and pqsE mutants [51].
2.10 AQs and Iron PQS is capable of chelating ferric iron, an essential bacterial nutrient. A 3-hydroxy group is essential for AQ iron chelation- two or three molecules of PQS are capable of binding to ferric iron (Fe3+ ) at physiological pH ranges of 6–8 [50, 40]. C9-PQS also possesses this ability, however AQs lacking this group, such as HHQ are unable to bind Fe3+ [50, 40]. Addition of PQS to a culture of P. aeruginosa induces an iron starvation response, indicated by the up-regulation of siderophore-mediated iron transport systems such as the pyochelin biosynthetic cluster (pchDCBAEGF), the iron-pyochelin outermembrane receptor fptA and the pyoverdin genes pvdE and pvdS [50, 40]. These pch genes were up-regulated at 5, 11 and 20 h between 3 and 25-fold. Also, the genes pvdJAD encoding pyoverdine synthetases were up-regulated between 2 and 10-fold at 11 and 20 h [50]. Quantitative real time PCR showed that pvdA and pchE are up-regulated by 6 and 17-fold respectively upon addition of 20 μM PQS [40]. Also, in PAO1 wild type and pqsA mutants, as well as pqsE and pqsR mutants, addition of PQS, but not HHQ, strongly induced pyoverdine production [40]. There were also effects on the transcription of other genes. The pqsR gene has been shown to be up-regulated in low iron conditions [84] and in the PQS-deficient PAO1 pqsA mutant grown in iron replete media, lecA and pqsA (as well as pvdE and pvdS) were all strongly induced by the addition of 50 μM PQS. However when grown in iron deficient CAA medium, PQS, PQS:Fe (3:1) and HHQ strongly induced pqsA but C1 -PQS did not, therefore the induction of pqsA was not due to the iron chelating properties of PQS [40]. A mechanism for iron-dependent regulation of PQS has been proposed. In irondepleted growth conditions, the P. aeruginosa ferric uptake regulator (Fur) protein induces the production of two small regulatory RNAs, prrF1 and prrF2 [85]. Microarray and real-time PCR analysis have also indicated that under low iron conditions, prrF and prrF2 jointly repress the expression of antABC, responsible for anthranilate degradation [69]. PQS production was abolished under iron-limited
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conditions in a prrF1/prrF2 double mutant; however this could be restored by the provision of exogenous anthranilate, suggesting the loss of PQS production is a result of excessive anthranilate degradation from increased expression of antABC. Under low iron conditions, prrF and prrF2 therefore repress the expression of antABC, sparing anthranilate for PQS synthesis, a mechanism that may be selfreinforcing since addition of PQS leads to an iron-starvation response [81, 40]. Surprisingly, the provision of iron to the wild type did not reverse the above effect and lead to decreased PQS production, as instead an increase in PQS production was observed. This effect was also seen in the prrF1/prrF2 double mutant. It was speculated that the kynurenine pathway of anthranilate biosynthesis may be responsible for the provision of anthranilate in high iron conditions as the kynA and kynU genes were up-regulated and there was an increased utilisation of tryptophan [69]. PAO1 and a PAO1 pvdD/pchEF double mutant (no functioning high-affinity active iron acquisition system) were grown in iron-deficient CAA media in the absence or presence of PQS. PQS abolished growth of the double mutant (which grows similar to PAO1) [40]. Growth was restored upon provision of iron while mutants lacking either pyochelin or pyoverdin also grew normally since either siderophore is presumably capable of acquiring iron bound to PQS. Around 60% of PQS is associated with the cell envelope [40, 35] and therefore can potentially trap iron, however the above results suggest it can not deliver this trapped iron into the cell in the absence of a siderophore transport system. Consequently, PQS is likely to act as an iron trap in the cell membrane, rather than a siderophore per se [40]. The cell membrane of iron rich cells is visibly pink due to complexed Fe3+ , possibly stored in AQ-containing inclusion bodies [86, 87]. Since it has been shown that HHQ performs similar functions to PQS, such as induction of pqsA expression [40, 70], many of the specific effects seen upon addition of PQS may be due more in relation to its iron chelating effects. It is also probable that the production of PQS, with its iron-chelating effect brings a survival advantage when P. aeruginosa is growing with other competing microorganisms in iron-poor environments.
2.11 The Role of AQs in Infection P. aeruginosa is a major source of nosocomial infections and lung infections in patients with CF amongst others [88]. A role for AQs in establishing infection, regulating virulence factors and enhancing the severity of infection has been implicated in several studies. Clinical isolates from sputum in CF patients have been shown to produce HQNO, NQNO and UQNO plus their mono-unsaturated derivatives, HHQ [89] and also PQS [90]. Levels of PQS correlated with the population density of the sample. PQS was also found in isolates from infants with CF and early stage Pseudomonas infection [91]. Regulation of PQS production was irregular in these isolates as they were shown to produce PQS earlier in growth, during the log phase [91]. It has been shown that CF sputum is a good growth medium for P. aeruginosa and supports high population densities [57]. Differential regulation of PQS production in CF sputum media is also seen as the phnAB genes are induced 14–22-fold
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and the pqsABCDE operon between 17 and 19-fold compared to the wild type grown in a morpholinepropanesulfonic acid (MOPS) glucose medium [57]. This resulted in the production of five times more PQS (presumably with corresponding levels of other AQs) in the sputum. This induction was not due to AHLs but is possibly linked to the presence of aromatic amino acids in the medium, which is interesting given that P. aeruginosa has been shown to be able to use the aromatic amino acid tryptophan as a substrate to make anthranilate [55]. It is likely that AQs are at least partly utilised as antibiotics during early infection to kill any competing strains as AQs inhibit the growth of S. aureus and the yeast C. albicans [89]. AQs packaged in MVs were able to inhibit the growth of S. epidermidis [43] and mutants in kynAU were unable to kill S. aureus and a kynB mutant displayed reduced killing, presumably due to a lack of AQs [55]. HHQ and PHQ have antibacterial activity against V. anguillarum, S. aureus, C. albicans and V. harveyi [92]. PHQ also inhibited the growth of phytoplankton (Synechococcus), algae (Chaetoceros simplex, Cylindotheca fusiformis and Thalassiosira weissflogii) and impacts on particle-associated marine bacterial communities [93]. A clinical strain of P. aeruginosa inhibited metronidazole-resistant Helicobacter pylori in a cross streak assay due to production of HHQ and HQNO [94]. These findings may explain the absence, in many instances, of CF early colonisers such as S. aureus in lungs colonised by P. aeruginosa, which has been shown to out compete other co-habiting organisms [89]. Coupled with their iron chelating properties and effects on virulence factor expression, it is likely that AQs contribute to providing P. aeruginosa with a highly favourable environment in which to grow in the CF lung. Interestingly, Candida albicans produces farnesol, a sesquiterpene that reduces the expression of pqsA, possibly by causing aberrant binding of PqsR to the pqsA promoter [95]. This reduced both PQS and pyocyanin production and indicates that this type of inter-species competition can be reciprocal. Also, HQNO has been shown to induce formation of persistent small colony variants of S. aureus that may resist P. aeruginosa niche colonisation [96]. The importance of the AQs for regulation of virulence and contributing to the severity of infection has also been demonstrated. A phnAB mutant was 4-fold less virulent than the wild type in a wax moth larvae (Galleria mellonella) model [97]. In a nematode assay (C. elegans), mutations in a number of pqs genes (pqsC, pqsD, pqsE, pqsR, pqsH and phnA) resulted in severely reduced killing by P. aeruginosa to between 37 and 39% of the wild type levels [46]. A pqsR mutant also had only ∼ 35% mortality rate compared with that of the wild type in burned mice. This mutant also displayed reduced pyocyanin, elastase, exoprotein, PQS, 3-oxo-C12 HSL production [47]. Mutants in pqsA and pqsE also exhibit reduced virulence in mice with mortality rates similar to that of the pqsR mutant [51]. Most interestingly, in PA14, a pqsH mutant has been shown to possess wild type levels of virulence. This suggests that PQS is not essential for virulence and that virulence may be regulated via a PQS precursor, most likely HHQ [70]. Zaborina et al. [98] tested the ability of various opioid compounds such as synthetic U50,488 and the endogenous -opioid receptor agonist, dynorphin (released in the human small intestine), to enhance PAO1 virulence. Addition of U50,488 or dynorphin to a culture of PAO1 stimulated a dose-dependent increase in pyocyanin
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and enhanced expression of pqsA and lecA (although not pqsR), with corresponding increases in the concentrations of PQS, HHQ and HQNO. These compounds also enhanced the growth inhibitary activity of P. aerugnosa against Lactobacillus and C. elegans, probably due to the above effects. Dynorphin binds and enters bacterial cells. Intriguingly therefore, it appears that virulence factor and AQ production is up-regulated in response to host stress responses to P. aeruginosa. PQS also has immune modulatory activity. PQS suppresses T-cell proliferation and interleukin-2 cytokine release in human peripheral blood mononuclear cells (hPBMCs) activated with the pan-activating lectin, concanavalin A. PQS also induced TNF-α release from hPBMCs after activation with lipo-polysaccharide (LPS) at concentrations over 10 μM. It is therefore possible that AQs have a role in the dysregulation of the host immune response [99]. In the lung, P. aeruginosa forms biofilms which protect the bacterial cells from adverse environmental conditions including the host immune system and antimicrobial agents. AQs are important in maintaining and establishing the biofilm mode of life. Addition of 60 μM exogenous PQS to a growing culture of PAO1 enhanced biofilm development, possibly partly due to an increased effect on the expression of the lectin gene lecA [36] since this gene has been shown to play an important role in maintaining biofilm architecture [100]. P. aeruginosa releases extracellular DNA which can function as cell-to-cell interconnecting matrix component in biofilms [80]. This DNA presumably originates from the lysis of a bacterial subpopulation. DNA itself is an antimicrobial with cation-chelating properties and has been shown to cause disruption of the outer membrane of the bacterium (with the loss of cell contents and presumably more DNA) by chelating Mg2+ cations responsible for membrane stability [101]. Additionally, the chelation of Mg2+ by DNA induced the expression of the PhoPQ two-component system [101]. This system is responsible for increased antibiotic resistance of P. aeruginosa to cationic antimicrobial peptides (CAPs). CAPs are broad spectrum antimicrobials released from host immune cells that disrupt the outer membrane of bacteria, causing cell death. DNA is released maximally in late log phase and this correlates with maximal PQS release [36, 35]. Correspondingly, a pqsA mutant culture (that generates no AQs) produces low amounts of extracellular DNA, whereas a pqsL mutant culture (that overproduces PQS) contains increased amounts of extracellular DNA [78, 79]. A pqsA mutant does not form structured biofilms, as they are flat and thin, containing little extracellular DNA [78]. This biofilm also displayed increased sensitivity to detergents, possibly due to the loss of this DNA as a wild type biofilm treated with DNase became increasingly sensitive to detergents. Interestingly, there was a correlation between bacterial cell lysis and PQS levels which could explain the release of the extracellular DNA observed in biofilms [45]. A mutation in the pqsL gene produced pronounced lysis in bacterial colonies whereas colonies of pqsA and pqsR mutants displayed no lysis, but this could be restored by addition of synthetic PQS. A reason for this has been proposed as induction of prophage-lysis due to PQS [45]. P. aeruginosa chromosomal DNA contains the filamentous Pf4 prophage the deletion of which leads to a loss of bacterial autolysis and aberrant biofilm formation
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[102]. PQS can also function as a pro-oxidant and increases the sensitivity of P. aeruginosa to stressors such as peroxide and ciprofloxacin [79] which could in turns also be another possible cause of cell lysis and DNA release.
2.12 AQs in Other Bacteria As production of PQS in P. aeruginosa utilises anthranilate derived from the common tryptophan biosynthetic pathway, other bacterial species may be capable of making similar molecules. This is indeed the case and AQs have been found in several other microorganisms. HHQ and 2-pentyl-4-quinolone (PQ) have been discovered in the marine organism Pseudomonas bromoulitis [92], NHQ, UHQ and NQNO are produced by new species of Pseudomonas harvested from the surface of a marine sponge Homophymia [103], and PQ plus HHQ were found in a marine Alteromonas SWAT5 strain [93]. Diggle et al. [82] found that Burkholderia pseudomallei and Burkholderia thailandensis have complete putative pqsABCDE operons in their chromosomes with between 31 and 53% identity to that of P. aeruginosa. These were named hhqABCDE. The hhqA and hhqE genes were shown to complement PAO1 pqsA and pqsE mutants respectively and restore PQS, HHQ, lectin and pyocyanin production in the pqsA mutant and pyocyanin and lectin production in the pqsE mutant. A novel AQ bioreporter was used to identify HHQ in Pseudomonas putida and Burkholderia cenocepacia and was coupled with LCMS/MS to identify HHQ, NHQ, UHQ and HQNO in B. pseudomallei [36]. Disruption of AQ signalling in B. pseudomallei resulted in altered colony morphology and increased elastase synthesis but further work is required to uncover the true importance of these molecules to the organisms. 3-methyl derivatives of PHQ, HHQ and NHQ have also been found in several species of Burholderia such as B. thailandensis, Burkholderia ambifaria, B. pseudomallei and Pseudomonas cepacia (now probably a Burkholderia sp. of unspecified species) [104, 83]. Vial et al. [83] found that these methylated derivatives of AQs were the main species produced in these organisms (previously, Diggle et al. had not looked for these compounds) and termed them 4-hydroxy-3-methyl-2-alkylquinolines (HMAQs). The possible methyltransferase hmqG, part of the renamed hmqABCDEFG operon (formerly hhqABCDE) has been implicated in their biosynthesis. None of the above bacteria produced PQS and previously researchers had also looked for PQS in other strains of Pseudomonas such as Pseudomonas fluorescens, Pseudomonas syringae and Pseudomonas fragi [19] without success. The lack of a homologous pqsH gene suggests that these Pseudomonas species lack the sophistication of PQS signalling encountered in P. aeruginosa. In Burkholderia thailandensis, ambifaria and pseudomallei, the -3 position is largely methylated [83], which would presumably preclude hydroxylation by any pqsH analogue in these species and rule out any signalling pathway based upon PQS.
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2.13 Concluding Remarks The discovery of a non-AHL based QS system in P. aeruginosa, mediated via AQs and linked to the las and rhl QS systems provides a major insight into a complex regulatory network that plays a key role in infection via the regulation of virulence and biofilm maturation. Some AQs, such as PQS are also able to sequester iron and have multiple functionalities. AQ biosynthesis requires several proteins and occurs via a condensation reaction between anthranilate and β-keto fatty acids. Their production is up-regulated by both the las QS system and by AQs themselves, and down-regulated by the rhl QS system. AQs are present in bacteria other than P. aeruginosa, mainly other Pseudomonads and Burkholderiaceae, although the role in these organisms is not, at present, very well defined. The prevalence of AQs in bacterial species other than P. aeruginosa may provide fertile grounds for further research. At present there are few published studies of screens for AQ production in bacteria, yet AQs have been discovered in several of the limited numbers of species that have been investigated [82, 19, 83]. The functions of AQs and their mechanism of synthesis in these recently discovered species is similarly unclear and may prove interesting, although many possess homologues of the pqsABCDE genes from P. aeruginosa. A clinically relevant area of future research is the development of agents that block AQ signalling and the discovery of compounds that can disrupt the production of AQs (quinolone-quenching). A primary target for this inhibition would be the blocking of the binding of PQS and HHQ to PqsR with the objective of preventing pqsABCDE expression. This would have the double-effect of both blocking AQ synthesis and their effect on virulence factor expression. Another target would be to allow AQ synthesis but block the action of PqsE. Methyl anthranilate has been previously shown to inhibit AQ synthesis [58] and recently, halogenated homologues of anthranilate have been found to inhibit the production of AQs, possibly by competing with anthranilate for PqsA [49, 105]. This inhibition limited the systemic proliferation of infection in mice, thereby demonstrating that in principle, quinolone quenching could be a useful avenue of antibiotic research [105]. Other research has revealed that farnesol can inhibit the activation of pqsA by PqsR [95]. However, the activity of these compounds is generally in the millimolar range, hence more potent analogues would need to be discovered before any potential clinical use could be envisaged. In addition to the above, arguably, some of the biggest remaining mysteries of AQ signalling in P. aeruginosa involve the PqsB, PqsC, PqsE and PqsL proteins. While mutational analysis has revealed that these proteins are involved in the synthesis and effect (PqsE) of AQs [50, 36, 52, 46], the molecular and biochemical functions of these proteins are mainly deduced from their gene sequences. The role played by the PqsBC proteins in the biosynthesis of the more prominent AQs such as HHQ and PQS is not yet known, nor how PqsE signals the up-regulation of virulence factors such as pyocyanin. Likewise, it is not known how PqsL interacts with PqsABCD to produce N-oxides, since HHQ appears not to be a precursor of these molecules [18].
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AQ biosynthesis, signalling and regulation in Pseudomonas is an increasingly complex, diverse and interesting field with many questions still unanswered. From inconspicuous beginnings as potential antibiotic compounds, AQs have been demonstrated to be highly versatile molecules, central to the biology of P. aeruginosa.
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85. Wilderman, P.J., Sowa, N.A., FitzGerald, D.J., FitzGerald, P.C., Gottesman, S., Ochsner, U.A. and Vasil, M.L. (2004) Identification of tandem duplicate regulatory small RNAs in Pseudomonas aeruginosa involved in iron homeostasis. Proc. Natl. Acad. Sci. USA 101: 9792–9797. 86. Royt, P.W., Honeychuck, R.V., Pant, R.R., Rogers, M.L., Asher, L.V., Lloyd, J.R., Carlos, W.E., Belkin, H.E. and Patwardhan, S. (2007) Iron- and 4-hydroxy-2-alkylquinolinecontaining periplasmic inclusion bodies of Pseudomonas aeruginosa: a chemical analysis. Bioorg. Chem. 35: 175–188. 87. Royt, P.W., Honeychuck, R.V., Ravich, V., Ponnaluri, P., Pannell, L.K., Buyer, J.S., Chandhoke, V., Stalick, W.M., DeSesso, L.C., Donohue, S., Ghei, R., Relyea, J.D. and Ruiz, R. (2001) 4-hydroxy-2-nonylquinoline: a novel iron chelator isolated from a bacterial cell membrane. Bioorg. Chem. 29: 387–397. 88. Govan, J.R. and Deretic, V. (1996) Microbial pathogenesis in cystic fibrosis: mucoid Pseudomonas aeruginosa and Burkholderia cepacia. Microbiol. Rev. 60: 539–574. 89. Machan, Z.A., Taylor, G.W., Pitt, T.L., Cole, P.J. and Wilson, R. (1992) 2-Heptyl-4hydroxyquinoline N-oxide, an antistaphylococcal agent produced by Pseudomonas aeruginosa. J. Antimicr. Chemother. 30: 615–623. 90. Collier, D.N., Anderson, L., McKnight, S.L., Noah, T.L., Knowles, M., Boucher, R., Schwab, U., Gilligan, P. and Pesci, E.C. (2002) A bacterial cell to cell signal in the lungs of cystic fibrosis patients. FEMS. Microbiol. Lett. 215: 41–46. 91. Guina, T., Purvine, S.O., Yi, E.C., Eng, J., Goodlett, D.R., Aebersold, R. and Miller, S.I. (2003) Quantitative proteomic analysis indicates increased synthesis of a quinolone by Pseudomonas aeruginosa isolates from cystic fibrosis airways. Proc. Natl. Acad. Sci. USA 100: 2771–2776. 92. Wratten, S.J., Wolfe, M.S., Andersen, R.J. and Faulkner, D.J. (1977) Antibiotic metabolites from a marine Pseudomonad. Antimicrob. Agents. Chemother. 11: 411–414. 93. Long, R.A., Qureshi, A., Faulkner, D.J. and Azam, F. (2003) 2-n-Pentyl-4-quinolinol produced by a marine Alteromonas sp. and its potential ecological and biogeochemical roles. Appl. Environ. Microbiol. 69: 568–576. 94. Lacey, S.L., Mehmet, S. and Taylor, G.W. (1995) Inhibition of Helicobacter pylori growth by 4-hydroxy-2-alkyl-quinolines produced by Pseudomonas aeruginosa. J. Antimicr. Chemother. 36: 827–831. 95. Cugini, C., Calfee, M.W., Farrow, J.M., 3rd, Morales, D.K., Pesci, E.C. and Hogan, D.A. (2007) Farnesol, a common sesquiterpene, inhibits PQS production in Pseudomonas aeruginosa. Mol. Microbiol. 65: 896–906. 96. Hoffman, L.R., Déziel, E., D’Argenio, D.A., Lépine, F., Emerson, J., McNamara, S., Gibson, R.L., Ramsey, B.W. and Miller, S.I. (2006) Selection for Staphylococcus aureus small-colony variants due to growth in the presence of Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 103: 19890–19895. 97. Jander, G., Rahme, L.G. and Ausubel, F.M. (2000) Positive correlation between virulence of Pseudomonas aeruginosa mutants in mice and insects. J. Bacteriol. 182: 3843–3845. 98. Zaborina, O., Lepine, F., Xiao, G., Valuckaite, V., Chen, Y., Li, T., Ciancio, M., Zaborin, A., Petrof, E.O., Turner, J.R., Rahme, L.G., Chang, E. and Alverdy, J.C. (2007) Dynorphin activates quorum sensing quinolone signaling in Pseudomonas aeruginosa. PLoS. Pathog. 3: e35. 99. Hooi, D.S., Bycroft, B.W., Chhabra, S.R., Williams, P. and Pritchard, D.I. (2004) Differential immune modulatory activity of Pseudomonas aeruginosa quorum-sensing signal molecules. Infect. Immun. 72: 6463–6470. 100. Diggle, S.P., Stacey, R.E., Dodd, C., Cámara, M., Williams, P. and Winzer, K. (2006) The galactophilic lectin, LecA, contributes to biofilm development in Pseudomonas aeruginosa. Environ. Microbiol. 8: 1095–1104. 101. Mulcahy, H., Charron-Mazenod, L. and Lewenza, S. (2008) Extracellular DNA chelates cations and induces antibiotic resistance in Pseudomonas aeruginosa biofilms. PLoS. Pathog. 4: e1000213.
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102. Rice, S.A., Tan, C.H., Mikkelsen, P.J., Kung, V., Woo, J., Tay, M., Hauser, A., McDougald, D., Webb, J.S. and Kjelleberg, S. (2009) The biofilm life cycle and virulence of Pseudomonas aeruginosa are dependent on a filamentous prophage. ISME. J. 3: 271–282. 103. Bultel-Poncé, V.V., Berge, J.P., Debitus, C., Nicolas, J.L. and Guyot, M. (1999) Metabolites from the sponge-associated bacterium Pseudomonas species. Mar. Biotechnol. (NY) 1: 384–390. 104. Moon, S.S., Kang, P.M., Park, K.S. and Kim, C.H. (1996) Plant growth promoting and fungicidal 4-quinolinones from Pseudomonas cepacia. Phytochemistry. 42: 365–368. 105. Lesic, B., Lépine, F., Deziel, E., Zhang, J., Zhang, Q., Padfield, K., Castonguay, M.H., Milot, S., Stachel, S., Tzika, A.A., Tompkins, R.G. and Rahme, L.G. (2007) Inhibitors of pathogen intercellular signals as selective anti-infective compounds. PLoS. Pathog. 3: 1229–1239.
Chapter 3
Cell-Surface Signalling in Pseudomonas María A. Llamas and Wilbert Bitter
3.1 Introduction Pseudomonads are known for a high proportion of regulatory genes in their genomes [1]. This is not only due to the number of two-component regulatory systems, but they also contain a large number of different cell-surface signalling (CSS) systems. P. aeruginosa PAO1 for instance contains 14 of these systems [2]. CSS systems are generally composed of three different components, an alternative sigma factor of the extracytoplasmic function (ECF) family, a sigma factor regulator (or anti-sigma factor) located in the cytoplasmic membrane and an outer membrane receptor [3, 2] (Fig. 3.1). The outer membrane receptor belongs to the protein family of TonBdependent receptors, which are mostly involved in the transport of iron-siderophore complexes across the outer membrane. To accomplish this task these receptors need to be energized by a protein complex in the cytoplasmic membrane. This protein complex is composed of TonB, ExbB and ExbD, of which the TonB protein is the one that actually makes contact with the outer membrane receptor, hence the name TonB-dependent receptors [4, 5, 6]. Coupling with the cytoplasmic membrane is necessary because the iron-siderophore complex has to be actively transported across the outer membrane, where there is no other source of energy available. Not all TonB-dependent receptors are involved in CSS, only a subfamily known as TonB-dependent transducers [5]. This subfamily can be easily distinguished from other TonB-dependent receptors on the basis of an N-terminal extension of 70–80 amino acids. This extension is usually accompanied with an extension of the signal sequence, although the function of this long signal sequence is yet unknown [7]. Fluorescent pseudomonads are entangled in a fierce competition for iron, as can be judged from the large structural variation in their siderophores between different
M.A. Llamas (B) Department of Medical Microbiology, VU University Medical Centre, Amsterdam, 1081 BT, The Netherlands e-mail:
[email protected]
J.L. Ramos, A. Filloux (eds.), Pseudomonas, DOI 10.1007/978-90-481-3909-5_3, C Springer Science+Business Media B.V. 2010
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Fig. 3.1 Structure of the P. aeruginosa pyoverdine CSS system. Schematic illustration of the interactions of FpvA with the periplasmic domain of FpvR and the TonB protein, and between the amino-terminal region of FpvR and region 4 of both ECF sigma factors FpvI and PvdS (in vivo each FpvR molecule probably interacts with only one of the ECF sigma factors at a time). Interaction of FpvI and PvdS, respectively, with the RNA polymerase (RNAP) core enzyme is shown by arrows. The FpvI–RNAP holoenzyme binds to the promoter of fpvA (PfpvA), whereas the PvdS–RNAP holoenzyme binds to the promoters of the pyoverdine biosynthetic genes (Ppvd), and the toxA (PtoxA) and prpL (PprpL) genes. These genes are also indirectly regulated by iron since the Fur protein loaded with Fe2+ represses pvdS and fpvI transcription under iron rich conditions. C, cytoplasm; CM, cytoplasmic membrane; OM, outer membrane; P, periplasm
species ands even between different strains [8]. These siderophores are generally known as pyoverdin or pseudobactin (in soil isolates) and are composed of a fluorescent chromophore, derived from a dihydroxyquinoline moiety, linked to a variable peptide chain of 6–12 amino acids (Fig. 3.2). These peptide chains are produced by nonribosomal peptide synthetases and contain normal and unusual amino acids in both the D- and L-configuration. In total, more than 50 different pyoverdine structures have been identified [10, 9]. Siderophore variation can even be used for strain/species typing in a technique called siderotyping [11]. Some pseudomonads also produce a second type of siderophore, such as the siderophore pyochelin
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Fig. 3.2 Pyoverdines I, II and III from P. aeruginosa. Structure of pyoverdine I is shown, which consists of a conserved fluorescent dihydroxyquinoline moiety linked to an octapeptide. This peptide chain contains amino acids in both the L- and D- configuration, as indicated, and part of the chain is cyclized. At the bottom part of the figure the peptide chains of all three different pyoverdines of P. aeruginosa are shown. Only pyoverdine I is cyclized (as indicated). Figure is adapted from [9]
of P. aeruginosa. Pseudomonads not only produce their own siderophore(s), but are also highly specialized in the utilisation of heterologous siderophores. For this they have an impressive amount of different TonB-dependent receptors [12, 1], and it seems common sense to produce these receptors only when the heterologous siderophores are present in the direct surrounding of the bacterium. CSS systems are responsible for this task; the same receptor that is used to transport the ironsiderophore complex is also the receptor for the signal transduction pathway used to upregulate the synthesis of this receptor. In addition, also other genes can be upregulated such as a periplasmic transport system for the siderophore complex. Both CSS systems and TonB-dependent outer membrane transport systems are difficult to study in vitro, because components of both the cytoplasmic and outer membrane are involved in this process. However, in recent years the elucidation of the structure of components involved has helped to direct biochemical experiments, which resulted in a general picture for siderophore transport and CSS. In this chapter we will first discuss the various components involved in CSS based on the system for which most information is available, the Fe-pyoverdine CSS system of P. aeruginosa. Subsequently, other CSS systems of pseudomonads will be discussed.
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3.2 Structure of the Cell-Surface Signalling System 3.2.1 TonB and the TonB-Dependent Transducer The signal transduction pathway of CSS starts with binding of the inducing signal, which is usually the iron-siderophore complex, to the outer membrane receptor. As mentioned previously, this receptor is also involved in transport of the Fesiderophore complex over the membrane. For P. aeruginosa’s own siderophore pyoverdine this outer membrane receptor is mainly FpvA (Fig. 3.1), although the presence of a second pyoverdine receptor, FpvB, has been also reported [13]. The nomenclature of FpvA is somewhat confusing. As mentioned above, there is considerable species and strain diversity in siderophore production among the pseudomonads, including P. aeruginosa. In total three highly different pyoverdines have been identified in the various P. aeruginosa strains, designated group I, II and III pyoverdines (Fig. 3.2), of which only one is produced by a specific strain [14, 15, 9]. These different pyoverdines are recognized by different outer membrane receptors, which are all called FpvA [16, 17]. However, it should be realized that the group I, II and III FpvA receptors are only distantly related. The most-studied P. aeruginosa strains, such as PAO1 and PA14, are from the type I group and also the known FpvA structures are all from type I receptors. Therefore we will focus on this group of FpvA receptors. All integral outer membrane proteins of Gram-negative bacteria form a β-barrel and the TonB-dependent receptors are no exception [6]. In fact, members of this protein family form with 22 strands the largest β-barrel observed thus far (Fig. 3.3). β-barrels are formed by antiparallel β-sheets that are connected via short periplasmic turns and larger surface exposed loops. The C-terminal portion of the TonBdependent receptors forms this structure (Fig. 3.3). The size of the β-barrel of TonB-dependent receptors is much larger than those of the outer membrane porins and would therefore allow the diffusion of large molecules. However, the channel in the barrel is occluded by a plug domain (Fig. 3.3), which is an important characteristic of this protein family. Obviously, the plug domain must play a crucial role in the facilitated transport of siderophores across the outer membrane, although its exact role is still a matter of debate. What is known is that the plug domain is involved in pyoverdine binding, both the apo- and the holo- (Fe-bound) form [19–21, 22]. The siderophore binds with its chromophore group to the top part of the plug domain [20]. Three tyrosine residues are involved in this interaction. Because the chromophore group is conserved in all pyoverdines this plug-siderophore interaction will probably be similar for all pseudobactin/pyoverdine receptors. The variable peptide chain of the siderophore, which will determine the specificity of the receptor, is pointed upwards and interacts with residues of the β-barrel and the extracellular loops. Especially loop number 7 plays an important role in the binding of pyoverdine to FpvA [20, 21]. After binding to the siderophore receptor a channel in the receptor will have to be formed. For this, two possibilities can be envisaged, the first is the so-called transient pore model
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Fig. 3.3 Structure of the P. aeruginosa FpvA TonB-dependent transducer. The FpvA receptor is shown in the apo form, i.e. without (Fe)-pyoverdine. The C-terminal β-barrel is shown in green, the plug domain in red, the TonB box in yellow and the signalling domain in blue. The signalling domain is specific for TonB-dependent receptors involved in CSS-dependent signal transduction. At the bottom a linear presentation of FpvA with the different domains and the border of these domains based on amino acid residues of the precursor form of FpvA. Upon binding of Fepyoverdine to the top of the plug domain, the TonB box and the signalling domain dissociate. Structure was adapted from [18]. Ss, signal sequence
where the plug stays within the barrel but shows a conformational change which opens a channel for passage of the Fe-pyoverdine complex [23]. This channel has to be approximately 20 Å to allow for the transport of the bulky siderophore complex. The second model is the ball-and-chain model, in which the plug domain dislodges into the periplasm thereby creating a large open channel. In this model the plug domain can be removed either as a folded structure with a hinge-like connection to the β-barrel or by (partial) dissociation of the plug domain [23]. Different experiments have been performed to prove one of these models, but thus far with somewhat conflicting results. A double-cysteine approach was for instance used to create artificial disulfide bonds in the TonB-dependent receptor FhuA of Escherichia coli [24]. By doing so, the plug domain was tethered to the β-barrel. This crosslinking had no clear effect on siderophore transport, indicating that the plug stays within the β-barrel during transport. Another study used an alternative approach to
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study the same mechanism for the FepA receptor of E. coli [25]. They produced multiple single cysteine mutations in FepA and then analyzed whether these different cysteine residues could be labelled with fluorescein maleimide under different circumstances. A residue located within the plug domain, which is usually buried within the β-barrel, could be labelled with this compound, but only in the presence of both the cognate siderophore and a functional TonB cytoplasmic membrane complex. These data indicate that the plug domain is in fact translocated from the barrel during transport. Recently, a synthesis between these two different models has been proposed based on simulation experiments using the E. coli vitamin B-12 receptor BtuB [26]. Here it was shown that a large force had to be applied to pull the plug out of the β-barrel, making this option less plausible. However, only a limited force is sufficient to partially unfold the bottom part of the plug domain, which could result in the opening of a channel. This simulation is in line with both experimental observations, namely that residues within the bottom part of the plug are (temporarily) exposed to the periplasm and that tethering the top part of the plug to the β-barrel has no effect on iron-siderophore transport. Furthermore, it has also been shown in another study that tethering the bottom part of the plug domain to the β-barrel does abolish outer membrane transport [27]. In both models, there must be energy available to allow the conformational changes needed for iron-siderophore transport. As mentioned in the introduction, the energy needed for this process is supplied by the TonB-ExbB-ExbD cytoplasmic membrane complex. These proteins are all integral cytoplasmic membrane proteins of which both ExbD and TonB have large periplasmic domains. TonB proteins have a highly characteristic proline-rich domain that contains a large number of Pro residues alternating with either Glu or Lys residues [28, 29]. This domain forms an extended structure that is thought to allow TonB to span the periplasm and contact the outer membrane receptors [30]. The C-terminal domain of TonB is well structured and is shown to interact with a specific region of the TonB-dependent outer membrane receptors through β-sheet strand exchange. In this interaction three β-sheet strands of TonB interact with one β-sheet of the TonB-dependent receptor [18, 31, 32]. This interacting β-strand of only 6 amino acids is located at the Nterminal side of the outer membrane receptor and is generally known as the TonB box [33]. How TonB exactly energizes the receptor is unknown, but simulation experiments have shown that, despite the small interaction surface, the interaction between TonB and the receptor is relatively stable, which would allow a mechanical coupling [26]. All the TonB-dependent receptors have the different domains discussed above, i.e. the β-barrel, the plug and the TonB box. However, TonB-dependent receptors involved in CSS have an extra extension of approximately 70–80 amino acids. This extension contains the so-called signalling domain (Fig. 3.3). Because this domain is absent in normal TonB-dependent receptors not involved in CSS, it is not surprising that this domain is not involved n substrate transport. The significance of this domain was shown in a classical experiment where the N-terminal extension of the
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Pseudomonas putida TonB-dependent transducer PupA was exchanged with that of PupB [34]. As a result the genes that were normally induced by the PupB-dependent CSS system were not anymore induced by their normal signal, but only in the presence of Fe-siderophore complex bound by the PupA receptor. This induction was dependent on the presence of an active TonB-ExbB-ExbD complex. Recently, it was shown that the structure of the signalling domain consists of a novel protein fold of two alpha-helices that are sandwiched in between two beta-sheets [35, 36]. The structure of this domain is conserved in all bacterial TonB-dependent transducers. Crystals of intact TonB-dependent transducers usually do not show this domain, due to the flexible hinge connecting this domain with the plug domain. However, recently a new structure of the FpvA receptor was reported where also the signalling domain was resolved. From this analysis it was clear that the signalling domain is subjected to major changes upon the binding of the Fe-pyoverdine complex [18]. In the apo form, i.e. without the Fe-siderophore signal, the signalling domain associates with the single β-strand of the TonB box. This means that in this conformation TonB is probably not able to contact the outer membrane receptor. In contrast, the structure of the holo-FpvA receptor showed the signalling domain without the TonB box attached to it (which was not identified in this structure). This means that binding of Fe-pyoverdine complex to FpvA destabilizes the interaction between the TonB box and the signalling domain, after which these domains are probably free to interact with their partners in the cytoplasmic membrane, respectively TonB and the sigma factor regulator. The interaction between the TonB box and the signalling domain in the absence of an inducing signal could perhaps also explain some intriguing observations made by mutational analysis. Point mutations within the signalling domain have been identified that completely block ferric-citrate transport [37]. However, as discussed previously, the signalling domain can de deleted in the ferric-citrate receptor FecA without affecting transport. This apparent controversy could be explained if these signalling domain point mutations would stabilize the interaction with the TonB box, which could therefore not anymore interact with the TonB protein. TonB-dependent transducers generally bind both the iron-free and the iron-bound siderophores, but only upon interaction with Fe-siderophore complex a number of conformational changes are induced [38, 39]. Structural analysis has shown that the differences between the apo and holo form of the receptor are very small. For instance, binding of the Fe-pyoverdine leads to a number of subtle changes in the plug domain [36]. However, these small changes result in more pronounced changes at the periplasmic site, such as the dissociation of the TonB box from the signalling domain [18]. This order of event can than explain why the TonB complex is triggered to interact with the receptor. The major remaining question is how exactly TonB does this job. Furthermore, it is not yet clear why a functional TonB complex is needed for signal transduction through the CSS system, but if the pulling mechanism of TonB is correct it could be envisaged that an active TonB protein pulls the signalling domain away from the outer membrane receptor to allow the interaction with the sigma factor regulator.
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3.2.2 ECF Sigma Factor Regulator The least understood protein within CSS system is the sigma factor regulator in the cytoplasmic membrane. The function of this protein is to couple the signal perceived by the TonB-dependent transducer to the ECF sigma factor in the cytoplasm. For the pyoverdine signalling system of P. aeruginosa this protein is FpvR, which is, like the other sigma factor regulators involved in CSS, an integral cytoplasmic membrane protein with a single transmembrane domain [40]. The large periplasmic C-terminal part of FpvR interacts with the FpvA receptor in the outer membrane, whereas the short cytoplasmic tail (residues 1–92) should bind the ECF sigma factor. Biochemical studies support this hypothesis. The isolated cytoplasmic tail of FpvR [41] and also the tail of other sigma factor regulators have been shown to interact with the ECF sigma factors [42, 43]. Currently, there is no structural data available for any member of this protein family. The function of the sigma factor regulator is not completely understood. It is generally known as an anti-sigma factor, which is based on the fact that overexpression of only the ECF sigma factor results in the constitutive induction of the CSS system [44]. In accordance with this, overexpression of the sigma factor regulator results in a strongly reduced induction upon the presence of the extracellular signal [40]. For an anti-sigma factor it is furthermore expected that a knockout of this gene also results in constitutive expression. This is observed for a knockout of the fpvR and pupR sigma factor regulators in P. aeruginosa and P. putida, respectively [40, 45], albeit that the induction level in P. putida is below the normal response. However, for the ferric-citrate CSS system in E. coli the regulator FecR is in fact essential for ECF sigma factor activity. Constitutive expression is only achieved if the cytoplasmic part of FecR (FecR1-85 ) is expressed [3]. This fragment is able to bind to the ECF sigma factor FecI and the conserved Trp residues in this domain play an essential role in this process [43]. Apparently, binding of this domain of FecR is needed for FecI activity and/or stability. Subsequently, it is shown that FecR itself is prone to cleavage, probably by the cytoplasmic membrane protease RseP, which releases the cytoplasmic tail of FecR [3]. This mechanism could explain how the signal perceived at the periplasmic site is transmitted: binding of the signalling domain of the receptor induces a conformational change in the membrane part of the sigma factor regulator. This domain is then susceptible to proteolytic cleavage, which releases the cytoplasmic domain with the attached ECF sigma factor. Similar data have been found for the haem CSS system of Bordetella bronchiseptica [46]. This scenario is not uncommon, also other ECF sigma factors that are not involved in CSS are regulated via proteolytic conversion of their sigma factor regulators [47]. The periplasmic stress response sigma factor σE is for instance regulated by the cytoplasmic membrane protease DegS, whereas the alginate sigma factor AlgU is regulated by MucP [47]. The situation for CSS systems in Pseudomonas is not studied in the same detail, but seems to be different from the Fec system in E. coli. First of all, as mentioned above, in pseudomonads ECF sigma factors are active in the absence of the sigma factor regulator. This means that the cytoplasmic tail is apparently
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not needed for activity. This is also shown in reconstitution experiments, expression of the ECF sigma factor fpvI in E. coli without fpvR results in high activity, whereas coexpression of fpvR almost completely blocks activity [40]. These results all point to a straightforward anti-sigma factor role of FpvR. A second difference between the Fec system and the pyoverdine CSS system is that the ECF sigma factors of P. aeruginosa are more stable in the absence of the sigma factor regulator FpvR. Overexpression of fpvR results in increased degradation of the ECF sigma factor PvdS and possibly also FpvI [41, 48]. This means that FpvR not only retains the ECF sigma factor at the cytoplasmic membrane in an inactive form but possibly also delivers it to a specific endoprotease. Future experiments will have to show which protease is involved and what the exact role of FpvR is in CSS systems.
3.2.3 ECF Sigma Factor The pyoverdine CSS system of P. aeruginosa is in several aspects an unusual and unique CSS system. Whereas most CSS systems are induced by heterologous siderophores or other iron sources such as citrate and heme, here the inducing signal is its own siderophore. This situation is unusual but might be more common in pseudomonads, because also in P. putida the homologous pseudobactin receptor contains a signalling domain [49]. Secondly, most CSS systems induce genes involved in the uptake of the inducing signal, i.e. heterologous siderophores or other sources of iron such as haem and citrate. These genes not only encode the outer membrane receptor but can also include the periplasmic transport system. However, pyoverdine CSS induces not only pyoverdine uptake, but also pyoverdine synthesis [50] and, most importantly, also the synthesis of two virulence factors, exotoxin A and the endoprotease PrpL [40, 51]. Because the signal is produced by the bacterium itself and regulates its own production and several other genes, one could argue that pyoverdine is in fact an autoinducer. The inducing signal is not completely produced by the host cell, because it contains an extra molecule, i.e. the iron atom. However, the AI-2 autoinducer is also not directly produced by the host cell and also carries a borate ion [52]. A third and final unusual characteristic of the pyoverdine CSS is that the sigma factor regulator FpvR does not interact with one but with two ECF sigma factors: PvdS and FpvI [40, 51]. The fpvI gene is located adjacent to fpvR, although in the reverse orientation. Normally the genes encoding the ECF sigma factor and the sigma factor regulator are located in the same operon. FpvI is, like all other ECF sigma factors, a small sigma factor, when compared to the other members of the σ70 superfamily [40, 47]. Sigma factors usually contain four structural domains, while ECF sigma factors, such as FpvI, miss domain 3 completely and also the 1.1 domain is missing, which leaves mainly domain 2 and 4 separated by a linker region. These domains probably form the minimal requirements for a functional sigma factor. Domain 2.4 and 4.2 are both involved in recognition of the promoter region, they bind the –10 and the –35 region respectively [47]. Domains 2.1, 2.2 and 4.2 are involved in binding the core
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RNA polymerase complex, whereas domain 2.3 is responsible for melting of the DNA to form the open complex. A mutation approach was used to determine which residues of FpvI are crucial for interaction with the cytoplasmic tail of FpvR [41]. This analysis showed that mainly alteration in domain 4 affected this interaction. FpvI binds probably only to the promoter region of the fpvA gene and induces the expression of this outer membrane transducer. PvdS is the second ECF sigma factor that is controlled by FpvR. As might be expected the residues of FpvR that are involved in FpvI binding are also important for PvdS binding, with the only difference that the interaction with PvdS seems to be more sensitive to small alterations [41]. The pvdS gene is not located near the fpvR gene, but next to a pyoverdine biosynthesis locus as a single gene. PvdS regulates the expression of more than 20 genes, which are mainly genes involved in pyoverdine biosynthesis and in addition genes encoding other virulence factors such as toxA and prpL [53, 54]. The binding site for this sigma factor has been studied in detail, which showed that the consensus binding sequence for PvdS in P. aeruginosa is TAAAT-N16 -CGT. Furthermore, point mutation in the –10 region showed a stronger effect on promoter activity as compared to the –35 region [55]. Both PvdS and FpvI compete with each other for binding to FpvR and, upon activation, for the core RNA polymerase. However, for the latter they also have to compete with other sigma factors, such as the major sigma factor RpoD (σ70 ). Microarray experiments showed that overexpression of several different ECF sigma factors of CSS systems resulted in the upregulation of specific genes, but not to a general effect on cell growth or gene regulation [44]. This indicates that increased amounts of these ECF sigma factors do not significantly affect RpoD binding to the core RNA polymerase and therefore probably these ECF sigma factors have a much lower affinity for this complex. This was also shown in biochemical experiments, where it was determined that under maximum inducing conditions the cellular amount of PvdS is relatively high, up to 60% of the RpoD level [56]. However, isolation of the RNA polymerase holo complex from these cells showed a 20-fold higher amount of RpoD bound to RNA polymerase as compared to PvdS. Similar data were obtained with the sigma factors of E. coli, where it was shown that the affinity of the CSS sigma factor FecI for RNA polymerase was 7-fold lower as compared to RpoD [57]. This competition for the core RNA polymerase complex also explains why deletion or overexpression of algQ, which is coding for an RpoD anti-sigma factor, affects the induction of pyoverdine genes [58].
3.3 P. aeruginosa Cell-Surface Signalling Systems P. aeruginosa not only contains a CSS system for its own siderophore pyoverdine, but contains an impressive total of 14 of these regulatory systems (Table 3.1). As mentioned above, the sigma factors involved in CSS belong to the ECF subfamily of sigma factors. P. aeruginosa contains in total 19 ECF sigma factors, 14 of these (74%) are iron starvation sigma factors involved in CSS [53, 44, 2]. P. aeruginosa also possesses a high number of TonB-dependent receptors, among which a
54% similar to E. coli FecI
–
FemI
– – FpvI PvdS
FoxI HasI FecI 64% similar to E. coli FecI
PA1300
PA1363
PA1912
PA2050 PA2093 PA2387 PA2426
PA2468 PA3410 PA3899 PA4896
PA2466 (FoxA) PA3408 (HasR) PA3901 (FecA) PA4897
PA2057 and PA2070 PA2089 and PA2590 PA2398 (FpvA) PA2398 (FpvA)
PA1302 (57% similar to heme utilisation protein Hxu of Haemophilus influenzae) PA1365 (68% similar to the alcaligin E receptor AleB of Ralstonia eutropha) PA1910 (FemA)
PA0470 (FiuA) PA0674
PA0151
TonB-dependent transducer
PA number in the P. aeruginosa genome annotation project (http://www.pseudomonas.com)
PA2467 (FoxR) PA3409 (HasS) PA3900 (FecR) PA4895
PA2051 PA2094 PA2388 (FpvR) PA2388 (FpvR)
PA1911 (FemR)
PA1364
PA1301
PA0471 (FiuR) PA0676 (PigE)
PA0150
Sigma factor regulator
(carboxy)Mycobactin uptake Metal uptake Siderophore uptake Pyoverdine uptake Production of pyoverdine, exotoxin A and PrpL endoprotease Ferrioxamine uptake Haem uptake Ferric citrate uptake Pyocin production
Alcaligin uptake
Haem uptake
Ferrichrome uptake Virulence
Metal uptake
Function
[59] [60] [61] [44]
[44] [44] [40] [51]
[44]
[44]
[59] (Llamas et al., in preparation) [44]
[44]
Reference
Cell-Surface Signalling in Pseudomonas
a
PA0472 PA0675
48% similar to PvdS FiuI PigD
ECF protein name
PA0149
ECF sigma factora
Table 3.1 P. aeruginosa CSS systems (adapted from ref [44])
3 69
70
M.A. Llamas and W. Bitter
number of TonB-dependent transducers [12, 5]. This gives some indication of the range of iron-chelates that are present in the environments where P. aeruginosa is found. Although the majority of these CSS systems are involved in iron uptake, there are some exceptions. At least nine of the P. aeruginosa CSS systems seem to control iron uptake, six of them regulate the uptake of iron via siderophores (FiuI, FemI, FpvI, FoxI, PA1363 and PA2093,), two are hypothesized to regulate iron uptake via haem (HasI and PA1300), and one via citrate (FecI) [61, 40, 44, 59, 60] (Table 3.1). Furthermore, another iron starvation sigma factor, PA4896, could be involved in the uptake of iron through siderophores, but also controls the production of pyocins probably in response to the presence of a specific siderophore [44]. Finally, PvdS does not regulate the uptake of iron, but the production of the siderophore pyoverdine itself [51, 62] (Table 3.1). There are also two P. aeruginosa iron-starvation sigma factors, PA0149 and PA2050, that seem to regulate the uptake of a metal ion(s) different than iron, probably zinc or manganese [44]. Finally, the PA0675 (PigD) iron starvation sigma factor regulates the expression of genes encoding secreted proteins and components of secretion systems (Llamas et al., in preparation). This CSS system is the most unusual in P. aeruginosa and seems to be involved in the regulation of P. aeruginosa virulence (see below). Most P. aeruginosa CSS pathways are also present in at least one of the other Pseudomonas species sequenced to date, such as the saprophytes Pseudomonas putida and Pseudomonas fluorescens, the insect pathogen Pseudomonas entomophila, and the plant pathogen Pseudomonas syringae (Table 3.2). The fact that these bacteria share most of these systems suggests that they use the same siderophores. There are three systems specific of P. aeruginosa, two siderophore transport systems, for alcaligin and an unknown siderophore, and one for metal ion transport (Table 3.2). This may reflect the different habitats that these bacteria encounter, which seems to imprint a characteristic profile on the repertoire of signalling systems they possess. We will discuss some of these CSS systems in P. aeruginosa in more detail.
3.3.1 Pyoverdine-Mediated Signalling Pathway P. aeruginosa mutants that are unable to synthesize pyoverdine have a greatly reduced ability to cause disease in animal models [63, 64]. The synthesis and uptake of pyoverdine is both regulated by the pyoverdine CSS system, as discussed previously, and by the availability of iron through the Fur protein [65]. The ferric uptake regulator (Fur) protein is a Fe2+ -dependent transcriptional repressor of iron-responsive genes. The promoters of these genes contain the so-called Fur box or iron box that enables the binding of the Fe2+ -Fur complex. In pseudomonads, Fur represses the transcription of most CSS systems, including the pvdS and fpvI genes.The pyoverdine-mediated signalling system, in particular the PvdS sigma factor, is present in all Pseudomonas species sequenced to date (Table 3.2). Using a combination of promoter trapping, gene fusion, RT-PCR and bioinformatics analysis, a recent study has shown that PvdS has a ‘core’ regulon associated
Regulation of virulence Regulation of heme uptake Regulation of alcaligin uptake Regulation of mycobactin/ carboxymycobactin uptake Regulation of metal uptake
PA0674-0675-0676
PA23872388/PA2398 (Fpv system)
PA20502051/PA2057/ PA2070 PA2089/PA20932094/PA2590
PA1912-1911-1910 (Fem system)
PA1363-1364-1365
PA1300-1301-1302
Regulation of siderophore uptake Regulation of pyoverdine uptake
Regulation of metal uptake Regulation of ferrichrome uptake
PA0149-0150-0151
PA0472-0471-0470 (Fiu system)
Function
P. aeruginosa
–
PFL4080/ PFL2903/ PFL4092
–
PP4208/PP3555/ PP4217
–
–
–
PP3086-30853084 and PP3577-35763575
–
–
PFL27452746-2747 PFL13731372-1371 –
PFL09880989/PFL0995 PFL57045705-5706
P. fluorescens
–
PP4608-46074606 PP0352-03510350
P. putida
–
–
PSEEN3663/ PSEEN2995
–
PSYR25802581-2582
–
–
PSYR47314730 PSPTO04440445 –
–
P. syringae
–
–
–
PSEEN21072108-2109 PSEEN43314332-4333 –
PSEEN45674566-4565 PSEEN51315132-5133
P. entomophila
Table 3.2 Presence of the P. aeruginosa CSS systems in other Pseudomonas species
–
–
–
–
–
–
–
Pmem_26732674-2675
–
P. mendocina
3 Cell-Surface Signalling in Pseudomonas 71
PA4897-4896-4895
PA3410-3409-3408 (Has system) PA3899-3900-3901 (Fec system)
Regulation of siderophore uptake
Regulation of pyoverdine, exotoxin A and PrpL endoprotease synthesis Regulation of ferrioxamine uptake Regulation of haem uptake Regulation of ferric citrate uptake
PA2426/PA2388/ PA2398 (PvdS system)
PA2468-2467-2466 (Fox system)
Function
P. aeruginosa
–
PFL5380-53795378 PA4611-46124613
–
PSEEN47684767-4766 PFL09840983-0982
PFL01270126-0125
PFL4190/ PFL2903/ PFL4092
PP4244/PP3555/ PP4217
PP0162-01610160
P. fluorescens
P. putida
Table 3.2 (continued)
PSYR10401039-1038 PSPTO12091208-1207 –
PSEEN45714572-4573
PSEEN06180619-0620
–
–
PSYR1943/ PSYR1961 PSPTO1286/ PSPTO2151
P. syringae
–
PSEEN25312530-2529
PSEEN1814/ PSEEN2995
P. entomophila
–
–
–
Pmem_2873
P. mendocina
72 M.A. Llamas and W. Bitter
3
Cell-Surface Signalling in Pseudomonas
73
with pyoverdine synthesis that is conserved among the fluorescent pseudomonads, and an ‘extended or accessory’ regulon that varies from species to species [55]. The genes from the ‘extended’ regulon have diverse functions, presumably to meet the specific needs of these organisms in the environments they encounter.
3.3.2 Ferric Citrate-Mediated Signalling Pathway The ferric citrate-mediated signalling pathway has been extensively studied in E. coli. It was observed already long time ago that this bacterium could transport ferric citrate via an inducible citrate-dependent iron transport system [66]. This finding was, however, surprising since E. coli cannot grow in citrate as sole carbon source. In E. coli there is no co-transport of iron and citrate into the cytoplasm [67]. When supplied as a ferric complex, at least 10 times more iron enters the cell as compared to citrate. This finding raised the question of how an inducer that does not enter the cell can activate a transport system. The fact that iron, and not citrate, is transported into the cytoplasm [68] clearly showed that the ferric citrate complex induces the transport system acting from outside the cell. The ability of citrate to mediate iron acquisition in P. aeruginosa has also been known for some time [69]. Like E. coli, citrate-mediated iron uptake in P. aeruginosa requires prior exposure to citrate, which induces the expression of an outer membrane protein [70]. However, until recently there was no experimental evidence linking specific genes to ferric citrate utilisation in this bacterium. P. aeruginosa contains a gene cluster (PA3899-PA3901) that shows more than 50% identity to the E. coli FecIR regulatory proteins and to the FecA outer membrane receptor (Table 3.1). The PA3901 protein, which encodes a putative TonB-dependent transducer, has been shown to be necessary for iron citrate utilisation [61]. In this study it was shown that a P. aeruginosa mutant devoid of pyoverdine is not able to form a well-structured biofilm. This defect could be restored by adding iron citrate to the medium, but only if PA3901 was present, which confirms the involvement of this protein in iron citrate utilisation [61]. Based on homology it is to be expected that the P. aeruginosa PA3899-PA3900 genes encode the regulatory proteins involved in PA3901/fecA expression, although this has not been experimentally tested. The homology of PA3900 with FecR is significantly lower as compared to the other proteins of the putative Fec CSS system, which could indicate that this sigma factor regulator has a different function, perhaps more in line with FpvI/PvdS. P. aeruginosa fecA gene is not clustered with homologues to the E. coli periplasmic transport system. Overall, the P. aeruginosa genome shows a striking lack of obvious periplasmic and cytoplasmic membrane transport components for ferric siderophores, which suggests that iron-siderophore complexes in this bacterium are dissociated in the periplasm and a common iron carrier be then responsible for iron uptake into the cell [71]. This would mean that iron and citrate are independently transported into P. aeruginosa cytoplasm, and that P. aeruginosa may have a transport system specific for citrate. This could explain why this bacterium is able to transport citrate more efficiently than E. coli. There are two genes, PA5468 and PA5476 annotated
74
M.A. Llamas and W. Bitter
in the P. aeruginosa genome as probable citrate transporter. It is possible that the P. aeruginosa FecIRA signalling system does not only control fecA expression, but also the expression of these putative transporters. Further experiments are needed to prove this hypothesis.
3.3.3 Iron-Siderophore-Mediated Signalling Pathways Pyoverdine and iron citrate are not the only sources of iron that P. aeruginosa can used. This bacterium is also able to incorporate many heterologous siderophores of bacterial or fungal origin [12, 17]. For this they have an impressive amount of different TonB-dependent receptors [12]. Some of these receptors are produced under iron limiting conditions, but most of them need additional conditions, such as the presence of their cognate siderophores, to be expressed [59, 62]. Proteomic analysis has revealed that the heterologous siderophores ferrichrome and ferrioxamine B induces the synthesis of the TonB-dependent transducer FiuA and FoxA, respectively [59] (Table 3.1). These receptors are involved in both sensing and transmitting the siderophore-mediated signal that induces their own synthesis, and in the uptake of these siderophores [59]. An alignment of the sequences of the putative foxA promoter with that of the E. coli fecA promoter has revealed the putative FoxI recognition sequence as -35 GAAAAT and -10 TGTCGG [72]. Moreover, the specific interaction of FoxI with its sigma factor regulator FoxR has been also demonstrated experimentally [42]. FiuA and FoxA proteins share high sequence identity (40%). In fact, FiuA can functionally replace FoxA for the uptake of ferrioxamine B in the absence of this main ferrioxamine B receptor and vice versa [61, 59]. However, the expression of the ferrichrome FiuA receptor is not induced by ferrioxamine B [59]. Whether the FiuA and FoxA receptors are the only genes controlled by ferrichrome and ferrioxamine B, respectively, is not known. A protein, FoxB, has been identified that is involved in both ferrichrome and ferrioxamine B cytoplasmic membrane transport [73]. The P. aeruginosa foxB gene is located directly downstream of foxA, although it seems to form a different transcriptional unit. Since this protein mediated the transport of both ferrichrome and ferrioxamine B, it is possible that its expression is regulated by both sigma factors FiuI and FoxI. Orthologues to the P. aeruginosa ferrichrome and ferrioxamine B-mediated signalling pathways are present in most of the Pseudomonas species (Table 3.2). P. aeruginosa also contains a TonB-dependent receptor for the uptake of (carboxy)mycobactin, the FemA receptor [44] (Table 3.1). Mycobactin and carboxymycobactin are produced by Mycobacterium species [74]. Whereas mycobactin is located at the cell surface, carboxymycobactin is secreted by the cell, and it is the main extracellular iron binding agent for some pathogenic mycobacteria such as Mycobacterium tuberculosis and M. avium [74]. Although the FemA receptor is the main P. aeruginosa mycobactin receptor, these siderophores can also be taking up by other P. aeruginosa receptors since their uptake is not completely abolished in a
3
Cell-Surface Signalling in Pseudomonas
75
femA mutant [44]. The expression of the FemA mycobactin receptor is probably regulated through FemI and FemR, whose genes are located adjacent to the mycobactin receptor (Table 3.1) [44]. The FiuIRA, FoxIRA and FemIRA signalling systems show mechanistically high similarities to the well-known E. coli FecIRA CSS system [68]. However, whereas the sigma factor regulator FecR is required for maximal activity of its sigma factor [75], mutants in fiuR, foxR and femR constitutively express the FiuA, FoxA and FemA receptors, respectively, showing that they are not needed for their corresponding sigma factor activity ( [59] and unpublished data).
3.3.4 Haem-Mediated Signalling Pathway As many other pathogens, P. aeruginosa is able to use free haem or haem complexed to haemoglobin as a source of iron. Haem from intracellular host haemoglobin becomes available as an iron source after lysis of erythrocytes. P. aeruginosa secretes several factors that have the potential to facilitate this process such as phospholipase C and rhamnolipids, which both have haemolysin activity, and exotoxin A, which has broad cytotoxic activity towards eukaryotic cells [76]. P. aeruginosa possesses two distinct haem-acquisition systems, the Phu and the Has systems [60] (Fig. 3.4A). In general haem acquisition systems can be separated in three categories [77]: the fist one comprise uptake systems similar to those for siderophores that involve a specific TonB-dependent receptor and an ABC transport system; the second one consist of an outer-membrane receptor, an extracellular haem binding protein (haemophore) and an ABC export system; and the third one involves a haem-binding outer-membrane lipoprotein. The P. aeruginosa Phu system belongs to the first category of haem-acquisition systems [60]. It is composed by six proteins: a TonB-dependent outer membrane haem receptor, PhuR, two periplasmic binding proteins PhuS and PhuT, and three cytoplasmic membrane proteins: PhuU with permease activity, PhuV with ATPase activity, and PhuW (Fig. 3.4A). The Phu system seems to be regulated directly via Fur and not through a CSS system. The TonB-dependent receptor involved in this process, PhuR, also does not contain the N-terminal signalling domain (Fig. 3.4B). Although the growth of a P. aeruginosa phuR mutant in haemoglobin as sole iron source is severely impaired, it is not completely abolished [60], which implicated the presence of other haem-acquisition system(s). One of these systems is the P. aeruginosa Has system, which belongs to the second category of haemacquisition systems. The hasA gene is coding for the extracellular haem-binding protein, the HasA haemophore [78]. Downstream of P. aeruginosa hasA gene is an operon of three genes encoding factors with high homology to ABC-exporter proteins required for HasA secretion [60] (Fig. 3.4A). Immediately upstream of hasA and clustered in the same operon is a gene encoding a TonB-dependent transducer, the hasR gene [60] (Fig. 3.4A). By analogy with the well-studied haemophoremediated signalling pathway of Serratia marcescens [79] HasR probably forms a
76
M.A. Llamas and W. Bitter
Fig. 3.4 (a). Genetic organization of the P. aeruginosa Has (grey arrows) and Phu (black arrows) haem-acquisition systems. The arrows represent the different genes, their relative sizes, and the transcriptional orientation. The promoter regions are also indicated (bented arrows). Four of them contain a putative Fur box (b). N-terminal extension of mature P. aeruginosa HasR compared to mature P. aeruginosa PhuR. The numbering of amino acid residues is indicated
CSS system with the ECF sigma factor HasI and the regulator HasS encoded by the adjacent operon (Fig. 3.4A) (Table 3.1). This haemophore-dependent haem-uptake system seems to be the high-affinity transport system for haem, whereas the Phu system would be a lower-affinity haem-acquisition system [78, 60]. The P. aeruginosa genome contains another locus that is probably involved in haem transport, encoded by PA1300-1301-1302. Also this locus seems to be regulated via a CSS system, although experimental details are lacking.
3.3.5 Other CSS Pathways Involved in the Regulation of Metal Uptake The regulons of three other P. aeruginosa iron starvation sigma factors, PA0149, PA2050 and PA2093, have recently been determined by microarray analysis using cells overexpressing the respective sigma factors [44]. These analyses have shown that these sigma factors regulate the expression of TonB-dependent transducers encoded by the same locus (Table 3.1). Interestingly, both sigma factors PA2050 and PA2093 also control the expression of a second TonB-dependent receptor, PA2070 and PA2590 respectively (Table 3.1). Since all of these receptors have a signalling domain, it is likely that these receptors are also involved in the CSS signalling pathway, although this has not been experimentally demonstrated yet.
3
Cell-Surface Signalling in Pseudomonas
77
Microarray analysis also revealed that PA0149 and PA2050 not only control the transcription of their cognate outer membrane transducer(s), but also that both sigma factors regulate the expression of another locus, PA2403-PA2410 [44]. Interestingly, these genes, which are located within the pyoverdine locus of P. aeruginosa, are also regulated by the PvdS sigma factor [62]. This tightly controlled locus is coding for a cytoplasmic membrane transport system. Therefore, possibly this transport system is not unique for pyoverdine, but is involved in the transport of a range of siderophores or other high-affinity metal scavengers. These three ECF sigma factors also control the expression of PA2384, which encodes a Fur-like protein that was shown recently to be important or the expression of pyoverdine and pyochelin genes [80].
3.3.6 Regulation of Pyocins by a CSS System Although most CSS systems regulate the uptake of iron, one of the iron starvation sigma factors, PA4896, controls also the expression of the bacteriophage-like R2/F2 pyocins and pyocin S5 of P. aeruginosa PAO1 [44]. Pyocins are P. aeruginosa bacteriocins that are normally effective against the same or closely related species. Three different classes of pyocins have been identified: S-type, R-type and F-type [81]. S-type pyocins are colicin-like proteins with DNase activity, whereas R-type and F-type resemble phage tails that cause depolarization of the cytoplasmic membrane and inhibit active transport. The reason why P. aeruginosa have a CSS system involved in pyocin synthesis regulation is still unknown. Since the outer membrane transducers of these regulatory systems are normally involved in both (siderophore) uptake and signal transduction, it is possible that P. aeruginosa uses this system to ‘steal’ siderophores from other bacteria and simultaneously uses it to sense the presence of these bacteria and kill them. An orthologue of this CSS system is present in P. entomophila (Table 3.2). This bacterium also contains a gene cluster encoding a bacteriophage-like pyocins (PSEEN4129 to PSEEN4184). It would therefore be very interesting to determine if the P. entomophila CSS system is also involved in the synthesis of pyocins. Moreover, the identification of the signal activating this P. aeruginosa and P. entomophila signalling system would provide a clue to determine the real function of this interesting system.
3.3.7 Regulation of Virulence by a CSS System One of the most interesting P. aeruginosa CSS systems is the one formed by PA0674-PA0675 and PA0676, which encodes a putative receptor, an ECF sigma factor and a sigma factor regulator, respectively (Table 3.1). The first interesting difference with all other CSS systems is that the PA0674 receptor gene forms a single operon with the ECF-encoding gene, whereas the sigma factor regulator-encoding gene is located in a different transcriptional unit. Another obvious difference with
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M.A. Llamas and W. Bitter
other CSS systems is the size of the receptor protein. This protein (PA0674) is considerably smaller (23 KDa) than that of other CSS outer membrane receptors (75–85 KDa). The N-terminal half of this protein is homologous to the signalling domain of TonB-dependent transducers, and therefore might interact with the sigma factor regulator. However, the PA0674 receptor does not have the C-terminal β-barrel domain typical of TonB-dependent receptors. It is thus possible that, in contrast to the traditional receptors that have a role both in signalling and in the uptake of their ligands, this protein is only involved in signalling. The third important characteristic of this CSS system is that although there is a functional Fur box in the promoter region upstream of PA0674-0675 [82], this CSS system does not seem to be mainly involved in the regulation of iron uptake. Microarray analysis showed that this CSS system, designated PUMA3, regulates the expression of genes encoding secreted proteins and components of secretion systems (Llamas et al., in preparation). These include components of a type II secretion system (the Hxc system) involved in the secretion of a lowmolecular weight alkaline phosphatase [83], and a putative two-partner secretion system (TPS). TPS systems mediate the secretion of high-molecular-weight surfaceexposed proteins that are often involved in cell adhesion and pathogen dissemination [84]. Interestingly, this CSS system seems to be induced by a host signal [85]. Although the role of most PUMA3-induced genes has not been established yet, this CSS system seems to be involved in the regulation of P. aeruginosa virulence. In fact, the P. aeruginosa PUMA3-induced strain, by overexpression of the ECF sigma factor, is more virulent than the wild-type strain in zebrafish (Danio rerio) embryos (Llamas et al., in preparation). The PUMA3 system is, however, not the first example of a CSS system involved in the positive regulation of virulence upon interaction with the host. The plant pathogen Ralstonia solanacearum, the causal agent of bacterial wilt, uses a CSS network to regulate the expression of a type III protein secretion system in response to a non-diffusible plant signal [86]. This means that, although the involvement of CSS in virulence seems to be more an exception than a rule [5], CSS can be used for broader purposes than iron uptake only. Homologues of the P. aeruginosa PUMA3 receptor PA0674 are present in the genome of P. fluorescens and P. entomophila (Table 3.2), as well as in Rhodopseudomonas palustris and some Burkholderia species. In all these species, the PA0674 homologue is associated with the two other components of PUMA3. Both P. fluorescens and P. entomophila contain an Hxc-like type II secretion system immediately downstream of PUMA3, and P. fluorescens also a TPS-like system. In R. palustris, the PUMA3 cluster is repeated three times in the same genetic locus of the genome. Two of these clusters are followed by a homologue of the secreted component of the TPS system (the large exoprotein), and the third one by the second component of the TPS pathway. In B. cepacia homologues of some of the hxc-like genes are located upstream of PUMA3, interrupted by a large exoprotein-like gene. In this bacterium, the second component of the TPS pathway is located downstream of PUMA3. This gene association further suggests a functional interaction between PUMA3 and especially the TPS system.
3
Cell-Surface Signalling in Pseudomonas
79
3.4 Cell-Surface Signalling Systems in Other pseudomonas 3.4.1 P. putida P. putida KT2440, like P. aeruginosa PAO1, contains in total 19 ECF sigma factors [87]. Thirteen of these P. putida ECF sigma factors show high similarity to the iron starvation sigma factor FecI of E. coli. Except PP4208 and PP4244, which encode for homologues to P. aeruginosa FpvI and PvdS proteins, respectively, all P. putida ECF sigma factors are clustered with a sigma factor regulator, and except PP0704, also with a TonB-dependent transducer (Table 3.3). This suggests the involvement of these sigma factors in a CSS pathway. The protein encoded by the PP4244 gene, which is located within a siderophore biosynthetic locus, is almost identical to the PfrI sigma factor characterized in P. putida WCS358 as an activator protein that regulates pseudobactin 358 biosynthesis under iron restricted conditions [91]. Pseudobactin 358 is taken up by the PupA TonB-dependent outer membrane receptor [49]. PupA contains an N-terminal extension, and although its role in signalling has not been analyzed, by analogy with the P. aeruginosa pyoverdine-mediated CSS system, it is likely to be involved in a pseudobactin 358-mediated CSS system that allows the activation of PfrI. Another CSS system of this P. putida strain has been studied in more detail, the PupIRB system [34]. This CSS system was, together with the E. coli FecIRA system, the first CSS system reported. P. putida WCS358 is able to use the heterologous siderophores pseudobactin BN7 and BN8 produced by other Pseudomonas strains, and the uptake of these siderophores is regulated by the PupIRB CSS system. The sequenced P. putida KT2440 strain also seems to contain a Pup CSS system encoded by the PP0667-0668-0669 genes [87] (Table 3.3). This system is not present in P. aeruginosa. The product of the P. putida KT2440 PP4208 gene shows high sequence identity to P. aeruginosa FpvI (68%) (Table 3.3). However, unlike in P. aeruginosa, this fpvI gene is not clustered with an FpvR homologue. This situation is in fact more common in Pseudomonas species, P. aeruginosa is the exception (Table 3.3). In most species fpvI is clustered with pyoverdine synthetic genes, whereas fpvR is located in a completely different locus of the genome. This isolated position of fpvR is unusual since the genes for ECF sigma factors and their cognate sigma factors regulators are normally clustered together (Table 3.3). The functional characterisation of these FpvR proteins would be interesting, but has not been performed yet. One very interesting P. putida ECF sigma factor is the one encoded by the PP2192 gene (Table 3.3). PP2192 is a hybrid gene which encodes the sigma factor in the N-terminal part and the sigma factor regulator in the C-terminal part. If this hybrid gene is functional, it has implications to the mechanism of signalling, because a permanent interaction would occur and signalling would need to occur solely via conformational changes. Alternatively, proteolysis of the linker region between the sigma and the sigma factor regulator domain could liberate the sigma factor, as has been suggested for FecR in E. coli. Analysis of this hybrid protein would help in understanding the role of the sigma factor regulators, especially of these that seem to function as both chaperones and anti-sigma factors.
PMEN2873
P. mendocina ymp
P. syringae pv. syringae B728a
PA2426/PvdS (41%) –
PMEN2673
P. stutzeri A1501
PA3899/FecI (71%)
–
PA2426/PvdS (84%)
PSYR1040
PSYR1107
PSYR1943
PMEN4508
PA0472/FiuI (73%)
PST4176
Pseudomonas sp.
PA1912/FemI (52%)
ECF sigmaa
% Identitity with P. aeruginosa ECF
–
PSYR1106 (F)
PSYR1039 (F)
PMEN4509 (F)
–
PMEN2674 (F)
PST4175 (F)
Sigma factor regulatorb
PSYR1961 (F)
PSYR1105 (F)
PSYR1038 (F)
PMEN4510 (F)
–
PMEN2675 (F)
PST4174 (F)
TonB-dependent transducerb, c
Table 3.3 Pseudomonas sp. CSS systems
Haem/haemoglobin receptor (65% to P. syringae PSYR1105) Ferric citrate receptor FecA (66% to P. aeruginosa PA3901/FecA) Haem/haemoglobin receptor (66% to P. aeruginosa PhuR) Ferripyoverdine receptor (38% to P. aeruginosa FpvA)
Siderophore receptor (64% to P. putida PP0160 and 57% to P. aeruginosa PA2466/FoxA) Ferrichrome receptor FiuA (59% to P. aeruginosa PA0470/FiuA) –
Transducer functiond
[88, 55]
[88]
[88]
–
–
–
–
Reference
80 M.A. Llamas and W. Bitter
P. entomophila L48
P. syringae pv. tomato DC3000
PA0472/FiuI (78%) PA0472/FiuI (77%)
PSYR4731
PSPTO2358 (50% to P. aeruginosa FpvR) PSEEN0275 (F) PSEEN0618 (F)
PSPTO2133 PA2426/PvdS (97% to (85%) PSYR1943)
–
PA4896 (80%)
PSEEN0274
PSEEN0619
PSEEN0620 (R)
PSEEN0276 (F)
PSPTO2151 (F)
PSPTO1284 (F)
PSPTO1206 (F) PSPTO1207 (F)
PSPTO1204 (F) PSPTO1208 (F)
PSPTO1285 (F)
–
–
PSYR2582 (F)
PSPTO0445 (F)
PSYR4730 (F)
PSYR2581 (F)
PSPTO1286 – (88% to PSYR1107)
PSPTO0444 (92% to PSYR4731) PSPTO1203 – PA3899/FecI PSPTO1209 (68%) (92% to PSYR1040)
PA1912/FemI (57%)
PSYR2580
Siderophore receptor (74% to P. aeruginosa PA4897)
Siderophore receptor
Siderophore receptor Ferric citrate receptor FecA (65% to P. aeruginosa PA3901/FecA) Haem/haemoglobin receptor (67% to P. aeruginosa PA4710/PhuR) Ferripyoverdine receptor (89% to PSYR1961)
–
Siderophore receptor (41% to P. aeruginosa PA1910/FemA) –
–
–
[88, 55]
[88]
[88] [88]
[88]
[88]
[88]
3 Cell-Surface Signalling in Pseudomonas 81
Pseudomonas sp.
% Identitity with P. aeruginosa ECF
–
PvdS (90%)
PA0675 (69%)
PA2468/FoxI (51%)
PA2387/FpvI (72%)
–
ECF sigmaa
PSEEN1037
PSEEN1814
PSEEN2108
PSEEN2531
PSEEN3663
PSEEN4328
PSEEN2995 (54% to P. aeruginosa FpvR) PSEEN4327 (F)
PSEEN2530 (F)
PSEEN2995 (54% to P. aeruginosa FpvR) PSEEN2109 (F)
PSEEN1038 (F)
Sigma factor regulatorb
PSEEN4326 (F)
–
PSEEN2529 (F)
PSEEN2107
–
PSEEN1039 (F)
TonB-dependent transducerb, c
Table 3.3 (continued)
Siderophore receptor (43% to P. aeruginosa PA1910/FemA)
45% identical to P. aeruginosa PA0674 Siderophore receptor (42% to pseudobactin BN7/BN8 PupB receptor and 41% to P. aeruginosa FpvA) –
Siderophore receptor (92% to PP0865 and 66% to P. aruginosa PA3268) –
Transducer functiond
–
–
–
–
–
–
Reference
82 M.A. Llamas and W. Bitter
PA1300 (57%)
–
PA0149 (68%)
PA3899/FecI (68%)
PA3410/HasI (60%)
PA0472/FiuI (75%)
PSEEN4331
PSEEN4420
PSEEN4567
PSEEN4571
PSEEN4768
PSEEN5131
PSEEN5132 (F)
PSEEN4767 (F)
PSEEN4572 (F)
PSEEN4566 (F)
PSEEN4421 (F)
PSEEN4332 (F)
PSEEN5133 (F)
PSEEN4766 (F)
PSEEN4573 (F)
PSEEN4565 (F)
PSEEN4422 (F)
PSEEN4333 (F)
Haem/heme receptor (58% to P. aeruginosa PA1302) Haem/haemoglobin receptor (87% to P. putida PP1006 and 68% to P. aeruginosa PA4710/PhuR) Siderophore receptor (66% to P. aeruginosa PA0151) Ferric citrate receptor FecA (93% to P. putida PP4613/FecA and 70% to P. aeruginosa PA3901/FecA) Haem receptor HasR (51% to P. aeruginosa PA3408/HasR) Ferrichorme receptor (67% to P. aeruginosa PA0470/FiuA) –
–
–
–
–
–
3 Cell-Surface Signalling in Pseudomonas 83
P. putida KT2440
Pseudomonas sp.
PA2468/FoxI (70%)
PA0472/FiuI (79%)
–
PA2468/FoxI (52%) PA1912/FemI (51%) –
–
– PA1912/FemI (53%)
PP0352
PP0667
PP0704
PP0865
PP1008
PP2192 PP3086
% Identitity with P. aeruginosa ECF
PP0162
ECF sigmaa
PP2192 PP3085 (F)
PP1007 (F)
PP0866 (F)
PP0703 (F) and/or PP0700 (F)
PP0668 (F)
PP0351 (F)
PP0161 (F)
Sigma factor regulatorb
PP2193 (F) PP3084 (F)
PP1006 (F) (no N-terminal extension)
PP0867 (F)
–
PP0669 (F)
PP0350 (F)
PP0160 (F)
TonB-dependent transducerb, c
Tabke 3.3 (continued)
Siderophore receptor (77% to P. aeruginosa PA3268) Haem/haemoglobin receptor (67% to P. aeruginosa PA4710/PhuR) Siderophore receptor Siderophore receptor (39% to P. aeruginosa PA1910/FemA)
Ferrioxamine receptor (60% to P. aeruginosa PA2466/FoxA) Ferrichrome receptor (66% to P. aeruginosa PA0470/FiuA) Pseudobactin BN7/BN8 receptor (38% to PupB)
Transducer functiond
[87] [87]
[87]
[87]
[87]
[87]
[87]
[87]
Reference
84 M.A. Llamas and W. Bitter
P. fluorescens Pf-5
PA3899/FecI (66%)
PP4611
PA2468/FoxI (53%) PA1912/FemI (52%)
PA0149 (58%)
PP4608
PFL0145
PA2426/PvdS (86%)
PP4244/PfrI
PA2468/FoxI (72%)
PA2387/FpvI (68%)
PP4208
PFL0127
PA1912/FemI (52%)
PP3577
PFL0146 (F)
PFL0126 (F)
PP4612 (F)
PP4607 (F)
PP3555 (55% to P. aeruginosa FpvR)
PP3555 (55% to P. aeruginosa FpvR)
PP3576 (F)
PFL0147 (F)
PFL0125 (F)
PP4613 (F)
PP4606 (F)
PP4217
PP4217
PP3575 (F)
Siderophore receptor (41% to P. aeruginosa PA1910/FemA) Ferripyoverdine receptor (49% P. aeruginosa PA4168/FpvB) Ferripyoverdine receptor (49% P. aeruginosa PA4168/FpvB) Siderophore receptor (66% to P. aeruginosa PA0151) Ferric citrate receptor FecA (68% to P. aeruginosa PA3901/FecA) Ferrioxamine receptor (63% to P. aeruginosa PA2466/FoxA) Siderophore receptor –
–
[87]
[87]
[87]
[87]
[87]
3 Cell-Surface Signalling in Pseudomonas 85
Pseudomonas sp.
PA0149 (65%)
PA1300 (55%)
#x2013; PA3899/FecI (53%) PA1912/FemI (51%) –
PA3899/FecI (50%) PA1912/FemI (48%) PA2468/FoxI (47%)
PFL0988
PFL1373
PFL2291 PFL2363
PFL2529
PFL2393
PA3899/FecI (65%)
% Identitity with P. aeruginosa ECF
PFL0984
ECF sigmaa
PFL2391 (F)
PFL2527 (F)
PFL2528 (F)
PFL2293 (F) PFL2365 (F)
PFL1371 (F)
PFL0995 (F)
PFL0982 (F)
TonB-dependent transducerb, c
PFL2392 (F)
PFL2292 (F) PFL2364 (F)
PFL1372 (F)
PFL0989 (F)
PFL0983 (F)
Sigma factor regulatorb
Tabke 3.3 (continued)
Ferric citrate receptor FecA (69% to P. aeruginosa PA3901/FecA) Siderophore receptor (64% to P. aeruginosa PA0151) Haem receptor (60% to P. aeruginosa PA1302) Siderophore receptor Siderophore receptor (69% to P. aeruginosa PA4897) Ferripyoverdine receptor (68% to P. aeruginosa PA2398/FpvA) Siderophore receptor
Transducer functiond
–
–
– –
–
–
–
Reference
86 M.A. Llamas and W. Bitter
–
PA1912/FemI (50%) PA0472/FiuI (47%) PA2468/FoxI (46%) PA1912/FemI (52%) PA3899/FecI (48%) PA2468/FoxI (46%) PA0472/FiuI (45%) – –
PA2387/FpvI (71%)
PFL3156
PFL3313
PFL3610 PFL4041
PFL4080
PFL3483
PA0675 (73%)
PFL2746
PFL2903 (51% to P. aeruginosa FpvR)
PFL3611 (F) PFL4040 (F)
PFL3484 (F)
PFL3314 (F)
PFL3155 (F)
PFL2747 (F)
PFL4092
PFL3612 (F) PFL4039 (F)
PFL3485 (F)
PFL3315 (F)
PFL3154 (F)
PFL2745 (F)
Ferripyoverdine receptor (62% to P. aeruginosa PA2398/FpvA) Siderophore receptor Siderophore receptor (75% to P. aeruginosa PA3268) Ferripyoverdine receptor (53% to P. putida PP4217, and 46% P. aeruginosa PA4168/FpvB)
53% identical to P. aeruginosa PA0674 Siderophore receptor (77% to P. putida PP2193) Siderophore receptor
–
– –
–
–
–
–
3 Cell-Surface Signalling in Pseudomonas 87
PFL5379 (F)
–
PA3410/HasI (61%)
PA0472/FiuI (77%)
PFL4625
PFL5380
PFL5704
PFL5705 (F)
PFL2903 (51% to P. aeruginosa FpvR) PFL4626 (F)
Sigma factor regulatorb
PFL4190/PbrA PA2426/PvdS (91%)
% Identitity with P. aeruginosa ECF
PFL5706 (F)
PFL5378 (F)
PFL4627 (F)
PFL4092
TonB-dependent transducerb, c
Haem/haemoglobin receptor (66% to P. aeruginosa PA4710/PhuR) Haem/haemoglobin receptor HasR (75% to P. aeruginosa PA3408/HasR) Ferrichrome receptor (66% to P. aeruginosa PA0470/FiuA)
Transducer functiond
b
Number in the Pseudomonas genome annotation project (http://www.pseudomonas.com) The orientation of the sigma factor regulator and transducer genes to the ECF sigma factor is indicated (F, forward; R, reverse) c The identity of the transducer protein to other Pseudomonas receptors is indicated d All transducers, except PP1006, contain an N-terminal extension
a
Pseudomonas sp.
ECF sigmaa
Table 3.3 (continued)
–
–
–
[89, 90]
Reference
88 M.A. Llamas and W. Bitter
3
Cell-Surface Signalling in Pseudomonas
89
3.4.2 P. fluorescens Genome analysis of P. fluorescens Pf-5 reveals the presence of an impressive total of 28 ECF sigma factors. Twenty of these display substantial sequence similarity with iron starvation sigma factors and seems to be part of a CSS system (Table 3.3). By homology with other Pseudomonas ECF sigma factors, it seems that most of these P. fluorescens iron starvation sigma factors are involved in the regulation of iron uptake (Table 3.3). However, only two of these sigma factors have been analysed. P. fluorescens contains a homologue to the P. aeruginosa PvdS sigma factor that in the strain Pf-5 is encoded by the PFL4190 gene (Table 3.3). The function of this sigma factor has been analysed in another P. fluorescens strain, the M114 strain, where it has been called PbrA (for pseudobactin regulation activation) [92]. Under iron-restricted conditions, P. fluorescens M114 produces and secretes a siderophore, the pseudobactin M114. PbrA is needed for full transcription of pseudobactin M114 biosynthetic genes, but this sigma factor also regulates the expression of a metalloprotease gene, aprA [89]. This gene encodes for the major extracellular protease produced by P. fluorescens M114. Expression of aprA is, however, not PvdSdependent in P. aeruginosa [62]. This discrepancy is not so strange since PvdS seems to have a ‘core’ regulon common to all Pseudomonas species, which contains genes involved in pyoverdine/pseudobactin synthesis, and an ‘extended’ regulon, which is specific for each species [55]. P. fluorescens also contains a homologue of the P. aeruginosa FpvI ECF sigma factor, encoded in strain Pf-5 by the PFL4080 gene (Table 3.3). In P. fluorescens, like in P. aeruginosa, this sigma factor controls the expression of the TonB-dependent receptor gene fpvA [90]. In addition, and very unexpectedly, P. fluorescens FpvI also controls the expression of a fpvR homologue (the PFL2903 gene in strain Pf-5) located in a different locus of the P. fluorescens genome [90]. Since in P. aeruginosa FpvR controls the activity of both PvdS and FpvI sigma factors, it would be interesting to determine the effect of the FpvI-dependent fpvR expression in P. fluorescens on the activity of these sigma factors and on pyoverdine (pseudobactin) production.
3.4.3 P. syringae and P. entomophila Despite the fact that P. syringae has a genome similar in size as compared those of other Pseudomonas species (6–7 Mbp), only 10 ECF sigma factors have been identified in P. syringae phatovars sequenced to date [88]. This is significantly lower than the 19 ECF sigma factors present in P. aeruginosa and P. putida [87, 2], and the 28 ECF sigma factors predicted in P. fluorescens. Only five of these P. syringae ECF sigma factors show high homology to iron starvation sigma factors and seems to be part of a CSS system [88] (Table 3.3). Since it seems to be a correlation between the number of CSS systems and bacterial lifestyles, the reduced number of CSS systems in P. syringae may indicates an adaptation to its specialized lifestyle as a plant pathogen. All P. syringae CSS systems have a counterpart in one of the other
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M.A. Llamas and W. Bitter
Pseudomonas, and they seem to be mainly involved in the regulation of different iron transport systems (Table 3.3). P. entomophila contains fourteen putative CSS systems (Table 3.3), none of these have been characterized yet. Most of these systems have homology to P. aeruginosa and P. putida CSS systems suggesting a similar role in P. entomophila. As usual in Pseudomonas, most of the P. entomophila CSS systems seem to be involved in the regulation of iron uptake.
3.5 Concluding Remarks Iron is an essential element for most microorganisms, but the struggle for this element is probably nowhere in nature more clear than in the Pseudomonads. They have evolved a staggering amount of iron-uptake systems to be able to win the competition against their neighbours. A number of these uptake systems are regulated by CSS systems, which are cleverly making use of the TonB-dependent outer membrane receptors involved in iron transport. Recently a number of details of the CSS systems have been unravelled, especially for the signal transduction process at the outer membrane. Structural data of the signalling domain of the TonB-dependent transducers and the interaction with the TonB protein has greatly accelerated this research. More unclear is the role of the sigma factor regulator in the cytoplasmic membrane. This protein with only a single transmembrane domain has to transduce somehow the signal from the periplasm to the ECF alternative sigma factor in the cytoplasm. Although this protein clearly represses the function of the ECF sigma factor in the absence of the inducing signal, it is also required for ECF sigma factor activity in a number of CSS systems. Recent data shows that protease(s) probably play a role in this process, but the nature of this protease and its exact function still have to be clarified. Although most CSS systems are involved in the acquisition of iron, some of them seem to have a different or alternative function. For instance, CSS systems can regulate the production of bacteriocins in addition to iron uptake systems, which could be used to reduce the competition. Furthermore, the pyoverdine CSS system of P. aeruginosa not only controls siderophore biosynthesis and uptake, but regulates also the production of two virulence factors. In this respect, this pyoverdine is in fact similar to the well-known autoinducers, adding an extra layer of complexity to the already elaborate autoinducer system in this organism. Furthermore, sequence data indicate that this siderophore-autoinducer situation might also be present in a range of other fluorescent Pseudomonads. But there is more to CSS systems as was shown by the very recent discovery of the unusual CSS system PUMA3 in P. aeruginosa. This system, which has a remarkable receptor and an aberrant gene organisation, seems to be solely involved in regulation of virulence. Our future experiments will be directed to unravel this latest addition to CSS systems and to find its inducing signal. It can be concluded that CSS systems play an important and underestimated role in gene regulation in Pseudomonads, a role that is beyond the acquisition of iron.
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56.
57.
58.
59.
60.
61. 62.
63. 64.
65.
66. 67. 68. 69. 70. 71. 72. 73.
74. 75.
M.A. Llamas and W. Bitter DC3000 reveals regulon members and insights regarding PvdS function in other pseudomonads. Mol. Microbiol. 68: 871–889. Tiburzi, F., Imperi, F. and Visca., P. (2008) Intracellular levels and activity of PvdS, the major iron starvation sigma factor of Pseudomonas aeruginosa. Mol. Microbiol. 67: 213–227. Maeda, H., Fujita, N. and Ishihama., A. (2000) Competition among seven Escherichia coli sigma subunits: relative binding affinities to the core RNA polymerase. Nucleic. Acids Res. 28: 3497–3503. Ambrosi, C., Tiburzi, F., Imperi, F., Putignani, L. and Visca., P. (2005) Involvement of AlgQ in transcriptional regulation of pyoverdine genes in Pseudomonas aeruginosa PAO1. J. Bacteriol. 187: 5097–5107. Llamas, M.A., Sparrius, M., Kloet, R., Jimenez, C.R., Vandenbroucke-Grauls, C. and Bitter., W. (2006) The heterologous siderophores ferrioxamine B and ferrichrome activate signaling pathways in Pseudomonas aeruginosa. J. Bacteriol. 188: 1882–1891. Ochsner, U.A., Johnson, Z. and Vasil, M.L. (2000) Genetics and regulation of two distinct haem-uptake systems, phu and has, in Pseudomonas aeruginosa. Microbiology 146(Pt 1): 185–198. Banin, E., Vasil, M.L. and Greenberg., E.P. (2005) Iron and Pseudomonas aeruginosa biofilm formation. Proc. Natl. Acad. Sci. U.S.A 102: 11076–11081. Ochsner, U.A., Wilderman, P.J., Vasil, A.I. and Vasil., M.L. (2002) GeneChip expression analysis of the iron starvation response in Pseudomonas aeruginosa: identification of novel pyoverdine biosynthesis genes. Mol. Microbiol. 45: 1277–1287. Meyer, J.M., Neely, A., Stintzi, A., Georges, C. and Holder., I.A. (1996) Pyoverdin is essential for virulence of Pseudomonas aeruginosa. Infect. Immun. 64: 518–523. Takase, H., Nitanai, H., Hoshino, K. and Otani., T. (2000) Impact of siderophore production on Pseudomonas aeruginosa infections in immunosuppressed mice. Infect. Immun. 68: 1834–1839. Leoni, L., Ciervo, A., Orsi, N. and Visca., P. (1996) Iron-regulated transcription of the pvdA gene in Pseudomonas aeruginosa: effect of Fur and PvdS on promoter activity. J. Bacteriol. 178: 2299–2313. Frost, G.E. and Rosenberg., H. (1973) The inducible citrate-dependent iron transport system in Escherichia coli K12. Biochim. Biophys. Acta 330: 90–101. Hussein, S., Hantke, K. and Braun., V. (1981) Citrate-dependent iron transport system in Escherichia coli K-12. Eur. J. Biochem. 117: 431–437. Braun, V. (1997) Surface signaling: novel transcription initiation mechanism starting from the cell surface. Arch. Microbiol. 167: 325–331. Cox, C.D. (1980) Iron uptake with ferripyochelin and ferric citrate by Pseudomonas aeruginosa. J. Bacteriol. 142: 581–587. Harding, R.A. and Royt., P.W. (1990) Acquisition of iron from citrate by Pseudomonas aeruginosa. J. Gen. Microbiol. 136: 1859–1867. Poole, K. and McKay., G.A. (2003) Iron acquisition and its control in Pseudomonas aeruginosa: many roads lead to Rome. Front. Biosci. 8: d661–d686. Enz, S., Mahren, S., Menzel, C. and Braun., V. (2003) Analysis of the ferric citrate transport gene promoter of Escherichia coli. J. Bacteriol. 185: 2387–2391. O’Cuiv, P.O., Keogh, D., Clarke, P. and O’Connell., M. (2007) FoxB of Pseudomonas aeruginosa functions in the utilization of the xenosiderophores ferrichrome, ferrioxamine B, and schizokinen: evidence for transport redundancy at the inner membrane. J. Bacteriol. 189: 284–287. Ratledge, C. and Dover., L.G. (2000) Iron metabolism in pathogenic bacteria. Annu. Rev. Microbiol. 54: 881–941. Ochs, M., Veitinger, S., Kim, I., Welz, D., Angerer, A. and Braun., V. (1995) Regulation of citrate-dependent iron transport of Escherichia coli: fecR is required for transcription activation by FecI. Mol. Microbiol. 15: 119–132.
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76. Sadikot, R.T., Blackwell, T.S., Christman, J.W. and Prince., A.S. (2005) Pathogen-host interactions in Pseudomonas aeruginosa pneumonia. Am. J. Respir. Crit. Care Med. 171: 1209–1223. 77. Wandersman, C. and Delepelaire., P. (2004) Bacterial iron sources: from siderophores to hemophores. Annu. Rev. Microbiol. 58: 611–647. 78. Letoffe, S., Redeker, V. and Wandersman., C. (1998) Isolation and characterization of an extracellular haem-binding protein from Pseudomonas aeruginosa that shares function and sequence similarities with the Serratia marcescens HasA haemophore. Mol. Microbiol. 28: 1223–1234. 79. Rossi, M.S., Paquelin, A., Ghigo, J.M. and Wandersman., C. (2003) Haemophore-mediated signal transduction across the bacterial cell envelope in Serratia marcescens: the inducer and the transported substrate are different molecules. Mol. Microbiol. 48: 1467–1480. 80. Zheng, P., Sun, J., Geffers, R. and Zeng., A.P. (2007) Functional characterization of the gene PA2384 in large-scale gene regulation in response to iron starvation in Pseudomonas aeruginosa. J. Biotechnol. 132: 342–352. 81. Michel-Briand, Y. and Baysse., C. (2002) The pyocins of Pseudomonas aeruginosa. Biochimie 84: 499–510. 82. Ochsner, U.A. and Vasil., M.L. (1996) Gene repression by the ferric uptake regulator in Pseudomonas aeruginosa: cycle selection of iron-regulated genes. Proc. Natl. Acad. Sci. U.S.A 93: 4409–4414. 83. Ball, G., Durand, E., Lazdunski, A. and Filloux., A. (2002) A novel type II secretion system in Pseudomonas aeruginosa. Mol. Microbiol. 43: 475–485. 84. Jacob-Dubuisson, F., Locht, C. and Antoine., R. (2001) Two-partner secretion in Gramnegative bacteria: a thrifty, specific pathway for large virulence proteins. Mol. Microbiol. 40: 306–313. 85. Frisk, A., Schurr, J.R., Wang, G., Bertucci, D.C., Marrero, L., Hwang, S.H., Hassett, D.J. and Schurr., M.J. (2004) Transcriptome analysis of Pseudomonas aeruginosa after interaction with human airway epithelial cells. Infect. Immun. 72: 5433–5438. 86. Aldon, D., Brito, B., Boucher, C. and Genin., S. (2000) A bacterial sensor of plant cell contact controls the transcriptional induction of Ralstonia solanacearum pathogenicity genes. EMBO. J. 19: 2304–2314. 87. Martinez-Bueno, M.A., Tobes, R., Rey, M. and Ramos., J.L. (2002) Detection of multiple extracytoplasmic function (ECF) sigma factors in the genome of Pseudomonas putida KT2440 and their counterparts in Pseudomonas aeruginosa PA01. Environ. Microbiol. 4: 842–855. 88. Oguiza, J.A., Kiil, K. and Ussery., D.W. (2005) Extracytoplasmic function sigma factors in Pseudomonas syringae. Trends Microbiol. 13: 565–568. 89. Maunsell, B., Adams, C. and O’Gara., F. (2006) Complex regulation of AprA metalloprotease in Pseudomonas fluorescens M114: evidence for the involvement of iron, the ECF sigma factor, PbrA and pseudobactin M114 siderophore. Microbiology 152: 29–42. 90. Moon, C.D., Zhang, X.X., Matthijs, S., Schafer, M., Budzikiewicz, H. and Rainey., P.B. (2008) Genomic, genetic and structural analysis of pyoverdine-mediated iron acquisition in the plant growth-promoting bacterium Pseudomonas fluorescens SBW25. BMC Microbiol. 8: 7. 91. Venturi, V., Ottevanger, C., Bracke, M. and Weisbeek., P. (1995) Iron regulation of siderophore biosynthesis and transport in Pseudomonas putida WCS358: involvement of a transcriptional activator and of the Fur protein. Mol. Microbiol. 15: 1081–1093. 92. Sexton, R., Gill, P.R., Jr., Callanan, M.J., O’Sullivan, D.J., Dowling, D.N. and O’Gara., F. (1995) Iron-responsive gene expression in Pseudomonas fluorescens M114: cloning and characterization of a transcription-activating factor, PbrA. Mol. Microbiol. 15: 297–306.
Chapter 4
Second Messenger c-di-GMP Signaling in Pseudomonas aeruginosa Massimo Merighi and Steve Lory
4.1 Introduction Signal transduction in living cells refers to the conversion of various chemical or physical “input” stimuli into signals that ultimately lead to one or more “outputs” in the form of biological responses. A signal transduction pathway consists of a cascade of interactions between signals and receptors transmitted to effector proteins that undergo various post-translational and allosteric modifications to affect gene expression, protein activity or protein turnover. During their transmission the signals can be amplified by the activities of intermediate effectors or they can be integrated with cellular signals originating from different stimuli. Ultimately some basic cellular function is affected, often at the transcriptional level. The nomenclature established for eukaryotic signal transduction, refers to primary messengers as input signals, while second messengers transmit information intracellularly following the interaction of the input signal with a particular receptor. In eukaryotes, the primary messengers are often hormones, while a second messenger can be any one of the diverse array of molecules including, but not limited to, cyclic nucleotides (cAMP and cGMP), phosphatidylinositol diacylglycerol, inositol triphosphate, arachidonic acid, calcium ions, as well as gases like nitric oxide and carbon monoxide [1]. In bacteria, a variety of primary signals are known, including ions, metabolites, peptides, oxygen, carbon monoxide, light, NADH, FAD and ATP. Perception of these signals is mediated by receptor proteins and many of them belong to defined families of domains often fused to various transmitter modules of the two-component signal transduction family. A comprehensive analysis of sensory transduction modules and primary signals in bacteria can be found in a recent review and will not be discussed here further [2]. A number of prokaryotic regulatory pathways utilize second messenger purine nucleotides such as cAMP and, less frequently, cGMP. The nucleotides ppGpp and pppGpp also act as intracellular signals in the stringent response, an M. Merighi (B) Department of Microbiology and Molecular Genetics, Harvard Medical School, Boston MA 02115, USA e-mail:
[email protected] J.L. Ramos, A. Filloux (eds.), Pseudomonas, DOI 10.1007/978-90-481-3909-5_4, C Springer Science+Business Media B.V. 2010
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adaptive mechanism facilitating bacterial survival in various stress conditions, including nutrient limitation and osmotic challenge and during the growth into stationary phase transition. More recently, bis- (3 -5 )-cyclic dimeric guanosine monophosphate (c-di-GMP) has been recognized as a ubiquitous second messenger in bacteria. The general topic of c-di-GMP signaling has been the subject of several excellent reviews [3–11]. In this chapter, we will focus primarily on cyclicnucleotide signaling in Pseudomonas aeruginosa with an emphasis on the diverse cellular functions regulated by c-di-GMP.
4.2 Purine Nucleotides as Second Messengers 4.2.1 Stringent Response and (p)ppGpp. A well-studied signaling pathway is activated in response to cellular stress and is mediated by a second messenger pair of guanosine penta and tetraphosphate, pppGpp and ppGpp, respectively. Under conditions that restrict bacterial growth, referred to as the stringent response, the activation of the RelA protein results in the transfer of pyrophosphate to GTP to give pppGpp, followed by conversion of a portion to ppGpp. Reversal of the stringent response is accomplished by degradation of these nucleotides by the SpoT esterase. The most frequently encountered condition that triggers the stringent response is cellular stress and the main function of accumulated pppGpp and ppGpp is to modulate energetically-costly processes including the synthesis of components of the translational machinery [12]. The role of pppGpp and ppGpp has not been studied as extensively in P. aeruginosa as in Escherichia coli. The stringent response in P. aeruginosa very likely plays a similar role in survival and/or adaptation under conditions where cellular growth is affected by changing environmental conditions. Moreover, the regulatory role of pppGpp and ppGpp in P. aeruginosa appears to be broad and includes a control over the expression of various virulence phenotypes [13]. This effect may be related to the observation that an increase in pppGpp and ppGpp levels by overexpression of relA leads to enhanced expression of RpoS and, consequently, activation of quorum sensing even in the absence of high cell density. Therefore, pppGpp and ppGpp could play a role in bacterial response to stress conditions in colonized tissues. Recently, the stringent response was shown to be required for the successful adaptation of P. aeruginosa to anaerobic energy stress [14]. It is conceivable that pppGpp and ppGpp play a central role in adaptation of this organism to a wide range of niches including nutrient limiting natural environments or those encountered during infection in hosts with active immune defense mechanisms.
4.2.2 cAMP and cGMP Signaling Cyclic adenosine-3, 5 -monophosphate (cAMP) has been known to regulate a number of metabolic functions and, in different bacterial species, it has been implicated
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in the control of diverse cellular processes ranging from virulence to bioluminescence [15]. In Enterobacteriaceae, cAMP is synthesized from ATP by a single adenylate cyclase, the product of the cyaA gene. However, genomes of many microorganisms encode multiple putative paralogues of adenylate cyclases. Based on their sequence similarities, purine nucleotide cyclases were proposed to group into three different classes [16], and a fourth was recently proposed [17]. The enterobacterial cyclase (also found in most Gram-negative bacteria) belongs to the Class I purine cyclases and in. E. coli this enzyme is regulated by protein components of the sugar transport system and is inhibited when glucose is abundant [18]. The class II family is a relatively heterogeneous group of enzymes and they include several bacterial toxins secreted by pathogens such as Bacillus anthracis (Edema Factor, EF), Bordetella pertussis (CyaA), and Pseudomonas aeruginosa (ExoY) [19]. The largest group of cyclases are the enzymes in Class III that have been subdivided into additional families [20, 21]. The modular Class III adenylate cyclases often consist of fused regulator and catalytic domains, and, in many instances, they also contain putative transmembrane segments. The sequenced genomes of P. aeruginosa strains carry genes encoding a single class I enzyme (CyaA), a Class II adenylate cyclase exoenzyme Y, secreted by the type III secretion system, and a Class III enzyme CyaB [22]. cAMP is an important regulator of virulence in P. aeruginosa, primarily through its control over the expression of the type III secretion system [23]. Both CyaA and CyaB contribute to the maintenance of cAMP levels in the cells, however P. aeruginosa strains with a mutation in cyaB are more substantially attenuated in an acute mouse infection model than those lacking cyaA [24]. The major known function of cAMP in bacteria is to control transcription of genes following binding to a regulatory protein, referred to in E. coli as CRP, the cAMP receptor protein (it is also known as the catabolite activator protein, CAP). In E. coli, the cAMP-bound form of CRP forms a homodimer and regulates transcription initiation from more than 100 different promoters [25]. All bacteria with adenylate cyclases also express CRP orthologues and therefore the signal transduction mechanism may be conserved in all microorganisms, although targets of the cAMP-CRP complex may be different. In P. aeruginosa, the best characterized CRP homologue is the virulence factor regulator (Vfr), a transcription factor that controls the expression of quorum sensing, exotoxin A production, twitching motility mediated by type IV pili, the type II and type III secretion systems and flagellar biogenesis [26]. Vfr is 67 % identical to E. coli CRP and this level of sequence conservation is sufficient for Vfr to complement a crp mutation in E. coli [26]. Interestingly, in P. aeruginosa, Vfr does not appear to be involved in catabolite repression control. The repression of catabolic pathways for sugars and hydrocarbons when other preferred carbon sources are present in the culture medium involves the global regulatory protein Crc (for catabolite repression control) and is independent of cAMP [27, 28]. The occurrence of cyclic guanosine 3 , 5 -monophosphate (cGMP) is less common among prokaryotes but it has been described in Cyanobacteria, where it is as abundant as cAMP. All known guanylate cyclases of eukaryotic or cyanobacterial origin belong to class III of the purine nucleotidyl cyclase family [29]. Although
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cGMP participates in a multitude of signaling pathways in eukaryotic cells, its biological function in bacteria has been less well defined. Early reports suggested that exogenously added cGMP affected various biological functions in rhizobia [30, 31]. More recently, cGMP was shown to regulate certain ion channels in Mesorhizobium loti, Synechocystis and Magnetospirillum magnetotacticum [32–35] and cyanobacterial phytochrome systems, where it binds to GAF (cGMP-specific and -regulated cyclic nucleotide phosphodiesterase, Adenylyl cyclase, and E. coli transcription factor FhlA domains) [36, 37].
4.3 Discovery of c-di-GMP in Bacteria In 1982, Benziman and coworkers reported that production of cellulose by the membrane-associated cellulose synthase complex in Acetobacter xylinum (later renamed Gluconacetobacter xylinus) was stimulated by GTP and an additional soluble factor generically termed cellulose synthase activator [38–40]. This soluble factor, synthesized from GTP was tentatively identified as a cyclic oligonucleotide composed of GMP. It was degraded by a calcium-inhibitable activity that co-localized with washed membranes [41]. The activating cofactor was further characterized in a follow up study as a heat-resistant, alkali labile, alkaline phosphatase-resistant, snake venom phosphodiesterase-sensitive cyclic oligoguanylate [42]. Its structure was later determined by NMR as Bis- (3 -5 )-cyclic dimeric guanosine monophosphate (c-di-GMP) [43] (Fig. 4.1). The authors also proposed a model whereby c-di-GMP acts as an intracellular messenger regulating the biosynthesis of cellulose in vivo. The regulatory effect is accomplished by controlling the intracellular concentrations of c-di-GMP through the antagonistic activities of two classes of enzymes, diguanylate cyclases (DGCs) and c-di-GMP phosphodiesterases (PDEs). Two PDE activities were experimentally identified, one hydrolyzing c-di-GMP to linear pGpG and the other hydrolyzing pGpG to GMP. Within the cell, c-di-GMP was shown to be tightly associated with a c-diGMP binding protein, a membrane protein that exhibited saturable and reversible c-di-GMP binding with high affinity [44]. Benziman’s group later identified in G. xylinus several proteins with diguanylate cyclase and phosphodiesterase activities by a reverse genetics approach [45]. Conserved amino acids comprise a motif in DGC and PDE proteins and were designated as the GGDEF (Pfam PF09900) and EAL (pfam PF0053) domains, respectively, after the most conserved amino acids in each domain [46, 45]. Computational analysis suggested that GGDEF domain proteins were related to adenylate cyclases [42]. Biochemical confirmation for PDE activity of an EAL domain protein was first obtained for G. xylinus phosphodiesterase A1 [47]. Direct biochemical data obtained from studies of enzymatic activity of recombinant proteins and genetic evidence based on correlation of various in vivo phenotypes with intracellular levels of c-di-GMP has provided strong support for the role of the GGDEF and EAL domains in c-di-GMP metabolism [48–75].
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Fig. 4.1 c-di-GMP synthesis, activity and degradation. Diguanylate cyclases (DGC) carrying GGDEF domains catalyze the synthesis of c-di-GMP from GTP. Purified phosphodiesterases (PDEs) with EAL domain catalyze the hydrolysis of c-di-GMP to linear pGpG, although conversion of this linear dinucleotide to GMP can be demonstrated with crude enzyme preparations. PDEs containing the HD-GYP family domains hydrolyze c-di-GMP directly to GMP. c-di-GMP acts through receptors (proteins or riboswitches) and these control various cellular functions, often in a reciprocal fashion
In addition to the EAL domain, other proteins with a second domain have been shown to catalyze the hydrolysis of c-di-GMP. This so called HD or HD/HQ-GYP domain, named after conserved residues in the protein (Pfam PF01966), is found in a number of eukaryotic cyclic nucleotide phosphodiesterases as well as in several prokaryotic dGTP triphospho-hydrolases and in SpoT, the ppGpp hydrolase [76]. In the plant pathogen Xanthomonas campestris, the HD domain is the C-terminal output domain of response regulator RpfG, which is activated by its cognate histidine kinase RpfC and the RpfC/RpfG signal transduction pathway has been implicated in virulence [77, 78].
4.4 Distribution of Genes Encoding Enzymes for c-di-GMP Metabolism in Pseudomonas aeruginosa One of the most striking features of enzymes involved in c-di-GMP turnover is their abundance and redundancy in individual bacteria. Inspection of genome sequences in the Pfam database (http://pfam.sanger.ac.uk/) currently reveals more than 10,000 GGDEF proteins distributed in 648 species, and almost 6,000 EAL proteins distributed in 585 species. They are encoded in bacterial genomes, but they are also present in the genomes of two Archea and several Eukarya (sea
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anemone, Dictyostelium discoideum and rice). The majority of genomes contain coding sequences for multiple paralogues of these two proteins, ranging, from none in Helicobacter, Bacteroidetes, Chlamydiales and Fusobacteria to 96 in Vibrio vulnificus [9]. In most instances, the GGDEF and EAL catalytic domains are fused to various signaling domains and occasionally, to single or multiple transmembrane segments. These modular arrangements suggest that their enzymatic activities are regulated. Since the output of the signaling pathways is likely the same (synthesis or degradation of the diffusible molecule c-di-GMP), this raises a question of specificity of their signaling activities. The various models explaining this functional redundancy will be discussed later in this chapter (Section 4.6). A number of genes encoding proteins with GGDEF and EAL domains were identified in P. aeruginosa during screens for alterations in phenotypes related to biofilm formation or altered colony morphology. These studies also implicated c-di-GMP in antibiotic-resistant phenotypic variant conversion [79], twitching motility [80, 60], flagellar development and biofilm formation [81], and in the induction of a hyperbiofilm/type III secretion defective phenotype caused by a mutation in the sensor kinase RetS [82]. Individual proteins with GGDEF or EAL domains were also implicated in the formation of tobramycin-inducible biofilms [83], as positive regulators of pellicle formation, biofilm and pel/psl gene expression induction [57], as regulators of swarming and biofilm production [63, 69], alginate biosynthesis [56, 84], cup pili formation [81, 85] and type III secretion dependent cytotoxicity [86]. In 2006, Kulasekara and colleagues [86] carried out a comprehensive analysis of proteins with GGDEF and EAL domains, using transposon insertion mutants as well as over-expression clones [64]. Examination of the available annotated genomes of P. aeruginosa strains showed that this organism has the potential to express 17 different proteins with a GGDEF domain, 11 proteins with an EAL domain and 17 proteins that contain both of these domains (Table 4.1). A schematic representation of the domains within these proteins is shown in Fig. 4.2. Some of the genes for Table 4.1 Distribution of the proteins with GGDEF, EAL and HD domains encoded in the genomes of sequenced P. aeruginosa strains. +, presence; or –, absence PAO1
PA2192
C3719
PA7
PA14
LESB58
CS2
GGDEF PA0169 PA0290 PA0338 PA0847 PA1107 PA1120 PA1851 PA2771 PA2870 PA3177 PA3343 PA3702 (WspR)
+ + + + + + + + + + + +
+ + + + + + + + + + + +
+ + + + – + + + + + + +
+ + + + + + + – + + + +
+ + + + + + + + + + +
+ + + + + + + + + + + +
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Table 4.1 (continued) PAO1
PA2192
C3719
PA7
PA14
PA4332 (SadC) PA4396 PA4843 PA4929 PA5487
+
+
+
+
+ + – +
+ + – –
+ + + +
+ + + +
+ + – +
+ + + +
+ + + – + +
+ + + + + +
+ + + + + +
+ + + + + +
+ + + + + +
+ + + + + +
+ + + + +
+ + – + +
+ + + + –
+ + + + +
+ + + + +
+ + + + +
+
+
+
+
+
+
+
+
+
+
+
+
+ + + –
+ + + –
+ + + PSPA7_4938
+ + + –
+ + + +
+ + + –
EAL PA0707(ToxR) PA2133 PA2200 PA2818(Arr) PA3825 PA3947(RocR) – – – – –
+ + + – + + – – – – –
+ – + – + + – – PACG_02185 – –
+ – + + + + + + – PSPA7_1020 PSPA7_2617
+ + + – + + PvrR PA14_15435 – – –
+ + + – + + – – – – –
+ + + – + + – – – – –
HD PA2572 PA4781 PA4383
+ + +
+ + +
+ + +
+ + +
+ + +
+ + +
GGDEF/EAL PA0285 PA0575 PA0861 PA1181 PA1433 PA1727 (MucR) PA2072 PA2567 PA3258 PA3311 PA4367 (BifA) PA4601 (MorA) PA4959 (FimX) PA5017 PA5295 PA5442 –
LESB58
CS2 +
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Fig. 4.2 Domain organization of putative DGCS and PDEs and their distribution in the sequenced P. aeruginosa genomes. Shown is the domain organization of P. aeruginosa proteins with GGDEF and EAL domains and the three studied HD-GYP proteins, based oan an analysis of the Pfam database.a , putative allosteric inhibitory I-site in GGDEF domain lacks conserved residueshas mutations making it very likely non-functional;b , conserved GGDEF residues are mutated substituted and the domain likely lacks diguanylate cyclase
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EAL and/or GGDEF proteins are not found in all strains, suggesting that they can be lost during the evolution of a particular strain or they are located on mobile genetic elements. In particular, genes encoding for EAL domain proteins appear less conserved across the available sequenced genomes of P. aeruginosa. A more detailed discussion of several individual GGDEF and EAL domain-containing proteins discovered in specific genetic screens will be given later in this chapter (Section 4.7). In proteins with modular organization, the additional domains are located always N-terminal to the GGDEF/EAL domains and most of them have been previously shown to play a role in sensing of environmental signals or participate in signal transduction [87]. These include domains that contain the PAS and PAC motifs (currently regrouped and renamed as the PAS fold domain) implicated in heme binding, and the 7TMR-DISMED2/7TMR-DISM_7TM (7 transmembrane receptor with diverse intracellular signaling module) domain that is found in association with signaling proteins that recognize carbohydrates and probably function as sensors of extracellular ligands [88]. Other domains that are associated with GGDEF/EAL domain containing proteins include the CHASE (Cyclases, Histidine kinase Associated Sensory Extracellular) domain and the response regulator receiver domains of two-component transcription and chemotactic factors that are probably involved in regulating the enzymatic activities of the adjacent DGC or PDE domains [89]. Additional domains such as the HAMP (Histidine kinases, Adenylyl cyclases, Methyl binding proteins, Phosphatases) domain found as a linker between extracellular sensory domains and intracellular signaling domains including two-component sensor kinases [90] are also associated with proteins that contain GGDEF/EAL domains. Finally, some of these proteins contain the MHYT domain (named after its conserved amino acid motif), predicted to bind oxygen, carbon monoxide or nitric oxide. MHYT domains are often found at the amino-terminus of proteins that are part of signal transduction pathways, including those involved in c-di-GMP metabolism [91]. The sequenced genomes of P. aeruginosa encode 11 proteins with the HD-GYP domain and three have been studied in greater detail (Ryan et al., Environmental Micro, in press). Two of these (PA4108 and PA4781) contain a highly conserved HD-GYP domain, and one protein (PA2572) has a variant of the consensus sequence (HQ-GYP). PA4108 and PA4781 were shown to have PDE activity while PA2572 appears to be enzymatically inactive. In PA4178 and PA2572, the HD-GYP domains are adjacent to a response regulator receiver domain and therefore the enzymatic activities of these two proteins may be regulated by phosphorylation. P. aeruginosa mutants for genes PA4178, PA2572, or PA4108 were strongly attenuated in virulence in the Galleria mellonella model, with a lethal dose of 50–100 fold higher than the wild-type. However, specific phenotypes related to the production of virulence factors varied. For example, a mutation in PA4108 (but not PA2572 and PA4781) abolished the production of the cytotoxin exoenzyme S, while loss of the two functional PDEs, PA4108 and PA4781 eliminated the ability of P. aeruginosa to swarm, without affecting swimming motility or rhamnolipid production. The phenotypes associated with the enzymatically-inactive PA2572 are particularly
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intriguing, suggesting that this regulatory protein has evolved from an ancestral PDE and this evolutionary feature may be found in other enzymes, including those with altered EAL and GGDEF domains.
4.5 Structure-Function Relationships of c-di-GMP Diguanylate Cyclases and Phosphodiesterases Structural analysis performed on GGDEF domain proteins showed that they are composed of a five-stranded central β-sheet surrounded by helices with a topology identical to that of the catalytic core of adenylate cyclase and the “palm” domain of DNA polymerases [50]. The DGC domain is also functionally related to adenylate cyclases and DNA polymerases; they are all enzymes that catalyze the formation of a 3 -5 phosphodiester bond [7, 92]. The mode of substrate binding at the active site and mechanism of catalysis have been inferred based on structural studies of the Caulobacter crescentus DGC PleD [93, 94] and residues important for catalysis have been identified by site-directed mutagenesis of various DGCs [67]. The mode of binding of substrate resembles that of adenylate cyclase. [94]. Dimerization appears to be important for product synthesis, with each monomer binding one molecule of GTP [50, 94]. In several putative DGC proteins the GG(D/E)EF motif is not well conserved [49] and where examined, no DGC was observed. However, for one protein with a degenerate GGDEF domain, GTP binding was demonstrated, and therefore it could play a role in regulating the activity of the adjacent EAL domain [52]. Models for two EAL domains are present in the Protein Data Bank (http://www.rcsb.org/) archive (Bacillus subtilis 1uze and Thiobacillus denitrificans 2r6o). Based on these structural data, a homology model of P. aeruginosa RocR has been generated [71]. Computational docking and site-directed mutagenesis followed by biochemical analysis of the PDE RocR suggested a catalytic mechanism for c-di-GMP hydrolysis by EAL proteins [71]. A large number of acidic residues (glutamate and aspartate) are present around the catalytic site, where a single Mg2+ is coordinated. In this model, the Mg2+ would play substrate activation and deprotonation of the nucleophilic hydroxide ion [71]. A conserved glutamate not included in the signature EAL motif is essential for catalysis (E352 in RocR) and appears to act as the general base catalyst for the nucleophilic attack of the c-di-GMP phosphodiester bond [71]. More recently a detailed analysis of Bacillus subtilis YkuI (a protein comprised of a EAL and a PAS domain) in complex with c-di-GMP-Ca2+ has been published [95]. The study identifies the EAL motif as part of the active site, with E33 being involved in cation coordination. The structure of the catalytic site suggested a mechanism for the nucleophilic attack of the phosphodiester bound also involving a non-motif residue, E209, as the general base. The authors also propose specific EAL-EAL and EAL-PAS dimer interactions as important for regulation of EAL activity [95].
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4.6 Models for c-di-GMP Signaling: Specificity of c-di-GMP Activity and the Redundancy of Phosphodiesterases and Diguanylate Cyclases One of the interesting paradoxes of signal transduction involving the secondary messenger c-diGMP is the redundancy of the enzymes responsible for its synthesis and degradation. In several occasions, distinct phenotypes have been observed in mutant bacteria carrying deletions of genes encoding individual DGCs and PDEs. One plausible hypothesis explaining the specificity of c-di-GMP generated or hydrolyzed by any single enzyme is based on a model that c-di-GMP functions within local complexes formed between the biosynthetic or degradative enzymes and cognate “adapter”, a c-di-GMP receptor. The adapter-c-di-GMP complex interacts with specific effectors directing the specific response. Individual PDEs function in an analogous fashion, interacting with specific c-di-GMP-adapter complexes, locally hydrolyzing c-di-GMP causing a reversal of the c-di-GMP regulatory activity. The essential features of this model are shown in Fig. 4.3. Based on the presence of signal transduction modules in DGCs and PDEs, their enzymatic activities are very likely controlled by conformational changes that follow the perception of environmental signals by individual regulatory domains. In the model depicted in Fig. 4.3, the activated DGC binds to its cognate partner, an adapter, which is a c-di-GMP binding protein controlling a specific cellular function. Each enzyme may function in concert with a single adapter, or they may recognize several adapters with different effector targets, allowing signal amplification.
Fig. 4.3 Lock and key model of c-di-GMP signalling
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The consequence of activation and binding of the GGDEF protein to the adapter is the localized synthesis of c-di-GMP, which is preferentially bound by the cognate adapter, before the signaling molecule can diffuse throughout the cell. The bound c-di-GMP facilitates the interaction of the adapter with components of various cellular processes (effectors) controlled by c-di-GMP. At this point, the DGC can dissociate from the adaptor, while c-di-GMP remains bound. The process is reversed when a PDE receives a different signal: it binds to a specific adapter and hydrolyzes the bound c-di-GMP, resulting in the de-activation of the effector functions of the pathway. In some cases, the same protein can contain an EAL and a GGDEF domain. This arrangement is quite common and is found in 17 P. aeruginosa proteins (Fig. 4.2). The localized synthesis and acquisition of c-diGMP by the binding protein depends on correct stoichiometry between enzymes and adapters, and unregulated synthesis or degradation of c-di-GMP by over-expression of diguanylate cyclases or phosphodiesterases results in activation of multiple adapters and effectors. Finally, it is conceivable that some of the enzymaticallyinactive GGDEF or EAL domain-containing proteins which lack catalytic residues can bind to adapters. These enzymatically-inactive proteins exert their regulatory effects through protein-protein interactions alone and not by altering directly local c-di-GMP levels. In addition to protein receptors that function as adapters, c-di-GMP can control gene expression by interacting with cis-acting regulatory RNAs. In a number of different bacterial species, c-di-GMP levels are sensed by folded segments of RNA found at 5 ends of certain transcripts, referred to as riboswitches [96]. Binding of c-di-GMP toriboswitches regulates gene expression by influencing transcriptional termination or translation initiation. The consequence of c-di-GMP binding to a riboswitch may be induction of expression of linked open reading frames, as has been shown for these RNA elements in front of the tfoX gene of Vibrio cholerae, or repression of genes, presumably by promoting transcript degradation, as exemplified by the c-di-GMP sensing riboswitches in the 5 untranslated regions of several Bacillus cereus and Clostridium difficile or V. cholerae gbpA genes [96]. The regulatory specificity of c-di-GMP action could be achieved by recognition of an RNA sequence by the DGC and PDE, suggesting that some of these enzymes may possess sequence or structure-specific RNA binding domains. A third level of regulation of specificity may involve temporal and environmental control of GGDEF and EAL domain protein expression at a transcriptional and post-transcriptional level. [97] In Salmonella enterica, different GGDEF domain protein (AdrA or STM1987) control cellulose biosynthesis under different growth conditions (LB with no salt or C-rich medium poor in trace elements) [53, 98]. In Yersinia pestis, temperature levels control the expression of the GGDEF domain protein HmsT [97]. In Thermotoga maritima cell density-dependent expression of GGDEF and EAL domain proteins and consequente modulation of exopolysaccharide production was observed by microarray analysis [99]. Perry et al. demonstrated that at least five of the 29 GGDEF/EAL genes are growth phase-induced under the control of the stationary phase sigma factor (σS ) and they exhibit differential
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control by additional environmental and temporal signals. In P. aeruginosa, microarray analysis has shown that most PDE and DGC are expressed at all times and in many different media [64].
4.7 Diguanylate Cyclases and Phosphodiesterases of P. aeruginosa In the following section we will examine the literature describing the major examples of diguanylate cyclases and phosphodiesterases. A summary view of the various c-di-GMP pathways in Pseudomonas aeruginosa is shown in Fig. 4.4.
Fig. 4.4 Synoptic model of c-di-GMP signaling in Pseudomonas aeruginosa. The model summarizes the interactions between cellular diguanylate cyclases, phosphodiesterases, diguanylate receptors, and various effectors modulating the subset of phenotypes indicated outside the cell outline. Black arrows indicate functional activating (arrow head)/repressing (thick bar) relationships inferred from mutant analysis, blue arrows indicate functional relationships demonstrated by ectopic expression of the DGC/PDE; red arrows indicate transcriptional effects. GGDEF DGC, diguanylate cyclases carrying a GGDEF domain; EAL PDE, phosphodiesterases carrying an EAL domain; HD/YN-GYP PDE, phoshodiesterase carrying a HD/HY-GYP domain; DGR, diguanylate receptor, collectively indicating various classes of adaptor proteins, including PilZ (Alg44), PelD, FleQ, for which direct binding of c-di-GMP was shown; SCV, small colony variant. For ToxR PDE activity has not yet to been demonstrated. Black arrows connecting squares show possible connections between enzymes and phenotypes based on mutant analysis. See text for detailed discussion of the relationships between the various DGCs, PDEs, c-di-GMP receptors and effectors
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4.7.1 The WspR and the Chemotransducing System Controlling Surface Properties of P. aeruginosa Genomes of all Pseudomonas species contain a cluster of genes encoding proteins that share significant similarities with chemotaxis (Che) proteins responsible for sensing and directing microorganisms towards favorable sources of nutrients. This system includes a protein with a GGDEF domain, thus implicating c-di-GMP in a chemosensory response. In Pseudomonas fluorescens and P. aeruginosa, certain transposon mutations in genes of this cluster (the wspA, wspB, wspC, wspD, wspE, wspF and wspR genes) result in the loss of swimming and swarming motility, cause an autoaggregative phenotype in liquid, and while growing on solid agar surfaces, colonies lack the characteristic spreading edge. This colony appearance is referred to as the Wrinkly spreader phenotype (Wsp) [100, 101]. The chemotactic response in bacteria is initiated by binding of the attractant to transmembrane chemoreceptors, the methyl-accepting chemotaxis proteins (MCP proteins). The soluble cytoplasmic adapter protein CheW and the CheA kinase bind to transmembrane receptor proteins, resulting in autophosphorylation of the CheA sensor kinase at its histidine residue. The CheA∼P then transfers its phosphate to the aspartate residue on the receiver domain of the response regulator CheY. Phosphorylated CheY interacts with the flagellar motility apparatus altering its rotation and consequently, the swimming behavior of the bacteria and their chemotactic response in the gradient of a specific attractant. In addition to CheY, the phosphorylated CheA can transfer a phosphate to CheB, activating its methylesterase activity, which, together with methyltransferase CheR, controls the methylation state of the MCPs. The reversal of methylation by CheB is an important step in restoring the ability of the chemoreceptor to become responsive to new stimuli. Although Pseudomonas species carry a set of genes specifying a complete chemotactic sensory system linked to flagellar motility, experimental evidence strongly suggests that the Wsp proteins participate in a specialized form of chemosensory signal transduction incorporating many mechanistic features of the Che signal transduction pathway. The Wsp system consists of a predicted MCP protein (WspA), two paralogues of CheW (WspB and WspD) and a hybrid histidine sensor kinase (WspE). Additional components of the chemosensory system include homologues of the methylesterase CheB and methyltransferase CheR, the Pseudomonas WspF and WspC proteins, respectively. The major difference between the two systems is in the response regulators. WspR resembles CheY in the presence of the phosphoacceptor receiver domain however the output is not alteration of flagellar rotation, but most likely, the synthesis of c-di-GMP by the adjacent GGDEF domain (Fig. 4.8). Studies of phenotypes of mutants for individual wsp genes strongly suggested that their products function in signal transduction by mechanisms that are analogous to the activities of the Che proteins. In P. aeruginosa, a mutation in wspF (the gene for the putative WspA-specific methyl esterase) but not in wspA, wspE, or wspR, resulted in enhanced biofilm formation and the wrinkly colony appearance
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[57] and loss of swarming motility [102]. Moreover, mutations in the wspA or wspR genes led to the reversion of the wrinkly colony appearance induced by the wspF mutation [103]. These phenotypes are consistent with the predicted role of WspF in maintaining WspA in a locked inactive form and, consequently, limiting phosphorylation of WspR until a proper chemical stimulus activates the MCP protein and signaling. Biochemical studies have provided additional support for a model whereby the primary function of the signal transduction pathway mediated by the Wsp proteins is to control the DGC activity of WspR. The concentration of the cellular pool of c-di-GMP in the P. aeruginosa wspF mutant is elevated [57]. Purified recombinant WspR protein is enzymatically active and converts GTP to c-di-GMP. The input domain of WspR is very likely phoshorylated, and this leads to enhanced catalytic activity of the adjacent domain [57]. A mutant protein with a substitution of a glutamate for the phospho-acceptor aspartate, known to mimic the phosphorylated state of the aspartate in the receiver domains of a number of two-component response regulators, resulted in an enzyme with a hundred-fold more DGC activity than the wild-type WspR, while substitution of the phosphoacceptor aspartate to asparagine eliminates enzymatic activity (Matewish and Lory, unpublished). Two studies addressed the putative targets of the Wsp regulatory pathway. Insertion mutants that suppressed the phenotypes of the wspF mutant included not only all of the other genes in the wsp cluster, but also PA2130 (cupA3) encoding a component of the CupA fimbriae, and PA2128, the homologue of a Salmonella gene encoding aggregative fimbriae. These surface proteins may represent some of the biofilm-promoting factors controlled by the Wsp pathway [100]. A subsequent microarray analysis of the wspF transcriptome demonstrated that the product of this gene, possibly due to the increase in c-di-GMP levels, is responsible for an increased steady-state level of transcripts encoding the component of the Pel and Psl polysaccharide biosynthetic machinery [57]. These two polysaccharides have been implicated in biofilm formation and altered colony morphology. However, transcriptional profiling of the wspF mutant did not show altered cupA3 and PA2128 transcript levels, suggesting that the expression of these two proteins may not be regulated by WspF. A recent study showed that pel, but not psl, gene products are required for the various autoaggregative phenotypes observed in the wspF mutant [102]. The hypothesis that the production and utilization of c-di-GMP by the Wsp system is compartmentalized was examined by Guvener and Harwood [103]. A hybrid protein consisting of WspA and yellow fluorescent protein (WspA–YFP) was fully functional, and when its subcellular location was examined, the protein was found in patches located on the periphery of the cell. A fusion of YFP to WspR was also functional, but it failed to form clusters and appeared dispersed in the cytoplasm of wild-type cells grown in liquid media. However, in P. aeruginosa with a wspF mutation, which leads to constitutive phosphorylation of WspR, the fusion protein formed clusters, suggesting that phosphorylation of the WspR receiver domain results in the clustering of WspR. The substitution of an asparagine for the phospho-accepting aspartate residue prevented clustering of WspR-YFP in
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the wspF mutant, confirming that the association of multiple WspR molecules was dependent on phosphorylation. Clustering of the WspR–YFP was dependent on growth conditions, and similar clusters of the hybrid protein were observed in wildtype bacteria grown on agar surfaces, suggesting that activation of the Wsp pathway, and consequently, enhanced phosphorylation of WspR, occurs on solid medium, or generated by surface-grown bacteria. Clustering of WspR did not require the presence of a functional GGDEF domain, as a WspR–YFP variant with an inactivating amino acid substitution in this domain still formed clusters in the wspF mutant. These studies, showing discrete localization of Wsp proteins, including the WspR DGC implies a localized action of c-di-GMP, in a specific location within the cell. Identification and localization of the c-di-GMP receptors would provide a final proof for the model. There are several likely candidates, including the PelD component of the PEL polysaccharide biosynthetic machinery of P. aeruginosa [104]. The P. aeruginosa WspR is only the second DGC protein that has been studied at the structural level [105], the other being the C. crescentus PleD [50]. WspR is a multi-domain DGC consisting of an N-terminal phosphoreceiver domain and a C-terminal GGDEF domain separated by a long helical stalk (Fig. 4.5). The purified WspR with its bound inhibitor c-di-GMP crystallizes as a tetramer [105]. In this structure (Protein Data Base code 3BRE), the GGDEF domains are oriented such
Fig. 4.5 Structure of the WspR with its allosteric I-site. Left. The structural model of the WspR dimer was derived from the atomic coordinates deposited in the PDB database (code 3BRE) and rendered with PyMol. One monomer is shown as a helix and strand cartoon, the other monomer (in red) is shown with a surface representation. The catalytic A-site is indicated without showing the conserved residues. The C2-symmetry related “head-to-head” dimer providing the residue R198’ to the allosteric I-site is not shown. A detailed rendering of the allosteric I-site, with the two residues of the conserved RxxD motif is shown on the right
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that the catalytic C-sites face each other in the anti-parallel dimer. This arrangement allows each subunit to bind one molecule of GTP followed by cyclization, possibly going through a short-lived linear dinucleotide. The enzymatic activity of WspR is subject to feedback inhibition by c-di-GMP. Based on the cumulative genetic and biochemical evidence, feedback inhibition occurs when c-di-GMP binds to the regulatory I-site, containing a R242 xxD245 motif and located adjacent to the catalytic site (Fig. 4.5), leading to a rearrangement of the two globular domains around the stalk. (See also the discussion of the I-site in Section 4.8.) In the inhibited tetramer complex the catalytic sites are obstructed by the stalk of the anti-parallel monomer, presumably blocking the cyclase activity. Furthermore, the two dimers are interlocked head-to-head, with four molecules of c-di-GMP (one intercalated dimer per GGDEF domain pair) bound to the cyclase domain (Fig. 4.5). Size-exclusion chromatography and multi-angle light-scattering analysis suggested that, in solution, WspR can exist in one of three stable forms: a tetramer, a globular dimer, and an elongated dimer [105]. Based on the results of several structural and functional studies, and interpretations of enzymatic and physical properties of engineered mutant proteins, the authors propose that while in the form of globular active dimers the proteins assume a parallel configuration (Fig. 4.5), and that in the elongated inactive dimer the WspR molecules are likely anti-parallel (Fig. 4.6). This is reminiscent of the two dimeric crystal structures isolated for the mouse PDE2A [105].
Fig. 4.6 WspR oligomerization cycle of activation and inhibition. Model of WspR activation/inhibition by oligomerization as proposed by [105]
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One of the unanswered questions from the models based on the structural work is the role of the N-terminal phosphoreceiver domain. Dimerization of WspR appears to be independent of the presence of the conserved phospho-acceptor aspartate residue in the receiver domain. De and colleagues [105] stated that the enzymatic activity of the protein is not affected by unspecified mutations of the phosphoacceptor domain. Moreover, in their hands, treatment of WspR with artificial phosphate donor acetyl phosphate or creating a phosphate mimic by a reaction of aspartate with berillium fluoride does not enhance its enzymatic activity and the model does not account for any conformational changes that may occur following phosphorylation of the aspartate in the receiver domain and resulting in enhanced enzymatic activity. This differs from published experimental evidence showing a stimulatory effect of treatment of WspR with acetyl phosphate [57]. Moreover, a mutation that changes the phosphoacceptor aspartate to alanine inactivates the enzyme and the substitution of glutamate for the same asparte leads to increased activity of WspR in vitro and in vivo. The WspR mutant with this aspartate to glutamate substitution in the receiver domain is a stable tetramer and it appears to be the enzymatically most active form of the protein (M. Matewish, unpublished). Clearly, the role of the phosphoreceiver domain in controlling the activity of WspR needs to be further studied.
4.7.2 SadC and BifA: Other DGCs and PDEs Involved in Biofilm Formation The most conserved function of c-di-GMP in P. aeruginosa is the participation of this regulatory cyclic dinucleotide in pathways that favor the biofilm lifestyle of the bacteria. Under certain conditions, mutations in numerous DGCs reduce biofilm formation; overproduction of the same enzymes lead to a so-called “hyperbiofilm” phenotype in various in vitro assays. Although biofilm formation has been primarily assayed in a variety of artificial laboratory systems, the consistency of these phenotypes suggests that c-di-GMP controls the same pathways in natural environments as well. Biofilm formation is a multistep process, beginning with reversible attachment of motile planctonic bacteria to solid surfaces, which over time becomes permanent and irreversible. Microcolonies are formed by proliferation of the adherent organisms and attachment of additional bacteria from the media. This is followed by the development of a mature biofilm (a macrocolony). Screening for phenotypes associated with specific stages of biofilm formation, O’Toole and co-workers identified DGCs and PDEs that coordinate the production of a number of biofilm-promoting factors. These included SadC, (PA4332) a membrane-associated protein with a DGC activity [69] (Fig. 4.2). This protein plays a role in biofilm formation during the transition from a transient to a permanent attachment state (Sad, surface adhesion defective). In particular a sadC mutant is characterized by a hyper-swarming phenotype and by a defect in biofilm formation [69] (Fig. 4.4). Consistent with this observation, ectopic expression of sadC leads to an increase in biofilm formation,
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which is dependent on the synthesis of the Pel polysaccharide. However, unlike overeproduction of WspR, which leads to increased transcription of the genes of the pel operon [106], overproduction of SadC appears to enhance pel expression at a post-transcriptional level [69]. Regulation of production of the Pel polysaccharide by SadC may be the result of a specific interaction of this DGC with its cognate c-di-GMP receptor. PelD, a c-di-GMP-binding protein involved in Pel polysaccharide biosynthesis, has been identified and this protein may serve as the specific adapter (see Fig. 4.3) capable of distinguishing between c-di-GMP produced by SadC and other DGCs expressed by P. aeruginosa [104]. Another P. aeruginosa cytoplasmic membrane protein, BifA (PA4367; biofilm formation), also controls c-di-GMP levels and biofilm formation, but with effects opposite to those regulated by SadC [63]. BifA contains adjacent DGC and PDE domains (Fig. 4.2), although only a PDE activity can be detected in vivo and in vitro. Deletion of the entire bifA gene, or substitutions in the conserved residues within the EAL and GGDEF domains, lead to enhanced biofilm formation and reduced swarming [63]. Analyses of enzymatic activities of BifA and phenotypes of bifA mutants strongly argue for a sole PDE activity, although the GGDEF domain is required for the biological function of this protein. BifA is an example of a family of proteins with GGDEF domains (the GGDQF sequence is present in BifA) that have putatively evolved from ancestral, enzymatically-active DGCs. Deletion of bifA results in an increase in the cellular pool of c-di-GMP and the production of the Pel polysaccharide with concomitant abolishment of swarming motility. During the development of a mature biofilm, it is necessary for bacteria to become less motile in order for the microorganisms to become irreversibly attached to a substratum, and for subsequent building of a three-dimensional biofilm structure. This stage of bioflm formation is very likely regulated by BifA, which appears to control the frequency of flagellar reversal. The receptor c-di-GMP responsible for controlling the on/off swarming state of the bacteria is unknown. However, genetic analysis points to a pathway, where c-di-GMP levels, controlled by BifA or SadC, affect, directly or indirectly, the activity of the SadB protein. This in turn may transmit the information to downstream effectors such as the Pel biosynthetic machinery and the flagellar/chemotaxis apparatus [63].
4.7.3 MucR and Alginate Regulation MucR (PA1727) is a hybrid GGDEF/EAL domain-containing protein (Fig. 4.2) that has been recently implicated in the production of the alginate polysaccharide by mucoid P. aeruginosa. The mucR gene is not linked to any other alginate biosynthetic genes but its deletion in the mucoid P. aeruginosa strain PDO300 causes a ∼40-fold reduction in alginate production, while mucR overexpression leads to a 7-fold increase [56]. MucR is a cytoplasmic membrane protein carrying in its carboxy-terminal region the GGDEF and EAL domains, while the N-terminal portion contains three tandem domains with MHYT motifs and several transmembrane
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Fig. 4.7 Model of RocR regulation of cup genes and virulence phenotypes
segments [56] (Fig. 4.7). MHYT domains are found on proteins involved in sensing oxygen and nitric oxide. Since alginate production is stimulated by anaerobic conditions and requires c-di-GMP, MucR was hypothesized to function as a sensor of low oxygen tension or the presence of other gases such as nitric oxide in the lungs of cystic fibrosis patients where it produces c-di-GMP directed towards alginate production. In strain PA14, MucR was previously shown to be an active DGC and its overproduction resulted in a hyper-biofilm phenotype [64], wrinkly colony appearance on solid medium, pellicle formation at the air-liquid interface, increased autoaggregation, formation of a structured biofilm in flow cells, and inhibition of swarming motility [56]. The motility defect was very likely caused by interference of elevated c-di-GMP levels with the synthesis of flagellin subunits [56]. MucR is hypothesized to regulate alginate production (Fig. 4.4) under anaerobic conditions by interacting with the PilZ-receptor Alg44 [84], a component of the alginate biosynthetic apparatus, and catalyzing localized synthesis of c-di-GMP (Fig. 4.8A).
Fig. 4.8 Model of diguanylate receptors and their relation with the cognate diguanylate cyclases and effectors. (a). Regulation of alginate polymerization and/or secretion by the DGC MucR and the c-di-GMP receptor Alg44. The PilZ protein Alg44 was shown to localize in the inner membrane where it regulates alginate production upon binding of c-di-GMP. MucR is a membrane-bound DGC that stimulates alginate production via Alg44. (b). Regulation of Pel polysaccharide synthesis by the DGC WspR and the the c-di-GMP receptor PelD. WspR also regulates the transcription of the pel genes
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4.7.4 MorA, PA1120 and the Regulation of the CupA Fimbrial System MorA (PA4601) is a hybrid GGDEF/EAL domain-containing cytoplasmic membrane-localized protein [64] carrying three PAS sensory domains at its aminoterminus (Fig. 4.2). PAS domains are conserved sensory signal transduction modules that have been implicated in sensing redox levels, energy charge, light, and oxygen via specific interactions with various ligands (heme, FAD, NAD, ATP etc.) [107] [2]. MorA (Motility regulator) was initially identified in Pseudomonas putida as a membrane-localized negative regulator of flagellar development. When compared to wild-type P. putida, deletion of the morA gene resulted in a hyperflagellated phenotype, enhanced motility and chemotaxis, and decreased biofilm formation. The motility phenotype of the P. putida morA mutant was attributed to over-expression of flagellin during early phases of growth, leading to an increase in the total number of flagella [81]. Mutants of P. aeruginosa PAO1 lacking morA also display a biofilm defect, but were not defective in flagellin production or motility. A screen of a transposon library of the P. aeruginosa strain 20265, a variant displaying an SCV (Small Colony Variant) phenotype for revertants of this colony morphology, identified morA and PA1120 [85]. The SCVs are frequently isolated from patients with cystic fibrosis and they show increased antibiotic resistance, have a propensity to form robust biofilms, aggregate in liquid medium, and are defective in both twitching and swimming motility. PA1120 is a cytoplasmic membranelocalized GGDEF domain-containing protein with an additional HAMP domain (named for their presence in histidine kinases, adenylyl cyclases, methyl-accepting chemotaxis proteins, and phosphatases) found in numerous bacterial signal transduction proteins [85]. Mutation in morA or PA1120 genes in an SCV strain result in the loss of autoaggregation, a large colony morphology, a decrease in the synthesis of CupA fimbriae, and a slight increase in swimming and twitching motility [85]. Higher levels of c-di-GMP were also detected in this SCV strain compared to the isogenic strain with normal colony morphology. A mutation in PA1120 also reduced cellular levels of c-di-GMP compared to the SCV parent [85]. Surprisingly, overexpression of PA1120 did not restore the small colony phenotype on solid media, and could only complement the loss of cupA expression, suggesting that additional layers of complexity determine the SCV phenotype. In a separate study, neither DGC nor PDE activity could be detected in extracts of P. aeruginosa expressing high levels of morA, while PA1120 was shown to be an active DGC in vivo [64]. It appears that many, but not all, traits associated with naturally-occurring SCVs can be attributed to c-di-GMP regulation of CupA synthesis. Since changes in c-diGMP levels do not completely restore swimming or twitching motility of SCVs, it is apparent that in addition to CupA, other factors play a role in the morphological changes associated with these colony variants.
4.7.5 PvrR In P. aeruginosa another protein involved in c-di-GMP metabolism has been implicated in the formation of compact colonies. PvrR (Phenotype variant regulator;
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Fig. 4.2) was identified in a screen of a cosmid library for genes that would ectopically suppress the rough small colony variant (RSCV) phenotype of P. aeruginosa strain PA14. The RSCV phenotype includes small size and rough colony appearance on agar plates, an increase in autoaggregation and attachment to solid surfaces when grown in liquid, increased surface hydrophobicity, altered biofilm morphology, and production of antibiotic-resistant biofilms. Multicopy expression of PvrR in SCVs of P. aeruginosa strain 20265 can suppress CupA expression and reduce c-di-GMP levels [85], suggesting that the SCV and RSCV phenotypes possibly represent the same physiological phenomenon. PvrR is an active PDE protein composed of an N-terminal phosphoreceiver module and a C-terminal EAL domain [64] (Fig. 4.2). The gene for PvrR is located on a mobile genetic element, the pathogenicity island 1 (PAPI-1) [108], and among sequenced P. aeruginosa genomes it is found only in PA14 and PA7. The RSCV phenotype appears in P. aeruginosa grown in the presence of antibiotics with a frequency as high as one in ten depending on the environmental conditions. RSCVs are also commonly observed in clinical CF isolates from individuals undergoing antibiotic therapy for chronic infections. Overexpression of pvrR in both RSCV and wild-type cells results in repression of attachment to inert surfaces and a 6-fold reduction in the frequency of RSCVs. However, deletion of pvrR caused a modest increase in resistance to kanamycin, but the pvrR mutants did not produce the RSCV phenotype. PvrR may act upstream of the colony morphology switching mechanism [79]. In the PAPI-1 mobile genetic element, the pvrR gene is flanked by rscC and PA14_59780, two genes encoding sensor histidine kinases. It is conceivable that the PDE activity of this enzyme is regulated by the phosphorylation (and dephosphorylation) of its receiver domain catalyzed by the products of the adjacent genes, whose activities may be environmentally regulated.
4.7.6 Arr and Regulation of Antibiotic Induced Bifilm Formation Arr (Aminoglycoside response regulator) (PA2818) is a PDE with a simple modular arrangement, containing at its carboxy-terminus the EAL domain and two transmembrane segments at the N-terminal half of the protein, possibly exposing a periplasmic loop (Fig. 4.2). Arr was discovered during screening of an ordered collection of mutants [109], with transposon insertions in genes with predicted GGDEF and EAL domains, for defects in tobramycin-induced biofilm formation [83]. The arr gene is located on a unique mobile genetic element. It is found in the genome of strains PAO1 and PA7 and is absent from the genomes of the other sequenced strains. Mutation in the conserved residues of the EAL domain (E297A substitution) caused a reduction in cellular c-di-GMP levels and abolished the function of Arr in tobramycin-induced biofilm formation. More recently, Arr was shown to be involved in the formation of biofilms induced by hydroxyurea and nalidixic acid, chemical agents that inhibit DNA replication [110]. The effect of the Arr PDE on biofilm formation is an important exception to the general paradigm that correlates biofilm formation with high levels of intracellular c-di-GMP.
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4.7.7 FimX and the Function of Type IV Pili FimX (PA4959) is a modular protein with a GGDEF and an EAL domain (Fig. 4.2). It also contains an amino-terminal phosphoreceiver domain followed by a PAS domain [80, 60, 64]. P. aeruginosa with a mutation in the fimX gene shows reduced twitching motility although this defect is not as strong as mutation in the pilA gene, encoding the major subunit of the pilus organelle. Among the environmental conditions that promote twitching in wild-type P. aeruginosa, the fimX mutants no longer respond to tryptone and mucin, suggesting that FimX plays a role is sensing or transducing environmental signals [80]. Deletion of the fimX gene resulted in a reduction in the number of type IV pili on the bacterial surface, although the expression of the pilin subunits was unaffected. In P. aeruginosa, type IV pili are found at the single pole of the rod-shaped cell and FimX appears to play a role in this polar localization process. In a P. aeruginosa mutant where the FimX protein lacks the phospho-receiver domain, the assembly of pili on the cell surface is only marginally affected but the pili are no longer restricted to the pole of the cell and are found over the entire surface of the bacterium. Studies with fluorescently-tagged proteins have shown that FimX itself is localized to a single pole of the cell, a process that requires the phospho-receiver domain of the protein [60]. Restriction of tagged FimX to one pole requires both the GGDEF and EAL domains. FimX has a detectable PDE activity in vitro, but the EAL domain (and the GGDEF domain) lacks many of the residues conserved in other EAL domains. However, both of these domains are essential for FimX function in assembly and polar localization of pili. FimX with substitutions in the critical residues in the EAL and GGDEF domains displays an unusual localization pattern, appearing at both poles of the cell [60]. Therefore, even in the absence of a conserved GGDEF domain and lack of DGC activity in vitro, this domain has a biological function, analogous to the EAL and GGDEF domains of BifA [63].
4.7.8 RocR/SadR and Transcriptional Regulation of Expression of CupB, CupC and the Type III Secretion System Phenotypic screens for mutants affected in the control of expression of biofilmpromoting factors in P. aeruginosa identified a linked group of three genes encoding a sensor kinase/response regulator pair of the two-component regulatory system family and a protein with an EAL domain. The same system was independently discovered in two laboratories [86] [111]. Kulasekara and colleagues identified these genes during screening of a mariner transposon library of P. aeruginosa PAK for activation of expression of silent genes within each of the three cup operons [64, 86]. The products of the cup genes belong to the chaperone/usher family of fimbriae and, when expressed, they accelerate the formation of biofilms on inert surfaces. Several
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insertions lead to activation of cupB- or cupC-lacZ fusions, but none affected cupAlacZ expression [86]. The locus identified by mariner mutanegenesis was named roc (regulator of cup). The EAL-containing protein was named RocR, the sensor kinase RocS1 and the response regulator RocA1. While screening a transposon mutant library of P. aeruginosa PA14, Kuchma and colleagues [111] identified a cluster of genes involved in the development of mature biofilms (referred to as sad, for surface attachment defective). They named the sensor kinase SadS, response regulator SadA and the EAL-containing protein SadR. RocR/SadR is a protein with a modular organization found in other response regulators, where a phosphorreceiver domain is fused to an EAL domain. RocR contains a functional PDE [64] and its enzymatic activity is expected to be modulated by phosphorylation of the adjacent receiver domain, but this modification has yet to be experimentally demonstrated. Transcriptional profiling and genetic analysis using deletion mutants and over-expression concluded that the sadRS locus represses type III secretion genes while it positively regulates the expression of fimbriae encoded in the cupB and cupC operons, but not cupA [111]. Therefore, this two-component system functions in a manner analogous to other molecular switches that control the reciprocal expression of virulence factors utilized by P. aeruginosa during acute or chronic infections [82]. Orthologues of RocR can be found in other bacterial species and have been studied in V. cholerae (VieA) and in Bordetella species (BvgR) [112, 113]. In each case, their respective structural genes are linked to genes encoding a twocomponent systems, which include a response regulator and sensor kinase (BvgS in Bordetella and VieS in V. cholerae) suggesting that these proteins are the kinases responsible for phosphorylation, and possible modulation of the PDE activities, of RocR, VieA, and BvgR [114, 115]. Each of the sensor kinases displays a similar modular organization. At their respective amino-termini they contain two tandem domains found in periplasmic solute binding proteins, flanked by transmembrane segments. RocS1 (and BvgS) also contains a cytoplasmic PAS domain adjacent to the second transmembrane segment. PAS domains are modular domains that function in sensing oxygen and/or redox potential of the cell [107]. This is followed by the cytoplasmic transmitter region containing the sensor kinase module. The next conserved domain in RocS1 (and in VieS and BvgS) is a receiver domain, containing the aspartic acid that serves as an acceptor of phosphate from the auto-phosphorylated histidine in the HisKA domain. These proteins also include at their carboxy-termini a conserved histidine-containing phosphotransfer domain (Hpt), which is an acceptor of phosphate from the aspartate of the receiver domain. From the Hpt domain phosphates can be transferred to acceptor aspartate residues on the receiver domains of response regulators, in P. aeruginosa, RocA1 and RocR. The known phenotypes associated with RocR appear to be antagonistic to the expression of those cellular functions controlled by the RocS1/RocA1 twocomponent system. Although not proven, RocS1 (SadS) is the likely kinase responsible for phosphorylation of the aspartic acid residues in RocA and RocR.
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As shown in the model in Fig. 4.7, the two proteins likely compete for phosphorylation by the RocS1 histidine kinase. It is conceivable that bacteria growing under conditions favoring the expression of biofilm are exposed to signals which interact with the periplasmic solute binding domains of RocS1, and activate the histidine autokinase activity of RocS1. Preferential binding of RocR to the Hpt domain of RocS1 would result in the efficient phosphorylation of RocS1, whilst at the same time interfering with the phosphorylation of RocA1. Since phosphorylated RocA1 is required for biofilm formation, including the expression of cupB and cupC genes, the inability of RocA1 to become activated would limit its transcriptional activity. In contrast, phosphorylated RocR1 would be enzymaticaly active and interact with those adapter proteins or potential riboswitches that were modulated by bound c-di-GMP. An interaction between the response regulator domain of both RocR and RocA1 and the Hpt domain of RocS1 has been demonstrated using a bacterial two-hybrid system [64], suggesting that both of these proteins, under certain conditions, can serve as substrates for phosphorylation or de-phosphorylation catalyzed by RocS1. Reversal of the process, favoring the expression of RocA1 transcribed genes, may involve a change of the selectivity or kinetic preference of RocS1 to preferentially phosphorylate RocA1 at the exclusion of RocR, dampening its PDE activity.
4.7.9 ToxR/RegA and Regulation of Expression of Exotoxin A ToxR (also known as RegA) is encoded in PAO1 by the open reading frame PA0707. ToxR is a 259 amino acid EAL domain-containing protein. It was initially identified as a regulator of exotoxin A (ToxA) production in P. aeruginosa implying that c-di-GMP adversely affects the expression of this potent protein toxin. The toxR gene appears to be co-regulated with the toxA gene, with the expression of both being subject to control by a transcriptional factor (PtxR) and an iron-dependent regulatory mechanism. Cell fractionation studies have shown that in both P. aeruginosa [116] and E. coli [117], ToxR associates with the membrane fraction despite the absence of obvious transmembrane domains. Although ToxR protein was initially believed to function as a transcription regulator [118], it was not possible to demonstrate an interaction of this protein with DNA regulatory sequences adjacent to the toxA gene. This is consistent with the absence of a recognizable DNA-binding motif in the protein. Moreover, the predicted EAL domain takes up nearly 90% of the ToxR sequence. Nevertheless, several lines of evidence suggest that ToxR enhances transcription of the toxA gene by an as yet unknown mechanism. In the hypertoxigenic strain PA103, an adjacent gene, regB, encodes a 75 amino acid polypeptide which positively affects exotoxin A production when co-expressed with toxR. Among the sequenced strains, only the genome of PA14 contains an intact regB open reading frame, while the others carry a pseudogene, with a mutation in the initiation codon, and it is likely that RegB is not synthesized in those strains. The function of RegB is also not known, however it could conceivably modulate the activity of ToxR.
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4.8 C-di-GMP Receptors There is little doubt that c-di-GMP exerts all of its regulatory activities by binding to various adapter proteins, regulatory domains of modular proteins or cis-acting regulatory RNAs (riboswitches), and that these interactions are translated to cellular responses. During the past several years, a number of c-di-GMP receptors have been identified and linked to specific physiological processes ranging from polysaccharide biosynthesis and direct regulation of gene expression, to motility. In some cases, multiple families of c-di-GMP proteins have been identified in the same organism, which greatly increases the potential diversity of signaling interactions that changing levels of c-di-GMP can coordinate. However, studies of c-di-GMP receptor interactions have also pointed to a diversity of structurally unrelated proteins or riboswitches that can bind c-di-GMP with great affinities, suggesting that c-di-GMP can be recognized by a number of different “folds” in proteins and regulatory RNAs. The properties of some of the characterized P. aeruginosa c-di-GMP binding proteins will be reviewed here (Fig. 4.8).
4.8.1 Allosteric Inhibitory I-Sites in the GGDEF Domain A number of genetic and biochemical approaches were used to show that DGCs contain, within their GGDEF domains, a c-di-GMP binding site (referred to as the I-site), whose occupancy results in non-competitive inhibition of enzymatic activity. Based on mutational analyses the core motif consists of the sequence RXXD and is found near the c-di-GMP binding site, with D located 6 amino acids upstream from the first G of the GG(E/D)EF sequence. The first example of a binding site for c-di-GMP was provided by the analysis of the crystal structure of the Caulobacter crescentus DGC PleD [50, p. 237]. The structure of the enzyme-product complex showed the presence of several c-di-GMP molecules tightly bound to the dimer. One molecule was located at the catalytic A-site where each dimer provides half of the functional catalytic pocket. Binding of c-di-GMP at this site involves various charged/polar residues, mainly glutamate, lysine, aspartate, asparagine (PleD E371, K332, D344, and N335). The second binding pocket functions as a tight and specific allosteric inhibition site for c-di-GMP and was located at the interface between the GGDEF domain and a nonfunctional receiver-like domain of PleD. The c-di-GMP binds via a multitude of interactions, including a set of arginines and aspartates, located in the GGDEF and in D2, one of the two receiver domains of each protein monomer (PleD R390, R359 in the GGDEF domain and R178 in the D2 domain, D362 in the GGDEF domain and R148 in the D2 domain). The residues found in the PleD I-site are highly conserved in DGC enzymes and it has been proposed that non-competitive allosteric inhibition is a common trait of DGCs [51]. Analysis of PleD mutant proteins showed that R359 and D362 are required but not sufficient for c-di-GMP binding and feedback inhibition. These two conserved residues need to be located in the appropriate sequence context of the I-site loop, as the presence
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of specific intervening residues, such as RNRD or RGQD, will abolish c-di-GMP binding [51]. In P. aeruginosa, most of the 34 GGDEF domains contain a canonical RxxD I-site motif, with the exception of PA4601, PA0285, PA5442, PA5017, PA4367, PA4396, PA4959, PA2567, where none of the conserved residues are present. The GGDEF domain of PA3258 lacks the arginine in RxxD. Interestingly, the hybrid GGDEF/EAL proteins PA4396, PA4959 and PA2567 also lack most of the conserved GGDEF residues (only the conserved F is present) and none of the proteins without the conserved I-domain were enzymatically active in vitro [86]. These findings suggest that the loss of the catalytic residues through mutational decay was accompanied by the corresponding mutations in the regulatory I-domain. These proteins may be representing inactive relics or DGCs, regulating the PDE function of the adjacent EAL domain via binding of GTP, as shown for CC3396, a hybrid GGDEF/EAL protein with only PDE activity [52]. No GGDEF domain protein in P. aeruginosa has intact I-site but degenerate A-site. In other bacteria this class of GGDEF domains have been shown to act as c-di-GMP-binding effectors [119]. The crystal structure of WspR (PA3702) has been solved [105] (Fig. 4.5) and together with the crystal structure of PleD, has served the basis for testing several hypotheses on the mechanism of catalysis of c-di-GMP formation and the structure of the allosteric I-site, which represents an example of a c-di-GMP-binding motif. The features of the c-di-GMP-binding site of the WspR were discussed in Section 4.7.1. However, a direct comparison of the two proteins is complicated by the unusual domain organization of PleD. The second receiver domain (D2) in PleD, not found in any other DGC, appears to play a role in the binding of c-di-GMP during allosteric regulation of its activity. This effect may be unique to PleD [105]. As discussed in Section 4.7.1, WspR proteins crystallize as tetramers composed of two head-to-head dimers (Figs. 4.5 and 4.6). Two c-di-GMP stacked dimers bind at each of the I-sites comprised of the canonical motif R242 xxD245 from one monomer (Fig. 4.5), and of residue R198, located in the GGDEF domain of a symmetry-related monomer (not shown in Fig. 4.5). This mode of binding is reminiscent to that observed in BeF3 activated PleD crystals [94], even though WspR shows an overall different quaternary structure [105]. Site-directed mutants lacking either the R242 or R198 side chains at the I-site do not bind c-di-GMP but are still highly active enzymes [105]. The common feature amongst these various I-sites is the presence of a network of charged residues like arginine and aspartate making ionic and hydrogen-bonding contacts with c-di-GMP, a common feature in many protein-nucleotide interactions, including those in DNA-transcription factor complexes [120].
4.8.2 The c-di-GMP Catalytic Site of EAL and HD-GYP Domains Since the hydrolysis of c-di-GMP by enzymes with the EAL or HD-GYP domains is preceded by binding of the substrate to the catalytic sites, all PDEs formally represent a family of c-di-GMP binding proteins. Currently, there is little structural
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information available on PDEs that would allow the identification of the structural features of the c-di-GMP biding pocket in these proteins. So far, only structures for two EAL domain proteins are available in the Protein Database (Bacillus subtilis 1uze and Thiobacillus denitrificans 2r6o), solved as part of high-throughput structural genomics efforts, and both are apo-proteins. Modeling of the primary sequence of RocR’s EAL domain into these crystal structures followed by sitedirected mutagenesis and computational docking analysis has lead to the proposal that together with the conserved residue E175, another five charged/polar residues cluster around a Mg+2 ion in the crystal structure (N233, E265, D295, K316, E352) [71]. It is conceivable to hypothesize a role in c-di-GMP binding for those EAL domain containing proteins that have degenerate EAL motifs (e.g. KVL in PA1433 and KLS in ToxR), and/or that do not show any catalytic activity [64]. Such proteins may constitute a regulatory subclass of c-di-GMP binding EAL domains. Two recent studies confirmed this hypothesis, demonstrating a role for c-di-GMP binding by proteins with degenerate catalytic domains. The E.coli YcgF protein, a blue-light sensor, contaisn an EAL domain with extensive substitutions in conserved catalytic residues, it lacks PDE activity but is capable of binding c-di-GMP in vitro. Simarly, the P. fluorescens LapD a protein involved in biofilm formation contains degenerate and enzymatically inactive GGDEF and EAL domains, yet it binds c-di-GMP [121][122]. Therefore, YcgF and LapD represent c-di-GMP receptors recently evolved from domains involved in the metabolism of this regualotry dinucleotide.
4.8.3 PilZ Domains and the Role of Alg44 in Alginate Biosynthesis Using computational analyses, Amikam and Galperin identified a c-di-GMP binding motif in a family of proteins that contain a so-called PilZ domain [123]. The PilZ domain (see Fig. 4.9) (Pfam07238) is a conserved sequence of 118 amino acids, with similarity to the C-terminal portion of BcsA, the c-di-GMP-regulated G. xylinus cellulose synthase [124, 42, 43]. Based on this observation, Amikam and Galperin used PSI-BLAST searches to identify similar PilZ domain-containing proteins in other bacterial genomes. The PilZ domain was named after a P. aeruginosa protein (PilZ) required for type IV pili biogenesis and twitching motility [80, 124]. PilZ domains in Salmonella enterica (YcgR), C. crescentus (DgrA, DgrB), Vibrio cholerae (PlzD), and P. aeruginosa have been shown experimentally to bind c-di-GMP with submicromolar affinities [125, 84, 126–128]. PilZ domains can be found in numerous bacterial species. The current Pfam database shows that there are 1,574 PilZ domain-containing proteins distributed in 391 bacterial species. Proteins with PilZ domains can be modular and are organized in 48 distinct architectures. They include single domain PilZ proteins (68% as single PilZ domain proteins), proteins with tandem PilZ domains, PilZ domains linked to GGDEF/EAL domains or PilZ domains linked to other enzyme domains (protein kinases, glycosylases, glycosyl transferases, proteases). Other PilZ domains are linked to sensory domains (PAS, MCP, GAF, Heme binding, AMP binding,
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Fig. 4.9 Structural models of PilZ proteins. (a) Structural model for Pseudomonas aeruginosa PA4608 derived from one of the 16 NMR structures deposited in PDB code 1YWU. (b) Structural model of Vibrio cholerae PlzD (VCA0042) dimer derived from the x-ray diffraction atomic coordinates deposited in PDB 2RDE. Both were visualized and rendered using PyMol. The conserved residues of the PilZ family are indicated in the PA4608 structure (in parenthesis are the corresponding coordinates for Alg44 residues). Shown in the PlzD model are the two arginine side chains (yellow patches) providing the double guanidino network of chemical bonds interacting with c-di-GMP (shown in blue sticks); the YcgR domain is involved in the dimerization of PlzD
nitrate/nitrite sensing), signaling domains (phosphoreceiver, Ser/Thr Kinase), or DNA-binding domains. The genomes of the same organisms usually contain genes for putative DGCs, although in some sequenced genomes, such as Staphylococcus aureus [129], Anabaena sp. strain PCC 7120 [130], and Mycobacterium species [131] PilZ coding sequences are absent. In Staphylococcus aureus this may not come as a surprise, as the sole DGC protein does not seem to have DGC activity and the phenotypes it controls are c-di-GMP-independent [132, 58] The genomes of all P. aeruginosa strains sequenced to date carry genes for eight proteins with PilZ domains [84] (Fig. 4.8A). Measurements of c-di-GMP binding to recombinant maltose-binding protein-PilZ fusion proteins identified seven of these as functional receptors for the dinucleotide. Interestingly, the only protein unable to bind c-di-GMP in vitro is PilZ (PA2960) [124]. Conceivably, native PA2960 may function as a receptor for c-di-GMP in P. aeruginosa, although its inability to bind the dinucleotide in vitro is consistent with the absence of certain highly conserved residues found in other PilZ domains [84]. Over-expression of the enzymatically active P. aeruginosa c-di-GMP PDEs leads to suppression of production of the alginate exopolysaccharide by mucoid strains, while over-expression of DGCs resulted in an increase in alginate production by an already mucoid strain, demonstrating a role for c-di-GMP in regulating alginate biosynthesis. The gene for Alg44, one of the members of the P. aeruginosa
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PilZ family, is located in a cluster encoding determinants for the production of the intracellular alginate and its deletion results in mutants unable to synthesize the polysaccharide [123, 133, 134, 84, 135]. Alg44 is a bipartite modular type II membrane protein [136]. It contains a cytoplasmic PilZ domain at its aminoterminus, that is separated from the carboxy-terminal periplasmic NolF domain by a short hydrophobic stretch of amino acids (Fig. 4.8A). The NolF-AcrA-EmrA domain is found in putative coupling proteins of various small molecule or protein transporters, where they link the ATP-binding cassette-containing inner membrane proteins to the outer membrane protein components of the transport system [137]. Binding of c-di-GMP to the PilZ domain of Alg44 requires several conserved amino acids found in other members of this family, and these are located on the exposed loop of the domain. This region of the protein was also shown to be one of the two c-di-GMP-binding surfaces of the PilZ domain of C. crescentus DgrA [125]. Specifically, the introduction of single alanine substitutions at positions 21, 44, or 46 significantly reduced the binding of c-di-GMP to the PilZ domain of Alg44. A mutant form of Alg44 PilZ in which the arginines at positions 17 and 21 were substituted with alanines showed the most significant impairment in c-di-GMP binding. The same amino acid substitutions, when introduced into full-length Alg44 significantly reduced the amount of alginate produced by the bacteria, providing evidence that binding of c-di-GMP to this region of the protein is required for the function of Alg44 in alginate biosynthesis. The critical role of the various amino acids in the exposed loop of Alg44 for c-di-GMP binding was supported by genetic studies using V. cholerae PilZ domaincontaining protein PlzD. Alanine substitutions in PlzD for residues corresponding to the R17, R21, D44 and S46 of Alg44 resulted in a loss of c-di-GMP binding [126]. Similarly, using the PilZ domain containing protein DgrA, involved in the control of flagellar motility in C. crescentus, Christen and colleagues [52] have shown that ultraviolet light-mediated crosslinking of c-di-GMP to DgrA was abolished when two arginines (corresponding to R17 and Q18 of Alg44) were changed to alanines. However, c-di-GMP could be efficiently crosslinked to a variant of DgrA with a mutation in the aspartate corresponding to D44 of Alg44. Since the binding of c-di-GMP to Alg44, and not its membrane localization, is affected by substitutions in the residues that abolish interaction of Alg44 with its dinucleotide ligand, Alg44 functions in alginate production as a cytoplasmic regulator of biosynthesis and/or export of this polysaccharide from the cell. Currently, the structures of three different PilZ domains have been determined. The structures of P. aeruginosa PA4804 and C. crescentus DgrA were solved using solution NMR and the structure of V. cholerae PlzD was determined both as an apoprotein, and bound to c-di-GMP, by X-ray crystallography [138, 127]. In PA4608, the PilZ domain is organized in six-stranded antiparallel β-barrel flanked by partially structured helices, while in PlzD the terminal helix is followed by two short β -strands (Fig. 4.8). The first helix is less structured assuming a predominantly random coil conformation. Even if PilZ domains do not seem to have a conserved surface, they contain tight clusters of residues that may together provide a binding site for c-di-GMP. Specifically, residues that are conserved in the
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family of motifs RxxxR and D/NxSxxG (i.e. R9, R13, D35, S37, G40) in PA4608 are located at the top of the barrel, suggesting that this is the likely binding site. However, a more definitive model of the c-di-GMP binding site was deduced from the crystal structure of V. cholerae PlzD, where a comparison of the apoprotein and c-di-GMP-bound forms was made [138]. In PlzD, c-di-GMP is bound in a cis-conformation with the planes of both guanine bases oriented parallel to one another, as seen in the allosteric I-site of GGDEF domains. C-di-GMP contacts seven of the nine strongly conserved residues in the PilZ domain, including three in a long N-terminal loop that undergoes a conformational switch as it wraps around c-di-GMP. In particular, the location of the guanidino groups and relative sets of contacts of the arginine side chain of the RxxxR motif with c-di-GMP in PilZ proteins is reminiscent of the R178 and R359 guanidino groups present in the allosteric I-site of PleD, indicating a common thread in c-di-GMP protein interactions.
4.8.4 PelD and Allosteric Control of Pel Polysaccharide Production The search for a mechanism explaining the effect of altering cellular c-di-GMP levels on Pel polysaccharide production in P. aeruginosa resulted in the identification of the cytoplasmic membrane protein PelD (PA3061) as a receptor for c-di-GMP [104] (Fig. 4.8B). PelD is encoded by the fourth gene in the pel operon, which expresses the structural genes required for Pel polysaccharide production. The transcription of this operon is increased in P. aeruginosa containing mutations in genes affecting various surface properties and bioflm formation including retS, wspF, and fleQ, as well as from the production of the Pel polysaccharide is stimulated by an artificial increase in c-di-GMP levels resulting from over-production of several DGCs [82] [57, 106]. Direct binding assays showed that c-di-GMP binds to the cytoplasmic portion of PelD [104]. This part of PelD has a predicted secondary structure resembling those found in the GGDEF domain of the C. crescentus DGC PleD. Specifically, a loop of PleD with the exposed allosteric inhibition I-site responsible for binding of c-di-GMP during allosteric regulation of its enzymatic activity, shares several structural features with a portion of PelD that contains the residues corresponding to the RxxD motif. In PelD this sequence is R367 xLD370 . Site-directed mutagenesis showed that these residues, and a more C-terminal arginine residue (R402), are required for c-di-GMP binding and Pel polysaccharide production [104]. PelD is a 455 amino acid protein composed of an N-terminal four-helix transmembrane segment domain followed by a large cytoplasmic domain (Fig. 4.8B). Genes for PelD orthologues, and genes for the Pel biosynthetic machinery, are present in all sequenced P. aeruginosa genomes, and in the genomes of several other organisms including P. fluorescens, and Ralstonia, Marinobacter, Pseudoalteromonas, Geobacter and Alkanivorax species. PSI-BLAST and Pfam analyses show only weak homology between the PelD cytoplasmic domain and the cyclic nucleotide binding domain GAF, spanning amino acid ∼180 to ∼310,
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and therefore does not include the R367 xLD370 motif. Mutagenesis of conserved arginine, histidine, and serine residues in the PelD region corresponding to the putative GAF domain do not affect c-di-GMP binding or Pel production, indicating that the GAF domain of PelD is likely not implicated in c-di-GMP binding. To date, none of the crystal structures of the PelD orthlogues have been solved.
4.8.5 FleQ and the Control of Transcription of Pel Polysaccharide Biosynthetic Genes In P. aeruginosa, an increase in c-di-GMP concentrations, as a consequence of constitutive synthesis of the dinucleotide in a wspF mutant, suppresses motility, leads to an increase in the expression of the Pel and Psl polysaccharides and consequently, a wrinkly colony morphology (see Section 4.7.1) [57]. A mutation in the gene encoding the Class I flagellar transcriptional regulator FleQ (PA1097), produced wrinkly colonies resembling those displayed by the wspF mutant [106]. Based on the analysis of the fleQ transcriptome, they concluded that in addition to its regulatory role in flagellar gene expression, FleQ is a negative regulator of the pel and psl operons, acting at the same level as c-di-GMP, and therefore may function as a receptor for c-di-GMP. FleQ activates flagellar gene expression, and its activity is antagonized by another regulatory protein, FleN, which binds to FleQ forming high molecular weight complexes in the presence of ATP (Fig. 4.8B). Gel-shift analyses were used to demonstrate that c-di-GMP acts as a regulatory molecule by abolishing the binding of the FleQ/FleN/ATP complexes to the pelA promoter [106]. Interestingly, c-di-GMP does not have a similar effect on the transcription of flagellar genes, nor does it affect the binding of FleQ to the promoter of the fleSR operon, which is the main FleQ target in the flagellar regulatory cascade. This suggests that the ability of elevated c-di-GMP concentrations to inhibit flagellar motility is not due to a block in the transcription of flagellar components but by interfering with flagellar assembly or function. The P. aeruginosa FleQ is a modular protein with similarities to transcription regulators that activate transcription from promoters transcribed by σ54 -containing RNA polymerase. It is composed of an N-terminal FleQ domain (pfam 06490), a domain related to Che-like phosphoreceivers but lacking the conserved aspartate, a central domain belonging to the AAA+ superfamily of ATPases, and a C-terminal helix-turn-helix DNA-binding domain (Fig. 4.8B). Deletion of the N-terminal FleQ domain enhanced the ability of this truncated protein to bind the pelA promoter, but this FleQ mutant was still able to bind c-di-GMP. These results suggested that the c-di-GMP binding site was not within the FleQ domain. The sequence of the truncated FleQ has no obvious sequence homology to the known c-di-GMP binding proteins such as the allosteric I-sites of DGCs, catalytic sites of PDEs, PleD or any members of the PilZ family [106]. It is conceivable that c-di-GMP interacts with the residues in the Walker A motif (GxxxxGK[S/T]) [139] of the AAA+ domain [140, 141], normally involved in ATP binding, or alternatively, c-di-GMP binds to an entirely novel protein motif.
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4.9 Concluding Remarks It is not surprising that bacteria have evolved complex regulatory mechanisms to coordinate the expression of various niche-specific factors. Among these, modified nucleotides play an important role and c-di-GMP has emerged as a key component of a signal transduction pathway, that controls the production of surface and secreted proteins and polysaccharides potentially at all levels, including transcription, translation, localization and activity. Particularly important are the c-di-GMP-influenced signaling and regulatory pathways in P. aeruginosa because, compared to most other microorganisms, it is capable of adapting to, and proliferating in, a wide range of environments, including those found in immunocompromised humans. Elevated intracellular c-di-GMP levels facilitate the expression and activities of factors promoting the establishment and maintenance of biofilm communities in chronic diseases, such as infections of individuals with cystic fibrosis, while they have a repressing effect on the expression of factors important for acute infections. These activities of c-di-GMP highlight its central role in coordinating the expression of virulence determinants in a pathogen with the ability to cause a multitude of infections in a wide range of hosts. Ever since the analysis of prokaryotic genomes pointed to the existence of multiple diguanylate cyclases and phosphodiesterases fused to diverse modules, the determination of the molecular basis and specificity of action of c-di-GMP became the “holy grail” of research in the field of prokaryotic signaling. The same question will have to be addressed by those studying the action of cAMP in organisms other than Enterobacteraceae, where genes encoding numerous paralogues of adenylate cyclases, exceeding those found in the human genome, have been identified in various annotated genomes and these are also enzymes with different fused regulatory modules (Class III cyclases). The diversity of c-di-GMP receptors potentially represents another mechanism for amplification and diversification of the c-di-GMP signal, particularly because of what appears to be a large potential of different molecular structures that can bind this nucleotide. These include PilZ, a conserved family of domains also found in multiple paralogs in many organisms where these domains are fused to unique regulatory modules or transmembrane segments. Individual regulatory proteins, such as PelD and FleQ of P. aeruginosa, represent c-di-GMP- binding proteins of unrelated primary and, very likely, tertiary sequences, as do the regulatory I-sites of most DGCs and the c-di-GMP-binding pocket of PDEs. Finally, the discovery of c-di-GMP binding riboswitches further points out that the receptors for this dinucleotide are not restricted to proteins. Equally poorly understood is the basis of phenotypes associated with different levels of cellular c-di-GMP. In P. aeruginosa, two c-di-GMP binding proteins associated with the production of alginate and Pel polysaccharide have been identified, and these suggest an as yet unidentified role for the c-di-GMP/receptor complex in regulating the production of these polysaccharides, while the discovery of FleQ and its role in the transcription of the Pel biosynthetic genes suggests specific function that may be experimentally tested. In most instances, c-di-GMP associated phenotypes such as formation of biofilm, unusual colony morphology, motility, and protein
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secretion, suggest the participation of this regulatory dinucleotide in many general cellular functions without providing important clues that would allow specific placement of c-di-GMP into a defined molecular process. What is apparent from the work in various organisms, and highlighted in this review for P. aeruginosa, is that our understanding of the molecular mechanisms that regulate c-di-GMP metabolism and activity is at a very early stage. Even the cataloging of the molecular components that play a part in regulating c-di-GMP levels has not been completed. The identification and role of environmental signals, and their interactions with the sensing modules (found on DGCs PDEs, and c-di-GMP receptors), is very likely only in its infancy. Elucidation of the rules that allow diverse and potentially redundant proteins to control specific cellular processes through the production of an identical cyclic dinucleotide, will undoubtedly represent a major accomplishment in prokaryotic cellular and molecular biology and there is little doubt that P. aeruginosa represents a model organism where some of the key questions related to c-di-GMP activity will be answered. Acknowledgments We thank Simon Dove and Debbie Hines-Yoder for critical reading of the manuscript. This work is supported by the NIH grant AI021451.
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106. Hickman, J.W. and Harwood, C.S. (2008) Identification of FleQ from Pseudomonas aeruginosa as a c-di-GMP-responsive transcription factor. Mol. Microbiol. 69: 376–389. 107. Zhulin, I.B., Taylor, B.L. and Dixon, R. (1997) PAS domain S-boxes in Archaea, Bacteria and sensors for oxygen and redox. Trends Biochem. Sci. 22: 331–333. 108. He, J., Baldini, R.L., Deziel, E., Saucier, M., Zhang, Q., Liberati, N.T., Lee, D., Urbach, J., Goodman, H.M. and Rahme, L.G. (2004) The broad host range pathogen Pseudomonas aeruginosa strain PA14 carries two pathogenicity islands harboring plant and animal virulence genes. Proc. Natl. Acad. Sci. USA 101: 2530–2535. 109. Jacobs, M.A., Alwood, A., Thaipisuttikul, I., Spencer, D., Haugen, E., Ernst, S., Will, O., Kaul, R., Raymond, C., Levy, R., Chun-Rong, L., Guenthner, D., Bovee, D., Olson, M.V. and Manoil, C. (2003) Comprehensive transposon mutant library of Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 100: 14339–14344. 110. Gotoh, H., Zhang, Y., Dallo, S.F., Hong, S., Kasaraneni, N. and Weitao, T. (2008) Pseudomonas aeruginosa, under DNA replication inhibition, tends to form biofilms via Arr. Res. Microbiol. 159: 294–302. 111. Kuchma, S.L., Connolly, J.P. and O’Toole, G.A. (2005) A three-component regulatory system regulates biofilm maturation and type III secretion in Pseudomonas aeruginosa. J. Bacteriol. 187: 1441–1454. 112. Merkel, T.J., Barros, C. and Stibitz, S. (1998) Characterization of the bvgR locus of Bordetella pertussis. J. Bacteriol. 180: 1682–1690. 113. Tischler, A.D. and Camilli, A. (2004) Cyclic diguanylate (c-di-GMP) regulates Vibrio cholerae biofilm formation. Mol. Microbiol. 53: 857–869. 114. Beier, D., Deppisch, H. and Gross, R. (1996) Conserved sequence motifs in the unorthodox BvgS two-component sensor protein of Bordetella pertussis. Mol. Gen. Genet. 252: 169–176. 115. Camilli, A. and Mekalanos, J.J. (1995) Use of recombinase gene fusions to identify Vibrio cholerae genes induced during infection. Mol. Microbiol. 18: 671–683. 116. Zimniak, L., Dayn, A. and Iglewski, B.H. (1989) Identification of RegA protein from Pseudomonas aeruginosa using anti-RegA antibody. Biochem. Biophys. Res. Commun. 163: 1312–1318. 117. Storey, D.G., Frank, D.W., Farinha, M.A., Kropinski, A.M. and Iglewski, B.H. (1990) Multiple promoters control the regulation of the Pseudomonas aeruginosa regA gene. Mol. Microbiol. 4: 499–503. 118. Wozniak, D.J., Cram, D.C., Daniels, C.J. and Galloway, D.R. (1987) Nucleotide sequence and characterization of toxR: a gene involved in exotoxin A regulation in Pseudomonas aeruginosa. Nucleic Acids Res. 15: 2123–2135. 119. Duerig, A., Abel, S., Folcher, M., Nicollier, M., Schwede, T., Amiot, N., Giese, B. and Jenal, U. (2009) Second messenger-mediated spatiotemporal control of protein degradation regulates bacterial cell cycle progression. Genes Dev. 23: 93–104. 120. Luscombe, N.M., Laskowski, R.A. and Thornton, J.M. (2001) Amino acid-base interactions: a three-dimensional analysis of protein-DNA interactions at an atomic level. Nucleic Acids Res. 29: 2860–2874. 121. Tschowri, N., Busse, S. and Hengge, R. (2009) The BLUF-EAL protein YcgF acts as a direct anti-repressor in a blue-light response of Escherichia coli. Genes Dev. 23: 522–534. 122. Newell, P.D., Monds, R.D. and O’Toole, G.A. (2009) LapD is a bis-(3 ,5 )-cyclic dimeric GMP-binding protein that regulates surface attachment by Pseudomonas fluorescens Pf0-1. Proc. Natl. Acad. Sci. USA 106: 3461–3466. 123. Amikam, D. and Galperin, M.Y. (2006) PilZ domain is part of the bacterial c-di-GMP binding protein. Bioinformatics 22: 3–6. 124. Alm, R.A., Bodero, A.J., Free, P.D. and Mattick, J.S. (1996) Identification of a novel gene, pilZ, essential for type 4 fimbrial biogenesis in Pseudomonas aeruginosa. J. Bacteriol. 178: 46–53. 125. Christen, M., Christen, B., Allan, M.G., Folcher, M., Jeno, P., Grzesiek, S. and Jenal, U. (2007) DgrA is a member of a new family of cyclic diguanosine monophosphate receptors
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M. Merighi and S. Lory and controls flagellar motor function in Caulobacter crescentus. Proc. Natl. Acad. Sci. USA 104: 4112–4117. Pratt, J.T., Tamayo, R., Tischler, A.D. and Camilli, A. (2007) PilZ domain proteins bind cyclic diguanylate and regulate diverse processes in Vibrio cholerae. J. Biol. Chem. 282: 12860–12870. Ramelot, T.A., Yee, A., Cort, J.R., Semesi, A., Arrowsmith, C.H. and Kennedy, M.A. (2007) NMR structure and binding studies confirm that PA4608 from Pseudomonas aeruginosa is a PilZ domain and a c-di-GMP binding protein. Proteins 66: 266–271. Ryjenkov, D.A., Simm, R., Romling, U. and Gomelsky, M. (2006) The PilZ domain is a receptor for the second messenger c-di-GMP: the PilZ domain protein YcgR controls motility in enterobacteria. J. Biol. Chem. 281: 30310–30314. Brouillette, E., Hyodo, M., Hayakawa, Y., Karaolis, D.K. and Malouin, F. (2005) 3 ,5 -cyclic diguanylic acid reduces the virulence of biofilm-forming Staphylococcus aureus strains in a mouse model of mastitis infection. Antimicrob. Agents Chemother. 49: 3109–3113. Neunuebel, M.R. and Golden, J.W. (2008) The Anabaena sp. strain PCC 7120 gene all2874 encodes a diguanylate cyclase and is required for normal heterocyst development under highlight growth conditions. J. Bacteriol. 190: 6829–6836. Kumar, M. and Chatterji, D. (2008) Cyclic di-GMP: a second messenger required for longterm survival, but not for biofilm formation, in Mycobacterium smegmatis. Microbiology 154: 2942–2955. Shang, F., Xue, T., Sun, H., Xing, L., Zhang, S., Yang, Z., Zhang, L. and Sun, B. (2009) The Staphylococcus aureus GGDEF protein GdpS influences protein A gene expression in a c-di-GMP-independent manner. Infect. Immun. 77: 2849–2856. Maharaj, R., May, T.B., Wang, S.K. and Chakrabarty, A.M. (1993) Sequence of the alg8 and alg44 genes involved in the synthesis of alginate by Pseudomonas aeruginosa. Gene. 136: 267–269. Mejia-Ruiz, H., Guzman, J., Moreno, S., Soberon-Chavez, G. and Espin, G. (1997) The Azotobacter vinelandii alg8 and alg44 genes are essential for alginate synthesis and can be transcribed from an algD-independent promoter. Gene 199: 271–277. Remminghorst, U. and Rehm, B.H. (2006) Alg44, a unique protein required for alginate biosynthesis in Pseudomonas aeruginosa. FEBS Lett. 580: 3883–3888. Oglesby, L.L., Jain, S. and Ohman, D.E. (2008) Membrane topology and roles of Pseudomonas aeruginosa Alg8 and Alg44 in alginate polymerization. Microbiology 154: 1605–1615. Binet, R., Letoffe, S., Ghigo, J.M., Delepelaire, P. and Wandersman, C. (1997) Protein secretion by Gram-negative bacterial ABC exporters–a review. Gene 192: 7–11. Benach, J., Swaminathan, S.S., Tamayo, R., Handelman, S.K., Folta-Stogniew, E., Ramos, J.E., Forouhar, F., Neely, H., Seetharaman, J., Camilli, A. and Hunt, J.F. (2007) The structural basis of cyclic diguanylate signal transduction by PilZ domains. EMBO J. 26: 5153–5166. Walker, J.E., Saraste, M., Runswick, M.J. and Gay, N.J. (1982) Distantly related sequences in the alpha- and beta-subunits of ATP synthase, myosin, kinases and other ATP-requiring enzymes and a common nucleotide binding fold. EMBO J. 1: 945–951. Confalonieri, F. and Duguet, M. (1995) A 200-amino acid ATPase module in search of a basic function. Bioessays 17: 639–650. Neuwald, A.F., Aravind, L., Spouge, J.L. and Koonin, E.V. (1999) AAA+: A class of chaperone-like ATPases associated with the assembly, operation, and disassembly of protein complexes. Genome Res. 9: 27–43.
Part II
Life Styles
Chapter 5
Emergence of Pseudomonas aeruginosa in Cystic Fibrosis Lung Infections Joanna B. Goldberg
5.1 Introduction Pseudomonas aeruginosa is typically considered an opportunistic bacterial pathogen, only affecting individuals with a compromised immune system. Despite the presence of a seemingly intact immune system, patients with the disease cystic fibrosis (CF) are uniquely susceptible to lung infections with P. aeruginosa. Chronic lung infections with this bacterium are the major cause of morbidity and mortality in this patient population. This chapter reviews the literature on the emergence of P. aeruginosa in chronic lung infections in CF, an understanding of which may identify strategies for future drug development.
5.2 Cystic Fibrosis Cystic fibrosis (CF) is the most common lethal genetic disease effecting Caucasians. The incidence is about 1 in 3,200 in this ethnic group with lower rates in other groups (1:15,000 in African Americans and 1:31,000 in Asian Americans) [1]. In the United States, there are approximately 30,000 individuals living with CF. Historically the disease was detected by a salty taste on the skin of infants. This phenomenon is now known to reflect the abnormal chloride channel dysfunction that remains one of the criteria for diagnosis even today. At these early times, this symptom was associated with an early death. Other manifestations of the disease reflect the obstruction of organs by viscous secretions. Pancreatic insufficiency, for example, results in the inability to absorb food, making it difficult to grow normally and maintain a healthy body weight. The thickened and sticky secretions of the lung set up an environment for recurrent pulmonary infection and inflammation, resulting in respiratory failure, which is the most common cause of morbidity and mortality.
J.B. Goldberg (B) University of Virginia, Charlottesville, VA, USA e-mail:
[email protected]
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The clinical presentation of CF today is dominated by a chronic cough as well as episodes of increased amount and purulence of sputum, dyspnea, anorexia, and weight loss. While a low-grade fever can accompany these exacerbations, high fever is unusual and sepsis and bacteremia are virtually unheard of, despite the overwhelming number of bacteria in the airways of these patients (108 CFU/ml of sputum) [1].
5.2.1 CFTR Gene The genetic defects responsible for the disease are mutations in the CF transmembrane conductance regulator (CFTR). The CFTR gene, identified in 1989 [2], is located to the long arm of chromosome 7. Now genetic diagnosis can be established by the recognition of a mutation on each allele of the CFTR gene. CFTR is a 1,480 amino acid membrane-bound glycoprotein with a molecular mass of 170,000. It is a member of the ATP-binding cassette (ABC) superfamily of proteins. The protein is comprised of two, six-span membrane-bound regions each connected to a nucleotide-binding domain, which binds ATP. Between these two units is an R-domain, which is comprised of many charged amino acids. The R-domain is a unique feature of CFTR within the ABC superfamily. The protein is a cAMP-regulated chloride channel. Mutated CFTR results in chloride transfer defects that affect the ion composition and mucus secretion of epithelial cells.
5.2.2 CFTR Mutations There are over 1,600 mutations described in the CFTR gene. These have been divided into classes based on the type of mutation and its effect [3]. Class I mutation are characterized by nonsense, frameshift, or splice mutations that result in truncated and mostly non-functional CFTR. Class II mutations, which include the most prevalent deletion of the phenylalanine at position 508 (F508), effecting about 70% of all CFTR mutant alleles, are defective in appropriate trafficking leading to abnormally folded proteins. This abnormally folded CFTR is recognized by cellular proteases and degraded, resulting in the absence of mature protein at the apical membrane. Class III mutations, including the prominent G551D mutation, have CFTR on the apical cell surface, but are defective in the regulation of CFTR. These first 3 classes of mutations are associated with more severe disease, worse lung function, and pancreatic insufficiency. Class IV and V mutations are less severe and associated with milder pulmonary disease and pancreatic sufficiency, and are due to defective conductance and reduced CFTR synthesis, respectively [1].
5.2.3 CFTR Modifiers Even among patients who carry the same CFTR mutations. a wide variation in lung function can be observed [4]. Although many genes have been suggested as potential
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modifiers of this variation, relatively few have been consistently associated with particular CF manifestations. For the most part, studies have found an association between decreased lung function and immune or inflammatory genes (reviewed in [4]). Recently Gu et al. scanned a cohort of CF patients for genetic modifiers and identified the histone-deacetylase-dependent transcriptional co-regulator IFRD1 as a modifier of CF lung disease severity [5]. They showed that intratracheal infection of a Ifrd1-/- mouse with a mucoid CF isolate of P. aeruginosa was associated with delayed bacterial clearance and decreased neutrophil-induced inflammation compared to similarly infected wild-type mice. These investigators also noted that IFRD1 was expressed during terminal neutrophil differentiation. The identification of genes that modify CF lung disease not only reveals the pathogenic pathways that are necessary for understanding this complex disease, but also allow for the potential for increased specificity of diagnosis and additional therapeutic targets.
5.2.4 Role of CFTR in Cellular Functions As mentioned, CF patients are not generally regarded as immunodeficient. However, their response to bacterial infection may be dyregulated [6]. In particular, CF patients have defective mucocillary clearance due to the inability of mutated CFTR to effectively secrete chloride from respiratory epithelial cells leading to a dehydrated, highly viscous airway surface liquid layer, which facilitates colonization with bacterial pathogens. Ceramide accumulation has also been noted in the lungs of cftr-deficient mice and in epithelial cells of CF patients. This results in a high rate of respiratory epithelial cell death, and the DNA released from the dying cells facilitates bacterial adherence. Ceramide accumulation produces a proinflammatory response that precedes bacterial infection [7]. The mechanism by which CFTR mediates ceramide accumulation is not known, but it may be related to the acidification of intracellular vesicles. Airway epithelial cells are the first line of pulmonary defense against microbial pathogens. The role of these cells in the control of P. aeruginosa infection in CF has recently been reviewed [8]. Normal functional CFTR has been recognized on epithelial cells as the receptor for the internalization of P. aeruginosa. This results in the endocytosis and subsequent removal of the bacteria from the airways [9]. When CFTR is mutated, there is less uptake of this bacterium and therefore an accumulation of P. aeruginosa in the airways. Other groups made similar observations and have shown that peptides corresponding to the receptor domain of CFTR block invasion of P. aeruginosa into different epithelial cell lines [10], while contradictory results have been found by others investigators [11]. Consistent with the observations of CFTR functioning as a receptor is that P. aeruginosa is never visualized bound to or internalized by epithelial cells from CF patients [12–15]. The bacterial ligand for this interaction is the outer core region of the P. aeruginosa LPS, which is lost during chronic infection, suggesting that this CFTR-P. aeruginosa interaction
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is relevant only to early stages in infection [16]. During the later stages of infection it is likely that other epithelial cell functions that are defective in CFTR mutant cells are responsible for the ensuing chronic infection [8]. There is some controversy as whether epithelial cells from CF airways are intrinsically hyperinflammatory, but these cells do sense and respond to the presence of bacteria and their products [17]. However the role of other cellular receptors, such as the Toll-like receptors (TLRs) and their importance in responding to P. aeruginosa in the CF infection remains somewhat unclear. A neutrophil-dominated exaggerated inflammatory response is a characteristic feature of CF lung disease. This airway inflammation in CF begins early in childhood, however it is still unclear whether it is a consequence of the mutation of CFTR or occurs as a response to infection. Activated neutrophils produce large amounts of proteases and reactive oxygen species. This “frustrated phagocytic” response is associated with diminished mucociliary clearance from the respiratory tract, the activation of epithelial cell signaling pathways, and the dysregulation of inflammatory signals, leading to the hyperinflammatory response, which contributes to further airway damage [18].
5.3 Epidemiology of Infections in CF As chronic respiratory infections are the cause of progressive and irreversible lung damage and eventually the most common cause of death in patients with CF, recognizing and understanding the microbes that are present in this site is of critical importance. As with other newborns, infants with CF have lungs that are sterile. But unlike normal babies, those with CF are colonized with pathogenic microbes in infancy. As previously mentioned, this is likely due to defective chloride secretion and sodium hyperabsorption in CF leading to dehydration of the airway surface liquid, breakdown of mucociliary transport, and increased adhesion of mucins to the airway surface, which creates an environment conducive to microbial colonization. The bacterial pathogens that have been most frequently identified are Haemophilus influenzae and Staphylococcus aureus, followed by P. aeruginosa, and then in some cases by members of the Burkholderia cepacia complex. The prevalence of these particular pathogens is distinctive and changes with the age of the individual (Fig. 5.1). Chronic colonization of the CF airway by P. aeruginosa, and particularly mucoid strains, worsens prognosis [19], is coincident with subsequent lung function decline [20], and hastens death [21]. However it is now appreciated that the CF airway is a complex and diverse ecosystem (reviewed in [22, 23]).
5.3.1 Viral Infections Whereas research on pathogens in CF has largely focused on the bacteria inhabiting the airways, it is becoming increasingly evident that viral respiratory infections also
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Fig. 5.1 2008 Annual Data Report from the Cystic Fibrosis Foundation (Besthesda, Maryland) patient registry, showing the prevalence of respiratory infections in CF patients by age. Colonization and infection rates follow an age-specific trend with respect to specific bacterial pathogens. Reprinted with permission
have a negative impact on CF lung function. In a study comparing viral respiratory infections between children with or without CF, there was no significant difference in the frequency of illness, yet the children with CF had longer periods of lower respiratory tract symptoms and these episodes were more severe than in children without CF. These CF children also had increases in lower respiratory symptoms after upper respiratory tract infections [24]. Many different viruses have been detected in CF patients; respiratory syncytial virus (RSV) and influenzae A and B most frequently [25]. The relevance of these early viral infections to subsequent bacterial infection has been suggested by a number of provocative studies. It has been shown that RSV is responsible for increased hospitalized times [26], that about two thirds of new bacterial infection are found during viral season [27], and that this (predominantly P. aeruginosa) colonization occurs within 3 weeks of an upper respiratory tract infection [28]. These clinical findings have been mimicked in the laboratory: both influenza [29] and RSV infection [30] have been shown to increase P. aeruginosa adherence to respiratory epithelial cells. Thus, there is now the suggestion that more active intervention be taken to curb these viral infections and perhaps decrease the onset of subsequent bacterial infections.
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5.3.2 Bacterial Infections With respect to the bacterial pathogens, S. aureus is found most frequently in pulmonary infections in infants and young children with CF. The impact of these infections and whether to treat them with prophylactic antibiotics is a topic of intense interest [31]. There is the implication that such infections are not the precursor to inflammation but that the inflammation evident in the CF lung environment is due exclusively to the abnormal function of CFTR; therefore there may not be a benefit from prophylactic treatment for S. aureus. However even if this is true, any infection may provoke a prolonged inflammatory response suggesting that such treatment may be useful. There is also evidence that anti-staphyloccoal prophylaxis actually leads to more frequent infection with P. aeruginosa [32]. More recently, Levy et al. [33]. found that a positive S. aureus culture was actually associated with a lower risk of mucoid P. aeruginosa. A trial that includes both anti-staphylococcal prophylaxis as well as anti-pseudomonal antibiotics will likely be required to clarify this. However, given that (a) the presence of S. aureus was associated with increased lower airway inflammation, and (b) the presence of both P. aeruginosa and S. aureus has an additive effect on markers of lower airway inflammation [34], prophylactic treatment of S. aureus may be well warranted. In vitro studies have suggested that S. aureus may play a more active role in the ensuing infection by P. aeruginosa. Mashburn et al. [35]. monitored the transcriptome of P. aeruginosa during co-culture with S. aureus. They noted decreased transcription of P. aeruginosa iron-regulated genes under these conditions, suggesting that the presence of S. aureus increased the availability of iron. These authors also showed that that iron acquisition required the lysis of S. aureus by P. aeruginosa. Together these results suggests that prior infection of CF patients with S. aureus may lead to an environment that P. aeruginosa can take advantage of to increase the bio-availability available iron, which is generally limiting in the human airway [36]. On the other hand, P. aeruginosa has also been shown to suppress S. aureus respiration and thereby protect this Gram-positive bacterium from killing by the aminoglycoside antibiotic, tobramycin [37]. Thus, these co-infections are complex and these polymicrobial interactions may lead to distinct outcomes that depend not only on the nature of the bacteria themselves but also the condition of the host. The impact of H. influenzae infection in CF is similarly not without controversy. Nontypeable H. influenzae (NTHi) can infect patients with CF, especially early in childhood. In bronchoalveolar lavage fluid (BALF) samples from young, asymptomatic patients Starner et al. [38] found evidence that NTHi formed biofilms in this environment. These bacteria were also shown to form biofilms on airway epithelial cells in vitro and exhibited increased chemokine and cytokine secretion, leading the authors to suggest that they may play a role in the infectious process. The increase antibiotic resistance normally associated with biofilm formation occurred in the context of these biofilms. However thus far there would appear to be no data showing that colonization with NTHi is related to clinical deterioration in CF. In fact, the
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finding that the patients studied by Starner et al. [38] were asymptomatic suggests that these infections may not be responsible for disease. P. aeruginosa is generally considered the most important pathogen in CF lung disease. Interestingly, patients with other chronic infections such as chronic bronchiectasis and advanced stage of chronic obstructive pulmonary disease are also highly susceptible to P. aeruginosa infection [39–41]. This suggests that the ability of P. aeruginosa to persist and evolve in the lung environment represents a trait critical to each of these disease states. In most CF patients, there are at least 3 stages of P. aeruginosa infection (Fig. 5.2) [42]. The prevalence of positive respiratory tract cultures increases with age. In some centers from 10 to 30% in patients ages 0–5 years to 80% in those 18 years and older have P. aeruginosa infections [43], but these numbers can vary and some centers show less infections [44]. In a longitudinal study of infections in children with CF, Li et al. found patients with (1) no P. aeruginosa, (2) initial nonmucoid P. aeruginosa, that can be intermittent, and (3) mucoid P. aeruginosa (Fig. 5.3) [20]. Early P. aeruginosa acquisition is an independent risk factor for mortality in CF [45, 46], and the presence, especially of mucoid strains, is associated with more rapid clinical deterioration [19, 20, 43]. The phenotypes and genotypes of these strains will be described in more detail below. Other bacteria that are considered emerging pathogens in respiratory infections in CF are members of the Burkholderia cepacia complex (Bcc), which includes at least 17 different species. These bacteria, which are quite antibiotic resistant, can cause infections ranging in severity from asymptomatic to acute or chronic [47]. Of most concern is that these bacteria appear to be readily transmissible between patients and can cause a rapid onset of acute pulmonary deterioration with bacteremia, necrotizing pneumonia, leukocytosis, and death within weeks to months, referred to as “cepacia syndrome”. Bcc infections generally occur subsequent to
Fig. 5.2 Schematic diagram of the evolution of P. aeruginosa airway infection in patients with CF. Reprinted with permission [42]
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P. aeruginosa infections. However whether this is due to CF patients living longer and being given intensive courses of antibiotics over longer periods of time and/or alterations in the lung environment due to chronic P. aeruginosa colonization that promotes Bcc infection are topics of considerable interest. Similarly, non-tuberculous mycobacteria (NTM) are being increasingly recognized as a cause of infection in CF [48]. Whether this is due to the fact that clinical laboratories are getting more adept at isolating NTM or whether there is an increase in prevalence of this type of lung infection as patients are living longer is not known [49]. Recently added to the short list of bacteria recognized in CF airway disease are Stenotrophomonas maltophilia and Achromobacter xylosoxidans and species of the genera Ralstonia, Pandorea, and Inquilinus [50, 49]; their role in CF lung disease pathogenesis is still being determined.
5.3.3 Fungal Infections A review of the fungi present in respiratory sections of CF patients has just been published [51]. The main fungal species associated with CF are the filamentous fungi, Aspergillus fumigatus, Scedosporium apiospermum, and Aspergillus terreus, and the yeast, Candida albicans. Other Aspergillus species may also be isolated transiently from CF respiratory secretions. While the role of these fungal species in CF airway colonization is not entirely clear, they likely contribute to the development of a local inflammatory response, which may lead to decreases in lung function.
5.3.4 Newer Non-Culture Detection Methods For the most part, the identification of bacteria associated with lung infections in CF has been based on culturing BALF, sputum, or other specimens from patients. There are a number of limitations of culture techniques, which may not recover or identify all bacteria present in a sample, due to the fastidious nature of some microbes and/or the over abundance of prominent bacteria. With these problems in mind, a comprehensive study was undertaken by Harris et al. [52]. They used BALF from CF and disease control subjects and extracted and amplified 16S rRNA genes. These rDNAs were sequenced and compared to GenBank databases to determine the species and/or phylogenetic relation of the sequence to known species. As anticipated, the CF samples were dominated by sequences of known CF pathogens. The results of this study showed unexpected sequences in both the samples from CF and the disease control subjects. In the case of CF, a number of potential pathogens, including Lysobacter, Prevotella, Coxiellaceae, and Rickettsiales species were detected in some samples. More recently the bacterial diversity in sputum samples from CF patients was evaluated [53]. Through this analysis, 44 different bacterial species including
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5 putative new species were identified. In both of these studies [53, 52], some of the bacteria that were recognized are normally considered flora of the oral cavity; others were anaerobes. Both of these studies suggest that the detection of these unanticipated bacteria may provide an explanation as to why some infections fail to respond to standard therapy and may suggest new pathogens that may need to be considered in the clinical setting. Supporting the validity of this suggestion, members of the Streptococcus milleri group (SMG) are becoming increasingly recognized as dominant members of the microbial community of the CF lung. When these bacteria were treated, there was a resolution of pulmonary exacerbations and a return to clinical stability in these patients [54]. In addition to using 16S rRNA sequences to identify the inhabitants of the CF lung environment, the new field of metagenomics should be applied to this type of investigation. This culture independent method allows not only the recognition of the microbial communities in this niche, but sequencing of microbial DNA would reveal the genes present in this environment. Additional studies could include characterizing the transcriptome and proteome to define the metabolic network (the “microbiome”) of this community. The effect of prolonged antibiotic use and how this alters the flora of this environment would also be an important avenue of investigation.
5.4 Infection Models of CF P. aeruginosa is recognized as the major pathogen in CF, however the exact factors required for its virulence are still being uncovered. The general approach for such investigations includes constructing strains defective in the production of putative virulence factors. However once constructed, a system must be in place to evaluate the importance of these factors to infection in order to determine whether they might be reasonable targets for anti-infective therapies. Both tissue culture and animal infection models have been used.
5.4.1 Tissue Culture Models There are numerous immortalized tissue culture epithelial cell lines available that have been used to assess CF cell biology. These have been useful in testing P. aeruginosa mutants as well. The read-outs for such analysis include bacterial attachment to and invasion into cells as well as cytotoxicity and induction of apoptosis (reviewed in [55]). With respect to CF, there are CFTR-/- cell lines available, as well as corrected cell lines. Some of these are polarized, while others are not. However immortalized cell lines do not typically maintain proper cellular differentiation or other epithelial cell properties. Primary cells can address some of these issues, but they have limited availability and may have different attributes depending on the patient from whom they were obtained. Finally, there is the general concern that considering CF simply as an epithelial cell defect ignores the complexity of this disease.
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5.4.2 Animal Models For the most part, the majority of P. aeruginosa infection models reflect acute diseases. For animal models of chronic P. aeruginosa infection and their value in studying infections in CF, readers are directed to a new review that nicely summarized the state of the art [55]. Rats have been used for many of these studies. In particular, P. aeruginosa has been embedded in agar beads or alginate to avoid the physical elimination and elicitation of an evasive host response and mimic bacteria encased in a biofilms, and inoculated intratracheally [56]. However, most studies are performed with mice. In general, mice have advantages compared with other animal models, including small size and rapid reproduction rate. There are also a number of knock-out strains of mice available to dissect host pathways involved in susceptibility or resistance. They also allow monitoring of various aspects of the infectious process, unlike tissue or cell culture models. Of particular interest are the available CFTR-transgenic or knockout mice [57, 58]. A few studies have shown that these mice can be highly susceptible to chronic P. aeruginosa infection in the lung [59, 58]. Even with these useful mouse models there are anatomic and immunologic differences that must be considered when evaluating infections with CF pathogens. Also, while some non-mammalian models (such as plants, flies, worms, and zebrafish) are even more genetically tractable and represent good models of some P. aeruginosa infectious processes [60], they are less similar to humans than are mammalian models and therefore may have less relevance to CF.
5.4.3 CF Pig Investigators at the University of Iowa have successfully generated a model of CF in newborn pigs [61]. This animal is considered an improved model, as the porcine anatomy and physiology, size, and life span are more similar to humans. In CFTR-/- pigs, no CFTR transcript or protein was detected. Compared to their littermates, the CFTR-/- mice exhibited a number of abnormalities that are typically seen in CF newborns including defective chloride transport, the presence of meconium ileus, exocrine pancreatic destruction, and focal biliary cirrhosis. The lungs of the neonatal CFTR piglets appeared similar to their littermates, in terms of airway epithelial cells and submucosal gland architecture. Interestingly, these animals also did not show any evidence of abnormal infection or inflammation up to 12 h after birth, arguing against the presence of inflammation prior to infection [62]. However whether the pig will develop spontaneous infections with the same array of microbes and whether the disease that follows mirrors the deterioration that is seen in humans with CF remains to be seen. If such clinical presentations are seen, the pig model may represent the ideal system for studies of not only for the physiological manifestations of CF, but also to the relevant bacterial pathogenesis and vaccine development.
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5.5 Phenotypes of P. aeruginosa Isolates from CF Infections The lungs of CF patients are colonized with bacteria within the first few years of birth. In the case of P. aeruginosa, each individual patient appears to acquire a clone that is retained throughout his or her lifetime [63]. Throughout the course of this infection, these clones show a consistent almost predictable pattern of phenotypic alteration that affects growth and expression of particular virulence factors. The phenotype and genotype of P. aeruginosa strains are distinctly different between those found in the environment (or from acute or early infections) and those from chronic CF lung infections. Thus, the interaction of P. aeruginosa in the CF lung environment can be overly simplified as a two-step process, where colonizing bacteria initially express virulence traits required for acute infection, while the subsequent chronic-colonizing form decreases expression these factors, including exoenzymes [64], the motility factors, flagella and pili [65, 66], and the lipopolysaccharide O antigen [67], to promote persistent infection (Fig. 5.4).
Acute Infection (Environmental/Initial)
Chronic Infection
Secreted toxins and enzymes Flagella-mediated motility
Defective LPS OprF
Type IV pili
Overexpression of alginate
Low level of alginate Complete LPS
Type III secretion apparatus and effectors
Biofilm mode of growth
Fig. 5.4 Composite cartoon showing some phenotypes characteristic of P. aeruginosa isolates from acute infections, which are similar to those found in the environment and initial infections in CF patients vs. those from chronic lung infections in CF
5.5.1 Mucoidy and Alginate Production The most apparent difference between early and later isolates is that strains from chronic CF lung infections produce copious amounts of an exopolysaccharide, alginate [68, 69], which gives these strains a mucoid phenotype (Fig. 5.5). The presence of P. aeruginosa with this phenotype in sputum, is essentially pathognomonic for
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Fig. 5.5 P. aeruginosa isolates from sputum samples of an individual with CF. Bacteria are shown after overnight growth on rich agar media. On the left is strain 383, which shows the nonmucoid phenotype that is typical of strains in the environment, in acute infections, and in early lung infections in CF patients. On the right is strain 2192, which shows the mucoid phenotype due to the over production of the exopolysaccharide alginate. The mucoid phenotype is pathognomonic of strains isolated from chronic lung infections in CF patients
CF. The conversion to the mucoid phenotype likely is an adaptation that allows bacteria to survive in the hostile CF lung environment [70]. While specific environmental conditions found in the CF lung induce the conversion to the mucoid phenotype, little is known about what these conditions are or how to control them. P. aeruginosa isolates from early lung infections are usually nonmucoid, motile, and antibiotic sensitive, suggesting that they are acquired from the environment, however their source has not been identified. The mucoid phenotype emerges and results in these bacteria becoming resistant to phagocytosis, antibiotics, reactive oxygen species, and other immune effectors, such as IFN-γ. Mucoidy also gives these bacteria the ability to form biofilms in the lung (reviewed [71, 72]), and predicts a shortened survival [19]. This adaptation results in a chronic infection that can last for years. However, it is becoming more appreciated that not all P. aeruginosa within the CF lung of an individual are identical and that during chronic infection strains may undergo clonal expansion leading to altered colonial morphology and virulence [73]. The alginate genes are normally silent in isolates from the environment and acute infections. Readers are directed to an outstanding recent review on the genetics of P. aeruginosa alginate biosynthesis [72]. The key regulator is the alternative sigma factor AlgT (also known as AlgU or σ22), which induces the expression of both itself and the alginate biosynthetic locus, by binding at the consensus σ22 promoter sites. The algT gene is in an operon with four other genes, encoding MucA, MucB, MucC, and MucD. MucA is an inner membrane protein that appears to be a major regulator; it is an anti-sigma factor that binds to and sequesters AlgT and thereby
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inhibits its ability to initiate transcription. Interestingly, mutations in mucA are those most typically found to be responsible for the conversion to the mucoid phenotype in CF isolates. One common mutation is mucA22, which is a deletion of a single base within a stretch of G residues that results in a truncated form of MucA [74]. A number of single nucleotide polymorphisms have been identified within the mucABD locus in clinical and environmental strains of P. aeruginosa [75]. Even within this varied group of strains, the mucA22 allele was most prevalent. It has been suggested that the deletion of a G residue in a homopolymeric tract is due to exposure of P. aeruginosa to reactive oxygen species in the context of biofilms in the CF lung environment [76]. Conversion of mucoid strains back to the nonmucoid phenotype in the laboratory environment is not due to the restoration of the wild-type mucA allele, but rather is due to spontaneous mutations in the algT gene [77], suggesting an advantage to this mechanism of alginate suppression in vitro. Gene expression studies have shown other genes regulated by mucA [78]. Many genes controlled by mucA were annotated as “hypothetical”. Some of these genes were up-regulated by mucA, while other genes were down-regulated [79]. In our in vitro analysis of two P. aeruginosa strains with phenotypes corresponding to those from early (383) and late (2192) CF infections (Fig. 5.5), we found differences in both the transcriptome [80] and the proteome [81]. At midexponential phase under laboratory conditions, we observed unique gene expression patterns between these two strains, with 3.4 % of the transcripts (188/5570) being expressed differentially. Of the 188 significantly varied (>1.8-fold) genes, 115 were upregulated in the nonmucoid strain, while 73 were upregulated in the mucoid strain. We observed that the majority of genes from each were described as “hypothetical” and “conserved hypothetical” proteins. Interestingly a comparison of the results of this microarray analysis to an ITRAQ (isobaric tag for relative and absolute quantitation) analysis of protein expression under these same conditions failed to find any strong correlation between these two detection systems (unpublished data). This lack of correlation between the transcriptome and the proteome suggests that in addition to transcriptional regulation, post-transcriptional control is likely critical for final protein expression. This may have relevance to the expression of phenotypes relevant during the emergence of chronic colonization. Even in our proteomic analysis of these two strains by 2-D gel electrophoresis followed by mass spectrometry, we found that a prominent outer membrane protein OprF was detected as a partial protein in the nonmucoid strain compared to the mucoid strain [81]. We speculated that the increased protease production of nonmucoid strains as compared to the mucoid strains [64] influenced the expression of this surface protein, which may also have implications in the expression of proteins from both the bacteria and the host in the context of infection.
5.5.2 Biofilm Formation As mentioned, alginate overproduction resulting in a mucoid phenotype is considered pathognomonic with lung infections in CF and is also generally been
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considered a hallmark of P. aeruginosa biofilms, which are defined as surfaceattached communities of cells encased within a self-produced extracellular matrix. It is now well appreciated that this mode of growth differs significantly from the typical planktonic liquid growth conditions utilized in the laboratory. The regulated developmental process of biofilm formation has been suggested to augment the ability of P. aeruginosa to maintain its life-long chronic lung infection. Years ago “microcolonies” or clusters of mucoid strains were detected in the lungs of CF patients [12, 82]. It is now appreciated that this “microcolony” formation is a step in biofilm development and that alginate is a critical component in these structures. However, it is now clear that even nonmucoid, alginate non-producing strains can make biofilms, indicating that the definition of biofilms being composed exclusively of alginate was overly simplistic [83]. Even nonmucoid strains from CF patients appeared to be capable of forming biofilms [84], indicating in P. aeruginosa biofilms can be formed from other polysaccharides, proteins, nucleic acids, or cellular components [85]. However, it has been found that mucoid strains have a more highly organized biofilm structure that are more resistant to antibiotics, such as tobramycin, than that formed by isogenic nonmucoid strains [86]. In the context of CF, nonmucoid biofilm-forming P. aeruginosa appear to be the progenitors of the chronic-colonizing mucoid biofilm-forming form. However it seems that the population of P. aeruginosa inhabiting the CF lung may include those with different phenotypes in different sites within this environment [87]. The mucus layer of the CF airway where P. aeruginosa grows can be considered microaerophilic or even anaerobic, with a steep hypoxic gradient [15]. Thus, the biofilms that are formed during these chronic infections are distinctly different from biofilms associated with attaching to medical devices, such as catheters [88]. There is a link between biofilm formation and induction of gene expression by quorum sensing (reviewed by [89]). In fact, the quorum sensing autoinducer molecules 3O-C12-homoserine lactone and C4-homoserine lactone have been found in the sputum of CF patients, which mirrors those found in biofilm-grown P. aeruginosa [90]. However, at this point there does not appear to be a direct link between quorum sensing and mucoidy in the CF lung.
5.5.3 Type III Secretion The mucA mutation has effects on other genes in addition to those involved in alginate production. In particular, global gene expression studies revealed that the mucA mutation results in the down regulation of type III secretion genes [79]. Type III secretion is a contact-dependent secretion/translocation pathway that allows proteins to be injected directly into the eukaryotic host cell. P. aeruginosa uses a Type III secretion system to inject effector proteins, ExoS, ExoT, ExoU, and ExoY. ExoS and ExoT function as activators for Rho-GTPase, which ribosylate host proteins. ExoU is a phospholipase and ExoY is an adenylate cyclase. All P. aeruginosa strains studied have the genes encoding the Type III secretion apparatus, but differ in the proteins they secrete [91]. It has been previously noted that carriage of exoU is
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accompanied by the absence of exoS, and that CF isolates carry the exoU gene less frequently and the exoS gene more frequently than do isolates from other sites of infection, as well as isolates from lung infections in patients without CF [92]. In the case of all acute infections, expression of type III effectors is associated with a poor prognosis. On the other hand, the prevalence of strains from CF patients that secrete type III effectors decreases with increasing age of the patient. In a survey of isolates from CF patients, Jain et al. looked for the expression of at least one type III secretion system effector or apparatus protein in a cohort of isolates from CF patients that were defined as those from “first infection”, “chronic infection in a patient less than 18 years old”, or “chronic infection in a patient greater than 18 years old”. As had been observed in their pilot study [93], there was an inverse correlation between type III secretion and duration of P. aeruginosa infection [94]. What mutations were responsible for the diminished expression of the type III secretion proteins were not investigated. However in an earlier study, Dacheux et al. found that among 12 noncytotoxic CF isolates, the defect in the secretion of type III effectors in 9 isolates could be complemented by supplying the transcriptional regulator, exsA, in trans [95]. The translocation of these effectors directly into epithelial cells and phagocytes suggests that they play a critical role in the interaction with host cells; this has also been seen in many animal models of acute infection (reviewed in [91]). On the other hand, the loss of this capacity is not required, and in fact may be detrimental to, the development of a chronic infection. Perhaps the decrease in the injection of effectors into the host cell may allow P. aeruginosa to cause less cellular injury and thereby evade or avoid the immune system.
5.5.4 Genetic Regulation A complex signaling network controls the genes associated with acute and chronic infections [96]. The gene, retS, encodes a hybrid sensor kinase/response regulator. Inactivation of this gene results in hyperbiofilm production and loss of cytotoxicity, due to a block in the secretion of type III effectors. Genomic profiling revealed that retS controls the expression of a number of virulence factors. Most notably, the retS mutation resulted in down regulation of gene whose products are required for early stages of colonization of the respiratory tract of CF patients and upregulation of genes associated with polysaccharides (other than alginate) that are components of the biofilm matrix [96].
5.5.5 Motility No one single mutation was found to be responsible for the nonmotile phenotype associated with chronic P. aeruginosa infections in CF, but providing multiple copies of the gene, rpoN, encoding the alternative sigma factor, on a plasmid, was found to provide partial complementation in some of nonmotile CF isolates [66]. In studies with one nonmotile CF isolate, FRD1, it was found that mutation of algT
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(algU), encoding the alternate sigma factor, σ22, resulted in expression of flagella via the transcription of fliC [97]. Further it was shown that algT repression inhibits fleQ and acts through the transcriptional regulator AmrZ [98, 99]. In addition to genetic regulation by the bacteria itself, studies using a fliC promoter-lacZ fusion, as readout for flagellin transcription, showed rapid reduction in transcriptional activity upon exposure of P. aeruginosa to mucus [100], suggesting environmental regulation. In addition, the innate immune effector neutrophil elastase has been shown to repress flagellin transcription [101]. Altogether these findings suggest that the nonmotile phenotype (and particularly the loss of the dominant surface structures, including flagella and pili) may provide a selective advantage in the CF lung environment, allowing P. aeruginosa to resist phagocytosis and detection by the innate immune system. In addition, it shows that this defect can come about by both bacterial and host derived mechanisms.
5.5.6 Lipopolysaccharide (LPS) Loss of the O antigen portion of LPS has also been recognized in strains isolated from chronic lung infections in CF patients [67, 102]. Strains from initial infections, acute infections, and from the environment are generally LPS-smooth, expressing long O antigen repeating units. Strains from chronic lung infections in CF patients are LPS-rough, expressing no, few, or short O antigen repeating units. This phenotype renders these chronic CF isolates non-typeable or poly-typeable by serogroup-specific antiserum, sensitive to killing by normal human sera, and avirulent in acute models of infection. Interestingly, these defects in O antigen biosynthesis resulting in a serum sensitive phenotype are not seen in other environmental conditions or infections. Based on complementation analysis, we showed that the defect in O antigen expression in CF isolates occurs, in some cases, by mutations in genes of the O antigen biosynthetic locus [103]. This mutational event is distinct for each strain and not necessarily linked to the alteration in expression of any particular regulatory factor, such as is seen with the production of the exopolysaccharide alginate. The data suggest that this selective transition from an LPS-smooth form to an LPS-rough form occurs in the lungs of patients with CF, and that this favors P. aeruginosa avirulence. In addition to changes in the immunodominant portion of the LPS, the O antigen, the lipid A (endotoxin) moiety of P. aeruginosa LPS is also altered during chronic lung infections. Strains isolated from either the upper or lower respiratory tract of young CF patients synthesized more highly acylated lipid A, while those from non-CF patients synthesized a penta-acylated form [104]. These different lipid A molecules interact with TLR4 leading to distinct responses: the penta-acylated LPS attenuates proinflammatory response, while the hexa-acylated form leads to a robust response. It is suggested that these different responses may result in selective advantages for P. aeruginosa during different infectious processes [105]. Additional aminoarabinose and palmitate substituents were also found to be CF-specific. A survey of different strains from CF patients revealed that the palmitate addition on
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the P. aeruginosa lipid A was always found, while the aminoarabinose was present in less than half of the strains [106]. Loss of the lipid A deacylase activity, presumably due to lack of expression of PagL (a 3-O lipid A deacylase), was only seen in strains from patients with severe lung disease [107]. These changes are likely responsible for the increase resistance to host innate immune defense that occurs during the progression of CF lung disease.
5.5.7 Phosphorylcholine (ChoP) Another attribute that has been associated with the chronic P. aeruginosa isolates from CF patients is the expression the phosphorylcholine (ChoP) epitope, a structure crucial for the virulence of several respiratory pathogens [108, 109]. In most P. aeruginosa strains, expression of this epitope is phase variable, being expressed at 25◦ C, but not at 37◦ C [110]. However, when ChoP expression was compared between P. aeruginosa isolates from acute and chronic infections, it was found that the expression of ChoP at 37◦ C was higher among strains from chronic infections. Since ChoP can mediate binding to host cells via platelet-activating factor receptor (PAFR), these results suggest that the expression of this epitope may represent a novel phenotype by which chronic infection isolates could mediate attachment to epithelial airway cells [111].
5.5.8 Response to Growth in the CF Lung In addition to an increase in alginate production, a number of other consistentlyobserved changes emerge in P. aeruginosa strains during chronic lung infections in CF. One of the early observations was that strains were often found to be auxotrophic [112]. Methionine auxotrophy was found to be most common single amino acid requirement. Other than their auxotrophic or prototrophic character, P. aeruginosa isolates colonizing the same CF patient were identical, a finding that suggests that auxotrophs are selected during the course of pulmonary infections in CF patients [113] and that the high amino acid content of sputum promotes this mutation [114]. The pressure that selects for this loss of metabolic capacity is not known. More recently, the composition of CF sputum has been reanalyzed and a synthetic CF sputum media (SCFM) has been developed. This media will be useful to model the CF sputum environment under standard laboratory conditions [115] and could be used to test how auxotrophy emerges. Other conditions that are thought to mimic those occurring during chronic late-stage CF is growth under reduced oxygen tension at a pH of approximately 6.5. Platt et al. [116] have monitored both the transcriptome and proteome of P. aeruginosa grown under these in vitro conditions. They found the up regulation of genes and proteins involved in anaerobic metabolism and suggested that these may represent potentially important targets for therapeutic intervention [116].
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In addition, the CF lung environment appears to promote altered regulation of basic P. aeruginosa metabolic activities, including carbon catabolism. Silo-Suh et al. [117] examined 10 CF isolates and showed that half of them had increased glucose-6-phosphate dehydrogenase activity. By monitoring sequential isolates, this increased activity was found to emerge and be maintained during chronic infection. Interestingly the gene encoding this enzyme, zwf, was critical for growth in lung sputum. The mechanism controlling this switch to increased zwf expression is not known but is apparently independent of the mucoid status of the strains [117]. Perhaps most importantly with respect to treatment for P. aeruginosa is the development of antibiotic resistance, which occurs in the CF lung environment. P. aeruginosa is intrinsically resistant to antibiotics due to its low outer membrane permeability and prominent efflux pumps that export antibiotics [118]. When P. aeruginosa is exposed to concentrations of antibiotics that are effectively subinhibitory, resistance can occur by classical mutational or acquired resistance mechanisms. In addition, during infection P. aeruginosa is exposed to host-derived antimicrobial peptides at the epithelial cell surface. Resistance to these may result in cross-resistance to similar peptide antibiotics.
5.6 Genomic Analyses of P. aeruginosa Strains Until recently, investigators have relied on phenotypic analysis of P. aeruginosa and made correlations with the clinical course of infections in CF. The release of the genome sequence of P. aeruginosa strain PAO1 [119] has revolutionized the study of this pathogen. PAO1 is a well-studied laboratory strain that, while it was not isolated from a chronic infection in a CF patient, has provided a standard for comparative analysis. Whole genome microarrays have been have been used to assess variation among a diverse set of P. aeruginosa strains. Wolfgang et al [120]. examined the genomes of 18 strains isolated from the most common human infections and environmental sources, including four respiratory isolates from very young children (<24 months) with CF. They showed that there was conservation of many genes including nearly all those encoding recognized virulence factors. They also determined that there was no obvious correlation between genome content and infection type [120]. A similar study was performed by Ernst et al. [121], who compared 20 P. aeruginosa strains to PAO1 by DNA microarray. Some of these were from CF patients <8 years of age. As had been seen previously, each patient carried a unique P. aeruginosa lineage. In strains from CF patients, some of the genetic islands recognized in PAO1 were either absent or divergent but there appeared to be no CF-specific pattern noted. Whole genome shotgun sequencing was used to study two P. aeruginosa isolates from 2 different CF patients [122]. An analysis of the O antigen biosynthetic locus and the mucA gene was performed on isolates collected over time from one of the two patients. The results of this showed the presence of an insertion element within the O antigen locus that resulted in a nontypeable phenotype. Also observed
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was the acquisition of the mucoid phenotype due to mutations in the mucA gene. Interestingly, the loss of O antigen expression was well-correlated with acquisition of the mucoid phenotype [122]. A sequence analysis of strains of P. aeruginosa isolated from the same CF patient at 6 months of age and at 96 months showed the genetic adaptation that occurred over time [123]. Whole genome shotgun sequencing revealed 68 mutations. Genes that were mutated included those involved in O antigen biosynthesis, type III secretion, twitching motility, exotoxin A regulation, multidrug efflux, osmotic balance, phenazine biosynthesis, quorum sensing, and iron acquisition [123]. As anticipated, the latter isolate showed phenotypes that were consistent with defects in the production of these virulence factors. As mentioned previously, these phenotypes are associated with decreased virulence in acute models of infection. The sequence of these 68 genes was determined for isolates from this same patient obtained between 6 and 96 months. Approximately one third of the mutations were present in the 60-month isolate and about one half of those were present in the 30-month isolate. This suggests that mutation accumulate in a sequential “parsimonious fashion”. A similar analysis with longitudinally collected isolates from 29 different CF patients was also performed on 24 genes. Interestingly one of the genes that was mutated in most of these isolates was the lasR gene. This has been considered somewhat paradoxical as lasR is known to be the principal quorum sensing regulator, which coordinates the expression of virulence factors and responds to the autoinducers found in the CF lung and responsible for biofilm formation [90]. However these findings suggest that there is a strong selective advantage to eliminate the expression of genes that are regulated by lasR [123]. To assess the clinical implications of lasR mutations, an analysis of 166 P. aeruginosa isolates from 58 CF patients was performed [124]. The lasR mutation was detected as a distinctive colony morphology that includes an iridescent sheen and colony flattening. Mucoidy was also detected by plate observations. These phenotypes were compared to the status of the patient at the time of the strain isolation. lasR mutation prevalence was comparable to that of the mucoid phenotype, but was detected at an earlier age. Lung function declined with age and was worse among patients with lasR mutant infection, similar to what has been observed for mucoidy [124], suggesting that this may be an additional marker for severity of infection. Two CF isolates from different origins have recently been sequenced [125]. Strain 2192 is an isolate from a chronically infected patient and has phenotypic attributes characteristic of the majority of CF isolates (Fig. 5.5). It was also the strain that we used for our proteomic and microarray experiments [81, 80]. The second strain is C3719, the so-called “Manchester epidemic strain” [126]. The comparison of 2192 and C3719 to other sequenced genomes revealed that there was extensive conservation of a set of genes shared by all strains, which represents about 90% of the genome and is referred to as the P. aeruginosa “core genome”. On the other hand, each strain carries a relatively small number (about 10%) of unique sequences, which are part of the “accessory genome”. The acquisition or loss of these genes occurs through horizontal gene transfer events and usually at the same location of
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the genome, referred to as regions of genomic plasticity [125]. From this analysis, it is apparent that P. aeruginosa can use genetic mechanisms to expand and alter its genetic material to survive in various environments including the unique niche of the CF lung. Until recently, P. aeruginosa was considered non-transmissible between CF patients and thus non-epidemic. In general, P. aeruginosa is presumed to be obtained from an environmental source. Except in rare cases of siblings who may have shared strains, for the most part, clones seemed to be exclusive for individual patients, and these evolve over time. However in 1995, typing at a pediatric CF center in Britain identified a widespread infection by a β-lactam resistant strain of P. aeruginosa. This strain, known as the Liverpool Epidemic Strain (LES), is the most frequent clone isolated from CF patients in England and Wales. Unlike the “Manchester epidemic strain”, LES has been shown to cause “superinfection” and has been associated with greater morbidity and mortality. This successful aggressive clone appears particularly adaptable to the CF lung environment. The earliest recognized clone, LESB58, has been recently sequenced [127]. This strain carries the gene cluster encoding the LPS O antigen serotype O6 genes, one of the most common serotypes. However similar to other chronic CF isolates, this strain is nontypeable. While not shown, it is likely that this strain is LPS-rough likely due to a mutation in the rmd gene, encoding GDP-mannose 4,6-dehydratase, within the LPS locus. This strain was also devoid of any form of motility, likely due to alterations in structural or regulator genes controlling these functions. This strain did not have mutations in mutT or mutS; indicating that it is not hypermutable. Therefore mutations in these genes are not responsible for increase in antibiotic resistance seen in this isolate. Interestingly however, subsequent isolates of this epidemic were found to be hypermutable [128]. This hypermutable phenotype of P. aeruginosa isolates from CF patients was initially recognized by Oliver et al. in 2000 [129], who proposed that these strains contribute to the expansion of variation by an increase in mutation rate. Similar observations have now been made by other investigators in the field (reviewed in [88]), which has revealed a complexity with respect to the hypermutability and pathoadaptation of P. aeruginosa. In a recent paper, the same strain collection that was followed by Smith et al. [123] was examined for hypermutation [130]. An analysis of the 90 strains that were collected longitudinally from 29 patients revealed that >15% were hypermutable, and mutators isolates were detected in >30% of patients. Most (>85%) of hypermutator strains were found to be defective in mutS or mutT genes. There was also a correlation between the number of mutator isolates and the number of mutations within the genes that were analyzed. These results indicate that mutators dramatically enhance the process of genetic adaptation and allow evolution of P. aeruginosa to the CF airway environment. The genome of LESB58 was found to contain many more genomic islands and prophage gene clusters than had been seen in other sequenced P. aeruginosa strains. To determine which genes were responsible for the enhanced virulence of this strain, the authors used an unbiased signature-tagged mutagenesis strategy to generate mutants. These mutants were tested in a rat agar bead model that mimics the chronic
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P. aeruginosa infection. Interestingly they found that some prophage genes were essential for virulence in this model, suggesting that the acquisition of these viral genes impacted the competitiveness of this strain [127].
5.7 Therapeutic Options A number of novel approaches have been proposed to combat P. aeruginosa infections in CF patients. These include the development of new antibiotics to target early infection. This type of infection control may help to keep “window of opportunity” open to eradicate P. aeruginosa from the airways [131]. This is especially important as early antibiotic treatment of P. aeruginosa postpones the onset of chronic infection [132]. The recent finding that strains are emerging that are transmissible between CF patients makes this approach even more critical.
5.7.1 Targeting Essential Genes In general, the majority of currently used antibiotics target essential bacterial gene functions including DNA, RNA, protein, and cell wall synthesis. With this as a backdrop, essential genes of P. aeruginosa have been suggested as appropriate targets for new antibiotics. Through transposon mutagenesis [133, 134] and signature-tagged mutagenesis studies [135], essential genes of P. aeruginosa have been recognized. A comparison of the genes inactivated in two transposon libraries that were constructed in different sequenced strains (PAO1 and PA14) revealed 335 P. aeruginosa candidate essential genes not disrupted in either library [134]. Those currently being investigated as drug targets include those involved in cell wall biogenesis: lpxC [136], murF [137], and murC [138].
5.7.2 Vaccines The development of immunotherapies to target P. aeruginosa in CF have showed R ) was tested in only moderate success. An early LPS-based vaccine (Pseudogen patients with CF. Not only was the organism not eliminated from the airways, but febrile responses were observed in 20–40% of patients [139]. This response was likely due to the presence of the endotoxin portion of LPS in this preparation. It has also been speculated that this vaccine may have increased immune complex formation in these CF patients, as they had already been infected with P. aeruginosa [140]. Another LPS vaccine trial in CF patients that were not colonized showed that neither the acquisition nor the course of disease was altered compared to the non-vaccinated control group [141].
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R A small open study with another LPS-based vaccine, Aerugen , that is composed of the O-polysaccharide of LPS, devoid of lipid A, conjugated to P. aeruginosa exotoxin A, was shown to be safe [142, 143]. It induced antibodies to the O-polysaccharides included in the vaccine and these antibodies were opsonic and promoted P. aeruginosa killing by human neutrophils. After 6 and 10 years, there was more infection noted in the non-vaccinated group compared to the vaccinated group. Also, lung function and weight was higher in the immunized CF patients compared to the non-immunized group [144]. A larger placebo-controlled trial of this same preparation was stopped by the manufacturer in 2006, since there was no difference between the two groups in the clinical parameters chosen to measure outcomes. A recent clinical trial of a P. aeruginosa flagella-based vaccine has also met with limited success [145]. This preparation was well tolerated and CF patients developed high serum IgG to the flagella subtypes that were included in the vaccine. The degree of protection against P. aeruginosa that was calculated from the relative risk was 34%. Unfortunately, the second primary endpoint, prevention of chronic P. aeruginosa infection, was not achieved, due a much lower than expected rate of colonization observed in the placebo control group [145]. This is likely due to the initiation of antibiotic treatment following an initial P. aeruginosa exposure in many CF patients enrolled in this trial [140]. It was interesting that among the P. aeruginosa strains isolated from the infected vaccinated group, most were “flagella-positive”, but were expressing flagella subtypes that were not included in the vaccine, suggesting the efficacy of this immunotherapy. At present, the production of this vaccine has been terminated. In fact, a recent report reviewing the literature on the state of the art concerning vaccines to prevent infections with P. aeruginosa in CF states, “vaccines against Pseudomonas aeruginosa cannot be recommended” [146]. Since alginate is a prominent antigen in chronic P. aeruginosa infections, it would appear to represent a viable target for vaccine development. However problems have been found when trying to use alginate as an immunogen. Opsonic, but not non-opsonic, antibodies to alginate protect animals against chronic endobronchial infection [147]. However, when purified alginate was injected into healthy volunteers only a small proportion produced an increased level of opsonic antibodies to alginate [148]. It has also been shown that antibodies to specific epitopes can mediate killing of mucoid strains in vitro [149], while antibodies to alginate produced during chronic lung infection do not [150]. Fully human monoclonal antibodies to alginate that are opsonic and protective against both mucoid and nonmucoid strains have been tested in a mouse model of acute pneumonia [151]. These may prove to be useful in passive administration to CF patients [140]. Further investigations with immunotherapies could be directed at other components expressed either early and/or later during the infectious process (Fig. 5.4); such vaccines could be multivalent. However there would likely continue to be the same problems in evaluating this type of vaccine in individuals with CF: the placebo control group may not develop infections at a rate that would be statistically different enough to warrant further development.
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5.7.3 Biofilm Inhibition Once an infection is established it is more difficult to treat. During chronic infection, P. aeruginosa strains become more antibiotic resistant and their biofilm mode of growth makes them more recalcitrant to treatment. It has been suggested that therapies that inhibit biofilm formation or induce their dispersion might augment antibiotic treatment. Therefore compounds that affect this mode of growth have been investigated. The chelation of iron with lactoferrin, has been shown to inhibit biofilm formation [152]. In addition, a number of different molecules have been found to induce biofilm dispersion including sublethal concentrations of nitric oxide (NO) [153], EDTA [154], and gallium [155]. Rhamnolipid produced by P. aeruginosa has been shown to induce its own detachment from biofilms [156]. Expression of this biosurfactant by P. aeruginosa is likely part of the natural process of biofilm development that provides the bacteria the ability to leave a particular location when conditions become unfavorable. Thus, a better understanding of the natural history of biofilm development may provide hints as to how to inhibit biofilm formation or induce the dispersion of its own biofilm and thus make this pathogen more susceptible to conventional treatment therapies. Care needs to be taken with this approach however, as the disruption of P. aeruginosa biofilms may induce a robust inflammatory response.
5.7.4 Targeting Defective CFTR Function Neutrophilic airway inflammation is a feature of CF lung disease. However it remains controversial whether the CFTR mutation itself causes the proinflammatory state of the CF lung or whether this inflammation is secondary to infection. In either case, it has been found that systemic administration of corticosteroids had a positive effect on decline in lung function in CF patients chronically colonized with P. aeruginosa. Unfortunately the long-term use of these agents was limited due to adverse effects [157]. Other anti-inflammatory agents are in various phases in the drug development pipeline (http://www.cff.org/research/ DrugDevelopmentPipeline/). R ), hypertonic saline, and drugs Lung therapies such as DNase (Pulmozyme that activate an alternative chloride channel, are being used or tested. These target the consequences of the defective CFTR including the build up of mucus, defective airway surface liquid, and defective chloride channel function, respectively [158–160]. People with CF who have severe lung disease often think about having a lung transplant. This is a complicated non-trivial procedure and the supply of good donor lungs for transplantation is quite limited. While the advantages of these surgeries are significant to the patient, there is also the concern of risk of infection after transplant. In particular, lung infections have been shown to be caused by pre-transplant pathogens retained in the upper airways of these patients or by the acquisition of new pathogens [161–163]. A more recent study of 60 CF and 60 non-CF patients
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undergoing lung transplant showed the normal lungs implanted into CF patients had significantly higher susceptibility to P. aeruginosa infections than those transplanted into non-CF patients. This finding suggests that defective innate immunity outside the lungs contributes to infection susceptibility [164]. With respect to the promise of gene therapy to restore functional CFTR and other correctors or potentiators that could reverse the effects of CFTR mutation, there has been limited progress to date [8]. Studies of CFTR and CFTR-gene modifiers have indicated a complex interplay between these proteins and environmental conditions that may influence expression. In addition to evaluating the effectiveness of these treatments on CFTR function, it will critical to determine how they impact the development of bacterial lung infections, which remain the major cause of death in these patients.
5.8 Conclusions In conclusion, the transformation of P. aeruginosa from the initial infecting agent to the long-term chronic colonizer in CF patients is due to its ability to adapt to the unique environment of the CF airway. These bacteria lose their invasive and inflammatory capacity due to defects in O antigen production, loss of flagella, defective type III secretion, and modification of lipid A. They also over produce alginate and exist in biofilms making them resistant to immune effectors. Altogether these alterations allow the bacteria to remain localized to the CF airways. However whether other as of yet unidentified factors that track with the expression of mucoidy are more important for this survival is not known. The specific conditions that exist in the CF lung to induce these changes also remain somewhat obscure. It would seem that a thorough understanding of P. aeruginosa lung infections at the molecular level, could increase the chance that new targets based on the recognition of these critical components would be found.
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Chapter 6
Insights into the Life Styles of Pseudomonas stutzeri Elena García-Valdés, Magdalena Mulet, and Jorge Lalucat
6.1 Introduction Pseudomonas stutzeri is a phenotypically well defined species. A combination of morphological, biochemical, and physiological traits allow for its easy discrimination from other Pseudomonas species and from most other bacterial species. Additionally, all P. stutzeri strains are located in the same phylogenetic branch as the other members of its genus when the sequences of several universal essential bacterial genes (16S rDNA, ITS1 region in the rrn operon, gyrB, rpoB, and rpoD) are considered. Notwithstanding these basic traits, the strains within this species are genetically diverse and make up at least 18 genomic groups (called genomovars (gv)) that correspond to different phylogenetic subbranches within the species. The diversity calculated in a multigenic sequencing analysis demonstrated that P. stutzeri is the species with the highest genetic diversity to be studied so far [1]. It may be speculated that this high diversity reflects the many and different ecological niches that are potentially occupied by members of this species, and that the genomic groups that have been described so far are evolving in a speciation process that may give rise to 18 novel species from a genomic point of view. A recent review article has been published that covers the basic biological properties that have been described for P. stutzeri since the discovery of the species more than 100 years ago [2]. In a quick search of the databases, more than 70 research articles in which P. stutzeri appears in the abstract have been published since 2006, indicating its relevance to microbiological research. P. stutzeri is a good model organism to understand bacterial phylogeny, taxonomy, and the speciation process. In addition, P. stutzeri is involved in many biotechnological processes, especially in environmentally-related processes.
J. Lalucat (B) Microbiologia, Departament de Biologia and IMEDEA (CSIC-UIB), Universitat de les Illes Balears, Palma de Mallorca, Spain e-mail:
[email protected]
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In recent years, the molecular and genomic methods for studying bacteria have added novel concepts to our knowledge of the evolution of bacteria and how they can adapt to the ecological niches they occupy. Genomic studies are essential for describing the core genes and pangenome of a bacterial species. The core genes are those common to all members of the species, and the pangenome comprises all the genes present in any strain of the species, but not necessarily present in the majority of strains. These considerations are also valid for the genus. For instance, the core genome of the genus Pseudomonas (after sequencing 15 complete genomes) seems to be limited to 1,997 genes, whereas the pangenome is greater than 20,000 genes [3]. The present chapter will focus on the novel molecular approaches used to understand the diversity of P. stutzeri, its evolution, and the ecological potential and life styles adopted by the strains of this species.
6.2 Phylogeny and Genomovars The basic combination of phenotypic traits that differentiate P. stutzeri from other species in the genus are its wrinkled, yellow-brown colony morphology, its denitrification capability under anaerobic conditions, its use of starch and maltose, and its negative reaction in the arginine dehydrolase test. Other biochemical properties, such as the substrates assimilation tests or the determination of cellular component profiles (whole-cell and outer-membrane proteins, lipopolysaccharides, and fatty acids) are extremely diverse. It is commonly accepted that members of the same species should share at least 70% binding in standardised DNA–DNA hybridisation and/or over 97% genesequence identity for the 16S ribosomal RNA. The strains of P. stutzeri are grouped at least in 18 gv that are numbered from 1 to 19 (the former gv6 was reclassified as a novel species, P. balearica). Members of the same gv have DNA–DNA hybridisation values greater than 70% (or the equivalent Tm value below 5◦ C). Two strains of different gv show a hybridisation value lower than 65% (or a Tm value higher than 5◦ C). There is a clear discontinuity between the genomic groups (Fig. 6.1). The similarity between gv is on the same order as the similarity expected between two bacterial species. The phylogenetic analysis of the 16S rDNA demonstrates a monophyletic origin of all strains that are phenotypically identified as P. stutzeri. The only exception is the Pseudomonas strain OX1, which was previously assigned to P. stutzeri. In a multigenic phylogenetic analysis, the OX1 strain was affiliated with a branch close to P. corrugata, which is distant from the P. aeruginosa intrageneric group and is considered a member of another species [4]. Individual phylogenetic analyses of six or three other housekeeping genes (ITS1, gyrB, rpoB, rpoD, catA, and nosZ) in representative strains of each gv resulted in the same clustering of P. stutzeri, again demonstrating a monophyletic origin of the 18 gv [5]. A concatenated analysis and a consensus analysis gave very similar tree topologies, maintaining the
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Fig. 6.1 Three-dimensional representation of P. stutzeri genomovars based on the Tm value of pairs of strains
groupings of strains; members of the same gv cluster together in the same phylogenetic group. In the consensus analysis, the minimal intragenomovar similarities ranged between 91.5 and 100%, and the intergenomovar similarities ranged between 88.6 and 95.2%. The closest species in the consensus matrix are: P. balearica (83.3–87.3%), P. mendocina (78.0–82.6%), P. aeruginosa (78.4–83.9%), and P. xanthomarina (84.3–91.5%). P. xanthomarina is the only described species that is affiliated within the P. stutzeri phylogenetic branch after a multigenic analysis. It shares some basic phenotypic traits with P. stutzeri, and could be considered a gv of P. stutzeri. However, additional biochemical tests (the inability to hydrolyse starch), physiological studies (the ability to grow at 4◦ C and in the presence of 8% NaCl), and chemotaxonomic analyses (fatty acid composition) justified the proposal of a novel species [6]. Figures 6.2 and 6.3 are three-dimensional distributions of the genomovars and the corresponding phylogenetic groupings of strains within the species. When P. stutzeri strains are compared pair-wise, a very good correlation can be detected between the phylogenetic consensus distances of three or six housekeeping genes and the DNA–DNA hybridisations [5]. The cut-off point proposed for the gv discrimination (70% DNA–DNA hybridisation or a 5ºC change in the Tm) corresponds to a similarity value of 95.2% in the consensus matrix of the partial 16S rDNA, gyrB, and rpoD genes. Recently, Goris et al. [7] studied the average nucleotide identity (ANI) of common genes in the analysis of complete genomes,
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Fig. 6.2 Consensus dendrogram of P. stutzeri strains based on phylogenetic analysis of partial sequences of the 16S rRNA, gyrB and rpoD genes. The bar indicates sequence divergence. Distance matrices were calculated by the Jukes-Cantor method. Dendrograms were generated by neighbor-joining
and found a similar result: the recommended cut-off value for bacterial species discrimination (70% DNA–DNA hybridisation) corresponded to an ANI value of 95 ± 0.5%. The analysis of only three housekeeping genes seems to sufficiently discriminate for the delineation of gv in P. stutzeri and to differentiate it from the closest-related species (Table 6.1).
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0,16 0,14
gv1
dsm50227 gv3
0,12 0,10
gv2
0,08 0,06 0,04 0,02
gv3
0,00 0,16 atc 0,14 0,12 c1 0,10 75 0,08 91 0,06 gv 0,04 2 0,02 0,00
0,00
0,02
0,04
0,06
0,08
0,10
0,12 0,14
0,16
v1
1256 g
ccug1
Fig. 6.3 Three-dimensional representation of 51 P. stutzeri strains based on the phylogenetic distances obtained in the consensus distance matrix. Each strain is represented by a point, whose coordinates are the distances to the reference strain of gv1 (X axis), gv2 (Y axis), or gv3 (Z axis). Circles indicate the clustering of strains of the same genomovar. Changing the reference axis allows the discrimination of the other genomovars
Table 6.1 Reference strains of P. stutzeri. Data obtained from [2, 5]
Strain CCUG11256
Other designations
19SMN4
ATCC17588 DSM5190 LMG11199 Stanier 221 Stanier 224 ATCC11607 LMG1228 DSM6084
DNSP21
DSM6082
ATCC17591 DSM50227
Taxonomy
Isolation
Origin, geographical location, physiological characteristic
Type strain gv1
Before 1966
Clinical, spinal fluid; Copenhagen
Refa . gv2 Ref. gv3
Clinical; Copenhagen Clinical; Copenhagen
Ref. gv4
1956 Before 1952 1988
Ref. gv5
1988
Marine sediment, naphthalene degrader; Barcelona, Spain Waste water, denitrifier; Mallorca, Spain
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Strain
Other designations
SP1402
Taxonomy
Isolation
DSM6083
Former ref. gv6; P. balearica
1988
DSM50238
ATCC17832
Ref. gv7
JM300
DSM10701
Ref. gv8
Before 1966 Before 1980
KC
ATCC55595 DSM7136
Ref. gv9
1990
Ref. gv10
1990
CCUG50544 DSM17089 CCUG50543 DSM17088 CCUG50542 DSM17087 CCUG50541 DSM17086 CCUG50538 DSM17082 CCUG50539 DSM17083
Ref. gv11
2002
Ref. gv12
2002
Ref. gv13
2002
Ref. gv14
2002
Ref. gv15
2002
Ref. gv16
2002
24a75
CCUG50540 DSM17084
Ref. gv17
2002
MT-1
CCUG50545 DSM17085
Ref. gv18
1997
CCUG46542
KMM235
Ref. gv.19
2005
CLN100 28a50 28a39 28a22 28a3 4C29 24a13
a Ref.,
Origin, geographical location, physiological characteristic Waste water, 2-methylnaphthalene degrader; Mallorca, Spain Soil, denitrifier; California Soil; California; natural transformation model organism Aquifer; California Chemical industry waste water; Germany Soil; Tel Aviv airport area, Israel Soil; Tel Aviv airport area, Israel Soil; Tel Aviv airport area, Israel Soil; Tel Aviv airport area, Israel Sea sediment; Dangast, Germany Soil contaminated with mineral oil; Espelkamp, Germany Soil contaminated with mineral oil; Espelkamp, Germany Marine sediment at 11,000 m depth, Mariana Trench Didemnum (marine ascidian), Maldives
reference strain.
6.3 Core/Pan Genome The core genome of a bacterial species is constituted by genes common to all members of the species. The phylogenetic analysis performed demonstrated that the housekeeping genes, and those genes considered to be species-specific, are monophyletic in their origin, and, in this way, part of the backbone of the species [1]. The other genes that code for accessory characteristics are responsible for the high
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phenotypic diversity observed in the species. The results of the DNA–DNA hybridisations indicate that strains of two genomovars only share approximately 50% of their genomes. At the moment, 18 genomovars have been described in the species, meaning that the global genome of the species (the pangenome) is much bigger than the core genome. The extremely phenotypic and genetic diversity of the species must rely on the variability of the accessory genes. The accessory genes are likely also responsible for the enormous adaptive capability of the species to occupy varied ecological niches. The multigenic analysis study performed with the housekeeping and speciesspecific genes clearly demonstrated the essentially clonal population structure of P. stutzeri. However, many other genes conferring relevant physiological characteristics may be acquired by lateral or horizontal gene transfer (HGT). Most strains of P. stutzeri have been isolated by their unusual metabolic capacities that are not present in all strains of the species. In many instances, the origin of these traits is considered to be the result of a HGT process. These traits have been found characterised by comparing the nucleotide sequences of the coding region of novel trait genes with the core genes of the species: discrepancies between the corresponding phylogenies, the mean GC-content, or the bias in the codon usage. These genes must be considered as accessories, but they are relevant for the survival in the specific niche of the ecosystem where the strain was isolated. These genes are added to the core genome and are constituents of the pangenome of the species. Figure 6.4 schematically shows the pan/core genome of P. stutzeri. For simplification, some genes conferring relevant properties to the strains are grouped with those providing:
Contaminants degradative genes: aromatic, aliphatic hydrocarbons Alternative electron acceptors in the respiratory chain: ClO3–, IO3–, Fe(III), Se(VI)
Metals and antibiotic resistance genes
CORE GENOME (housekeeping and species-specific genes) e.g. rrn, gyrB, rpoB, rpoD, nirS, nosZ, amy, Production of enzymes: Proteases Xylanases Laccase
Alternative energy sources: S2–, S2O32– Fe(II)
Niche-specific genes: Rhizosphere competence N2-fixation
PAN GENOME
Fig. 6.4 Schematic representation of the P. stutzeri Pan/Core genome. Only some relevant genes are indicated as examples
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(i) alternative final electron acceptors in the respiratory chain for anaerobic growth, (ii) catabolic biochemical routes for the degradation of unusual substrates, (iii) the production of enzymes that allow the use of polymeric or complex substrates, (iv) antibiotic properties or resistance to metals, (v) alternative energy sources allowing for chemolithoheterotrophic growth, and (vi) niche-specific genes for diazotrophic growth or maintenance in extreme environments. These properties will be discussed in next section that deals with the adaptation to ecological niches and differentiation into ecotypes. The best known mechanism for gene transfer and acquisition in P. stutzeri is transformation. Two other mechanisms exist (conjugation and transduction), but they have not been studied in detail. Bacteria become genetically transformed by taking up DNA from the environment and recombining it into their genome. P. stutzeri has been studied extensively by several groups for its natural transformation capability. Natural competence for transformation is a genetic and physiological property of particular strains. It has been demonstrated that the transformation efficiency is much higher when the exogenous DNA and the recipient are members of the same gv. Once the transforming DNA is taken up, it must be integrated into the chromosome by homologous recombination or, as we will consider later, by transposition. Many careful studies show that the ease with which genes recombine declines dramatically as their sequences diverge [8–11]. This constraint might be taken as a barrier to interspecific exchange and could be used as an upper limit in the delineation of a microbial species [12], or in the delineation of gv in the case of P. stutzeri. A difference in the genome size has been demonstrated between strains of P. stutzeri. A 20% difference exists between some strains, and the genome size ranges from 3.75 to 4.66 Mb in P. stutzeri (Table 6.2). On the other hand, the genome size must be maintained with an upper limit. The gain of new genes must be compensated with the loss of others that are unnecessary in the new niche. In this way, the genome size of P. stutzeri strains is maintained in the lower range of the sizes described for members of the genus Pseudomonas. A better picture on the acquisition and loss of genes will be reached when genomic studies on Pseudomonas have been extended to more species and more strains within the same species. For the moment, 15 Pseudomonas genomes have been sequenced: three P. aeruginosa, four P. putida, three P. syringae of three pathovars, two P. fluorescens, one P. entomophila (a species described by Vodovar et al. [15], close to P. putida), one P. mendocina, and one P. stutzeri. The genome of P. stutzeri A1501 is the first of the species to be sequenced completely. This strain was isolated from rice paddy soils and has been widely used as a crop inoculant in China because it is able to fix nitrogen, and is therefore of biotechnological interest. It belongs to gv1 and its genomic characteristics have been published by Yan et al. in 2008 [14]. Results on the general features of the genome in comparison to other Pseudomonas genomes are summarised in their publication. The single circular chromosome is 4,567,418 bp in length (much smaller than the genomes of the other Pseudomonas genomes), and encodes 4,146 probable proteins, 59 tRNA genes, and four rRNA operons. It carries six genes that encode
nd, not described.
Genomovar Genome size (Mb) rrn operons GC content Gene map
3.75–4.66 4 60.7–66.3
Species
1 4.31 4 65.0 Low resolution physical map
CCUG11256 1 4.56 4 63.8 Gene map available (genome sequence: GSE6572)
A1501
Low resolution physical map
2 4.29 4
ZoBell
3 4.5 4 63.4 Low resolution physical map
AN10
8 4.03 4 63.0 Low resolution physical map
JM300
Table 6.2 Basic known genomic characteristics of P. stutzeri and representative strains (taken from [2, 13, 14])
9 4.66 4 nd nd
KC
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site-specific recombinases of the phage integrase family. It contains 42 copies of repeat sequences that are grouped into 10 types, and the genome also encodes 57 transposases. Four distinct regions with atypical GC content that are flanked by tRNA genes have general characteristics of typical genomic islands, suggesting the recent transfer of genetic material into A1501. A fifth region containing the nif genes might also be considered a genomic island and will be discussed in the next sections.
6.4 Role of Mobile Genetic Elements Several mobile genetic elements have been detected in P. stutzeri. Their role in the chromosomal plasticity of the genome and in the evolution of P. stutzeri will be discussed here and in the section dedicated to their adaptation to ecological niches. Transposons (Tn) are mobile DNA sequence elements that can move from one site to another within the genome. Simple Tn or insertion sequences (IS elements) are found in nearly all bacterial genomes. They have inverted sequences at their ends and code for the enzyme transposase, which mediates the transposition. Composite Tn code for additional genes. IS elements have been described for many catabolic genes [16], and their presence has been taken as an argument in favour of the modular theory of evolution and construction of catabolic pathways for the degradation of organic contaminants. For example, in P. stutzeri AN10, IS elements have been found related to the naphthalene catabolic pathway [17, 18]. IS elements might also be involved in the modulation of gene expression, mostly by insertion within a gene that is inactivated. This situation has been demonstrated in the inactivation of the nahH gene in the naphthalene-degrading strain AN10 by ISPst9, a recently discovered ISL3-like element [19]. The precise excision of ISPst9 restores the gene activity. Integrons have been studied extensively in antibiotic resistant bacterial pathogens, but they may also have an important function in the adaptation of bacteria to novel habitats because they are also involved in the capture of metabolic islands. Integrons are large and complex mobile elements that can capture gene cassettes. In general, they consist of structural genes linked to a 59 bp element (an attC site) and assemble as cassettes at an attachment or capture site (attl) in the chromosome. Both are homologous and used to capture the gene cassette. A strong promoter that drives the expression of the captured genes lies next to the attl site. Class 1 integrons have an integrase gene at their 5 end. When the cassettes are shuffled in their position by excision and reintegration, they are called superintegrons because they contain multiple copies of the gene cassettes. Integrons in P. stutzeri were first described by Coleman and Holmes [20] in strain Q. This group was also the first to experimentally demonstrate that chromosomal integrons can capture gene-cassettes and express the cassette-associated genes [21]. In our laboratory, the screening of integrons in P. stutzeri strains has demonstrated that their presence in strain Q is not an exception (Scotta and Bennasar, personal communication).
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6.5 Adaptation to Ecological Niches; Ecotypes The extraordinary diversity of P. stutzeri is exemplified by its almost universal distribution. This species plays crucial roles in the carbon and nitrogen cycling in many habitats, and is also relevant in the metabolic transformations of phosphorous and sulphur compounds in specific situations. Some strains can use these molecules in their metabolic energy production chemolithotrophically as electron donors or as final electron acceptors in the anaerobic respiratory chain. Some well studied strains will be presented in the frame of the ecological niche that they are able to occupy.
6.5.1 Soil and Plant-Associated P. stutzeri plays a crucial role in the cycling of nutrients in the environment. It participates actively in the carbon, nitrogen, phosphorous, and sulphur cycles, and plays a key role in some ecosystems. The interaction of P. stutzeri with plants is of particular relevance. Microbial plant growth-promoting mechanisms that affect plants directly include fixing atmospheric nitrogen, increasing the availability of iron and phosphorous from the soil and providing these nutrients to the plant, and synthesising phytohormones. P. stutzeri has the enzymatic potential to be involved in all of these processes. P. stutzeri strains are easily isolated from soil samples by specific enrichment methods and have been studied in many instances by their interaction with plants in the rhizosphere. Those strains that are able to grow diazotrophically merit special attention. Their simultaneous capacity for nitrogen fixation and denitrification may be relevant for the overall nitrogen cycling in several ecosystems. Nitrogen fixation occurs in these strains at low oxygen tension under microaerobic conditions, as has been observed in many aerobic diazotrophs. Actually, only two recognised Pseudomonas species are known to be able to fix dinitrogen: P. stutzeri strains and only one representative of P. azotifigens. The two best studied N2 -fixing strains of P. stutzeri, A1501 (member of gv1) and CMT.9.A, have been isolated from the rhizosphere of rice in China and from the roots of a cultivar of Sorghum nutans in Germany, respectively. P. stutzeri A1501 survives in soil, colonises the root surface, and is able to invade the superficial layers of the root cortex. It has been reported to be an endophyte [22, 14]. The genome of strain A1501 has been sequenced recently, and the chromosomal location of the nif genes was clearly demonstrated. A plasmid has been identified in strain CMT.9.A that may carry some genetic information for nitrogen fixation. The origin of the nif genes in P. stutzeri strains can be explained by a plausible lateral gene transfer acquisition. This hypothesis seems to be supported by nifH phylogenies, the diagnostic gene for nitrogen fixation, and by the genome analysis of strain A1501. Some characteristics of the A1501 genome studied in detail by Yan et al. [14] have been indicated in a previous section, and some relevant comments on the nitrogen fixing genes and the rhizosphere competence will be considered in this
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section. All the genetic information for diazotrophic growth is clustered in a 49-kb region (PST1302-PST1359) consisting of 59 genes. The G+C content of this region was higher (66.8%) than the average of the entire genome (63.8%). This led Yan et al. to consider the hypothesis of a nitrogen fixation island that was acquired by lateral gene transfer. This same hypothesis has been discussed for other nitrogen fixers. Another very interesting result of the genomic analysis of strain A1501 is the identification of genes that presumably allow for the adaptation to the rhizosphere. A cluster of genes for cellulose biosynthesis that may play a role in the attachment to the plant surface was identified. In addition, other gene candidates that may act as plant growth promoters were identified, as were those that may prevent ethylene formation, which is an inhibitor of root elongation [14]. In this respect, Puente and Bashan [23] already described the isolation of a P. stutzeri strain able to fix nitrogen from the interior of the desert epiphyte Tillandsia recurvata, which grows on electrical cables and giant columnar cacti in the semiarid zone of Baja California, México. The authors state that this is the first study to indicate the possible endophytic growth between nitrogen-fixing bacteria and a bromeliad. Iron in the soil exists mainly in the oxidised form, which forms extremely insoluble oxide hydrates, carbonate, or magnetite. Therefore, most aerobic bacteria synthesise and secrete Fe(III)-complexing organic compounds called siderophores. Three types of siderophores are produced: catechol-type, hydroxamate-type, and dicarboxylic or tricarboxylic acids. Zawadzka et al. [24] and Mulet et al. [5] studied the siderophore production by P. stutzeri strains. Amonabactins (catechol-type) and proferrioxamine-type siderophores were detected. Interestingly, strains belonging to the same gv belong to the same siderophore group, but different gv can share the same type of siderophore. In approximately half of the strains studied, no siderophores were detected. It seems that strain A1501 is unable to synthesise siderophores, which are important for chelating iron in the rhizosphere because this strain does not have genes for siderophore production in its genome. However, A1501 possesses genes for siderophore receptors. In this way, it should receive iron from the siderophores produced by other soil bacteria in the rhizosphere [14]. A wetland was constructed by Aguilar et al. [25] for tannery wastewater remediation. The microbial community that was associated with Typha sp. and Scirpus americanus and involved in the sulphur cycle was investigated. Sulphur oxidising bacteria were isolated on media containing thiosulphate. In addition to other genera, Pseudomonas strains were isolated that were able to grow organotrophically, but were also able to oxidise reduced sulphur compounds (such as elemental sulphur or thiosulphate), thereby accumulating thiosulphate or tetrathionate during growth. The 16S rRNA phylogenies placed the Pseudomonas close to P. stutzeri in the same phylogenetic branch. The properties of N2 -fixation and sulphur compound oxidation are relevant for the ecosystem and may be provided by P. stutzeri strains. This is a good example of how bacteria associated with plants can effectively collaborate for the bioremediation of polluted environments.
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6.5.2 Groundwater The main property of the groundwater ecosystem is the scarce amount of nutrients, and food webs are almost heterotrophic due to the lack of light reaching the primary producers. Levels of available oxygen for aerobic respiration are also usually low. In deep subsurface environments, life is primarily restricted to growth under anaerobic conditions [26]. The biomass production depends upon the input of organic matter from the surface. Unpolluted groundwater systems have scarce trophic resources, and most of the organic matter entering the groundwater habitats is mineralised by autochthonous microorganisms adapted to oligotrophic conditions. Many subsurface environments may be considered extreme from a nutrient perspective. Some are suspended in water, but most are associated with solid surfaces. Besides the allochthonous natural organic carbon, groundwater might be anthropogenically polluted by organic or inorganic contaminants. These situations are of major concern for assessing the quality of aquifers, and many options exist for the bioremediation of polluted groundwater. Bacteria adapted to this kind of ecosystem must be selected for “in situ” bioremediation technical processes. Examples of P. stutzeri strains involved in these processes will be considered in this chapter. The P. stutzeri strain KC (ATCC 55595, DSM 7136) is a denitrifying bacterium that was originally isolated from an aquifer in Seal Beach, CA, USA [27]. It is the reference and single strain of gv9 [28]. Zawazka et al. [24] studied the siderophore production by strain KC in detail. Under iron-limiting conditions, strain KC induces genes for the production and secretion of pyridine-2,6-bis(thiocarboxylate) (PDTC), a molecule that can rapidly dechlorinate carbon tetrachloride (CCl4 ), thereby yielding CO2 and non-volatile compounds under anoxic conditions [27, 29, 13, 30, 31, 32]. It is highly likely that PDTC is a secondary siderophore in this strain, in addition to ferrioxamines. This activity is important for bioremediation applications in aquifer sediments because it is rapid, with half-lives of only a few minutes [33], and it occurs without the accumulation of chloroform. Lewis et al. [13] reported that a laboratory culture of strain KC spontaneously lost a 170 kb fragment containing the genes necessary for PDTC biosynthesis on a 25 kb fragment of the lost DNA. This fragment was not detected in other P. stutzeri strains, and the 25 kb fragment has a GC content of 59.8%, which is below the inferior limit of the GC content of the chromosome in this species, which ranges between 60.7 and 66.3% GC. This data suggests an HGT event. The spontaneous loss of a 170 kb fragment is in accordance with the known chromosome plasticity of P. stutzeri strains and with the idea that the size of the chromosome must be maintained within some limits in a species [34]. Strain KC has significance for environmental biotechnology because of its use in one of the first full-scale aquifer bioaugmentation applications [35]. Large volumes of strain KC were grown on-site and injected into a CCl4 -contaminated aquifer in Schoolcraft, MI, USA. The cells colonised the aquifer sediment, creating a biocurtain that efficiently removed CCl4 from the groundwater passing through it for over three years.
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Strain CCUG36651 was studied and isolated by Johnsson et al. [36] from a depth of 626 m below ground at the Äspö Hard Rock Laboratory, where research concerning the geological disposal of nuclear water is performed. Strain CCUG36651 was ascribed to gv 3 of P. stutzeri in a recent study [5]. Siderophores not only capture Fe(III), but also other metals, including thorium(IV), plutonium(VI), and uranium(VI). The possibility of mobilising radionuclides by complexing compounds from bacteria is an important research area in the context of nuclear waste disposal. It is still unknown whether such compounds are produced in aquifers under conditions relevant to a disposal site, which would be approximately 500 m underground in granite rock [37–39]. Strain CCUG36651 produces siderophores of the ferrioxamines type in aerobic conditions (as the other members of gv3 that have been studied), but not anaerobically with nitrate. Ferrioxamines may explain the solubilisation of metals under aerobic conditions. However, P. stutzeri was also effective in anaerobically solubilising actinides into the aqueous phase. This property was ascribed to the production of biological ligands, but the exact reason is still unknown [37, 36]. Strain CCUG36651 was able to grow when incubated aerobically and anaerobically with Fe-III-citrate without nitrate (http://www.ccug.se/). A similar situation was found by Naganuma et al. [38] in a screening for Fe(III) reducers. Two bacterial strains (KNA-6-3 and KNA-6-5) were isolated from deep groundwater (160–200 m below ground level) from the Tono uranium mine in central Japan in order to assess possible microbial responses to redox variation during subsurface waste disposal. The closest species of both strains in the 16S rRNA sequences and the phenotypic characterisation was P. stutzeri. Both strains were able to reduce Fe(III) weakly, but persistently.
6.5.3 Marine Environment For a strain to be considered of marine origin, it must have the physiological characteristic of requiring, or at least tolerating, NaCl. There are not many true Pseudomonas species in marine habitats. Eleven Pseudomonas species have been suggested as possible inhabitants of marine environments, and four of them are novel species described since 2005. P. stutzeri has been considered a classical marine bacterium since the studies of ZoBell and Van Niel. In this section, we will consider isolates representative of the three main habitats: water column, marine sediment, and deep-sea. The free-living inhabitants of the water column are considered oligotrophs if they are not attached to detritus particles. Those organisms present in the sediment receive plenty of nutrients from the particles of the water column, and the species immediately below the surface are in anaerobic conditions. Organisms from deep-sea sediments are also exposed to extreme environments, such as the high pressure or, in some habitats, also extreme temperatures. Strain ZoBell (ATCC 14405) was isolated from the water column in the Pacific Ocean. For many years, it was considered to be a typical representative of marine bacteria, and it was identified as Pseudomonas perfectomarina [40]. In the actual
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taxonomy, it is considered a member of gv2 [41]. Antibodies against this strain have been used for the detection of P. stutzeri by immunofluorescence in sea water and intertidal sediment environments [42]. Considered to be the archetypal denitrifier, it has been studied extensively by W. Zumft and his research group in Karlsruhe (Germany). The genetics and biochemistry of the denitrification by P. stutzeri has been reviewed recently [2, 43]. In a screening of marine bacteria from highly contaminated Mediterranean sediment, several bacteria that were adapted to the mineralisation of naphthalene were isolated by García-Valdés et al. [44]. Two strains were classified as P. stutzeri in gv3: AN10 and AN11 [45]. These strains were further studied by their chromosomallyencoded metabolic pathway for naphthalene and salicylate degradation, which was considered to be an unusual characteristic of an otherwise plasmid-coded trait [46, 47]. The entire pathway of strain AN10 was cloned and sequenced [17, 18]. It is organised into four operons: one coding for the enzymes involved in the conversion of naphthalene to salicylate, two coding for the conversion of salicylate to pyruvate and acetyl-coenzyme A, and another operon with the regulatory gene. The presence of two different salicylate hydroxylases coded by nahG and nahW is of special significance [48]. Each enzyme presents activity against salicylate and its methyl and chloro derivatives, but the relative transformation rates differ. NahW transforms 3-chlorosalicylate more efficiently, while NahG is more efficient in transforming methylsalicylates. NahG is specific for P. stutzeri. Naphthalene degradation is not a general property of P. stutzeri. It is only found in some specialised strains isolated from contaminated sites. The genes are considered as accessories in the species, but they are relevant for the adaptation to the contaminated sediment. These strains belong to different gv, and the sequence analysis indicates that the degradative genes have been acquired by members of the species by lateral gene transfer after the differentiation of the species in its genomovars. However, the key enzymes in the naphthalene pathway (nahA, nahG, and nahH) of P. stutzeri strains are phylogenetically closer between each other compared to the enzymes from other Pseudomonas strains, suggesting a preferential transfer between members of the same species. This is an additional argument supporting the existence of a barrier for the interspecific exchange of genes, as mentioned above. Two other marine strains have been isolated from the deep ocean and studied for their adaptation to this extreme habitat. Strains MT-1 and HTA-208 were isolated from deep-sea samples at a 10,897 m depth at the Mariana trench, the deepest habitat studied so far. MT-1 has been studied extensively to decipher the bacterial adaptation to high pressure. Strain NF13 was isolated from a sediment sample by Janasch and collaborators [49, 50] in the Galapagos rift near a hydrothermal vent at a depth of 2,500–2,600 m. A relevant physiological characteristic of strain NF13 is its ability to chemolithotrophically oxidise sulphur compounds. Sulphur metabolism is crucial for the elements cycling in hydrothermal vent habitats. H2 S is assumed to be the predominant energy source in this ecosystem. Besides this capacity, strain NF13 is also able to grow diazotrophically. Nitrogen is usually a nutrient limiting factor in the ocean and confers to NF13 another relevant ecological advantage.
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Several strains were isolated by Sorokin et al. [51] from Black Sea samples taken at a depth of more than 100 m. These strains have the ability to oxidise thiosulphate to tetrathionate both aerobically and anaerobically with nitrate or nitrite. The strains belonged to gv3, gv4, and gv5. This property is not a general property of P. stutzeri and must be considered as an adaptation to the habitat. In order to understand the behaviour of iodine in the environment to accurately perform safety assessments on 129 I, Amachi et al. [52] isolated the SCT strain from marine sediment slurries. This bacterium is able to grow anaerobically with iodate. The predominant forms of iodine in the environment are iodate and iodide. Iodate is more thermodynamically stable, but significant quantities of iodide are observed in surface waters. The high amounts of iodide in seawater are assumed to be the result of the biological reduction of iodate by marine microorganisms and phytoplankton. There is a limited number of isolates that are able to reduce iodate. Strain SCT was classified as a member of P. stutzeri with a dissimilatory iodate-reducing capability, and its iodate reductase is induced by anaerobic growth conditions. Other authors [53] reported that nitrate reductases are also able to reduce iodate and proposed that iodate in sea water is reduced by the nitrate reductase of marine organisms. However, Amachi et al. [52] concluded that the nitrate reductase of SCT does not play an important role in the iodate reductase activity that was detected, and that the dissimilatory iodate-reducing bacteria present in nature play a relevant role in iodate transformation. Another habitat occupied by P. stutzeri in the ocean is represented by the other marine macro or microorganisms. Symbiosis (in the original sense, “living together”) with marine organisms has been described for strain CCUG46542 [6], the only member of gv19 [5], with an ascidian (Didemnum sp.). Another strain survives symbiotically with the dinoflagellate Alexandrium lusitanucum [54]. P. stutzeri is an intracellular endosymbiont of A. lusitanicum, which is known to produce saxitoxin, the etiological agent of paralytic shellfish poisoning. The presence of P. stutzeri enhances the synthesis of the toxin, but the mechanisms involved are still being studied. Mineralisation of unusual carbon compounds, denitrification, diazotrophy and metabolism of inorganic sulphur compounds seem to be the main ecological roles of the P. stutzeri strains in the ocean. It must be mentioned that no strains have been isolated that are able to grow autotrophically.
6.5.4 Clinical Samples P. aeruginosa is the only true human pathogen described in the genus. Several pathogenicity determinants have been studied in this species, and these should also be present in other potentially pathogenic Pseudomonas species. In the genomic study of strain A1501, Yen et al. [14] emphasised the absence of the genes related to virulence and pathogenicity in P. aeruginosa, such as type III/VI secretion systems, the synthesis of both types of quorum sensing molecules, alginate polymer synthesis, siderophores, and antibiotic biosynthesis pathways.
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P. stutzeri is frequently isolated from clinical samples. In a study on the prevalence of Pseudomonas in hospitals, 1–3% of the Pseudomonas isolated from wound pus, blood, urine, tracheal aspirates, and sputum were P. stutzeri. A summary of infections and antibiotic susceptibilities is provided in [2]. In humans, P. stutzeri is generally considered a contaminant, and in rare occasions appears as the only organism isolated from clinical materials. Most infections are iatrogenic and associated with the administration of contaminated solutions (dialysates, green soap, intravenous fluids, and povidone-iodine), medications, blood products, or with the presence of indwelling catheters in compromised patients (Clinical Microbiology Proficiency Testing, may 1997). For instance, P. stutzeri has a propensity to be associated with renal dialysis units and has caused bacteraemia in haemodialysis patients receiving contaminated dialysis solutions [55]. It seems that this species becomes an opportunistic pathogen when the host’s defence mechanisms are weakened. Infections by P. stutzeri respond effectively to treatment with antibiotics, including aminoglycosides, antipseudomonal penicillins, trimethoprim-sulfamethoxazole, and the third-generation cephalosporins [56]. P. stutzeri is ubiquitous in hospital environments and is an opportunistic but rare pathogen. However, due to its genomic plasticity and capacity to capture genes from the environment, it must be considered of relevance as a possible reservoir of antibiotic resistance genes. Yan et al. [57] described the emergence of IMP- and VIM-type metallo-β-lactamases (MBLs) in Pseudomonas sp. isolated in Taiwan. Carbapenems are antibiotics of last resort against Gram negative bacteria resistant to other β-lactam antibiotics. Three P. stutzeri isolates were studied and found to carry the chromosomally located genes blaIMP-1 , while one isolate carried blaVIM-2 . They concluded that the environmental isolates acquired the MBL genes under long-term selection pressure in the hospital environment.
6.5.5 Genomovar, Habitat, and Niche-Adaptation At present, more than 200 strains have been assigned to gv, but unfortunately most gv are represented only by few isolates. Most isolates are assigned to gv1 (92 strains), gv2 (11 strains), gv3 (53 strains), gv4 (5 strains), gv5 (6 strains), gv7 (8 strains), gv15 (8 strains), and gv19 (7 strains). The others are represented with only one, two, or three isolates each. It is difficult to find a correlation between the habitat from which the strain was isolated and the gv to which it was ascribed. However, we can generalise that most clinical isolates (88%) belong to gv1 and that most gv3 strains (98%) are of environmental origin (from soil, marshes, water, or intertidal sand), independent from where the sample was isolated. The isolates from marine or shallow waters that are adapted to saline habitats are not found in gv1. P. stutzeri has an almost universal distribution and can be isolated from many habitats. When the gv distribution of strains was studied in the same locality, strains could be assigned to various gv, indicating that the various gv can share the same habitat [58]. However, one gv seems to predominate, perhaps because it is the best adapted to the environment.
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It might also be speculated that the P. stutzeri ecotypes, specialised in characteristic ecological niches, are taxonomically distributed in specific gv. This is not always the fact. The physiological or biochemical adaptation to the use of a contaminant, for instance naphthalene, is distributed among 4 gv. They are distributed in gv1, gv3, gv4, and gv7. Another contaminant is chlorate. The ability to dechlorinate is only present in a few strains of the species, which belong to gv1, gv3, and gv5 [59]. In one strain (PK), the genomic organisation of the chlorite dismutase gene (cld) and the chlorate reductase operon are in close proximity to a transposase gene, suggesting the horizontal gene transfer of this peculiar characteristic [60]. These characteristics must be considered the result of the acquisition of accessory genes by the species. The presence of strains in many polluted habitats is a further indication of how it can evolve to exploit the new situations and metabolise the contaminant in many ways. This adaptation is a characteristic of the species and not of the gv.
6.6 Speciation: Gain and Loss of Genes – Future Outlook In the evolution of eukaryotes, a clear advantage over prokaryotes has been argued due to sexual reproduction, the concomitant recombination of genes, and diploidy. In the present chapter, we have demonstrated that both aspects are circumvented by bacteria, which attain similar evolutionary advantages. Gene rearrangements and genetic exchange are relevant in the evolution of P. stutzeri, and are accomplished by the natural transformation ability of many strains. Mobile genetic elements also contribute significantly to this chromosomal flexibility. Gene duplications provide two evolutionary advantages for the strains harbouring them: the double gene dose ensures a possible duplication of the enzyme molecules, and one gene can evolve and assume a novel function without losing the gene activity of the precursor gene. The salicylate hydroxylases of P. stutzeri are good examples of how prokaryotes can duplicate genes and have both evolve separately, thereby acquiring novel functions. At the same time, the chromosome size cannot grow indefinitely. Therefore, genetic mechanisms exist that limit the total number of genes. An example is the PDTC gene that has been mentioned. Another example was studied experimentally when strain AN10 was adapted in chemostat cultures to high concentrations of 4-chlorosalicylate. During the adaptation, the upper naphthalene pathway, which leads to salicylate, was lost in the adapted AN142 strain, which suffered chromosomal rearrangements [19]. These mechanisms drive the differentiation of the species in genomovars, and can be studied experimentally. We must consider the genomovar as an intermediate state in the speciation process. In this sense, it constitutes one of the better examples for studying the evolution of a bacterial species. The development of novel culture-independent methods to specifically study Pseudomonas populations, and P. stutzeri in particular, as well as the metagenomic studies undertaken in many different habitats, will help in improve our understanding of the ecology of the species. On the other hand, several genomes of P. stutzeri strains isolated from different habitats are currently sequenced, and the comparative genomic analyses will decipher the evolutionary history of this species.
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20. Coleman, N.V. and Holmes, A.J. (2005) The native Pseudomonas stutzeri strain Q chromosomal integron can capture and express cassette associated genes. Microbiology 151: 1853–1864. 21. Holmes, A.J., Gillings, M.R., Nield, B.C., Mabbutt, B.C., Nevalainen, K.M.H. and Stokes, H.W. (2003) The gene cassette metagenome is a basic resource for bacterial genome evolution. Environ. Microbiol. 5: 383–394. 22. Rediers, H., Bonnecarrere, V., Rainey, P.B., Hamonts, K., Vanderleyden, J. and De Mot, R. (2003) Development and application of a dapB-based in vivo expression technology system to study colonization of rice by the endophytic nitrogen-fixing bacterium Pseudomonas stutzeri A15. Appl. Environ. Microbiol. 69: 6864–6874. 23. Puente, M.E. and Bashan, Y. (1994) The desert epiphyte Tillandsia recurvata harbours the nitrogen-fixing bacterium Pseudomonas stutzeri. Can. J. Bot. 72: 406–408. 24. Zawadzka, A.M., Vandecasteele, F.P.J., Crawford, R.L. and Paszczynski, A. (2006) Identification of siderophores of Pseudomonas stutzeri. Can. J. Microbiol. 52: 1164–1176. 25. Aguilar, J.R.P., Cabriales, J.J.P. and Vega, M.M. (2008) Identification and characterization of sulfur-oxidizing bacteria in an artificial wetland that treats wastewater from a Tannery. Int. J. Phytoremediation 10: 359–370. 26. Pedersen, K. (1997) Microbial life in deep granitic rock. FEMS Microbiol. Rev. 20: 399–414. 27. Criddle, C.S., DeWitt, J.T., Grbic-Galic, D. and McCarty, P.L. (1990) Transformation of carbon tetrachloride by Pseudomonas sp. strain KC under denitrification conditions. Appl. Environ. Microbiol. 56: 3240–3246. 28. Sepúlveda-Torres, L.C., Zhou, J.Z., Guasp, C., Lalucat, J., Knaebel, D., Plank, J.L. and Criddle, C.S. (2001) Pseudomonas sp strain KC represents a new genomovar within Pseudomonas stutzeri. Int. J. Syst. Evol. Microbiol. 51: 2013–2019. 29. Dybas, M.J., Tatara, G.M. and Criddle, C.S. (1995) Localization and characterization of the carbon-tetrachloride transformation activity of Pseudomonas sp. strain KC. Appl. Environ. Microbiol. 61: 758–762. 30. Lee, C.H., Lewis, T.A., Paszczynski, A. and Crawford, R.L. (1999) Identification of an extracellular catalyst of carbon tetrachloride dehalogenation from Pseudomonas stutzeri strain KC as pyridine-2,6-bis(thiocarboxylate). Biochem. Biophys. Res. Commun. 261: 562–566. 31. Lewis, T.A. and Crawford, R.L. (1993) Physiological factors affecting carbon tetrachloride dehalogenation by the denitrifying bacterium Pseudomonas sp. strain KC. Appl. Environ. Microbiol. 59: 1635–1641. 32. Sepúlveda-Torres, L.C., Rajendran, N., Dybas, M.J. and Criddle, C.S. (1999) Generation and initial characterization of Pseudomonas stutzeri KC mutants with impaired ability to degrade carbon tetrachloride. Arch. Microbiol. 171: 424–429. 33. Tatara, G.M., Dybas, M.J. and Criddle, C.S. (1993) Effects of medium and trace metals on kinetics of carbon tetrachloride transformation by Pseudomonas sp. strain KC. Appl. Environ. Microbiol. 59: 2126–2131. 34. Mira, A., Ochman, H. and Moran, N.A. (2001) Deletional bias and the evolution of bacterial genomes. Trends Genet. 17: 589–596. 35. Hyndman, D.W., Dybas, M.J., Forney, L., et al. (2001) Hydraulic characterization and design of a full scale biocurtain. Ground Water 38: 462–474. 36. Johnsson, A., Arlinger, J., Ödegaard-Jensen, A., Albinsson, Y. and Pedersen, K. (2006) Solid-phase partitioning of radionucleides by complexing compounds excreted by subsurface bacteria. Geomicrobiol. J. 23: 621–630. 37. Essén, S.A., Johnsson, A., Bylund, D., Pedersen, K. and Lundström, U.S. (2007) Siderophore production by Pseudomonas stutzeri under aerobic and anaerobic conditions. Appl. Environ. Microbiol. 73: 5857–5864. 38. Naganuma, T., Sato, M., Hoshii, D., Amano-Murakami, Y., Iwatsuki, T. and Mandernack, K. (2006) Isolation and characterization of Pseudomonas strains capable of Fe(III) reduction with reference to redox response regulator genes. Geomicrobiol. J. 23: 145–155.
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Part III
Physiology, Metabolism and Markers
Chapter 7
Pyoverdine Siderophores as Taxonomic and Phylogenic Markers Jean-Marie Meyer
7.1 Introduction Classically, but not officially, Pseudomonas species are divided into two sub-groups according to their ability to produce the yellow–green, brightly fluorescent pigment named pyoverdine [1]. This pigment is easily detectable when growing the bacteria on King’s B medium [2]. The so-called fluorescent Pseudomonas and their counterpart, the non-fluorescent Pseudomonas, contain many species, i.e., 64 and 53, respectively, according to the present work and to a recent compilation [3]. These numbers, by their size, well illustrate the great diversity encountered within the Pseudomonas genus. Although some of them are able to grow in anaerobic conditions in the presence of nitrates (anaerobic respiration), Pseudomonas are characterized by a strict oxidative metabolism and, therefore, display a high iron requirement for satisfying cytochromes and other electron transport system biosynthesis. Thus, it is not surprising that many siderophores, i.e., bacterial iron transporters allowing the bacteria to get access to iron, have been so far described in fluorescent Pseudomonas [4–6], among them the previously cited pigment, pyoverdine, which has been the first compound described as acting as a siderophore in Pseudomonas [7, 8]. Surprisingly, while no marked phenotypic differences could differentiate fluorescent Pseudomonas one from each other at the level of their pyoverdine production, an intensive study on more than 3,000 bacterial isolates analyzed through the so-called siderotyping methods (see below), allowed us to recognize the existence of more than 110 different pyoverdines. This huge molecular diversity among compounds having all the same aspect in colour and fluorescence and an identical biological function, is correlated in the present chapter with the taxonomic diversity of their producing bacteria with, as the major conclusion, that pyoverdine is a very powerful taxonomic marker. The characterization through siderotyping of the pyoverdine J.-M. Meyer (B) Département Génétique moléculaire, Génomique et Microbiologie, UMR 7156 CNRS-Université de, Strasbourg, France e-mail:
[email protected]
J.L. Ramos, A. Filloux (eds.), Pseudomonas, DOI 10.1007/978-90-481-3909-5_7, C Springer Science+Business Media B.V. 2010
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produced by a given strain usually results in the identification of the strain at the species level, without the need of any other taxonomical method. Furthermore, when available, structure comparison of pyoverdines could result in the identification of groups of structurally closely related pyoverdines, which producer strains, as far as they have been subjected to some phylogenic studies, correspond to highly phylogenetically related strains. Thus, pyoverdines, as taxonomic as well as phylogenic markers, are molecules of great importance for the characterization and for a better knowledge of the fluorescent Pseudomonas.
7.2 Taxonomic Diversity Among Pseudomonas Pseudomonas are well distributed in nature: omnipresent in soils and waters [half of the natural isolates in mineral water is constituted by pseudomonads [9]], they are one of the major bacterial population under study in fundamental research as well as in more applied domains. Interests are multiple since some are known as human or animal pathogens (e.g., Pseudomonas aeruginosa, Pseudomonas entomophila, Pseudomonas otitidis, Pseudomonas plecoglossicida), others display pathogenicity towards plants or mushrooms (e.g., Pseudomonas syringae, Pseudomonas tolaasii, Pseudomonas salomonii). Moreover, among the numerous other Pseudomonas which are considered as innocuous, many of them, thanks to the great metabolic versatility which characterize these bacteria, have been selected for their biotechnological values as biocontrol or bioremediation agents, e.g., the well studied strains Pseudomonas putida KT2440 [10], Pseudomonas fluorescens CHA0 [11], Pf-5 [12], SBW25 [13], WCS374 [14] and others. Indeed, it is of primarily importance to well characterize such strains and first, to well define their respective taxonomic position. The question, surprisingly, still remains open for the strains cited above, since taxonomists are presently considering the two concerned species, P. fluorescens and P. putida, as not so well defined species, but rather, as complexes of species [15–17]. A strong heterogeneity among them was seen as soon as they were analyzed in depth, with already the recognition of several biovars [18–20]. Other species, e.g., Pseudomonas syringae or Pseudomonas stutzeri, have been split in several so-called genomovars, following studies based on genomic comparisons [21–23]. These few examples demonstrate the still particular difficulty in determining nowadays species affiliation among pseudomonads, despite all the sophisticated genomic methods, which have been developed these recent years. The most accurate and presently requested way for Pseudomonas species affiliation is to consider the so-called polyphasic taxonomical method [24], which consists in integrating the maximum of taxonomic tools for a precise characterization of a pseudomonad at the species level. None of the conventional phenotypic or even genotypic methods presently in use being self-sufficient for an accurate species characterization, the accumulation of data resulting from as many methods as possible, should conduct to a better species definition. The way could be considered as particularly successful since about two third of the 60 species described in the pres ent chapter and listed in Table 7.1, are resulting from taxonomical studies developed
Type-strain or sv. reference strain
ATCC 10145T CFBP 2063T DSM 15318T CFBP 3279T NCIMB 10068T DSM 6698T CFBP 11144T CFBP 3280T CFBP 11706T CFML 96-344T CFBP 2341T CFML 96-198T ATCC 9446T CFBP 2101T DSM 50332T DSM 14939T CFBP 5705T DSM 18900T L48 CFBP 6729T CFBP 3224T ATCC 13525T CFBP 2065T CIP 105469T CIP 106645T CIP 105274T DSM 13647T
Species name
P. aeruginosa P. agarici P. antarctica P. asplenii P. aurantiaca P. aureofaciens P. avellanae P. blatchfordae P. brassicacearum P. brenneri P. cannabina P. cedrina P. chlororaphis P. cichorii P. citronellolis P. congelans P. costantinii P. delhiensis P. “entomophila” P. extremorientalis P. ficuresectae P. fluorescensc P. fuscovaginae P. gessardii P. grimontii P. jessenii P. kilonensis
89 1 1 1 2 4 1 1 10 12 1 9 3 9 3 2 10 1 1 1 1 9 16 13 34 8 12
No. of strains analyzed see Table 7.2 aga 13525 asp 13525 see Table 7.2 syr 96-192 PL9 PflW syr see Table 7.2 13525 see Table 7.2 see Table 7.2 syr cos del ent 95-275 syr 13525 fus PflW see Table 7.2 9AW A6
Siderovar name see Table 7.2 Unknown 46 Unknown 46 see Table 7.2 31 Unknown 36 42 31 see Table 7.2 46 see Table 7.2 see Table 7.2 31 38 Unknown Unknown 60 31 46 Unknown 42 see Table 7.2 55 28
PVD type No. (from Table 7.4)
[34, 18] [26, 69, 74] [75, 69, 76] [69, 77] [78, 69, 65] [69, 65] [79, 80, 69] [69] [56, 52, 81] [82, 83, 69] [79, 80, 52] [84, 69] [85, 78, 65] [26, 69] [49, 69] [86, 69] [87, 59] [69, 46] [69, 43, 44] [88, 69] [79, 80] [75, 69, 18] [52, 42, 89] [83, 69, 90] [91, 69] [92, 52, 93, 94] [95, 69, 62]
References
Pyoverdine Siderophores as Taxonomic and Phylogenic Markers
1 1 1 3 1 1
3 1 2 2 1 1
1 1
1 2
4
No. of siderovarsb
Table 7.1 Pseudomonas speciesa analyzed through siderotyping
7 203
P. palleroniana P. panacis P. “pavonanceae” P. poae P. “protegens” P. proteolytica P. putidad P. reinekei P. rhodesiae P. salomonii P. savastanoi P. simiae
P. koreensis P. libanensis P. lini P. lurida P. marginalis P. mandelii P. meliae P. meridiana P. migulae P. monteilii P. moraviensis P. mosselii P. orientalis P. otitidis
Species name 1 6 9 3 8 43 1 1 10 10 1 11 6 11
KACC 10848T CFML 96-195T CFBP 5737T DSM 15835T CFBP 2038T CIP 105273T CFBP 3225T DSM 15319T CFML 95-321T CFML 90-54 CCM 7280T CIP 105259T CIP 105540T ATCC BAA-1130T CFBP 4389T KCTC 12330T KACC10796 P529/17 CHA0 DSM 15321T ATCC 12633T DSM 18361T CIP 104664T CFBP 2022T CFBP 1670T CCUG 50988T 11 1 1 3 10 1 7 1 7 107 1 4
No. of strains analyzed
Type-strain or sv. reference strain
1
1 1
4
1 1
1
1 1 1
1 1
2 2 2 3 1
No. of siderovarsb
Table 7.1 (continued)
13525 Pfl12 B10 13525 CHA0 Pflii see Table 7.2 SB8.3 rho 96-318 syr G84
SB8.3 see Table 7.2 see Table 7.2 see Table 7.2 see Table 7.2 SB8.3 syr 51 W 9AW 90-54 B10 mos 13525 oti
Siderovar name
46 49 29 46 15 5 see Table 7.2 7 52 58 31 Unknown
7 see Table 7.2 see Table 7.2 see Table 7.2 see Table 7.2 7 31 6 55 16 29 Unknown 46 Unknown
PVD type No. (from Table 7.4)
[21, 78, 52] [101, 69, 102] [69, 37] [86, 78, 69] [45, 103] [69, 104, 76] [42, 18] [105, 47, 69] [4, 106, 52, 38] [21, 52, 107] [79, 69] [69, 24]
[92, 96, 69] [97, 69] [57, 69, 52] [60, 69] [69] [105, 69, 52, 94] [26, 79, 69] [69, 76, 93, 98] [92, 69, 90] [99, 52, 38] [69, 37, 77] [58, 52, 42] [84, 69, 42] [100, 69]
References
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60e 6 17 7 1 8 6
ATCC 19310T
CFBP 11261T CFBP 2068T DSM 14937T KACC 10847T CIP 104663T CFBP 2107T
P. syringae
P. thivervalensis P. tolaasii P. trivialis P. umsongensis P. veronii P. viridiflava 1 1
1 1 1
1
No. of siderovarsb
thi tol 13525 SB8.3 13525 Syr
syr
Siderovar name
Unknown 53 46 7 46 31
31
PVD type No. (from Table 7.4)
[54, 108, 79, 80, 52] [56, 52, 38] [83, 52, 109] [86, 78, 69] [105, 96, 69] [110, 78, 52] [54, 79, 80]
References
P. amygdali, P. caricapapaya, P. flectens and P. tremae did not grow on CAA- or Succinate medium. Therefore, they could not be included in the siderotyping studies. Two species, P. delhiensis and P. reinekei, described in the literature as non-fluorescent Pseudomonas, did produce large amounts of pyoverdine when grown in CAA medium and, therefore, were included in the study. On the contrary, P. nitroreducens and P. plecoglossicida were seen to be devoid of any pyoverdine system and, therefore, were excluded from the study. b see Table 7.2 for details concerning species with several siderovars. c restricted to strains having identical siderotyping features as the type-strain P. fluorescens ATCC 13525 (see [52]). d restricted to strains belonging to the P. putida sensu stricto group (see [42]). e each strain representing a different pathovar.
a
No. of strains analyzed
Type-strain or sv. reference strain
Species name
Table 7.1 (continued)
7 Pyoverdine Siderophores as Taxonomic and Phylogenic Markers 205
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since 1996 and based on the polyphasic approach. It is not our goal in the present chapter to review the many phenotypic and genomic methods required for an accurate Pseudomonas species definition (see [3, 18, 19, 25] for recent compilations). Altogether, these methods represent, however, a very heavy investment in time and money. It is not surprising, therefore, that the taxonomical aspect in many recent analyses of natural Pseudomonas populations is dramatically failing, and that an in depth taxonomical analysis of the complexes of species like the Pseudomonas fluorescens and Pseudomonas putida groups, still remains to be done, although their heterogeneity is recognized since more than fourty years. There is no doubt that future studies on Pseudomonas will comfort this already huge heterogeneity which characterizes the genus, and that new tools allowing a better discrimination between species will be necessary to clarify the taxonomical position of these bacteria. Phylogenetic relationships between Pseudomonas species have been considerably improved with the development of DNA-sequence comparisons of highly conserved genes. First studies, based on 16S-RNA (rrs)-genes [26–28] resulted in a very efficient “cleaning” with the exclusion of more than 60 species, the genus becoming limited since that time to the Pseudomonas group I of Palleroni’s classification [29]. However, the efficiency of the method for species delineation, although still in use in many publications, is clearly disappointing. rrs genes appear to be too much conserved among Pseudomonas species, with very often similarity values close to 100% between strains clearly belonging to different species. Consequently, the same approach but with the use of many other less-conserved housekeeping genes studied in parallel (e.g., atpD, carA, gyrB, rpoB, rpoD) has been recently developed, resulting in a better species definition [23, 30–33]. It remains, however, that these sophisticated approaches are time-consuming, costly, and require heavy computer and software equipment not always available to every one facing a taxonomic identification problem within pseudomonads. The siderotyping procedure, based on very rapid, easy to perform, and not expensive analytical methods, allowing the characterization of the pyoverdines produced by fluorescent Pseudomonas, should be considered as a powerful substitute, as demonstrated in the following sections.
7.3 Pyoverdine Diversity Among Fluorescent Pseudomonas 7.3.1 The Pyoverdine Molecule As said above, fluorescent Pseudomonas are easily characterized by the production of pyoverdine. However, this production is not a constant feature and is highly related to a bacterial growth in iron-deficient conditions [7]. King’s B medium, which has been empirically designed [2], favours the production of pyoverdine since it contains as the main nutrient a peptone (DIFCO peptone no. 3) which is particularly poor in iron. Other iron-poor media have been since preferentially used for pyoverdine production, e.g., the fully synthetic succinate medium [7] or the semi-synthetic CAA medium, based on a Casamino-acid preparation first
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manufactory-treated for a low iron and chloride content (see [34] for details related to growth conditions and pyoverdine detection and purification). The fact that iron deficiency is involved for pyoverdine production and also the property for the molecule to strongly bind iron (III), were strong indicators for a siderophoric function of the molecule [8]. Thirty years later, it can be said that pyoverdine is one of the most sophisticated siderophore molecule presently identified within the microbial world. Its structure responds to a general scheme involving a chromophore based on a quinolein residue to which are attached a peptide chain as well as a dicarboxylic acid or its amide form (Fig. 7.1). While the peptide chain and the dicarboxylic acid residue are subjected to many variations, the chromophore is highly conserved and defined commonly as (1S)-5-amino-2,3dihydro-8,9-dihydroxy-1H-pyrimido-[1,2-a]quinoline-1-carboxylic acid [35]. The only variation, which has been observed in a few compounds named for that reason “isopyoverdines” [36] concerns the position of the free carboxyl group, which occurs usually on the C1 of the chromophore, while it is on the C3 in isopyoverdines. The free NH2 group of the chromophore allows the attachment of a dicarboxylic acid chain, usually succinic acid, malic acid or their amide forms as the most frequent ones, but also fumaric acid or α-ketoglutaric acid in some pyoverdines [4].
α-thr
gly O
HO
H N
HN
D-ser
O
O OH
HN
D-thr NH
HO
N
NH
OH
O
O
gln
D-AcOHOrn
O
O N
HO
HN
+ NH O
H2N
O
HO
O
chromophore
NH
gly O
peptide part
O HN
NH
HO
O
OH
NH2
N H
O
dicarboxylic acid chain
N
D-cOHOrn
D-ser Fig. 7.1 The succinamide form of the pyoverdine of Pseudomonas costantinii [87], illustrating the three parts of a pyoverdine molecule and the presence of unusual amino acids in its peptide part. See footnotes of Table 7.4 for the abbreviations used for the amino acid residues
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It is important to remember that several “isoforms” (usually from 2 to 4) of almost any pyoverdine can be isolated from a culture medium, isoforms which precisely vary one from each others by the nature of the dicarboxylic acid side chain. Such plurality is still not very well explained, resulting in part from discrete variations during the biosynthetic pathway, but also resulting from hydrolyses of the amide forms occurring during the bacterial culture and due mainly to pH variations during growth. Attached to the free carboxyl group of the chromophore on C1 (or C3 ), is the peptide chain. It forms the most important part of the pyoverdine molecule because of its participation, together with the chromophore, to the iron (III) complexation but also and mainly, because it demonstrates a great strain-dependent variability. Since the first pyoverdine structure elucidated close to thirty years ago [37], a total of 63 structurally different pyoverdines have been described so far, each presenting a particular peptide chain varying by the type and number (6–14) of aminoacyl residues [38]. Most of the common amino acids are represented with, however, some like proline, leucine, isoleucine, tryptophan, cystine, cysteine, methionine, which are never found, while valine and histidine have been detected only once or twice. Uncommon amino acids like the D-forms of any usual amino acid, diaminobutanoic acid (DAB), threo-β-hydroxy-aspartic acid, threo-β-hydroxy-histidine, Land D-ornithine, are present in several pyoverdines. Most important, L- and D-δN-hydroxy-ornithine residues, very often located in the middle of the peptide chain as well as at the end, participate to the iron complexation. To be able to do that, they are acylated on the hydroxylated-N group to form a hydroxamate function, most commonly by formylation or acetylation. For the residue located at the end of the peptide, the hydroxamate function is formed by an internal cyclisation of the residue to form 3-amino-1-hydroxy-piperidone-2. In some pyoverdines, the hydroxamate functions delivered by hydroxy-ornithine residues can be partially or totally replaced by hydroxycarboxyl groups, then furnished in replacement by one or two hydroxy-aspartic residues. The rrs genes whatever the case, the third group necessary to fully complex iron (III) is always given by the catecholate function present on the chromophore. Such mixed iron-complexes, with two hydroxamates and one catecholate as iron ligands, are already considered as very strong, with an absolute affinity constant of 1032 [7]. Thus, they are located below the pure tri-catecholates (1052 for the enterobactin of Escherichia coli) but are stronger in iron complexation than the entirely hydroxamate fungal siderophores (1022 –1024 ). Structure determination of pyoverdines is not an easy task and necessitates an important investment in time and in sophisticated equipments. Thus, to overcome redundancies among structural studies, analytical as well as biological methods of pyoverdine differentiation were developed under the generic name of “siderotyping” for “siderophore typing” (see [39], for a detailed description of the different methods of siderotyping). Among them, mostly for their rapidity and simplicity in analyzing many pyoverdines or many producing strains in one round, isoelectrophoresis and pyoverdine-mediated iron uptake were the two most used ones, and largely contributed to the development of efficient siderotyping of the fluorescent Pseudomonas.
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7.3.2 Isoelectrophoresis as a Powerful Siderotyping Method The method of choice for a rapid analysis of the pyoverdine molecules excreted by a given fluorescent Pseudomonas is by isoelectrophoresis. The method used for the discrimination of proteins has been first applied to pyoverdines by Koedam et al. [40]. A convenient device is the mini-isoelectrofocusing (IEF) gel apparatus from BioRad. Using two such devices, 140 bacterial strains can be analyzed for their pyoverdine production within one day. 5% polyacrylamide gels (10 × 6.5 cm; 0.4 mm thickness), containing commercially available ampholins (the larger pH 3–10 range is the most appropriate for pyoverdines) are freshly casted within 1 h, charged with 10–20 pyoverdine-containing samples and electrophoresed during 1.5 h, following the manufacturer’s recommendations (15 min at 100 V, 15 min at 200 V and 1 h at 450 V). Samples consist usually of 1 μL of a 20-fold concentrated (through lyophilization) CAA-culture supernatant, or 1 μL of an aqueous 5 mM purified pyoverdine solution. The detection of the fluorescent pyoverdine bands is obtained through exposure to UV light (350 nm). Figure 7.2 shows that usually two to three major fluorescent bands are visualised for each sample, corresponding to the different pyoverdine-isoforms present in the supernatant at the time of harvest. The values of the isoelectric pHs (pHi) can be estimated by comparison with pyoverdine bands of known pHi values, as illustrated by the lane 1 deposit in Fig. 7.2 (see [39] for a complete description of the pHi pyoverdine markers). Indeed, strains producing identical pyoverdines result usually in identical IEF-patterns, while strains with structurally different pyoverdines present different patterns. We will see below, however, that some exceptions could be encountered and, thus, that it is advised to comfort the siderotyping data obtained through isoelectrophoresis by the use of another method.
Fig. 7.2 Pyoverdine-isoelectrofocusing patterns of 8 strains of the Pseudomonas chlororaphis complex: Pseudomonas chlororaphis subsp. chlororaphis ATCC 9446T (lane 2), Pseudomonas chlororaphis subsp. aurantiaca ATCC 33663T (lane 3), Pseudomonas chlororaphis subsp. aureofaciens DSM 6698T (lane 4), Pseudomonas aurantiaca NCIMB 10066 (lane 5), Pseudomonas aureofaciens DSM 50082 (lane 6), Pseudomonas chlororaphis DSM 50135 (lane 7), Pseudomonas chlororaphis DSM 50136 (lane 8), Pseudomonas aureofaciens DSM 50139 (lane 9). Lane 1 represents pHi markers as defined in [39]
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7.3.3 Pyoverdine-Mediated Iron Uptake as a Complementary Powerful Siderotyping Method Pyoverdines could be very well differentiated by iron uptake studies done on ironstarved cells tested towards pyoverdine-59 Fe (III) complexes. The iron transport systems developed in Pseudomonas involve usually very specific membrane receptors [41]. Thus, in case of a positive transport with a pyoverdine of foreign origin, it is very likely that the foreign pyoverdine is identical to the homologous pyoverdine, especially also when their respective IEF-pattern are identical. The original method [8] was to incubate cells in a non-proliferating medium (usually, succinate medium with the nitrogen source omitted, for cells previously grown in succinate medium), in presence of the labelled iron-pyoverdine complex added at time 0. Cells were then quickly filtered through ultrafiltration membranes (0.45 μm porosity) following different incubation periods, and, after washing with fresh medium and wrapping filters in aluminium foils, radioactivity was determined by using a Gamma-counter. Modifications of the original method have been adapted in order to rapidly analyze numerous strains and to test them with a large number of pyoverdines. The present experimental procedure (see [42] for a recent description of the modified method) allows assays on 6 strains tested towards 35 pyoverdines each, within a day. Even so, the method started to become quite heavy when the number of structurally different pyoverdines recognised by siderotyping or chemical analysis reached 100 or more compounds. Then, the pyoverdine bank was split into four groups, one group containing all the pyoverdines characterized by an acidic IEF profile (band(s) with pHi < pH 6; acidic PVDs), another one with pyoverdines showing IEF profiles with acidic and neutral pHi bands (neutral PVDs), a third one with pyoverdines with neutral and alkaline PVD bands (alkaline PVDs), and a fourth group with pyoverdines presenting together acidic, neutral and alkaline bands (mixed pHi PVDs). In such a way, for analyzing a new isolate, the IEF profile of its pyoverdine was first established, and then the strain was tested for iron uptake towards its own pyoverdine and also towards the group of pyoverdines of foreign origin present in the bank and having the same type of IEF profile. Indeed, in case one foreign pyoverdine was as efficient in iron uptake as the indigenous pyoverdine, then the unknown strain was assigned to the siderovar of the strain producing the pyoverdine belonging to the bank (after verifying, indeed, their IEF profile identity). In a large majority of cases, consisting results were obtained by using such protocol, as verified by pyoverdine structure controls. However, as described below, a few exceptions were detected, with strains producing pyoverdines with identical PVD–IEF patterns and identical iron-uptake efficiency, but showing slight structural differences, e.g., the replacement of a neutral amino acid residue of the peptide chain by another neutral amino acid. A third siderotyping method, based on the pyoverdine molecular mass determination by FAB-Mass spectrometry could differentiate such pyoverdines, as recently demonstrated [38].
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7.4 Siderotyping of the Fluorescent Pseudomonas Species 7.4.1 Fluorescent Pseudomonas Species Analyzed Through Siderotyping A total of 55 validly published fluorescent Pseudomonas species represented by 618 strains have been investigated through siderotyping, with results presented in the present chapter. Three non-officially recognized species, namely Pseudomonas entomophila (one strain [43, 44]), Pseudomonas pavonanceae (one strain, given by S.W. Kwon) and Pseudomonas protegens (10 strains, [45]), have been added to the study. Also added two species, namely Pseudomonas delhiensis (one strain, [46]) and Pseudomonas reinekei (one strain, [47]) which, according to an apparent negative King’s B test, have been described as belonging to the non-fluorescent group of Pseudomonas species, while they were seen in our hands to produce large amounts of pyoverdine when grown in CAA medium. On the contrary, two species, namely Pseudomonas plecoglocissida (6 strains, [48]) and Pseudomonas nitroreducens (one strain, [49]), expected to belong to the fluorescent Pseudomonas group according to the literature, were not included in the study since none of their representatives was seen to produce pyoverdine when grown in iron-deficient media (King’s B, CAA medium or Succinate medium). All these species are listed by alphabetical order in Table 7.1. A supplement of 6 other strains, belonging to the species Pseudomonas amygdali (1 strain), Pseudomonas caricapapaya (1 strain), Pseudomonas flectens (1 strain) and Pseudomonas tremae (3 strains), have been also briefly studied. Contrarily to the other strains, they did not grow or grew very poorly in the iron-depleted media used for siderotyping (CAA medium or Succinate medium) and, therefore, they could not be characterized. Thus, the corresponding species were not included in Table 7.1. It should be highlighted that about two third of the species listed in Table 7.1 have been created since the last 12 years or less, thus corresponding to bacterial strains which have been intensively studied, as requested now for a complete taxonomical work, through numerical taxonomy [24]. The other one third of the species have been described since longer time, with some which have been defined more than hundred years ago, e.g., the type species Pseudomonas aeruginosa or the well distributed P. fluorescens and P. putida species [50, 51]. Such species have been not so well characterized because defined on the basis of phenotypical characters, only. We will see in the present chapter that, in most cases, the discrepancies observed between siderotyping and taxonomy are due to the weakness of the taxonomical approach previously done on these strains. It explains also why the strains selected in the present study for representing the well common species P. fluorescens and P. putida, have been restricted to a very small number of strains, in fact those presently known by previous siderotyping studies, to be closely related to the respective type-strains P. fluorescens ATCC 13525 and P. putida ATCC 12633 ([52, 42]: see these publications for a detailed description of
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the heterogeneity which characterizes these two « species » recognized, rather, as complexes of species, which remain to be taxonomically revisited, [15]). As shown in Table 7.1, the number of strains belonging to each species under studies, varies considerably from one species to another. Twenty out of the 60 species were represented by a unique strain: this because some species have been defined based on an unique representative and thus corresponding to the recently proposed group of “species proponenda” [53], or because only the type-strain remains presently available in the Collections’s catalogues or in the inventor’s laboratory, for some other species. Fortunately, for the 40 remaining species, we were able to analyze more than a single strain, the maximum number being 107 strains for Pseudomonas salomonii. As it will be shown below, the possibility to analyze through siderotyping many representatives within a species is crucial for reaching valid conclusions related to the number of siderotypes (i.e., number of different pyoverdines) found within one species. The general rule being that one pyoverdine corresponds to a given bacterial species (see below), which means that all strains belonging to that species are producing the same type of pyoverdine, it is indeed of primary importance to be able to analyze more than one strain for well defining the siderotyping characteristics of a given species. It is a particularly good reason for encouraging microbiologists involved in taxonomy to save the maximum of well defined strains among their own bacterial collections and not only the one defined as the type-strain. Siderotyping data illustrated in the present work were usually obtained from experiments done mostly on species type-strains. However, some exceptions occurred for species with type-strains defective in pyoverdine production, i.e., Pseudomonas monteilii CFML 90-60T replaced by P. monteilii CFML 90-54 or Pseudomonas poae DSM 14936T replaced by P. poae strain P529/17 or, indeed, for species presenting two or more siderovars (i.e., groups of strains producing different types of pyoverdines), each one being illustrated by the use of a siderovar reference strain (see Table 7.2). Altogether, the study concerned thus 64 species represented by a total of 638 fluorescent Pseudomonas strains.
7.4.2 Analysis of the Fluorescent Pseudomonas Species Through Siderotyping 7.4.2.1 Analysis of the Fluorescent Pseudomonas Species Through Isoelectrophoresis The Table 7.3 represents the siderotyping data – IEF profiles and siderotypes according to pyoverdine-mediated iron uptakes – of the 60 analyzed species enumerated in Table 7.1. For conveniency, a schematic representation of the experimental PVD–IEF profiles, as illustrated in Fig. 7.2, has been adopted, by represen ting each fluorescent band by a bar with a thickness proportional to the fluorescence intensity, on a pH scale from 3.5 to 9.5, starting from the left side (see inner panel
sv.1 sv.2 sv.3 sv.4 sv. 1 sv. 2 sv.1 sv.2 sv.3 sv.1 sv.2 sv.1 sv.2 sv.1 sv.2 sv.3 sv.1 sv.2 sv.1 sv.2 sv.1 sv.2
P. aeruginosa
P. lurida
P. lini
P. libanensis
P. grimontii
P. citronellolis
P. cichorii
P. cedrina
P. aureofaciens
Siderovar (sv.)
Species ATCC 15692 ATCC 27853 PaR (Pa6) PaR’ DSM 6698T D-TR133 CFML 96-198T CFML 96-214 CFML 96-213 CFBP 2101T CFBP 4404 DSM 50332T DSM 11735 CIP 106645T CFML 97-523 CFML 96-347 CFML 96-195T CFML 96-192 CFBP 5737T CFBP 5732 DSM 15835T 239/01
Reference strain of the siderovar 37/89 37/89 14/89 1/89 1/4 3/4 3/9 5/9 1/9 4/9 5/9 1/3 2/3 26/34 6/34 2/34 3/6 3/6 2/9 7/9 2/3 1/3
No. of strains/ No. of strains tested PA01 27853 PaR (Pa6) PaR’ CHA0 D-TR133 96-198 96-214 96-213 Cic1 Cic2 Cit1 Cit2 13525 Pfl12 Pfl18.1 96-195 96-192 B10 PL9 95-275 96-318
Siderovar name
Table 7.2 Fluorescent Pseudomonas species with multiple siderovars
39 43 41 40 15 14 unknown unknown unknown 30 unknown unknown unknown 46 49 48 9 unknown 29 36 60 58
PVD structure (No from Table 7.4)
[111, 34] [34, 112] [113, 34] [114] [69, 103] [85, 69] [69] [69] [69] [108, 69] [69] [69] [69] [78, 75] [101] [115] [38] [69] [37] [81] [116] [107]
References
7 Pyoverdine Siderophores as Taxonomic and Phylogenic Markers 213
sv.1 sv.2 sv.3 sv.1 sv.2 sv.3 sv.4
P. marginalis
P. putida sensu stricto
Siderovar (sv.)
Species
No. of strains/ No. of strains tested 5/8 2/8 1/8 1/7 3/7 2/7 1/1
Reference strain of the siderovar CFBP 2038T CFBP 4051 CFBP 4044 ATCC 12633T CFML 90-40 CFML 90-46 CFML 90-49
Table 7.2 (continued)
DSM 50106 G76 Pfl12 12633 90-40 90-44 90-51
Siderovar name
47 61 49 22 11 17 20
PVD structure (No from Table 7.4)
[38] [38] [101] [117] [118] [119] [120]
References
214 J.-M. Meyer
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Table 7.3 Fluorescent Pseudomonas species and their respective siderotyping characteristics
PVD-IEF profilea Species P. avellanae P. cannabina P. congelans P. ficuresectae P. meliae P. savastanoi P. syringae P. viridiflava P. cichorii sv. 1 P. putida sv. 3 P. marginalis sv. 2 P. koreensis P. mandelii P. reinekei P. umsongensis P. putida sv. 1 P. aureofaciens sv. 2 P. brassicacearum P. lini sv. 2 P. "entomophila" P. putida sv. 2 P. citronellolis sv. 1 P. fuscovaginae P. lini sv. 1 P. moraviensis P. "pavonanceae" P. monteilii P. brenneri P. gessardii P. costantinii P. mosselii P. meridiana P. thivervalensis P. proteolytica P. aeruginosa sv. 3 P. aeruginosa sv. 4 P. putida sv. 4 P. panacis P. grimontii sv. 2 P. marginalis sv. 3 P. aureofaciens sv. 1 P. "protegens" P. asplenii P. aeruginosa sv. 1 P. otitidis P. rhodesiae P. marginalis sv. 1
No of strains with profile/no tested
(PVD-isoform pHi values) 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0
1/1 1/1 2/2 1/1 1/1 1/1 51/60 6/6 4/4 2/2
2/2 1/1 42/43 1/1 1/1 1/1 3/3 10/10 7/7 1/1 3/3 1/1 14/16 2/2 1/1 1/1 7/10 12/12 13/13 10/10 10/11 1/1 6/6 1/1 9/14 1/1 1/1 1/1 6/6 1/1 1/1 10/10 1/1 27/37 7/11 7/7
5/5
PVD(-)
Siderotype according to iron uptake b syr syr syr syr syr syr syr syr syr/cic1 90-44 G76 SB8.3 SB8.3 SB8.3 SB8.3 12633 D-TR133 PL9 PL9 Ent 90-40 cit1 fus B10 B10 B10 90-54 PflW PflW cos mos 51W thi Pflii Pa6/PaR Pa6/PaR' 90-51 Pfl12 Pfl12 Pfl12 CHA0 CHA0 asp PA01 oti L25 DSM 50106
216
J.-M. Meyer Table 7.3 (continued)
P. antarctica P. veronii P. fluorescens (13525) P. aurantiaca P. chlororaphis P. grimontii sv. 1 P. orientalis P. palleroniana P. poae P. trivialis P. blatchfordae P. libanensis sv. 2 P. grimontii sv. 3 P. extremorientalis P. lurida sv. 1 P. lurida sv. 2 P. salomonii P. simiae P. citronellolis sv. 2 P. cedrina sv. 1 P. cedrina sv. 3 P. cedrina sv. 2 P. tolaasii P. agarici P. aeruginosa sv. 2 P. kilonensis P. delhiensis P. libanensis sv. 1 P. cichorii sv. 2 P. jessenii P. migulae
1/1 8/8 9/9 2/2 3/3 26/26 6/6 10/11 1/3 7/7 1/1 3/3 2/2 1/1 2/2 1/1 105/107 4/4 2/2 3/3 1/1 5/5 17/17 1/1 30/37 12/12 1/1 3/3 5/5 8/8 10/10
13525 13525 13525 13525 13525 13525 13525 13525 13525 13525 96-192 96-192 Pfl18.1 95-275 95-275 96-318 96-318 G84 cit2 96-198 96-213 96-214 tol aga 27853 A6 del 96-195 Cic2 9AW 9AW
a Sources
are mainly from 69, 52 and 42. is assessed according to the pyoverdine which mediates the most efficient iron incorporation, providing that it shows an identical IEF pattern compared to the native pyoverdine. Abbreviations for siderotypes are identical to the ones defined in Tables 7.1 and 7.2 for siderovars.
b Siderotype
of Table 7.3). Thanks to the easiness and rapidity of the isoelectrophoresis method, all available strains within a species have been analyzed. The total number of analyzed strains for each species and the number of strains among them showing the depicted IEF profile are indicated in Table 7.3 (second column). In most cases, 100% of the strains belonging to a same species, developed the same IEF profile. A few exceptions, however, concerned some species with a few strains apparently devoid of pyoverdine production, e.g., Pseudomonas mandelii (1 PVD(-) strain over 43 strains analyzed, Pseudomonas fuscovaginae (2 over 16), Pseudomonas salomonii (2 over 107). Two species, however, appeared to be particularly affected in pyoverdine production, namely P. syringae with 9 strains over 60 and P. aeruginosa (10 over 37, 7 over 37 and 5 over 14, for, respectively, the three main sv. 1, sv. 2 and
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sv. 3 siderovars (see Table 7.3) recognized among that species (Table 7.2). While it appears that the apparent lack of pyoverdine production could be rather due to a weakness in pyoverdine expression for some P. syringae strains [54], the high number of pyoverdine-deficient mutants, particularly observed in P. aeruginosa strains of cystic fibrosis origin, seems to be related to environmental conditions but not to mucoidy itself [55]. The 47 different IEF-profiles depicted in Table 7.3 well illustrate the diversity of pyoverdines among fluorescent Pseudomonas species. It can be noted that: (1) acidic, neutral, alkaline and mixed profiles with 1–4 isoform bands were obtained, thus representing the diversity already observed during studies on collection strains or natural isolates [52, 42]; (2) 49 species of the 60 (82%), presented each a unique PVD–IEF profile. While 20 of them were represented by a unique strain, thus being of not conclusive value, the 29 other species well comforted the major conclusion already reached in previous studies [34,42,52] that all PVD(+) strains belonging to a given species (or to a given siderovar for species with multiple siderotypes, see below) presented an identical PVD–IEF profile; (3) The remaining 11 species, enumerated in Table 7.2, were seen to develop among their respective strains several PVD–IEF profiles, allowing the grouping of strains within each species of as many siderovars as PVD–IEF profiles, i.e. from 2 to 4 per species. Siderovars among these 11 species are detailed in Table 7.2, with the name of the reference strain adopted for each siderovar, and the distribution of strains within the different siderovars for each species. Indeed, more strains remain to be analyzed for some of them (e.g., P. citronellolis, P. lurida, with only 3 strains so far analyzed) in order to assess the number of siderovars and strain distribution among these species; (4) Ranging the profiles from the more acidic first band to the more alkaline ones, as presented in Table 7.3, allows to recognize 34 distinct PVD–IEF profiles characterizing each a different species or siderovar. Moreover, 12 other PVD– IEF profiles could be thus easily recognized as being each common to several species or species siderovars, since presenting identical or very similar patterns. Slight differences could be observed for some, affecting the intensity of bands or the number of bands, with sometimes supplementary minor bands present for some and absent for others (e.g., the P. brassicacearum/P. lini sv. 2 group, see Table 7.3). For these groups in particular, the use of another siderotyping method was then required in order to confirm the grouping of strains as suggested by isoelectrophoresis. 7.4.2.2 Analysis of the Fluorescent Pseudomonas Species Through Iron Uptake Pyoverdine-mediated iron uptake experiments were involved for mainly two purposes. The first one was for controlling the grouping of strains first obtained according to the identity or similarity of their PVD–IEF patterns. To do so, each strain belonging to a PVD–IEF group was tested towards its own pyoverdine and
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towards the pyoverdine of the type- (or reference-) strain recognized among the group. Indeed, to match the first classification, strains among one group had to incorporate iron at an identical efficiency for both pyoverdines. The second purpose of iron uptake studies was to determine if the pyoverdine produced by strains within a group could correspond to an already identified pyoverdine. To do so, one strain was selected within the group, usually the type- or reference-strain, and tested for iron incorporation with the different pyoverdines already recognized through structure identification (63 compounds) or through siderotyping (the 63 compounds and 49 others with specific isoelectrophoresis behaviour and iron uptake specificity). As mentioned previously (see Section 7.3.3), heterologous pyoverdine-mediated iron incorporations were usually limited to the pyoverdines belonging to the same pHi group (acidic, neutral, alkaline or mixed pyoverdines), as the strain tested. Altogether, results of the pyoverdine-mediated iron uptake method correlated very well with the isoelectrophoresis data, allowing the confirmation in a large majority of cases of strain grouping within species and siderovars as well as species or siderovar grouping for those sharing an identical PVD–IEF pattern. A few exceptions have to be mentioned, however, relevant to four P. cichorii sv.1 strains and to one strain of P. aeruginosa. Based on identical siderotyping data (identical IEF patterns and cross-incorporations at 100% efficiency), these strains were first recognized as belonging, respectively, to the P. syringae group, or to the siderovar 3 of P. aeruginosa. As could be seen in Table 7.4, pyoverdine structure determination revealed, however, small differences between the pyoverdine No. 30 of P. cichorii CFBP 2101T (reference strain for the P. cichorii sv.1 group) and the pyoverdine No. 31 of P. syringae ATCC 19310T: a gly residue in the PVD(30) is replaced by a ser residue in PVD(31). Concerning the P. aeruginosa R’ strain, its pyoverdine contains one unique gln residue, while the reference strain of the P. aeruginosa sv. 3 (PaR strain) has two such residues (PVDs No. 40 and 41, respectively, in Table 7.4). The most appropriate siderotyping method, which could differentiate such very closely related pyoverdines is FAB-Mass spectrometry, as recently recommended [38]. On the contrary, a weak correlation between isoelectrophoresis and iron uptake concerns a group of 10 species (or species siderovars) which were seen to all recognize very efficiently the pyoverdine of P. fluorescens ATCC 13525, while their respective IEF-profiles were quite different by the number of isoforms and their respective pHi values (see Table 7.3). Structure identification done on most pyoverdines produced by these species [39] revealed a structural identity between these compounds, which supported the iron incorporation results. Therefore, the 10 species or species siderovars were grouped together in a same siderovar, although the differences in IEF patterns which are maybe due to preferential strain-specific isoform accumulations during bacterial growth. 7.4.2.3 Conclusive Remarks on Siderotyping By itself, the isoelectrophoresis siderotyping method allows a good discrimination between pyoverdines produced by fluorescent Pseudomonas. The method is
PVD N° 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36
Pyoverdine peptide chain structure Ala-AcOHOrn-Orn-Ser-Ser-Ser-Arg-OHAsp-Thr Ala-Lys-Dab-OHAsp-(Thr-Gly- OHAsp-Gly-Thr-Thr) Ala-Lys-Gly-Gly-OHAsp-(Gln-Dab)-Gly-Ser-cOHOrn Ala-Lys-Gly-Gly-OHAsp-(Gln-Dab)-Ser-Ala-cOHOrn Ala-Lys-Gly-Gly-OHAsp-Gln-Ser-Ala-Ala-Ala-Ala-cOHOrn Ala-Lys-Gly-Gly-OHAsp-Gln-Ser-Ala-Gly-aThr-cOHOrn Ala-Lys-Thr-Ser-AOHOrn-cOHOrn Ala-Orn-OHAsp-Dab-AOHOrn-Lys Ala-Orn-OHAsp-Ser-Orn-Ser-cOHOrn Asn-FOHOrn-Lys-(Thr-Ala-Ala-FOHOrn-Lys) Asp-Ala-Asp-AOHOrn-Ser-cOHOrn Asp-Arg-AOHOrn-Lys-Ser-Asp-cOHOrn Asp-(AOHOrn-Dab)-Thr-Ala-Thr-Thr-Gln-cOHOrn Asp-FOHOrn-Lys-(Thr-Ala-Ala-FOHOrn-Ala) Asp-FOHOrn-Lys-(Thr-Ala-Ala-FOHOrn-Lys) Asp-Lys-AcOHOrn-Ala-Ser-Ser-Gly-Ser-cOHOrn Asp-Lys-AOHOrn-Thr-Ser-Ser-Gly-Ser-Ser-cOHOrn Asp-Lys-Thr-OHAsp-Thr-aThr-cOHOrn Asp- Lys-OHAsp-Ser-Ala-Ser-cOHOrn Asp- Lys-OHAsp-Ser-Gly-aThr-Lys-cOHOrn Asp- Lys-OHAsp-Ser-Ser-Thr-Thr-Thr-cOHOrn Asp- Lys-OHAsp-Ser-Thr-Ala-Glu-Ser-cOHOrn Asp- Lys-OHAsp-Ser-aThr-Ala-Thr-Lys-cOHOrn Asp-OHbutOHOrn-Dab-Thr-Gly-Ser-Ser-OHAsp-Thr Asp-Orn-(OHAsp-Dab)-Gly-Ser-cOHOrn Lys-OHAsp-Ser-Ser-Ser-cOHOrn Lys-AOHOrn-Ala-Gly-aThr-Ser-cOHOrn Lys-AOHOrn-Gly-aThr-Thr-Gln-Gly-Ser-cOHOrn Lys-OHAsp-Ala-aThr-Ala-cOHOrn Lys-OHAsp-Thr-(Thr-Gly-OHAsp-Ser) Lys-OHAsp--Thr-(Thr-Ser-OHAsp--Ser) (Ser-Dab)-Thr-Ser-AOHOrn-cOHOrn Ser-Ala-AOHOrn-(Orn-Asp--AOHOrn-Ser) Ser-Ala Thr-Lys-Orn-AcOHOrn-Thr-Thr-Ala-Ser-Thr-Ala-Ala-cOHOrn Ser-AOHOrn-Ala-Gly-(Ser-Ala-OHAsp-Thr) Ser-AOHOrn-Ala-Gly-(Ser Ser-OHAsp-Thr)
Isolated from P. fluorescens Pf0-1 P. putida G172 P. fluorescens Pfl 1.3 P. fluorescens Pfl 17400 P. fluorescens Pfl ii P. fluorescens 51W P. fluorescens Ps4a (SB8.3) Pseudomonas sp. Ps 6-10 P. libanensis CFML 96-195 P. fluorescens Pfl "ng" P. putida BTP1/CFML 90-40 Pseudomonas sp. LBSA1 P. putida 3b P. chlororaphis D-TR133 P. fluorescens CHA0 P. monteilii CFML 90-54 P. putida CFML 90-44 P. putida CFML 90-33 P. putida CFBP 11370 P. putida CFML 90-51 Pseudomonas sp. HR6 P. putida ATCC 12633 P. putida L1 P. putida PutC P. putida G4R P. putida GS43 P. fluorescens PL8 P. fluorescens A6 Pseudomonas sp. B10 P. cichorii CFBP 2101 P. syringae ATCC 19310 P. putida Thai P. fluorescens G173 Pseudomonas sp. IB3 P. putida ATCC 39167 P. fluorescens PL9
Correlated by siderotyping to species n.c. n.c. n.c. n.c. P. proteolytica P. meridiana see Table 7.5 n.c. P. libanensis (sv.1) n.c. P. putida (sv.2) n.c. n.c. P. aureofaciens (sv.2) see Table 7.5 P. monteilii P. putida (sv.3) n.c. n.c. P. putida (sv.4) n.c. P. putida (sv.1) n.c. n.c. n.c. n.c. n.c. P. kilonensis see Table 7.5 P. cichorii (sv.1) see Table 7.5 n.c. n.c. n.c. n.c. see Table 7.5
Structure references 83 83 51 37 86 131 23 26 83 100 64 83 22 10 135 83 118 119 25 116 83 99 124 112 109 83 9 15 121 28 65 108 123 83 124 133
Table 7.4 Pyoverdines classified alphabetically according to the amino acids of their respective peptide chain (starting from the aa-residue attached to the chromophore group by its NH2 )
7 Pyoverdine Siderophores as Taxonomic and Phylogenic Markers 219
Pyoverdine peptide chain structure Ser-AOHOrn-Ala-Gly-aThr-Ala-cOHOrn Ser-AOHOrn-Gly-aThr-Thr-Gln-Gly-Ser cOHOrn Ser-Arg-Ser-FOHOrn-(Lys-FOHOrn-Thr-Thr) (Ser-Dab)-FOHOrn-Gln-FOHOrn-Gly (Ser-Dab)-FOHOrn-Gln-Gln-FOHOrn-Gly Ser-Dab-Gly-Ser-OHAsp-Ala-Gly-Ala-Gly-cOHOrn Ser-FOHOrn-Orn-Gly-aThr-Ser-cOHOrn Ser-Lys-Ala-AOHOrn-Thr-Ala-Gly-Gln-Ala-Ser-Ser-cOHOrn Ser-Lys-Ala-Ser-Ser- AcOHOrn-Ser-Ser-cOHOrn Ser-Lys-Gly-FOHOrn-(Lys-FOHOrn-Ser) Ser-Lys-Gly-FOHOrn-Ser-Ser-Gly-(Orn-FOHOrn-Ser) Ser-Lys-Gly-FOHOrn-Ser-Ser-Gly-(Lys-FOHOrn-Ser) Ser-Lys-Gly-FOHOrn-Ser-Ser-Gly-(Lys-FOHOrn-Glu-Ser) Ser-Lys-OHAsp-Ser-Orn-Ser-cOHOrn Ser-Lys-FOHOrn-(Lys-FOHOrn-Glu-Ser) Ser-Lys-FOHOrn-Ser-Ser-Gly-(Lys-FOHOrn-Ser-Ser) Ser-Lys-Ser-Ser-Thr-Ser-AcOHOrn-Thr-Ser-cOHOrn Ser-Lys-Ser-Ser-Thr-Thr-AcOHOrn-Ser-Ser-cOHOrn Ser- Lys-OHHis-aThr-Ser cOHOrn Ser-Orn-OHAsp-Ser-Ser-Ser-cOHOrn Ser-Orn-FOHOrn-(Lys-FOHOrn-Glu-Ser) Ser-Orn-FOHOrn-Ser-Ser-(Lys-FOHOrn-Ser) Ser-Ser-FOHOrn-(Lys-FOHOrn-Lys-Ser) Ser-Ser-FOHOrn-Ser-Ser-(Lys-FOHOrn-Lys-Ser) Ser-Ser-FOHOrn-Ser-Ser-(Lys-Ser-FOHOrn) Ser-Thr-Ser-Orn-OHAsp-(Gln-Dab)-Ser-aThr-cOHOrn Ser-Val-OHAsp-Gly-Thr-Ser-cOHOrn Isolated from P. fluorescens PL7 P. aureofaciens P.au P. aeruginosa PAO1 P. aeruginosa R' P. aeruginosa R (Pa6) P. fluorescens Pfl W P. aeruginosa Pa 27853 P. fluorescens 1547 P. fluorescens G153 P. fluorescens ATCC 13525 P. fluorescens DSM 50106 P. fluorescens Pfl18.1 P. fluorescens Pfl12 Pseudomonas sp. G85 Pseudomonas sp. CFML 96-188 P. rhodesiae CFML 92-104 P. tolaasii CFBP 2068 P. putida CFML 90-136 P. fluorescens 9AW Pseudomonas sp. 2908 Pseudomonas sp. D47 Pseudomonas sp. CFML 96-318 Pseudomonas sp. CFML 96-312 P. fluorescens CFML 95-275 Pseudomonas sp. G76 P. fluorescens Gwose P. fluorescens BTP2
Correlated by siderotyping to species n.c. P. costantinii P. aeruginosa (sv.1) P. aeruginosa (sv.4) P. aeruginosa (sv.3) see Table 7.5 P. aeruginosa (sv.2) n.c. n.c. see Table 7.5 P. marginalis (sv.1) P. grimontii (sv.3) see Table 7.5 n.c. n.c. P. rhodesiae P. tolaasii n.c. see Table 7.5 n.c. n.c. see Table 7.5 n.c. see Table 7.5 P. marginalis (sv.2) n.c. n.c.
Structure references 9 14 19 107 53 37 120 106 83 60 83 3 50 83 134 22 37 83 24 132 110 111 111 117 83 57 92
Abbreviations: usual amino acids, three letter code; aThr, allo-Thr; Lys, Lys linked by its -NH2; AOHOrn, δN-acetyl-δNhydroxy- ornithine; FOHOrn, δN-formyl-δN-hydroxy-ornithine; OHButOHOrn, δN-hydroxy-butyryl-δN-hydroxy-ornithine; cOHOrn, cyclo-hydroxy-ornithine (3amino-1-hydroxy-piperidone-2); OHHis, threo-β-hydroxy-histidine; OHAsp, threo-β-hydroxy-aspartic acid; Dab, diamino-butanoic acid. Parentheses indicate cyclic structures. D-amino acids are underlined. A brocken line means that the two enantiomers have been detected among the underlined residues but with no precise affectation. Stereochemistry remains to be done for pyoverdines with no amino acid underlined.
PVD N° 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63
Table 7.4 (continued)
220 J.-M. Meyer
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Pyoverdine Siderophores as Taxonomic and Phylogenic Markers
221
usually self-sufficient for successfully grouping strains which produce an identical pyoverdine. Moreover, there is no doubt that strains presenting well differentiate IEF patterns correspond to structurally well differentiated pyoverdines. However, due to close similarities between some PVD–IEF patterns (e.g., see Table 7.3, the P. entomophila and the P. citronellolis sv. 1 patterns, or the patterns of P. aeruginosa sv. 3 and P. panacis/P. grimontii sv. 2/P. marginalis sv. 3), and also because of the different IEF features described above for the P. fluorescens ATCC 13525 group, it is advised to confort the isoelectrophoresis conclusive data by another siderotyping method, e.g., the pyoverdine-mediated iron uptake, before grouping strains into siderovars.
7.4.3 Pyoverdines as Taxonomic Markers: Correlations Between Pyoverdines and Fluorescent Pseudomonas Species The Table 7.4 is listing the 63 pyoverdines whose structures are presently fully determined at the level of their respective peptide chains (for some, however, i.e., PVDs No. 1, 2, 9, 12, 16, 21, 26, 32, 34, 45, 47, 50, 52 and 61, in Table 7.4, the stereochemistry of the amino acyl residues remains to be done). Of the 63 different compounds, 21 are each correlated to a unique species or species siderovar, while 11 are each shared by several species or species siderovars. For clarity, these 11 siderotypes and their respective corresponding bacterial groups are listed in Table 7.5, as well as a supplementary pyoverdine correlating with one species and one species siderovar [PVD (96-192)], which structure is presently unknown. Almost half of these pyoverdines (31 over 63; marked n.c. for “not correlated” in Table 7.4) still remain to be correlated to a well-defined bacterial group. It should be highlighted that they all were originally isolated from strains very poorly or not identified at all, described in the literature as P. fluorescens or P. putida (11 strains each), or as Pseudomonas sp. (9 strains). Indeed, according to their siderotyping characteristics, these 31 strains could be the source of many novel species, which remain to be taxonomically defined. Correlations between species or species siderovars and pyoverdines are also indicated in Tables 7.1 and 7.2, where the siderotyping status of the 60 fluorescent Pseudomonas species can be thus easily found. Of the 49 species characterized by a unique, well-defined siderotype, 39 correspond presently to a structurally known pyoverdine, while for 10 other species, their respective pyoverdine structure remain to be determined. Moreover, among the multisiderovaric species (11 species resulting in 29 siderovars listed in Table 7.2), 22 siderovars correlate with structurally defined pyoverdines, while 7 siderovars remain with structurally unknown pyoverdines. More work has thus to be done to reach a complete knowledge on pyoverdine structures and their correlations with well taxonomically defined Pseudomonas species. Whatever, any fluorescent pseudomonad isolate, characterized by one of the 49 monosiderovaric siderotypes, or by one of the 29 siderotypes found within the 11 multisiderovaric species of Table 7.2, can be consequently
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J.-M. Meyer Table 7.5 Siderotypes shared by several fluorescent Pseudomonas species
Siderotype
PVD No (from Table 7.4)
Reference-strain of the siderotype
Species characterized by the siderotype
SB8.3
7
P. fluorescens SB8.3
CHA0
15
B10
29
P. fluorescens CHA0 Pseudomonas sp. B10
syr
31
P. syringae ATCC 19310T
PL9
36
PflW
42
13525
46
P. fluorescens C-TR1015 P. fluorescens CCM 2798 P. fluorescens ATCC 13525T
P. koreensis, P. mandelii, P. reinekei, P. umsongensis P. aureofaciens (restricted to sv. 1), P. ‘protegens’ P. lini (restricted to sv. 1), P. moraviensis, P. ‘pavonanceae’ P. avallanae, P. cannabina, P. congelans, P. ficuserectae, P. meliae, P. savastanoi, P. syringae (60 pathovars), P. viridiflava P. brassicacearum, P. lini (restricted to sv.2) P. brenneri, P. gessardii
Pfl12
49
P. fluorescens 12
9AW
55
96-318
58
95-275
60
P. fluorescens strain 9AW Pseudomonas sp. CFML96-318 P. fluorescens CFML95-275
96-192
unknown
P. libanensis CFML96-192
P. antartica, P. chlororaphis, P. aurentiaca, P. fluorescens sensu stricto, P. grimontii (sv. 1), P. orientalis, P. palleroniana, P. poae, P. trivialis, P. veronii P. grimontii (restricted to sv.2), P. panacis, P. marginalis (restricted to sv.3) P. jessenii, P. migulae P. lurida (restricted to sv.2), P. salomonii P. extremorientalis, P. lurida (restricted to sv.1) P. blatchfordae, P. libanensis (restricted to sv.2)
correlated with a precise bacterial group, each corresponding to a taxonomically well defined species. In case its siderotype is identical to one of the 12 siderotypes cited in Table 7.5, then a taxonomical differentiation of the strain in-between a very limited number of species should allow its attribution to a precise species:
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from 2 to 4 species for most of the siderotypes, to 8 or 10 different species for the most widespread siderotypes corresponding, respectively, to P. syringae and P. fluorescens ATCC 13525. It should be highlighted finally, that siderotyping has been already successfully involved in the definition of recently described new species, namely Pseudomonas brassicacearum and Pseudomonas thivervalensis [56], Pseudomonas lini [57], Pseudomonas mosselii [58], Pseudomonas salomonii and Pseudomonas palleroniana [21], Pseudomonas costantinii [59], Pseudomonas lurida [60] and Pseudomonas cedrina [61]. Furthermore, the assignation of a phenotypic cluster to a given species or, on the contrary, the rejection of a misidentified strain, as postulated by siderotyping, were positively verified by DNADNA-hybridization [52], proving that the method is particularly efficient for the detection of potential new species. At last, siderotyping also agrees in the recognition of Pseudomonas kilonensis as a well-individualized fluorescent Pseudomonas species, although the particularly high DNA-DNA hybridization level (> 70%) this taxonomical group presented with the P. brassicacearum species [62].
7.4.4 Pyoverdines, as Phylogenic Markers Among the methods used to define phylogenetic relationships between bacterial strains, DNA sequence comparison of the housekeeping gene rpoB is presently considered as one of the most accurate for Pseudomonas species, being three times more discriminative compared to the classical 16S-RNA gene sequencing method [30]. Therefore, in order to estimate the phylogenetic value of siderotyping, a comparative study involving rpoB sequencing and siderotyping has been developed on a collection of 22 strains recognized by phenotypic features to belong to the biovar 1 of the P. fluorescens species [63]. Pyoverdine isoelectrophoresis and pyoverdine-mediated iron incorporation studies done on the 22 strains and 9 reference strains resulted in the recognition of 9 siderovars, among them some correlating with well defined species, i.e., P. rhodesiae, P. mandelii, P. jessenii. The most important siderovar by the number of isolates corresponded precisely to the siderotype of the P. fluorescens type-strain (ATCC 13525) and was designed as the P. fluorescens sensu stricto siderovar. Two siderovars, each containing a single strain (CFBP12298 or CFBP11354), presented original features and therefore should correspond to novel pyoverdine structures. Interestingly, uptake studies revealed a strong cross-reactivity between the Pfl18.1, P. fluorescens sensu stricto, and CFBP12298 siderovars, while the P. rhodesiae siderovar presented a weak reactivity with the three others. All the other siderovars, i.e., those of Pseudomonas sp. B10, P. mandelii, P. jessenii, Pseudomonas sp. A214 and CFBP11354, presented each specific and independent siderotyping properties. This cross-reactivity observed for the three main siderovars of the P. fluorescens biovar 1 strain collection could be explain by structure similarities between their respective pyoverdines, as can be seen in Table 7.4 for the pyoverdines of P. fluorescens ATCC 13525 (No. 46) and P. fluorescens Pfl18.1 (No. 48), the pyoverdine of P. rhodesiae (No. 52) being
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somewhat different, although related since, like the other structures, having an internal cycle in its peptidic chain. On the contrary, the 4 pyoverdines corresponding to the 4 independent siderovars are structurally unrelated with any of the pyoverdines involved within the group (PVDs No. 29, 7, 55 and 35, respectively). The rpoB sequences of the 22 P. fluorescens bv. 1 strains and the 9 corresponding siderovar reference strains were obtained and compared through the two classical Fitch and Maximum-likelihood methods, resulting in the dendrograms presented in Fig. 7.6, together with the siderotyping data. Although some strain-specific delocalization is depending on the sequence analysis method, it could be stated that correlations between rpoB sequencing and siderotyping are satisfactory. The structural relationships between the cyclic pyoverdines characterizing the three major siderovars correlated well, altogether, to the phylogenetic relationships showed by the two dendrograms. Meanwhile, the 4 independent siderovars grouped separately, like they did in the phylogenetic dendrograms. Pyoverdines in Table 7.4 are ranked by alphabetical order according to the amino acids contained in their respective peptide chains, starting from the N-terminal
Strains
PVD-IEF profiles
Maximum-likelihood method
Fitch method
Sidérovar
P.fl18.1
P.fl 18.1
CFBP11347 79
CFBP11355 CFBP11356
ATCC 17563
ATCC 17563
Pfl 18.1 97
83
ATCC 17563
CIP 56-90
CIP 56-90
78
CFBP 12155
CFBP 12155 89
CFBP 11351
CFBP 11351 95
CFBP12298
unknown
CFBP 12152
CFBP 12152
CFBP 12298
CFBP 12298
99
CFBP12147
CFBP12155 CFBP12204
CFBP 12147
CFBP 12147
CFBP12152
CFBP 11355
CFBP 11355
P. fluorescens 56
sensu stricto
54 57
CFBP11345
P. rhodesiae
CFBP 11356
CFBP 12297
P.fluorescens ATCC 13525
58
CFBP 12300
CFBP 12204
CFBP11351 62
65 100
CFBP 12299
CFBP 12297
CFBP 12204
CFBP 12300
ATCC13525 97
CFBP 11356
CFBP 12299
CFBP12297
P.fluorescens ATCC 13525
P. rhodesiae
CFBP12299 P. rhodesiae
100
P. rhodesiae
80
CFBP12300
CFBP 11347
CFBP 11347
CFBP 11345
CFBP 11345
100
CFBP12156
CFBP 12156
CFBP 12156
CFBP 11363
CFBP 11363
CFBP11363
B10
CFBP 11357
CFBP11344 CFBP12180
95
CFBP11400
CFBP 12180
B10
100
Pseudomonas
CFBP 11354
CFBP 12180
sp. B10
CFBP 12206
CFBP 11354
P.sp. B10 CFBP11354
unknown
CFBP12206
P. mandelii
100
55
CFBP 11400
CFBP 11344
CFBP 11344
CFBP 11400
P. jessenii
CFBP 12206
CIP 105273
53 99
P. jessenii
CFBP 11365
P. jessenii
P. jessenii
P.mandelii
CFBP 11365
CFBP11365 CFBP 11357
P.mandelii
CFBP11357
Pseudomonas
P. sp. A214
sp. A214
A214
A214
P. syringae
P. syringae
Fig. 7.3 Comparison of siderotyping and rpoB sequencing for 22 strains formerly classified in the biovar 1 of the Pseudomonas fluorescens species complex, and 9 reference strains. The pyoverdines of Pfl18.1, Pfl ATCC 13525 and P. rhodesiae (PVDs No. 48, 46 and 52, respectively, in Table 7.4) are structurally related with internal cyclic peptide chains resulting in partial crossreactivity in iron-uptake, while the linear peptide chains of the other pyoverdines (Pseudomonas B10, P. mandelii, P. jessenii and Pseudomonas A214 (PVDs No. 29, 7, 55 and 35, respectively, in Table 7.4) do not cross-react and have no structural relationships
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residue attached to the chromophore. Such classification allows to group together pyoverdine molecules, which present structural relationships at the level of their peptide chains, among them those sharing receptor recognition sites and demonstrating cross-incorporation properties (pyoverdines regrouped within a frame in Table 7.4). Unfortunately, no comparative taxonomical studies have been presently done on strains involved in the biosynthesis of such structurally related pyoverdines. That phylogenetic relationships should exist between such strains could be highly suspected. From the comparison of a group of pyoverdines all starting their peptide chain by an aspartyl residue (pyoverdines No. 11–25 in Table 7.4), it can be deduced that most of their respective producers correspond to strains identified as belonging to the P. putida complex, among them four strains which have been recognized by DNA-DNA hybridization as belonging to the so-called P. putida sensu stricto species [42]. Interestingly, a phylogenic tree can be drawn [42] based on the structure comparison of the four corresponding pyoverdines (No. 11, 17, 20 and 22 in Table 7.4), suggesting effectively that pyoverdines could be used as relevant phylogenetic markers. Comforting this view, it can be added also that species grouped together in a same siderovar (e.g., the P. syringae- or the P. fluorescens ATCC 13525 groups in Table 7.5) are usually found close together in 16S-rDNA trees [26, 60, 64, 57, 21, 65, 66]. Moreover, a recent comparative study done on 85 phytopathogenic Pseudomonas strains corresponding to the fluorescent species P. salomonii, P. palleroniana, P. tolaasii, P. costantinii and P. fuscovaginae, showed a perfect matching between the phenotypic clusters reached by numerical taxonomy and siderotyping [3].
7.5 Conclusions Thanks to its easiness and rapidity, siderotyping is of particular interest as an efficient method for the characterization and identification of fluorescent Pseudomonas at the species level. Meanwhile, as already detailed [67, 68], the method can be of great help in environmental or ecological studies involving numbers of natural pseudomonads isolates. Based on a few phenotypic characters related to pyoverdine, i.e., isoelectrophoresis pattern, molecular mass or iron uptake capacity, siderotyping is, to our knowledge, the only taxonomical method which does not require a subjective borderline in order to delineate a taxonomical group. Once the siderotype of a given strain is defined, it should correspond to a precise siderovar and therefore, in most cases, to a defined fluorescent Pseudomonas species. Otherwise, it should represent a potential new siderovar and therefore a potential new species. Does the method, adjusted to other siderophores, be valid for other bacterial groups? A positive answer is strongly suggested by work done in my laboratory on bacteria closely related to fluorescent pseudomonads, e.g., non-fluorescent Pseudomonas [69, 52], or Burkholderia and Ralstonia species [6]. It should be also remembered that a few publications highlighted already some relationships between siderophores and taxonomic features in other major bacterial groups, i.e., enterobacteria [70], mycobacteria [71, 72] or Aeromonas spp [73].
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Acknowledgments The author deeply acknowledges the many collaborators who participated to the siderotyping development, especially Christelle Gruffaz and Gérard Seyer for their technical expertise and their valuable participation to student formation. W. Achouak, U. Behrendt, H. Budzikiewicz, B. Cámara, M. Champomier, J. Chun, P. Cornelis, J. Djacs, J.F. FernándezGarayzábal, M. Fischer-LeSaux, L. Gardan, D. Haas, D. Izard, S.W. Kwon, E. Lang, P. Lemanceau, P. Munsch, A. Peix, S. Shivaji, J. Sikorski, L. Tvrzová are acknowledged for the gift of strains.
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102. Park, Y.-D., Burm Lee, H., Yi, H., Kim, Y., Bae, K.S., Choi, J.-E., Jung, H.S. and Chun, J. (2005) Pseudomonas panacis sp. nov., isolated from the surface of rusty roots of Korean ginseng. Int. J. Syst. Evol. Microbiol. 55: 1721–1724. 103. Wong-Lun-Sang, S., Bernardini., J.J., Hennard, C., Kyslic, P., Dell, A. and Abdallah, M. (1996) Bacterial siderophores: structure elucidation, 2D 1 H and 13 C NMR assignments of pyoverdins produced by Pseudomonas fluorescens CHA0. Tetrahedron Lett. 37: 3329–3332. 104. Mohn, G., Taraz, K. and Budzikiewicz, H. (1990) New pyoverdin-type siderophores from Pseudomonas fluorescens. Z. Naturforsch. 45b: 1437–1450. 105. Budzikiewicz, H., Schröder, H. and Taraz, K. (1992) Zur Biogenese der PseudomonasSiderophore: der Nachweis analoger Strukturen eines Pyoverdin-Desferribactin-Paares. Z. Naturforsch. 47c: 26–32. 106. Coroler, L., Elomari, M., Hoste, B., Gillis, M., Izard, D. and Leclerc, H. (1996) Pseudomonas rhodesiae sp. nov., a new species isolated from natural mineral waters. Syst. Appl. Microbiol. 19: 600–607. 107. Schlegel, K., Fuchs, R., Schäfer, M., Taraz, K., Budzikiewicz, H., Geoffroy, V.A. and Meyer, J.-M. (2001) The pyoverdins of Pseudomonas sp. CFML 96-312 and CFML 96-318. Z Naturforsch 56c: 680–686. 108. Bultreys, A., Gheysen, I., Wathelet, B., Schäfer, M. and Budzikiewicz, H. (2004) The pyoverdins of Pseudomonas syringae and Pseudomonas cichorii. Z. Naturforsch. 59c: 613–618. 109. Munsch, P., Geoffroy, V., Alatossava, T. and Meyer, J.-M. (2000) Application of siderotyping for the characterization of Pseudomonas tolaasii and Pseudomonas reactans isolates associated with brown blotch disease of cultivated mushrooms. Appl. Environ. Microbiol. 66: 4834–4841. 110. Elomari, M., Coroler, L., Hoste, B., Gillis, M., Izard, D. and Leclerc, H. (1996) DNA relatedness among Pseudomonas strains isolated from natural mineral waters and proposal of Pseudomonas veronii sp. nov. Int. J. Syst. Bacteriol. 46: 1138–1144. 111. Briskot, G., Taraz, K. and Budzikiewicz, H. (1989) Pyoverdine-Type siderophores from Pseudomonas aeruginosa. Liebigs. Ann. Chem.: 375–384. 112. Tappe, R., Taraz, K., Budzikiewicz, H., Meyer, J.-M. and Lefèvre, J.-F. (1993) Structure elucidation of a pyoverdin produced by Pseudomonas aeruginosa ATCC 27853. J. Prakt. Chem. 335: 83–87. 113. Gipp, S., Hahn, J., Taraz, K. and Budzikiewicz, H. (1991) Zwei Pyoverdine aus Pseudomonas aeruginosa R. Z. Naturforsch. 46c: 534–541. 114. Ruangviriyachai, C., Fernandez, D.U., Fuchs, R., Meyer, J.-M. and Budzikiewicz., H. (2001) A new pyoverdin from Pseudomonas aeruginosa R . Z. Naturforsch. 56c: 933–938. 115. Amann, C., Taraz, K., Budzikiewicz, H. and Meyer, J.-M. (2000) The siderophores of Pseudomonas fluorescens 18.1 and the importance of cyclopeptidic substructures for the recognition at the cell surface. Z. Naturforsch. 55c: 671–680. 116. Sultana, R., Fuchs, R., Schmickler, H., Schlegel, K., Budzikiewicz, H., Siddiqui, B.S., Geoffroy, V. and Meyer, J.-M. (2000) A pyoverdin from Pseudomonas sp. CFML 95-275. Z. Naturforsch. 55c: 857–865. 117. Persmark, M., Frejd, T. and Mattiasson, B. (1990) Purification, characterization, and structure of pseudobactin 589A, a siderophore from a plant growth promoting Pseudomonas. Biochemistry 29: 7348–7356. 118. Jacques, P., Ongena, M., Gwose, I., Seinsche, D., Schröder, H., Delfosse, P., Thonart, P., Taraz, K. and Budzikiewicz, H. (1995) Structure and characterization of isopyoverdin from Pseudomonas putida BTP1 and its relation to the biogenetic pathway leading to pyoverdins. Z. Naturforsch. 50c: 622–629. 119. Sultana, R., Siddiqui, B.S., Taraz, K., Budzikiewicz, H. and Meyer, J.-M. (2001) An isopyoverdin from Pseudomonas putida CFML 90-44. Z. Naturforsch. 56c: 303–307. 120. Sultana, R., Siddiqui, B.S., Taraz, K., Budzikiewicz, H. and Meyer, J.-M. (2000) A pyoverdine from Pseudomonas putida CFML 90-51 with a Lys e-amino link in the peptide chain. Biometals 13: 147–152.
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121. Georgias, H., Taraz, K., Budzikiewicz, H., Geoffroy, V. and Meyer, J.-M. (1999) The structure of the pyoverdin from Pseudomonas fluorescens 1.3. Structural and biological relationships of pyoverdins from different strains. Z. Naturforsch. 54c: 301–308. 122. Budzikiewicz, H., Schäfer, M., Fernandez, D.U. and Meyer, J.-M. (2006) Structure proposal for a new pyoverdin from Pseudomonas sp. PS6.10. Z. Naturforsch. 61c: 815–820. 123. Poppe, K., Taraz, K. and Budzikiewicz, H. (1987) Pyoverdine type from Pseudomonas fluorescens. Tetrahedron 43: 2261–2272. 124. Sultana, R., Siddiqui, B.S., Taraz, K., Budzikiewicz, H. and Meyer, J.-M. (2001) An isopyoverdin from Pseudomonas putida CFML 90-33. Tetrahedron 57: 1019–1023. 125. Budzikiewicz, H., Fernandez, D.U., Fuchs, R., Michalke, R., Taraz, K. and Ruangviriyachai, C. (1999) Pyoverdines with a Lys -amino link in the peptide chain? Z. Naturforsch. 54c: 1021–1026. 126. Seinsche, D., Taraz, K., Budzikiewicz, H. and Gondol, D. (1993) Neue PyoverdinSiderophore aus Pseudomonas putida C. J. Prakt. Chem. 335: 157–168. 127. Salah-el-Din, A.L.M., Kyslic, P., Stephan, D. and Abdallah, M.A. (1997) Bacterial iron transport: structure elucidation by FAB-MS and by 2D NMR (1 H, 13 C, 15 N) of pyoverdin G4R, a peptidic siderophore produced by a nitrogen-fixing strain of Pseudomonas putida. Tetrahedron 53: 12539–12552. 128. Barelmann, I., Taraz, K., Budzikiewicz, H., Geoffroy, V.A. and Meyer, J.-M. (2002) The structures of the pyoverdins from two Pseudomonas fluorescens strains accepted mutually by their respective producers. Z. Naturforsch. 57c: 9–16. 129. Ruangviriyachai, C., Uria-Fernandez, D., Schäfer, M. and Budzikiewicz, H. (2004) Structure proposal for a new pyoverdin from a Thai Pseudomonas putida strain. Spectroscopy 18: 453–458. 130. Uría-Fernández, D., Fuchs, R., Schäfer, M., Budzikiewicz, H. and Meyer, J.-M. (2003) The pyoverdin of Pseudomonas fluorescens G173, a novel structural type accompanied by unexpected natural derivatives of the corresponding ferribactin. Z. Naturforsch. 58c: 1–10. 131. Vossen, W., Fuchs, R., Taraz, K. and Budzikiewicz, H. (2000) Can the peptide chain of a pyoverdin be bound by an ester bond to the chromophore ?- The old problem of pseudobactin 7SR1. Z. Naturforsch. 55c: 153–164. 132. Beiderbeck, H., Risse, D., Budzikiewicz, H. and Taraz, K. (1999a) A new pyoverdin from Pseudomonas aureofaciens. Z. Naturforsch. 54c: 1–5. 133. Ruangviriyachai, C., Barelmann, I., Fuchs, R. and Budzikiewicz, H. (2000) An exceptionally large pyoverdin from a Pseudomonas strain collected in Thailand. Z. Naturforsch. 55c: 323–327. 134. Weber, M., Taraz, K., Budzikiewicz, H., Geoffroy, V. and Meyer, J.-M. (2000) The structure of a pyoverdine from Pseudomonas sp. CFML 96.188 and its relation to other pyoverdines with a cyclic C-terminus. Biometals 13: 301–309. 135. Vossen, W. and Taraz, K. (1999) Structure of the pyoverdine PVD 2908 – A new pyoverdin from Pseudomonas sp. 2908. Biometals 12: 323–329. 136. Schäfer, M., Fuchs, R., Budzikiewicz, H., Springer, A., Meyer, J.-M. and Linscheid, M. (2006) Structure elucidation of cyclic pyoverdins and examination of rearrangement reactions in MS/MS experiments by determination of exact product ion masses. J. Mass. Spectrom. 41: 1162–1170. 137. Gwose, I. and Taraz, K. (1992) Pyoverdine aus Pseudomonas putida. Z. Naturforsch. 47c: 487–502. 138. Ongena, M., Jacques, P., Thonart, P., Gwose, I., Fernandez, D.U., Schäfer, M. and Budzikiewicz, H. (2001) The pyoverdin of Pseudomonas fluorescens BTP2, a novel structural type. Tetrahedron Lett. 42: 5849–5851.
Chapter 8
Metabolism of Acyclic Terpenes by Pseudomonas Jesús Campos-García
8.1 Introduction Pseudomonas is a highly versatile bacterial genus with the ability to grow on a wide diversity of organic compounds. The compounds that can be used as a carbon and energy source include root exudates of plants such as sugars, amino acids, and organic acids, animals in putrefaction steps, and other contaminant compounds [1]. Pseudomonas species are found in pristine waters, wild soils and extreme habitats. Some species of the genus Pseudomonas are pathogenic for humans [2], insects [3], nematodes [4] and plants [5]. The effectiveness of this genus in causing infection is likely due to a suite of well-regulated virulence factors and defense mechanisms such as multidrug resistance pumps and biofilm formation [6, 7]. The ability to assimilate acyclic terpenes as sole carbon and energy source is a characteristic of some Pseudomonas species. Terpenes are commonly toxic for microorganism by having easy entrance and intercalation in the cytoplasmic membrane, causing modification in membrane properties, unstabilization and even cellular lyses; in addition, they are recalcitrant due to the methyl branch in the β-carbon, that impedes the action for enzymes of the fatty acid oxidation pathway. Pioneer studies which aimed to the elucidation of the acyclic isoprenoid/terpene catabolic pathways date to the 60s when a new bacteria was isolated and named as Pseudomonas citronellolis by its ability to oxidize citronellol and farnesol [8]. This strain is able to grow on terpenes as citronellol, citronellal, citronellic acid, geranic acid, farnesol, dimethylacrylic acid, and isovaleric acid. Subsequently, other Pseudomonas species were also described as able to grow on citronellol as sole carbon source, i.e. P. aeruginosa, P. mendocina [9], and P. delhiensis [10]. The first pathway for acyclic terpenes degradation was proposed by Cantwell et al. [9], using metabolite identification in cultures grown on citronellol, geraniol or nerol. In addition, a combination of degradative pathways was obtained in J. Campos-García (B) Instituto de Investigaciones Químico-Biológicas, Universidad Michoacana de San Nicolás de Hidalgo, Edif. B-3, Ciudad Universitaria, CP 58030, Morelia, Michoacán, México e-mail:
[email protected]
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P. citronellolis when it was transformed with the OCT plasmid, rendering a strain with the ability to degrades a wider range of branched-chain hydrocarbons [11] and reports describing that acyclic terpenes metabolism was encoded in a conjugative 50-megaDa plasmid pSRQ50 from P. putida [12].
8.2 Terpenes/Isoprenoids Sources Terpenoids and their precursor isoprenoid hydrocarbons are among the oldest known and most ubiquitous organic chemicals on Earth, having been found in fossil remains from a variety of geological formations and sediments [13]. The reconstruction of biosynthetic pathways from such records indicates that the ability to synthesize isoprenoids, such as pristane and phytane and triterpenoids such as squalene and hopanes, was present in non-photosynthetic bacteria before the evolution of chlorophyll biosynthesis. The occurrence of triterpenoids in bacteria, and especially in non-photosynthetic bacteria, is thus of interest from both evolutionary and functional aspects. Particular emphasis is focused on the acyclic triterpenoid carotenoids and the cyclic hopanoids since these two groups of compounds are rapidly emerging as major classes of bacterial natural products [13].
8.2.1 Terpenes/Isoprenoids as Plant Components Plants produce a vast array of volatile compounds that mediate their interactions with the environment. A large portion of these compounds are terpenoids, also known as isoprenoids because all are synthesized through the condensation of C5 isoprene units (Fig. 8.1). Terpenes are the largest class of natural products known, finding widest structural and functional variety in plants where they play important physiological and ecological roles. Among the physiological roles, terpenoids serve as phytohormones, redox cofactors, photosynthetic pigments, photo-protectants, and anchors for membrane associated proteins [16]. The ecological functions of plant terpenoids include acting as attractants for pollinating insects or fruit dispersing animals, anti-herbivore defenses, anti-fungal defenses, agents of plant-plant competition, plant-to-plant communication and as signaling compounds in multitrophic interactions [14, 16]. Terpenoids are derived from either the mevalonate pathway, active in the cytosol/endoplasmic reticulum (responsible for the production of sesquiterpenes (C15 ) and triterpenes (C30 , e.g. sterols)), or the plastidial 2-Cmethyl-d-erythritol 4-phosphate (MEP) pathway (responsible for the production of isoprene (C5 ), monoterpenes, diterpenes (C20 , e.g. gibberellins), and tetraterpenes (C40 , e.g. carotenoids)) [14]. Both pathways lead to the formation of isopentenylpyrophosphate (IIP) and its allylic isomer dimethylallyl-pyrophosphate (DMAPP), the basic terpenoid biosynthesis building blocks (Fig. 8.1). The different monoterpenes and sesquiterpenes are generated through the action of terpenoid synthases.
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CO2 Photosynthesis 2(Acetyl-CoA) Acetyl-CoA acetyltransferase Acetoacetyl-CoA
HMG-CoA HMG-CoA Reductase
Citronellol, geraniol, nerol (C10)
Mevalonate MEP
Isopentenyl-pyrophosphate (IIP, C5 , isoprene)
DMAPP (C5)
Farnesol (C15)
Sesquiterpenes (C15) Triterpenes
Monoterpenes (C10)
Farnesyl-pyrophosphate (C15)
Sterols
Phytol (C20)
Diterpenes (C20)
Tetraterpenes (C40)
Geranyl-pyrophosphate (C10)
Squalene (C30)
Wax ester (phytoyl-farnesol) (C35)
Isoprenoid Wax ester (C40)
HMG-CoA Synthase
Farnesylated proteins
Geranyl-geranyl-P (C20) Ubiquinones
Geranylated proteins
Fig. 8.1 Biosynthetic pathways of acyclic terpenes/isoprenoids and biomolecules which contain them. The basic terpenoid biosynthesis building blocks isopentenyl-pyrophosphate (IIP) or dimethylallyl-pyrophosphate (DMAPP) are derived from mevalonate and the 2-C-methyld-erythritol 4-phosphate (MEP) pathways to produce monoterpenes, diterpenes, sequiterpenes, triterpenes. Additionally, the acyclic terpenes as citronellol, geraniol, and nerol are produced beginning from monoterpenes; or phytol from diterpenes until Wax esters. Adapted from [14, 15]
In Arabidopsis plants a set of 40 terpenoid synthases has been reported and is translated in the production of a high number of monoterpenes with diverse properties conferred to the plants [14]. The annual global emission by plants of volatile organic compounds such as isoprene, monoterpenes, and ethane has been estimated in the order of 1 billon tons [17].
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8.2.2 Terpenes/Isoprenoids as Petroleum Components The biodegradability of environmental pollutants, and hence the degree of persistence of contaminants in natural environments, is influenced by factors as the chemical structure, the presence of a viable microbial population able to degrade the contaminants, and environmental conditions suitable for microbial biodegradative activities. The most common pollutants of organic category concern to the petroleum hydrocarbons (a mixture of n-alkanes; other aliphatic; mono-, di-, and polyaromatic compounds; heterocyclic aromatics; and other minor constituents). A rough order of removal for aliphatic hydrocarbons (HC) has been established as follows (from most susceptible to least susceptible compounds): n-alkanes> alkylcyclohexanes> cyclohexanes> acyclic isoprenoid alkanes> bicyclic alkanes> C27–29 -steranes> C30–35 -hopanes> diasteranes> C27–29 -hopanes> C21–22 -steranes> tricyclic tepanes [18]. Linear, branched, and acyclic alkanes are saturated compounds that consist of carbon and hydrogen. Major sources of these alkanes are geochemical processes involving heat and/or pressure, resulting in the formation of crude oil from decaying plant and algal material. Depending on the source of crude oil, alkanes constitute about 20–50% of the petroleum, terpenes as petroleum components play an important role in its biodegradability, that will be influenced by the microorganisms involved in the degradation. In particular, the Pseudomonas genus has played a prominent role in oil and aliphatic hydrocarbons biodegradation [18].
8.2.3 Terpenes/Isoprenoids as Bacterial Components Isoprenoids are well suited as biological markers since they are often abundant and are widely distributed on earth. Also, the relatively stable isoprene skeletal unit is readily identified and allows compounds such as phytane and pristane to be used as tracers over long periods of geological time [19]. The C20 isoprenoid alcohol “phytol” produced from chlorophyll metabolism is generally considered as the most abundant acyclic isoprenoid compound in the biosphere. Neutral lipids biosynthesis is ubiquitous in nature and occurs in animals, plants, and microbes (Fig. 8.1). Microorganisms have been reported to synthesize triacylglycerols, polyhydroxyalkanoates, and wax esters (WEs). These lipids can be useful as carbon and energy storage under growth-limiting conditions. A class of WEs is produced by condensing long-chain isoprenoyl alcohol and isoprene fatty acid substrates (i.e. phytanoyl and farnesol or phytol esters) [15, Fig. 8.1]. It has been recently reported the formation of isoprenoid wax esters during aerobic growth of the marine bacteria Acinetobacter, Pseudomonas, and Marinobacter genus when grown on free phytol, pristane derivates, and farnesol widely distributed compounds in marine sediments; also when grown in environments where a carbon source as petroleum hydrocarbons and gluconate may be abundant over other nutrients [20, 15].
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8.3 Branched-Chain Alkane Degradation Alkanes, also known as paraffins, must be activated before they are degraded and used as carbon source. As alkanes are an excellent source of carbon and energy, many organisms have evolved the necessary mechanisms to exploit this resource. The first step in the aerobic degradation of these compounds is the introduction of oxygen atoms derived from molecular oxygen into the substrate. This critical step is carried out by oxygenases, enzymes that are highly interesting for bioremediation as well as for biocatalysis process [17]. Typically, alkanes are hydroxylated to the corresponding primary alcohol and further oxidized by alcohol and aldehyde dehydrogenases. Short-chain alkanes are metabolized via terminal, as well as by subterminal oxidation. The resulting fatty acids enter the β-oxidation cycle or are incorporated as cellular lipids [21]. In some cases, both ends of the alkane substrates are oxidized, which has been exploited for the production of dicarboxylic acids.
8.3.1 2-Methyl Branched-Chain Alkanes Degradation Degradation of 2-methyl branched-chain alkanes is generally represented by mechanism of pristine isoprenoid (2,6,10,14-tetramethyl-pentadecane) or the Wax ester degradation (composed by the ester of 6,10,14-trimethylpentadeca-2-one and phytol). In this case the mechanism of degradation involves the participation of several catabolic pathways as the alpha, beta and omega oxidations, producing metabolic intermediaries oxidized in the end of the molecules. The products are then hydrolyzed to compounds with two or three carbons less, and acetyl-CoA or propionyl-CoA are generated as final products (when the methyl radical is located in the α-carbon). Examples such as pristane catabolism by Brevibacterium erythrogenes [22] and catabolism of the 6,10,14-trimethylpentadecan-2-one and the Wax esters by marine bacteria as Acinetobacter sp., Pseudomonas nautical, and Marinobacter hydrocarbonoclasticus have been described [19, 20].
8.3.2 3-Methyl Branched-Chain Alkanes Degradation The acyclic terpenes are classified within the group of the 3-methyl branchedchain alkanes. These compounds are characterized by their low biodegradability in the environment. The main reason is the unability of the β-oxidation enzymes to hydrolyze compounds with methyl group in the beta-carbon (3-methyl radical) in contrast to 2-methyl branched chain alkanes which do not present this impediment. In consequence, these compounds show a high environmental persistence or recalcitrance. The strategy to degrade 3-methyl branched-chain alkanes (acyclic terpenes/isoprenoids) consists of an additional carboxylation enzymatic
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activity followed by the reaction known as beta-decarboxymethylation and concomitant oxidation [23]. This reaction consists in the modification of the methyl radical found in the beta-carbon, generally by a carboxylation reaction producing a beta-acetyl group and its subsequent hydrolysis as acetate; then occurs the elimination of the beta-methyl group rendering a substrate susceptible to be degraded by alpha or beta oxidation pathways. The prototype compounds that show these properties are the acyclic terpenes/isoprenoids such as citronellol (3,7-dimethyl-6-octen-1-ol), geraniol (3,7-dimethylocta-2,6-dien-1-ol) and farnesol (3,7,11-trimethyl-2,6,10-dodecatrien-1-ol) family. Microorganisms with the ability to degrade these compounds are principally related to Pseudomonas genus such as P. citronellolis, P. aeruginosa, P. mendocina, P. delhiensis and some isolates of P. fluorescens, as the Pf-5 strain [24].
8.4 Acyclic Terpenes Metabolism in Pseudomonas 8.4.1 Introduction to Acyclic Terpenes Catabolism Pioneering studies of acyclic isoprenoids degradation lead to the isolation of a strain of the Pseudomonas genus able to growth on citronellol, geraniol and farnesol as sole carbon source, was which named P. citronellolis [8]. Catabolism studies carried out by accumulation of metabolites, enzymatic and growth phenotypes, contributed with proposing the first acyclic terpenes/isoprenoids catabolic pathway [25, 9]. These early studies indicated that in P. citronellolis the acyclic isoprenoids pathway involves at least three unique enzymes: (i) the geranylCoA carboxylase (EC 6.4.1.5), which acts to activate the beta-methyl group of the substrate via carboxylation (specific only for cis-geranyl-CoA isomer), (ii) a hydratase, that converts the carboxylated product (isohexenyl-glutaconyl-CoA) to 3-hydroxy-3-isohexenylglutaryl-CoA, and (iii) an enzyme that was named as 3-hydroxy-3-isohexenylglutaryl-CoA lyase, which catalyzes the removal of the activated β-carboxymethyl group. The effect of these enzymes was described as the steps to replace a β-methyl substituent with a carbonyl oxygen, generating a suitable substrate for β-oxidation; in addition, it was suggested that possible identical reaction sequence occurs in other Pseudomonas species such as P. aeruginosa and P. mendocina [9]. Fall et al. [11] carried out the enzyme recruitment for the construction of Pseudomonas strains capable to utilize certain recalcitrant branched hydrocarbons by a combination of alkane and citronellol pathways, demonstrating that enzyme recruitment can provide a pathway for the degradation of otherwise recalcitrant branched hydrocarbons. Vandenbergh and Wright [12] identified a P. putida strain able to grow on citronellol or geraniol as sole carbon and energy sources, associating this ability to transmissible pSRQ50 plasmid. Although the identification and characterization of the enzymes related with the acyclic terpenes/isoprenoids catabolic pathway was proposed at the end of 1970 s, and genetic determinants associated to plasmids
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in 1983, the genes remained unidentified until 2004. Diaz-Pérez et al. [26], using the Himar1 transposon, identified in the P. aeruginosa PAO1 genome a gene cluster involved in acyclic terpenes catabolism which in first instance was named gny gene cluster (by geranyl-CoA metabolism), corresponding to the ORFs PA2010 to PA2016; these genes were later renamed as the liu gene cluster (by leucineisovalerate utilization). A homologous gene cluster, called the atu gene cluster (by acyclic terpene utilization) was identified additionally conformed by the ORFs PA2886 to 2893 [27, 28]. Complementary studies of these two research groups have contributed to the elucidation and knowledge of the genes and enzymes involved in the acyclic terpenes catabolism.
8.4.2 Acyclic Terpenes Catabolic Pathway The acyclic terpenes catabolic pathway in Pseudomonas has been described with detail in the P. aeruginosa and P. citronellolis species. It may be considered from a particular point of view as conformed by four steps: (i) upper oxidation-activation pathway, (ii) central acyclic terpenes pathway (ATU), (iii) β-oxidation coupling, and (iv) convergence with leucine/isovalerate pathway (LIU) (Fig. 8.2). 8.4.2.1 Upper Oxidation-Activation Pathway The upper pathway of the acyclic terpenes catabolism starts with the oxidation and activation of the terpenol molecule, that may be citronellol, geraniol or nerol compounds. In this step the terpenol is oxidized to the corresponding terpenal and terpenoic acid [9; 26–29, Fig. 8.3]. It has been suggested in these steps the participation of dehydrogenases for the oxidation of citronellol to citronellal, citronellic acid and geranic acid, respectively [30, 29], although this remains to be elucidated. Mutagenesis and protein identification in P. aeruginosa grown on citronellol suggest that the citronellol and citronellal dehydrogenases are probably encoded by the atuB or atuG genes; the proteins encoded by these genes show homology with shortchain dehydrogenases [29]. In P. aeruginosa the oxidation of geraniol to geranylate is a molybdenum-dependent step, catalyzed by enzymes different than those utilized for the oxidation of citronellol [31]. The respective acid derivative produced from dehydrogenation steps of acyclic terpenes (citronellic acid or geranic acid) is then activated by a successive reaction that involves the acyl-CoA synthase, producing the citronellyl-CoA or geranyl-CoA, respectively (Fig. 8.3). For these enzymatic steps there is not evidence for the encoding genes, however, informatics analysis suggests that the atuH gene encodes for the acyl-CoA synthase [29]. 8.4.2.2 Central ATU Pathway The next step for acyclic terpenes catabolism is the central pathway, whose enzymes involved are encoded in the atuRABCDEFGH gene cluster (Fig. 8.4). Starting with
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Acyl-CoA synthase
Upper oxidationactivation pathway
Terpenyl-CoA
Central acyclic terpenes pathway (ATU)
cis-geranyl-CoA CO2 acetate 7-methyl-3-oxo-6-octenyl-CoA
β-oxidation coupling
2 Acetyl-CoA 3-methylcrotonyl-CoA CO2
Convergence with leucine/isovalerate pathway (LIU)
Acetyl-CoA + Acetoacetate
Acetyl-CoA
Glyoxylate cycle
TCA cycle
Glyoxylate and TCA pathways
Fig. 8.2 General steps that comprise the acyclic terpenes/isoprenoids catabolic pathway in Pseudomonas species. Details are described in the text
citronellyl-CoA and trans-geranyl-CoA metabolites of the preceding step, these are oxidized or isomerized to cis-geranyl-CoA. This step involves the participation of the enzyme citronellyl-CoA dehydrogenase if the initial compound is citronellyCoA (Fig. 8.5). It has been proposed that citronellyl-CoA dehydrogenase is encoded by the atuD gene [24]. Studies using 2D gel electrophoresis showed that AtuD and the PA1535 ORF are expressed in cells grown on acyclic terpenes. Both AtuD (42.7 kDa) and PA1535 (42 kDa) recombinant proteins expressed in Escherichia coli and purified to homogeneity showed citronellyl-CoA dehydrogenase activity (Vmax 850 mU/mg) and high affinity to citronellyl-CoA (Km 1.6 μM); the enzymes were inactive with
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H Citronellal dehydrogenase
Citronellal
O–
Nerol dehydrogenase
Neral
O
Geranial dehydrogenase
Neral dehydrogenase
=
O
Geranylate
=
O
Nerylate
H
O–
=
Nerol CH 2-OH (cis-geraniol)
Geranial
O
=
=
Geraniol
Citronellyl-CoA
O
H
Geraniol dehydrogenase
S-CoA Citronellyl-CoA synthase
Citronellate
O
CH 2-OH
O
=
=
Citronellol dehydrogenase
Citronellol
O
=
O
CH2-OH
–
S-CoA Geraniyl-CoA synthase
Nerayl-CoA synthase
trans-Geranyl-CoA
O
=
Neryl-CoA S-CoA (cis-geranyl-CoA)
Fig. 8.3 Upper oxidation-activation pathway of acyclic terpenes step and putative enzymes. Adapted from [9; 26–29, 24]
atuR PA2885
atuA PA2886
atuB PA2887
atuC PA2888
atuD
atuE
PA288 9
PA289 0
PA2891
41 %
51 %
34 %
27 %
atuF
atuG PA289 2
atuH PA2893
46 %
liuR
liuA
PA2016
PA2015
liuB PA201 4
liuC PA201 3
liuD
liuE
PA2012
PA2011
Fig. 8.4 The atu and liu gene clusters of Pseudomonas aeruginosa PAO1. These genes are proposed to encode the enzymes involved in acyclic terpenes and leucine/isovalerate catabolic pathways, respectively. Gene names and its respective ORF’s of the PAO1 genome are shown. Amino acid identity of homologous proteins is indicated in percentage below the genes. Adapted from Aguilar et al. [28]
octanoyl-CoA, 5-methylhexen-4-enoyl-CoA and isovaleryl-CoA, suggesting a high substrate specificity for terpenoid molecule and essentiality for a functional ATU pathway. The PA1535 protein also revealed citronellyl-CoA dehydrogenase activity, although with significantly lower affinity than AtuD (Km 18 μM) and utilizing also octanoyl-CoA as substrate (Km 130 μM); it was concluded that the PA1535 ORF was not essential for ATU catabolism [24]. The next reaction is mediated by the geranyl-CoA carboxylase encoded by the atuC and atuF genes, a reaction that has been studied with detail and will described below. The following reaction of the central pathway is carried out by the isohexenylglutaconyl-CoA hydratase, converting the isohexenyl-glutaconyl-CoA produced by the geranyl-CoA carboxylase, to 3-hydroxy-3-isohexenylglutaryl-CoA (Fig. 8.5). This activity was referred since early studies by Seubert and Fass [25] and it was recently found that the enzyme is encoded by the atuE gene [26–28, 32, 29, 24]. However a mutant in the atuE gene did not show a deficiency of growth in terpenes, suggesting that the hydratase AtuE is important but not essential for growth on acyclic terpenes and that may be replaced by other isoenzymes [29]. The following reaction in the central pathway is carried out by the 3-hydroxy3-isohexenylglutaryl-CoA lyase (HIHG-CoA lyase), also called 3-hydroxy-
244
J. Campos-García O
=
AtuD/PA1535 S-CoA
Citronellyl-CoA O
= Geranyl-CoA isomerase
COO–
AtuC/AtuF O
S-CoA
trans-Geranyl-CoA
CO2
Citronellyl-CoA dehydrogenase
=
S-CoA
Geranyl-CoA Carboxylase (GCCase)
CO-S-CoA
Isohexenyl-glutaconyl-CoA
cis-Geranyl-CoA
CH3-COO-
Isohexenylglutaconyl-CoA hydratase
LiuE ? AtuA?
O
= CO-S-CoA
3-Hydroxy-3-isohexenylglutaryl-CoA Lyase (HIHG-CoA Lyase)
7-Methyl-3-oxo-6-octenyl-CoA
AtuE COO– OH
CO-S-CoA
3-Hydroxy-3-isohexenylglutaryl-CoA
Fig. 8.5 Central acyclic terpenes (ATU) pathway and enzymes involved. Metabolites from the upper oxidation-activation acyclic terpenes step converges in cis-geranyl-CoA compound that is carboxylated by the cis-geranyl-CoA carboxylase (GCCase). Hydratase and lyase reactions produce acetate and the 7-methyl-3-oxo-6-octenyl-CoA, substrate for β-oxydation pathway. Adapted from [25–28, 32, 29, 24, 33]
3-isohex-enylglutaryl-CoA:acetate-lyase. In early studies it was described that the activity of HIHG-CoA lyase occurs in Pseudomonas; isotopic marker assays and compounds identification suggested that the reaction products are free acetic acid and 7-methyl-3-oxo-6-octenyl-CoA [23, 25]. This metabolite is considered as the final product of the central ATU pathway. Although the gene and enzyme involved have not been characterized, it has been reported that the HIHG-CoA lyase could be encoded by the atuA gene [29]. On the other hand, a mutant strain in the liuE gene was unable to grow on both citronellol and leucine as sole carbon sources, suggesting that the liuE gene could encode for a bi-functional enzyme with both HMG-CoA lyase and HIHG-CoA lyase activities [28, 32, 33]. 8.4.2.3 β-Oxidation Coupling to the ATU Pathway Once that 7-methyl-3-oxo-6-octenyl-CoA is obtained by the central ATU pathway, it is integrated in the β-oxidation pathway [24, Fig. 8.6]. This compound is cleaved to 5-methylhex-4-enoyl-CoA and acetyl-CoA by a probable acyl-CoA thiolase activity of the complex of fatty acids oxidation (β-oxidation). Following, the 5-methylhex-4-enoyl-CoA is oxidized to (2E)-5-methylhex-2,4-dienoyl-CoA by the acyl-CoA dehydrogenase; afterwards it is hydrated by the enoyl-CoA hydratase to 3-hydroxy-5-methylhex-4-enoyl-CoA and then to the 5-methyl-3-oxo-4-hexenoylCoA metabolite by the 3-hydroxyacyl-CoA dehydrogenase, that is finally cleaved to 3-methyl-crotonyl-CoA metabolite plus acetyl-CoA by the acyl-CoA thiolase enzyme (Fig. 8.6). In this case, the 3-methyl-crotonyl-CoA is the convergent metabolic compound of the acyclic terpenes pathway with the leucine/isovalerate catabolic pathway [28, 32, 29, 24].
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8.4.2.4 LIU-Convengence to ATU Pathway In the LIU-convergence pathway are involved the enzymes encoded by the liuRABCDE gene cluster, required in leucine/isovalerate utilization which has been described in P. aeruginosa and P. citronellolis (Fig. 8.4). The leucine catabolic pathway involves first the transamination to 2-ketoisocaproic acid, and a second step yielding isovaleryl-CoA after a dehydrogenation reaction with the elimination of a CO2 molecule. Isovaleryl-CoA metabolite converges also from isovalerate catabolism by previously isovaleric acid activation by an acyl-CoA synthase (Fig. 8.7). The following step is a dehydrogenation to produce 3-methylcrotonylCoA (MC-CoA) catalyzed by the activity of the isovaleryl-CoA dehydrogenase, encoded by the liuA gene [24]. The MC-CoA produced is the metabolite of convergence on both leucine/isovalerate and acyclic terpenes catabolic pathways, which are constituted by four analogous reactions (dehydrogenation, carboxylation, hydration and cleaving) (Figs. 8.5 and 8.7). The carboxylation reaction is carried out by the 3-methyl-crotonyl-CoA carboxylase (MCCase) encoded by the liuB/liuD genes to produce 3-mehtyl-glutaconyl-CoA, which is converted to 3-hydroxy-3-methylgluratyl-CoA by the 3-methyl-glutaconyl-CoA hydratase, probably encoded by the liuC gene [28, 32, 29, 24]. Finally, the 3-hydroxy-3-methyl-glutaryl-CoA metabolite is cleaved to acetyl-CoA and acetoacetate by the 3-hydroxymethyl-glutaryl-CoA lyase (HMG-CoA lyase), which is encoded by the liuE gene [28, 32, 29, 24, 33]. Finally, the products acetate, acetyl-CoA and acetoacetate are channelled to central pathways as TCA and glyoxylate cycles (Fig. 8.2). Metabolites from both the acyclic terpenes and leucine catabolism converges in 3-methyl-crotonyl-CoA compound that is carboxylated by the 3-methyl-crotonyl-CoA carboxylase (MCCase). Hydratase and lyase reactions produce acetyl-CoA and acetoacetate as final products [25–28, 32, 29, 24, 33].
CoA-SH
Acetyl-CoA
O
=
=
O
FadA
=
O
FadE S-CoA
CO-S-CoA
S-CoA Acyl-CoA dehydrogenase
3-Ketoacyl-CoA thiolase
7-Methyl-3-oxo-6-octenyl-CoA
5-Methylhex-4-enoyl-CoA
(2E)-5-Methylhex-2,4-dienoyl-CoA
Enoyl-CoA hydratase Acetyl-CoA
CoA-SH
3-Ketoacyl-CoA thiolase
3-Hydroxyacyl-CoA dehydrogenase
O
S-CoA
5-Methyl-3-oxo-4-hexenoyl-CoA
FadN
OH
O
=
= S-CoA
3-Methyl-crotonyl-CoA
O
= =
FadA
O
FadB
S-CoA
3-Hydroxy-5-methylhex-4-enoyl-CoA
Fig. 8.6 Coupling β-oxidation to acyclic terpenes catabolic pathway. Putative enzymatic steps and proteins suggested were adopted of general fatty acid metabolism in bacteria. Adapted from Fujita et al. [21]
246
J. Campos-García NH3+
Transaminase
O Dehydrogenase
=
COO–
COO-
Leucine
2-Ketoisocaproate
=
COO-
Acetoacetate
3-Hydroxy-3-methylglutaryl-CoA Lyase (HMG-CoA Lyase)
LiuE
Isovaleryl-CoA
COOOH CO-SCoA
CO-SCoA
Isovaleryl-CoA Dehydrogenase
Isovaleryl-CoA synthase
Isovalerate
O
LiuA
CO-SCoA
3-Methyl-crotonyl-CoA
CO2
3-Methyl-crotonyl-CoA Carboxylase (MCCase) 3-methyl-glutaconyl-CoA Hydratase
LiuB/LiuD
COO CO-SCoA
LiuC
3-Hydroxy-3-methylglutaryl-CoA
3-Methyl-glutaconyl-CoA
Acetyl-CoA
Fig. 8.7 Leucine/isovalerate catabolic pathway (LIU) and putative enzymes encoded in the liu gene cluster involved. Adapted from [28, 32, 33, 29, 24]
8.4.3 Carboxylases Involved in Acyclic Terpene Catabolism 8.4.3.1 Common Properties of GCCase and MCCase Enzymes Key enzymes in both acyclic terpenes and leucine/isovalerate pathways are the geranyl-CoA carboxylase (GCCase) and 3-methylcrotonyl-CoA carboxylase (MCCase). These enzymes are encoded by atuC/atuF and liuB/liuD genes, respectively (Fig. 8.4). In SDS-PAGE the proteins showed relative molecular mass (Mr) for AtuC, AtuF, LiuB, and LiuD subunits of 63, 74, 63, and 78 kDa, respectively [28, 29, 31]. These carboxylases are made of two subunits: AtuC and LiuB are the β-subunits of GCCase and MCCase, respectively. These subunits show two main domains, the acyl-CoA-binding and the carboxybiotin-binding domains, involved in the transfer of a carboxyl group to the acyl-CoA substrate [34]. On the other hand, both AtuF and LiuD (α-subunits of these carboxylases) show four highly conserved domains in the acyl-CoA carboxylases family: (1) the ATP-binding site (GGGGKGM), (2) a CO2 fixation domain (RDCS), (3) the catalytic site of biotindependent carboxylase family (EMNTR), and (4) a biotin-carboxyl carrier domain (AMKM) [28, 32]. The optimal pH and temperature for both GCCase and MCCase enzymes are 8.5 and 37◦ C; similar values were reported for MCCases from mammalian, bacterial, and plant sources. Like MCCase from P. citronellolis, GCCase and MCCase from P. aeruginosa are inactivated by temperatures higher than 50◦ C [35]. 8.4.3.2 Kinetic Parameters of the GCCase When P. aeruginosa is grown on citronellol, both GCCase and MCCase enzymes are expressed. The kinetic parameters for both carboxylase activities were, K0.5 (affinity constant for non-Michaelis-Menten kinetics behavior) of 8.8 and 8.84 μM, Vmax of 627 and 591 nmol/min mg of protein for G-CoA and MC-CoA, respectively. Using G-CoA as substrate the enzymatic activity showed ATP and NaHCO3 dependence with a typical Michaelis-Menten kinetics for ATP, whereas with NaHCO3
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Metabolism of Acyclic Terpenes by Pseudomonas
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a sigmoidal kinetics was observed. The kinetic parameters for GCCase were Km 10 μM for ATP, and Vmax 423 nmol/min mg of protein; and for NaHCO3 were K0.5 1.2 μM, and Vmax 210 nmol/min·mg of protein [32]. Heterologous co-expression of AtuC/AtuF proteins showed both GCCase and MCCase enzymatic activities. In relation to the G-CoA substrate, this enzyme exhibited sigmoidal kinetics, but in relation to MC-CoA substrate it exhibited a Michaelis-Menten kinetics. The P. aeruginosa GCCase enzyme is able to utilize both G-CoA and MC-CoA as substrates (Fig. 8.8). The kinetic constants found with G-CoA substrate were K0.5 8.8 μM and Vmax 492 nmol/min·mg of protein; and with the MC-CoA substrate, Km 14 μM and Vmax 308 nmol/min·mg of protein. The catalytic efficiency indicated that the GCCase enzyme prefers G-CoA over MC-CoA as a substrate [32]. In P. citronellolis it was observed that GCCase is able to carboxylate about 15 different acyl-CoA substrates, including MC-CoA [36]. An interesting fact is that the plant GCCase shows a strict substrate preference, carboxylating GCoA but not MC-CoA [37]. This finding suggests that in Pseudomonas the GCCase may play a multifunctional role with participation in several catabolic pathways, behaving as a promiscuous enzyme. 8.4.3.3 Kinetic Parameters of Recombinant MCCase When P. aeruginosa is grown in leucine or isovalerate as sole carbon source the MCCase is expressed while the GCCase is absent. Under optimal conditions at pH 8.5 and 37◦ C, the MCCase enzyme showed a typical sigmoidal behavior with respect to MC-CoA [32]. The kinetic constants for MCCase from both recombinant and native P. aeruginosa were similar. For the recombinant enzyme, kinetic parameters were K0.5 9.8 μM and Vmax 279 nmol/min·mg of protein. On the other hand, the kinetic dependence on the ATP and NaHCO3 substrates showed a sigmoidal response, with apparent kinetic parameters for these substrates of K0.5 13 μM for ATP and Vmax 356 nmol/min·mg of protein; for NaHCO3 they were K0.5 0.8 μM and Vmax 178 nmol/min·mg of protein, suggesting an allosteric regulation of MCCase by ATP and NaHCO3 [32, Fig. 8.8]. The kinetic studies for both P. aeruginosa and P. citronelolis MCCases indicate that the enzyme specifically recognizes MC-CoA as its substrate and does not carboxylate the analogous substrate GC-CoA [32, 36]. A difference between the MCCases of P. aeruginosa and P. citronellolis (K0.5 9.8 μM over 43 μM) is the apparent 5-fold lower affinity for the substrate in the last bacterial species [32].
8.5 Regulation of the Atu and Liu Gene Clusters The atuR and liuR genes located upstream of the atuABCDFGH and liuABCDE gene clusters are suggested to function as the regulators of the expression of the atu and liu clusters, respectively [26, 29, Fig. 8.4]. Proteins encoded by both gene clusters have been reported that are expressed under citronellate or isovalerate growth conditions, suggesting that both clusters may be controlled in both metabolic pathway by a common regulation system. Expression assays revealed that the liu gene
248
J. Campos-García
A
3-Methylcrotonyl-CoA
300
Geranyl-CoA
0.012 0.010
300
0.008
200
0.006 0.004 0.002
100
250 0.05 0.04
200
0.03
1/v
Carboxylase Activity (nmol/min*mg)
Geranyl-CoA
400
1/V
(nmol/min*mg)
Carboxylase Activity
B
3-Methylcrotonyl-CoA 500
150
0.02 0.01
100
–0.1 –0.01
0.1 0.2 0.3 0.4 0.5 0.6
1/[S]
50
–0.10 -0.05 0.00 0.05 0.10 0.15 0.20
1/[S]
0 0
20
40
60
Acyl-CoA (µM)
80
0 100
0
20
40
60
80
100
Acyl-CoA (µM)
Fig. 8.8 Kinetic behavior of recombinantly produced P. aeruginosa GCCase and MCCase enzymes. Carboxylase activities of recombinant AtuC/AtuF (a) and LiuB/LiuD (b) proteins, purified and reconstituted. Insets show the Lineweaver-Burk analysis of carboxylase activities. Adapted from [32]
cluster was induced by acyclic terpenes (citronellol and citronellate) and repressed by glucose [26, 28, 27, 29]. The use of transcriptional fusions suggested that liu gene cluster was induced under leucine/isovalerate growth and also with citronellol, supporting that the liu genes participate in both catabolic pathways [28].
8.6 Atu/liu Homologous Clusters Among Pseudomonas Species Analysis of the genetic arrangement of the atu cluster revealed nine ORFs highly conserved in several Pseudomonas, namely atuRABCDEFGH (Fig. 8.9A). The sequential order of related genes was exactly the same for P. aeruginosa strains PAO1, PA14, and PA7, P. citronellolis, and P. mendocina. In P. fluorescens strains of Pf-5 and PfO-1 only seven and five homologous genes are found, respectively. In addition, potential atu clusters were also identified with high homology score and arrangement conservation in other bacteria as Hahella chejuensis KCTC 2396, Marinobacter aquaeolei VT8, Alcanivorax borkumensis SK2 and Acinetobacter baumannii AYE (Fig. 8.9A). Surprisingly in P. putida and P. syringae the homologous atu gene cluster were not found. This analysis confirms the fact that P. aeruginosa, P. citronellolis, and P. mendocina are able to utilize acyclic terpenes, as described above, and suggests that other bacterial species could also be able to use these compounds as carbon source. Additional phylogenetic analysis carried out for the AtuRABCDEFGH proteins and their homologous proteins from the liu cluster, LiuRABCDE, showed that the seven Pseudomonas strains which have been described to use acyclic terpenes were grouped in the same phylogenetic node (Fig. 8.9B, node 1). Other bacterial genus suggested to be able to use acyclic terpenes as carbon source were classified in a near but different node (Fig. 8.9B, node 2). While the LiuRABCDE proteins all were grouped in a distant root (Fig. 8.9B, node 3). This analysis clearly indicates
8
Metabolism of Acyclic Terpenes by Pseudomonas
A
atuR
atuA
atuB
249
atuC
atuD
PA2887
PA2888
PA2889
atuF
atuG
PA2890
PA14-2675
PA14-2673
PA14-2672
PSPA7-2268
PSPA7-2267
PSPA7-2266
DQ328853
atuE
PA2891
PA2892
atuH
PA14-2670
PA14-2669
PA14-2667
PA14-2665
PA14-2664
PSPA7-2265
PSPA7-2264
PSPA7-2263
PSPA7-2262
PSPA7-2261
DQ328855
DQ328856
DQ328857
DQ328858
Pmen-2692
Pmen-2693
Pmen-2694
Pmen-2695
Pmen-2696
PFL-4197
PFL-4198
PFL-4199
PAO1 PA2885
PA2886
PA14-2676
PSPA7-2269
PA2893
PA14 PA7 P. cit DQ767718
DQ328851
DQ328852
Pmen-2688
Pmen-2689
Pmen-2690
DQ328854
P. men Pmen-2691
Pf-5 PFL-4193
PFL-4194
PFL-4195
PFL-4196
PflO1_3946
PflO1_3947
PflO1_3948 PflO1_3949
PflO1_3950
PfO-1 H. chej HCH-05743
HCH-05745
HCH-05746
HCH-05748
HCH-05750 HCH-05751
HCH-05752
HCH-05753
HCH-05754
Maqu-2801
Maqu-2802
Maqu-2803
Maqu-2804
Maqu-2805 Maqu-2806
Maqu-2807
Maqu-2808
Maqu-2809
ABO-0990
ABO-0989
ABO-0988
M. aqu A. bor ABO-0992
ABO-0991
ABAYE0477
ABAYE0478
ABO-0987
ABO-0986
ABO-0985
A. bau ABAYE0479
ABAYE0480
ABAYE0481 ABAYE0482
ABAYE0483
node 1
node 2
node 3
90
80
70
60
50
40
30
20
10
(100 ) (99.8) (97.6) (85.1) (82.4) (72) (77) (64) (61.2) (62.3) (58.7) (79.7) (79.7) (79.7) (79.4) (77.7) (79.2) (80.1) (99.3) (100) (97.8)
LiuRABCDE
AtuPAO1 AtuPA7 AtuPA14 AtuP. cit AtuP.men AtuPf5 AtuPfO-1 AtuH.chej AtuM.aqu AtuA.bor AtuA.bau LiuPputF1 LiuPputKT LiuPent. LiuPf5 LiuPsyr LiuP.men LiuP.stu LiuPA14 LiuPAO1 LiuPA7
AtuRABCDEFGH
(%)
B
0
Amino acid substitutions (x100)
Fig. 8.9 Sequence analysis of products encoded in the atu and liu gene clusters. (a) Genetic arrangement of the atu homologous clusters from bacterial species compared to that of P. aeruginosa PAO1. (b) Phylogenetic tree of the Atu/Liu homologous proteins of bacterial species. The amino acid sequences of AtuRABCDEFGH (Atu) and LiuRABCDE (Liu) were joined and aligned using the ClustalW software and phylogenetic tree was carried out by Neighbour-Joining method. Identity between Atu and Liu PAO1proteins and homologous proteins are indicated in percentage. PAO1, Pseudomonas aeruginosa PAO1; PA14, P. aeruginosa PA14; PA7, P. aeruginosa PA7; P. cit, Pseudomonas citronellolis; P. men, Pseudomonas mendocina; Pf-5, Pseudomonas fluorescens Pf-5; PfO-1, P. fluorescens PfO-1; H. chej, Hahella chejuensis KCTC; M. aqu, Marinobacter aquaeolei; A. bor, Alcanivorax borkumensis SK2; A. bau, Acinetobacter baumannii AYE; PputF1, Pseudomonas putida F1; PputKT, P. putida KT2440; P.ent, Pseudomonas entomophila L48; Psyr, Pseudomonas syringae pv. syringae B728a; P.stu, Pseudomonas stutzeri A1501. Node 1, indicates Pseudomonas species with the ability of acyclic terpenes assimilation; node 2, indicates bacterial genus that could be able of acyclic terpenes assimilation; node 3, Pseudomonas species with the ability of leucine assimilation
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J. Campos-García
that both protein groups (encoded in the atu and liu gene clusters) belong to the same family and had evolved as two separated phylogenetic subfamilies. Interestingly, it was found that the LiuA, LiuB, LiuC and LiuD proteins from P. aeruginosa PAO1 are more similar to LiuABCD orthologous from the other Pseudomonas species (77–79%) than with the paralogous proteins AtuC, AtuD, AtuE and AtuF from P. aeruginosa (41.5%). In addition, phylogenetic analysis of AtuF, α-subunits of the GCCase, located this protein in a root with homologous proteins from the alpha-proteobacterias Bradyrhizobium japonicum and Rhodopseudomonas palustris, than itself from the Pseudomonadales located in the gamma-proteobacteria root. That finding suggests that the atu cluster from P. aeruginosa PAO1 probably was originated by horizontal transfer from bacteria of the Rhizobiales from the alpha-proteobacteria root [28]. Another interesting feature is that a liuE homologue gene is not present in the atu cluster or elsewhere in the PAO1 genome. This fact has led to suggest that LiuE may have a dual function, used in both the leucine and acyclic terpenes catabolic pathways [28, 32, 33].
8.7 Other Genes Involved in Acyclic Terpenes Catabolism Interestingly, other genes or proteins have been described to be related with acyclic terpenes utilization in P. aeruginosa and P. citronellolis; in most cases these genes are related to steps of metabolic pathways that indirectly have influence over acyclic terpenes metabolism. Some examples are: a four-gene cluster (the PA1339 to PA1342) that may function as an ABC transport system; this cluster was specifically expressed in cells grown on acyclic terpenes, and is probably involved in uptake of terpenes [38]. Additional genes expressed under acyclic terpenes growth conditions correspond to TCA (tricarboxylic acid cycle), whose function is involved in channeling final metabolites of acyclic terpenes pathway in the TCA and glyoxylate pathways have been identified such as: malate:quinone oxidoreductase (MqoB), glutamate synthase, acetyl-CoA-acetyltransferase, acetyl-CoA-acetoacetate transferase, and isocitrate lyase [39, 29, 40].
8.8 Conclusion Remarks The acyclic terpenes are ubiquitous organic compounds on Earth which may be synthetized as collateral products of the metabolism in plants, animals, and microorganisms. Pseudomonas is a bacterial genus able to metabolize the acyclic terpenes, and therefore it plays an important role in their biodegradation. Recent studies indicate that the acyclic terpenes metabolism involves the recruitment of several catabolic pathways that are mainly encoded in at least three genetic clusters. The acyclic terpenes metabolism has been described with detail in two bacterial species, P. citronellolis and P. aeruginosa. The two main gene clusters located in different genomic regions are the atuRABCDEFGH and liuRABCDE. These gene clusters encode the enzymes of the central acyclic terpenes pathway (ATU) and
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coupling with the leucine/isovalerate catabolism (LIU), respectively. Mutagenesis and enzymatic assays suggest that the atu and liu clusters products are essential for acyclic terpenes and leucine/isovalerate catabolism, confirming the metabolic relation between both pathways. Two of the best characterized enzymes are the geranyl-CoA carboxylase (GCCase) and the 3-methyl-crotonyl-CoA carboxylases (MCCase). Kinetic studies indicate that the GCCase enzyme is able to carboxylate both geranyl-CoA and 3-methyl-crotonyl-CoA substrates, while the MCCase enzyme prefer to carboxylate the 3-methyl-crotonyl-CoA substrate. In addition, the kinetic behavior of both GCCase and MCCase over the ATP and NaHCO3 co-substrates suggests that an allosteric regulation of the mechanism was present. Pseudomonas atu gene clusters from species with the ability to grow on acyclic terpenes (P. citronellolis, P. aeruginosa, and P. mendocina) were found grouped in a phylogenetic root, and probably other Pseudomonas species and other bacterial genera that their atu homologous genes were grouped in this subfamily are able to utilize them as carbon source. Protein alignment analysis showed that amino acid substitutions in ATU proteins were most abundant than in LIU proteins, suggesting an increased divergence in the atu gene cluster. Although the majority of genes involved in acyclic terpenes utilization have been characterized, some of them remain to be identified; i.e., genes involved in oxidation and activation of terpenols (citronellol, geraniol, nerol, farnesol), genes involved in the synthesis of acyl-CoA metabolites, and genes which direct the coupling with β-oxidation pathway. Acknowledgments Thanks to CONACYT (P-46547-Z) and C.I.C.-UMSNH (2.14).
References 1. Lugtenberg, B.J. and Weger, L.A. (1992) Plant root colonization by Pseudomonas spp, pp. 13–19. In E. Galli, S. Silver and B. Witholt (eds.), Pseudomonas molecular biology and biotechnology. ASM Press, Washington, D.C. USA. 2. Wagner, V.E. and Iglewski, B.H. (2008) Pseudomonas aeruginosa biofilms in CF infection. Clin. Rev. Allergy Immunol. 35: 124–134. 3. Kim, S.H., Park, S.Y., Heo, Y.J. and Cho, Y.H. (2008) Drosophila melanogaster-based screening for multihost virulence factors of Pseudomonas aeruginosa PA14 and identification of a virulence-attenuating factor, HudA. Infect. Immun. 76: 4152–4162. 4. Adonizio, A., Leal, S.M., Jr, Ausubel, F.M. and Mathee, K. (2008) Attenuation of Pseudomonas aeruginosa virulence by medicinal plants in a Caenorhabditis elegans model system. J. Med. Microbiol. 57: 809–813. 5. Starkey, M. and Rahme, L.G. (2009) Modeling Pseudomonas aeruginosa pathogenesis in plant hosts. Nat. Protoc. 4: 117–124. 6. Vila, J.,and Martínez, J.L. (2008) Clinical impact of the over-expression of efflux pump in nonfermentative Gram-negative bacilli, development of efflux pump inhibitors. Curr. Drug Targets 9: 797–807. 7. Anderson, G.G. and O’Toole, G.A. (2008) Innate and induced resistance mechanisms of bacterial biofilms. Curr. Top. Microbiol. Immunol. 322: 85–105. 8. Seubert, W. (1960) Degradation of isoprenoid compounds by microorganisms. Isolation and characterization of an isoprenoid-degrading bacterium, Pseudomonas citronellolis n. sp. J. Bacteriol. 79: 426–434. 9. Cantwell, S.G., Lau, E.P., Watt, D.S. and Fall, R.R. (1978) Biodegradation of acyclic isoprenoids by Pseudomonas species. J. Bacteriol. 153: 324–333.
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10. Prakash, O., Kumari, K. and Lal, R. (2007) Pseudomonas delhiensis sp. nov., from a fly ash dumping site of a thermal power plant. Int. J. Syst. Evol. Microbiol. 57: 527–531. 11. Fall, R.R., Brown, J.L. and Schaeffer, T.L. (1979) Enzyme recruitment allows the biodegradation of recalcitrant branched hydrocarbons by Pseudomonas citronellolis. Appl. Environ. Microbiol. 38: 715–722. 12. Vandenbergh, P.A. and Wright, A.M. (1983) Plasmid involvement in acyclic isoprenoid metabolism by Pseudomonas putida. Appl. Environ. Microbiol. 45: 1953–1955. 13. Taylor, R.F. (1984) Bacterial Triterpenoids. Microbiol. Rev. 48: 181–198. 14. Aharoni, A., Giri, A.S., Deuerlein, S., Griepink, F., de Kogel, W., Verstappen, F.W.A., Verhoeven, H.A., Jongsma, M.A., Schwab, W. and Bouwmeester, H.J. (2003) Terpenoid metabolism in wild-type and transgenic Arabidopsis plants. Plant Cell 15: 2866–2884. 15. Holtzapple, E. and Schmidt-Dannert, C. (2007) Biosynthesis of isoprenoid wax ester in Marinobacter hydrocarbonoclasticus DSM 8798: identification and characterization of isoprenoid coenzyme A synthetase and wax ester synthases. J. Bacteriol. 189: 3804–3812. 16. Phillips, M.A., Bohlmann, J. and Gershenzon, J. (2006) Molecular regulation of induced terpenoid biosynthesis in conifers. Phytochem. Rev. 5: 179–189. 17. Van Beilen, J.B. and Witholt, B. (2005) Diversity, function, and biocatalystic applications of alkane oxygenases, pp. 259–275. In B. Ollivier and M. Magot (eds.), Petroleum microbiology. ASM Press, Washington, D.C. USA. 18. Haiping, H. and Larter, S. (2005) Biodegradation of petroleum in subsurface geological reservoirs, pp. 1–121. In B.Ollivier, and M. Magot (eds.), Petroleum microbiology. ASM Press, Washington, D.C. USA. 19. Rontani, J.F., Gilewicz, M.J., Michotey, V.D., Zheng, T.L., Bonin, P.C. and Bertrand, J.C. (1997) Aerobic and anaerobic metabolism of 6,10,14- trimethylpentadecan-2-one by a denitrifying bacterium isolated from marine sediments. Appl. Environ. Microbiol. 63: 636–643. 20. Rontani, J.F., Bonin, P.C. and Volkman, J.K. (1999) Production of Wax esters during aerobic growth of marine bacteria on isoprenoid compounds. Appl. Environ. Microbiol. 65: 221–230. 21. Fujita, Y., Matsuoka, H. and Hirooka, K. (2007) Regulation of fatty acid metabolism in bacteria. Molec. Microbiol. 66: 829–839. 22. Pirnik, M.P., Atlas, R.M. and Bartha, R. (1974) Hydrocarbon metabolism by Brevibacterium erytrogenes: normal and branched alkanes. J. Bacteriol. 119: 868–878. 23. Pirnik, M.P. (1977) Microbial oxidation of methyl branched alkanes. Critical Rev. Microbiol. 5: 413–422. 24. Förster-Fromme, K., Chattopadhyay, A. and Jendrossek, D. (2008) Biochemical characterization of AtuD from Pseudomonas aeruginosa, the first member of a new subgroup of acyl-CoA dehydrogenases with specificity for citronellyl-CoA. Microbiology 154: 789–796. 25. Seubert, W. and Fass, E. (1964) Studies on the bacterial degradation of isoprenoids. V. The mechanism of isoprenoid degradation. Biochem. Z. 341: 35–44. 26. Diaz-Perez, A.L., Zavala-Hernandez, N.A., Cervantes, C. and Campos-Garcia, J. (2004) The gnyRDBHAL cluster is involved in acyclic isoprenoid degradation in Pseudomonas aeruginosa. Appl. Environ. Microbiol. 70: 5102–5110. 27. Hoschle, B., Gnau, V. and Jendrossek, D. (2005) Methylcrotonyl-CoA and geranyl-CoA carboxylases are involved in leucine/isovalerate utilization (Liu) and acyclic terpene utilization (Atu), and are encoded by liuB/liuD and atuC/atuF, in Pseudomonas aeruginosa. Microbiology 151: 3649–3656. 28. Aguilar, J.A., Zavala, A.N., Díaz-Pérez, C., Cervantes, C., Díaz-Pérez, A.L. and CamposGarcía, J. (2006) The atu and liu clusters are involved in the catabolic pathways for acyclic monoterpenes and leucine in Pseudomonas aeruginosa. Appl. Environ. Microbiol. 72: 2070–2079. 29. Förster-Fromme, K., Höschle, B., Mack, C., Armbruster, M.W. and Jendrossek, D. (2006) Identification of genes and proteins necessary for catabolism of acyclic terpenes and leucine/isovalerate in Pseudomonas aeruginosa. Appl. Environ. Microbiol. 72: 4819–4828.
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30. Campos-Garcia, J. and Soberon-Chavez, G. (2000) Degradtion of the methyl substituted alkene, citronellol, by Pseudomonas aeruginosa, wild type and mutant strains. Biotechnol. Lett. 22: 235–237. 31. Hoschle, B. and Jendroseek, D. (2005) Utilization of geraniol is dependent on molybdenum in Pseudomonas aeruginosa: evidence for different metabolic routes for oxidation of geraniol and citronellol. Microbiology 151: 2277–2283. 32. Aguilar, J.A., Díaz-Pérez, C., Díaz-Pérez, A.L., Rodríguez-Zavala, J.S., Nikolau, B.J. and Campos-Garcia, J. (2008) Substrate specificity of the 3-methylcrotonyl Coenzyme A (CoA) and geranyl-CoA carboxylases from Pseudomonas aeruginosa. J. Bacteriol. 190: 4888–4893. 33. Chavez-Aviles, M., Diaz-Perez, A.L., Reyes-de la Cruz, H. and Campos-Garcia, J. (2009) The Pseudomonas aeruginosa liuE gene encodes the 3-hydroxy-3-methylglutaryl-Coenzyme A lyase, involved in leucine and acyclic terpenes catabolism. FEMS Microbiol. Lett. 296: 117–123. 34. Kimura, Y., Miyake, R., Tokumasu, Y. and Sato, M. (2000) Molecular cloning and characterization of two genes for the biotin carboxylase and carboxyltransferase subunits of acetyl coenzyme A carboxylase in Myxococcus xanthus. J. Bacteriol. 182: 5462–5469. 35. Hector, M.L. and Fall, R.R. (1976) Multiple acyl-coenzyme A carboxylases in Pseudomonas citronellolis. Biochemistry 15: 3465–3472. 36. Fall, R.R. (1981) 3-Methyl-crotonyl-CoA and geranyl-CoA carboxylases from Pseudomonas citronellolis. Methods Enzymol. 71: 791–799. 37. Guan, X., Diez, T., Prasad, T.K., Nikolau, B.J. and Wurtele, E.S. (1999) Geranoyl-CoA carboxylase: a novel biotin-containing enzyme in plants. Arch. Biochem. Biophys. 362: 12–21. 38. Förster-Fromme, K. and Jendrossek, D. (2008) Biochemical characterization of isovalerylCoA dehydrogenase (LiuA) of Pseudomonas aeruginosa and the importance of liu genes for functional catabolic pathway of methyl-branched compounds. FEMS Microbiol. Lett. 286: 78–84. 39. Förster-Fromme, K. and Jendrossek, D. (2006) Identification and characterization of the acyclic terpene utilization gene cluster of Pseudomonas citronellolis. FEMS Microbiol. Lett. 264: 220–225. 40. Diaz-Perez, A.L., Roman-Doval, C., Diaz-Perez, C., Cervantes, C., Sosa-Aguirre, C.R., LopezMeza, J.E. and Campos-Garcia, J. (2007) Identification of the aceA gene encoding isocitrate lyase required for the growth of Pseudomonas aeruginosa on acetate, acyclic terpenes and leucine. FEMS Microbiol. Lett. 269: 309–316.
Chapter 9
Heavy Metal Resistance in Pseudomonads Esther Aguilar-Barajas, Martha I. Ramírez-Díaz, Héctor Riveros-Rosas, and Carlos Cervantes
9.1 Introduction The metabolic diversity of the genus Pseudomonas (and related bacterial species, called collectively pseudomonads) has attracted researchers to study this versatile microbial group. The ability to thrive in hostile environments, aided by a notable capacity to degrade or tolerate a wide variety of natural and synthetic compounds, results from the possession of highly adapted genomes. About 25 genomes from pseudomonad strains have been sequenced to date, representing eight different species from varied habitats. Genomic analyses confirm that pseudomonads evolved complex enzymatic strategies, delicate genetic regulatory switches, and efficient transport systems, to keep pace in ever-changing environments. These adaptive mechanisms include those conferring resistance to toxic compounds such as antibiotics and the ions derived from heavy metals and metalloids. Transport systems able to actively efflux metal ions out from the cytoplasm or the periplasmic space are a key strategy to withstand heavy metal toxicity. As with other bacteria, heavy metal resistance genes in pseudomonads may reside either in the chromosome or within plasmids. In this chapter, the information concerning the strategies used by pseudomonads to tolerate heavy metals is summarized. It should be noted that some of these resistance mechanisms have been assigned to pseudomonads only by the finding of homologous genes and operons when compared with characterized genes from sequenced genomes of different bacteria. In other cases, biochemical evidence for specific heavy metal resistance systems has been directly provided by the analysis of genes from pseudomonads. A first compilation on heavy metal resistance mechanisms in pseudomonads appeared almost two decades ago [1], but an overwhelming amount of information has accumulated since, notably by the advent of the genomic era. More recently, a monograph book covered the interactions of heavy metals with the wider microbial C. Cervantes (B) Instituto de Investigaciones Químico-Biológicas, Universidad Michoacana, Morelia, México e-mail:
[email protected]
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world [2]; reference to specific chapters will be given below for studies related to pseudomonads. For this review, the heavy metal protagonists have been divided into three main groups: (i) micronutrient cations (copper, cobalt, nickel and zinc), (ii) toxic cations (cadmium, lead and mercury), and (iii) toxic oxyanions (arsenate/arsenite, chromate and tellurite). A final recount includes other less-studied toxic ions (silver, tin, selenium) for which some information exists in pseudomonads. For each case, a brief account on the metal(loid) toxicity mechanisms is followed by a description of the resistance strategies reported in pseudomonads or (when missing) the best-studied systems uncovered in related bacteria. A scrutiny of the genomes of Pseudomonas aeruginosa [3, 4] and Pseudomonas putida [5, 6] already showed the presence of numerous metal resistance determinants, including members of the main transporter families: resistance-nodulation-cell division (RND), cation diffusion facilitator (CDF), major facilitator superfamily (MFS), and P-type ATPases able to efflux toxic metal cations or oxyanions. Regulatory systems for bacterial heavy metal resistance, as for other adaptive strategies, are of paramount importance for cell economy. Expression of the corresponding genetic determinants is subjected to delicate control mechanisms, commonly acting at the transcriptional level [7; reviewed in 8]. These regulatory systems will be mentioned in this review but, for space reasons, will not be detailed.
9.2 Micronutrient Cations As most living organisms, pseudomonads require the essential micronutrient cations derived from copper, cobalt, nickel and zinc, used mainly as enzyme cofactors and regulatory effectors. For these purposes, divalent cations form complexes with diverse ligands within the cells. Higher concentrations of these transition metals, however, may exert toxic effects on most cells as harmful complexes may be formed with varied biomolecules. This dual behavior has made it necessary for bacteria to develop strict homeostasis mechanisms in order to avoid metal toxicity, while allowing intracellular basal levels of the essential ions. Homeostasis commonly includes transmembrane uptake and efflux systems that carefully regulate intracellular cation levels. This review emphasizes on those pseudomonad systems devoted to tolerate the noxious effects of toxic divalent cations, not considering the physiological mechanisms for micronutrient acquisition and use. Also, systems for the homeostasis of essential but almost not toxic cations (i.e. iron and manganese) will not be considered.
9.2.1 Copper Copper is an essential metal, mainly required by aerobic cells as a cofactor for electron transport and redox enzyme systems [9]. Copper exists in the cytoplasm in the Cu(I) reduced state, being its ability to undergo redox Cu(II) to Cu(I) transformations partly responsible of its toxic properties. Additional toxicity effects derive
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from the ability of copper to displace other metals (i.e. Ni2+ , Co2+ , Mn2+ , Zn2+ ) from essential complexes as well as to unspecifically bind to biomolecules [10]. Bacterial copper transport and homeostasis has been widely studied in Escherichia coli and in Gram-positive Enterococcus hirae (reviewed in [9, 11–15]) and will not be treated here. Some copper resistance systems related to pseudomonads will be next described. The copABCD operon from Pseudomonas syringae plasmid pPT23D was one of the first bacterial copper-resistance systems analyzed [16]. pPT23D was found in a copper-resistant P. syringae pv. tomato strain isolated from copper-treated tomato fields [17]. Unlike other cation resistance mechanisms, the cop operon encodes a copper-sequestering system that prevents copper ions from entering the cytoplasm (Fig. 9.1A). CopA and CopC are periplasmic copper-binding proteins able to capture 11 and 1 copper atoms per polypeptide, respectively [18]. CopA also displays
Fig. 9.1 Mechanisms of resistance to essential cations in pseudomonads. A, Cop copper binding system. B, P-type ATPase CopA and RND complex CusCBA. C, nickel/cobalt resistance systems. RND complexes CnrCBA and NccCBA; MFS transporters RcnA and NreB. D, zinc metallothioneins (MT). E, zinc resistance systems. P-type ATPase ZntA and CDF transporter ZitB. P-type ATPases domains shown are: P, phosphorylation domain; N, nucleotide-binding domain; A, activator and phosphatase domain. The functions of the resistance systems are described in the text. Note that CopA in A is a periplasmic multi-copper oxidase of the COG2132 protein family whereas CopA in B is a P-type ATPase of the COG2217 protein family. These two non-homologous proteins received the same name
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a multi-copper oxidase activity, transforming Cu(I) to Cu(II), similar to that of CueO from E. coli [19] and may protect periplasmic enzymes from copper damage. CopC is probably a chaperone which delivers copper to CopD, an inner membrane protein with eight transmembrane segments. Outer membrane protein CopB also binds Cu2+ and is proposed to function in concert with CopD in Cu2+ uptake [20]. Copper-inducible expression of the cop operon is regulated by a chromosomallyencoded repressor and by a plasmid-borne two-component CopR/S system [21]. Chromosomal homologs of CopA and CopB have been identified in many pseudomonads [22], whereas CopC and CopD are less common and seem to be auxiliary determinants for optimal copper resistance. An additional gene, transcribed from a different promoter, encodes the small periplasmic CopE protein, which is related to PcoE from E. coli and to SilE from the Salmonella silver-resistance operon; as PcoE, CopE seems to bind Cu(I) and may function as a copper chaperone [9]. The other pseudomonad cop genes also show sequence similarity with the corresponding genes from the plasmid-mediated E. coli copper-resistance pco operon, although the latter system catalyzes the efflux of copper rather than its binding [23]. Other potential copper-resistant determinants studied in enterobacteria have been identified from the sequenced genomes of pseudomonads. For example, the genomes of P. aeruginosa [3] and P. putida [6] possess homologs of the CusCBA system, a proton-driven RND transporter which effluxes Cu+ (and Ag+ ) from the cytoplasm [24] (Fig. 9.1B), and of SilP, a P-type ATPase which extrudes Ag+ (and probably Cu+ ) [25]. Also, a homolog of the widespread P-type ATPase CopA [26], able to efflux copper from the cytoplasm, has been located in the P. aeruginosa genome [27] (Fig. 9.1B). A transcriptomics analysis of P. aeruginosa PAO1 showed that a P-type ATPase (ORF PA3920), three RND transporters (PA1436, PA2520, and PA3522), and two CDF family members (PA0397 and PA1297) were up-regulated in response to copper exposure [28]. One of the RND determinants encodes the czrCBA system [29], mentioned below because it confers Cd2+ , Zn2+ and Co2+ resistance by an efflux mechanism. The CDF systems encode homologs of the CzcD and RzcB transporters which confer resistance to divalent cations in other bacteria [30]. It is possible that some of these systems also efflux copper. These findings confirm that efflux systems constitute a major strategy for copper homeostasis as well as a main protection barrier for pseudomonads against copper toxicity.
9.2.2 Cobalt and Nickel Cobalt and nickel are similar transition metals of oxidation state II. They play essential roles for microorganisms as cofactors for many diverse metalloenzymes. Thus, uptake and homeostasis systems for these micronutrient divalent cations must exist in all bacteria [31–33]. Bacterial Co2+ and Ni2+ homeostasis systems have been studied with great detail in E. coli and to a lesser extent in other microorganisms [33]. As with copper, high levels of Co2+ and Ni2+ may exert toxic effects on microorganisms [23, 34]. The main mechanism of Co2+ and Ni2+ toxicity probably relates to their potential interference with iron (and possibly manganese) homeostasis. As
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for most divalent metal cations, the main tolerance bacterial strategies to cope with excess Co2+ and Ni2+ are usually associated with membrane efflux systems. Cobalt and nickel resistance systems have not been studied directly in pseudomonads. However, the identification of homologous genes for metal cation resistance in the genomes of species of Pseudomonas indicates that these bacteria have the potential to display tolerance mechanisms against Co2+ and Ni2+ . Co2+ resistance is usually accompanied by resistance either to Ni2+ , Cd2+ , or Zn2+ . Two RND systems from megaplasmids of Cupriavidus metallidurans (previously named Alcaligenes eutrophus and Ralstonia metallidurans), the cnrCBA and nccCBA operons, confer resistance to both Co2+ and Ni2+ [4, 35, 36]. The CnrCBA system from plasmid pMOL28 is formed by the three typical RND polypeptides: CnrA, an inner membrane transporter, CnrC, located in the outer membrane, and CnrB, a membrane fusion protein bridging the periplasmic space [37]. This tripartite complex functions as a chemiosmotic pump driven by the proton-motive force that effluxes the cations probably from the cytoplasm to the periplasm and then to the outside (Fig. 9.1C). Additional cnrYXH genes regulate the expression of the efflux pump [38, 39]. The NccCBA complex is structurally and functionally similar to CnrCBA and is also regulated by corresponding nccYXH genes [40] (Fig. 9.1C). Unlike Cnr, the Ncc system, besides Co2+ and Ni2+ resistances, also confers resistance to Cd2+ . The CzcCBA complex from plasmid pMOL30 of C. metallidurans, the first characterized RND system related to heavy metals, confers resistance to Cd2+ , Co2+ and Zn2+ [4, 41, 42] and will be described below in the cadmium section. A variant of this system, the Czn complex from Helicobacter pylori, has a distinct substrate specificity, exporting Cd2+ , Zn2+ and Ni2+ [43]. Also located in the Czc determinant is CzcD, a member of the CDF family, originally reported as a regulatory protein [44] but later found to confer low resistance to Co2+ , Cd2+ and Zn2+ [30]. Similar CDF transporters related to cation efflux, DmeF and FieF, have been identified in the C. metallidurans chromosome [45]. An interesting interplay between the Czc/Cnr RND systems and CDF proteins has been reported. CDFs seem to first export the cations from the cytoplasm to the periplasm and then RNDs pump them from the periplasm to the outside [45– 47]. Transporters of the MFS group have been also assigned functions in Co2+ or 2+ Ni efflux. This includes the first MFS protein found to be involved in metal transport, NreB of Achromobacter xylosoxidans, only transporting Ni2+ [48], and RcnA from E. coli which effluxes Co2+ and Ni2+ [49] (Fig. 9.1C). NreB and RcnA are histidine-rich polypeptides displaying a distinct topology of 12 and six transmembrane segments, respectively. The RcnR repressor regulates the expression of RcnA [50]. RcnA has been found to be also controlled by the global regulator Fur and was proposed to function as a connector of cobalt, nickel and iron homeostasis [51]. The P-type ATPase ZntA from E. coli, which confers cation resistance by the efflux of Zn2+ , Cd2+ and Pb2+ (see the sections of these metals below), is stimulated by Co2+ and Ni2+ [52] and may also efflux these ions although with little efficiency. The genomes of P. aeruginosa [3] and P. putida [6] contain structural and regulatory czc genes (two copies in P. putida) which are probably involved in the efflux of
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Co2+ and other divalent cations. Accumulation of Ni2+ as a resistance strategy has been reported in strains of P. aeruginosa [53] and P. putida [54], but the mechanisms involved have not yet been studied.
9.2.3 Zinc A widely distributed enzyme cofactor, zinc displays affinity for ligands possessing oxygen, nitrogen or sulfur. As mentioned for copper, toxicity of zinc is associated with its ability to replace other metals (i.e. Ni2+ , Co2+ , Mn2+ ) from enzymes or by forming complexes with other biomolecules. Zinc homeostasis has been studied with detail in several bacterial species [55]. Zinc occurs naturally as the divalent cation Zn2+ and the level of the metal is regulated by processes of Zn2+ uptake, sequestration by metallothioneins (MT), and efflux from the cytoplasm [56]. The pumps of Zn2+ efflux are usually not restricted to Zn2+ as a substrate, and may also catalyze transport of other divalent cations [56]. Zn2+ is exported across the cytoplasmic membrane by the P-type ATPase ZntA, described in E. coli [57], and by its closest homologue, CadA, first described in Staphylococcus aureus [58]. ZntA was the first example described of a specific Zn2+ transporting protein in E. coli [57], but now is known to transport a broad range of soft metal ions, including Cd2+ , Pb2+ , Ni2+ , Co2+ and Cu+ [59] (Fig. 9.1E). ZntA is a protein of 732 aminoacid residues with all the characteristics of a soft metal ion-translocating P-type ATPase, which include a cysteine-rich hydrophilic amino-terminal region that contains a single metal-binding motif GMDCAA C [56]. ZntA is regulated by ZntR (the zntR gene is located in another region of the chromosome in E. coli), that belongs to the MerR family of regulators. The expression of ZntA is induced by Zn2+ , Cd2+ and Pb2+ , being Cd2+ the more effective inducer [60, 61]. ZntR functions as a dimeric protein and tightly binds to its cognate promoter, PzntA , located upstream of the zntA start codon [56]. A well-characterized system of Zn2+ transport is the CzcCBA complex of C. metallidurans [4], described below in the cadmium section. Another protein that has been associated with zinc resistance is ZitB, a CDF transporter that mediates efflux of Zn2+ in E. coli [62] (Fig. 9.1E). ZitB is closely related to CzcD that transports Cd2+ , Zn2+ and Co2+ [30] (described in the cadmium section). The expression of the zitB gene leads to a significant increase in Zn2+ resistance and to reduced Zn2+ accumulation in zntA-disrupted E. coli cells [62]. It has been proposed that ZitB contributes to Zn2+ homeostasis at low concentrations of zinc, while ZntA is required for growth at higher concentrations [62]. Zinc-regulated genes have been analyzed in Pseudomonas fluorescens employing mutagenesis [63]. One of the genes identified was a zntA-like gene that was inducible by the presence of Zn2+ , Cd2+ , Pb2+ , Ni2+ , Hg2+ , and Ag+ ions. A mutant in this gene exhibited only hypersensitivity to Zn2+ , Cd2+ and Pb2+ , suggesting that it encodes a transporter for these cations. The P. putida strain S4 employs a dual strategy for zinc resistance [64]. One strategy is mediated by an inducible ATPase that effluxes the ion during the exponential phase of growth. The second mechanism is the accumulation of Zn2+ that can be stored by proteins in the outer membrane and the periplasm.
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In addition to membrane transport pumps, some bacteria produce metallothioneins (MT) [15]. MTs are small poly-thiol proteins that bind metal cations, lowering their free concentrations within the cytoplasm (Fig. 9.1D). The best characterized prokaryotic MT is SmtA from Synechococcus PCC 7942, which protects against elevated levels of Zn2+ [65, 66]. For a long time SmtA was the only prokaryotic MT identified [67], but currently other related bacterial MTs, called BmtA, have been described [68]. MTs were purified from P. putida and P. aeruginosa strains and found that they were associated with three to four Zn2+ atoms [68]. Additional BmtA-like proteins were identified in P. fluorescens strains pf01 and SBW25. Most bacterial MTs identified to date have been found in cyanobacteria and pseudomonads [68].
9.3 Toxic Cations This group of elements includes heavy metals with no known biological function and clear toxic effects over living cells: cadmium, lead and mercury. Potentially toxic metals which are irrelevant in biological terms, mainly by their presence at very low levels or in non available forms in most environments (i.e. gold, thallium, aluminum), will not be considered in this review. As with the essential cations, transmembrane efflux systems are also used by bacteria as key resistance mechanisms against toxic metal cations. Mercury represents a unique case for which intracellular sequestering followed by detoxification has evolved as a best suited bacterial tolerance strategy.
9.3.1 Cadmium and Lead Cadmium chemistry is closely related to that of essential zinc. Cadmium and lead commonly form cations of oxidation states II, although lead may also exist in the IV valence. Lead differs from cadmium and zinc in their chemical coordination properties. In contrast to zinc, cadmium and lead bind preferentially sulfur (soft) ligands. Due to their similarity, zinc homeostasis and cadmium and lead resistance mechanisms often overlap, as reflected by their sharing of uptake and efflux transporters and metal-responsive regulatory proteins [59]. Intracellular Cd2+ is maintained at low levels through the control of sequestration or efflux of the ion. Cd2+ can be effluxed from bacterial cells by at least three systems: the P-type ATPase CadA, a large single polypeptide, the CzcCBA system, a three-polypeptide chemiosmotic RND complex that functions as an ion/proton exchanger, and CzcD, a single CDF membrane protein acting as a chemiosmotic efflux pump [15]. The P-type ATPase CadA from pI258 plasmid of S. aureus is the most studied Cd2+ resistance system [58]. CadA homologs have been found in several bacterial species, including pseudomonads. The system is localized in the cadAC operon. CadA catalyzes the active efflux of Cd2+ , Zn2+ , or Pb2+ [58, 69], and contains all the characteristic domains of a P-type ATPase [70, 71] (Fig. 9.2A). CadC is a transcriptional regulator needed for full Cd2+ and Zn2+ resistance in S. aureus [72].
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Fig. 9.2 Mechanisms of resistance to toxic cations in pseudomonads. A, cadmium P-type ATPase CadA. B, multication RND complex CzcCBA. C, CDF transporter CzcD. D, cadmium pseudothioneins (PT). E, Pbr lead resistance system. F, mercury resistance Mer system. G, organomercurial resistance MerB enzyme. P-type ATPases domains shown are: P, phosphorylation domain; N, nucleotide-binding domain; A, activator and phosphatase domain. The functions of the resistance systems are described in the text
The best studied homologue of CadA, the above mentioned ZntA from E. coli, is 30% identical to CadA. The level of resistance to Cd2+ varies widely within species of Pseudomonas [73, 74]. P. putida 06909 possesses the cadA and cadR genes that are homologs to zntA and zntR from E. coli, respectively. CadA from P. putida 06909 confers a high level of resistance to Cd2+ , partial resistance to Zn2+ , and, unlike ZntA, does not confer Pb2+ resistance [75]. The level of Cd2+ resistance conferred by the cadA gene in P. putida 06909 is 17-fold higher than that conferred by zntA in E. coli. Homologous ORF’s PA3690 (cadA) and PA3689 (cadR) are also present in the genome of P. aeruginosa PAO1, but their function has not been elucidated yet. CadA sequences from P. putida and P. aeruginosa both have a histidine-rich N-terminal extension that is missing in other CadA sequences; this region is probably responsible for the higher level of resistance to Cd2+ of these strains [75]. CadR is a transcriptional regulator of the MerR family [76]. CadR represses its own
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expression in the absence of Cd2+ , it is induced in the presence of Cd2+ , and is necessary for full resistance to Cd2+ and Zn2+ [75]. P. putida KT2440 possesses the CadA1 and CadA2 transporters. CadA2 confers resistance to Cd2+ and Pb2+ in P. putida whereas CadA1 does not seem to confer metal tolerance in P. putida, but confers Zn2+ resistance when overexpressed in E. coli. CadA1 expression is inducible by Zn2+ . CadA2 is considered a housekeeping resistance mechanism against Cd2+ and Pb2+ [77]. CadA2 is constantly expressed at a high level even when Cd2+ is absent, but its expression increases in the presence of metals. The second Cd2+ resistance determinant, the CzcCBA system, actively transports Cd2+ , Zn2+ and Co2+ out of the bacterial cell [15] (Fig. 9.2B). One of the best characterized systems is the czc determinant from C. metallidurans [36]. The system is organized like other three-component RND transporter complexes. CzcA is a cation/proton antiporter located in the cytoplasmic membrane that effluxes toxic cations to the periplasm [78]. CzcA is essential for cation transport and is considered the core of the complex. CzcB is a membrane fusion protein which spans the periplasmic space, bringing the outer and inner membranes in close position. The third component, CzcC, is an outer membrane protein that effluxes cations from the periplasm to the outside [59]. The CzcCBA system catalyzes the efflux of both toxic and essentials cations and, for that reason, is tightly regulated by downstream and upstream regulatory regions. Czc homologues have been detected by Southern hybridization in several Pseudomonas strains, including P. aeruginosa PAO1 [29]. The system is annotated as czcCBA-like in the PAO1 strain genome and probably confers resistance to Zn2+ , Cd2+ , and Co2+ [3]. In the environmental isolate P. aeruginosa CMG103, the czrSRCBA gene cluster confers a high level of resistance to Cd2+ and Zn2+ [29]. In P. aeruginosa PT5 (a PAO1 derivative) cross-resistance between heavy metal and antibiotic pumps has been reported [74]. The two-component system CzcS-CzcR controls the expression of the Czc efflux pump and also regulates negatively the expression of the OprD porin, leading to carbapenem resistance. A czcCBA system is also functional in P. putida KT2440, which possesses two copies of the transporter [77]. CzcCBA1 confers Zn2+ resistance and its expression is induced by the metal; Cd2+ , and possibly Pb2+ , are also transported by CzcCBA1, but is less efficient when it acts as a Cd2+ or Pb2+ transporter. CzcCBA2 also confers Zn2+ resistance, but its expression is not induced by any metal. The third Cd2+ resistance determinant, CzcD, is an efflux pump that belongs to the CDF protein family [30] (Fig. 9.2C). The function of CzcD has been analyzed only in C. metallidurans but there are homologs in the genomes of P. aeruginosa PAO1 [3] and P. putida KT2440 [6]. CzcD is located in the cytoplasmic membrane and possesses at least six transmembrane helices. The level of resistance to Cd2+ , Zn2+ and Co2+ conferred by CzcD is lower as compared to that conferred by CzcA. CzcD is also involved in the regulation of the expression of the CzcCBA efflux system. A distinct cadmium resistance mechanism reported in Pseudomonas involves cadmium-binding proteins called pseudothioneins (PT) [79] (Fig. 9.2D). PTs have been identified in a P. putida strain adapted to grow in 3 mM cadmium.
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Pseudothioneins CdPT1, CdPT2 and CdPT3 are synthesized in different growth phases, being CdPT1 the major protein produced during the exponential phase. As metallothioneins, PTs are small cysteine-rich proteins (3.5–7 kDa). PTs have a lower cysteine content than mammalian metallothioneins (12–23% compared to 33%), but have in common that bind Cd2+ , Zn2+ and Cu2+ [79]. ZntA of E. coli and CadA of S. aureus also confer Pb2+ resistance in E. coli cells expressing the corresponding genes. A mutant strain with a disruption of the zntA gene showed hypersensitivity to Pb2+ ; expression of cadA from pI258 plasmid complemented the phenotype, indicating that cadA also confers Pb2+ resistance [69]. The only specific mechanism of Pb2+ resistance described so far is the pbr system, reported in C. metallidurans CH34 [80] (Fig. 9.2E). In contrast to the cad and znt systems, which only comprise the ATPase with a regulatory gene, the pbr system is constituted by several genes arranged in the divergent operons pbrUTR and pbrABCD. These operons encode proteins involved in three different processes: uptake, efflux and accumulation of lead, which together confer maximal Pb2+ resistance. The role of PbrU on lead resistance has not been elucidated. PbrT is a permease that takes up Pb2+ ; expression of pbrT alone in the absence of the pbrABCD genes results in Pb2+ hypersensitivity, due to an increase in Pb2+ uptake. The pbrR gene encodes the PbrR repressor that belongs to the ArsR/SmtB family of regulators. PbrR controls transcription of the pbr structural genes. PbrA is an inner membrane P-type ATPase, closely related to the CadA and ZntA ATPases, that effluxes Pb2+ to the periplasm. PbrB is an outer membrane lipoprotein that probably functions in removing Pb2+ from the periplasmic compartment, assisting PbrA for lead resistance [80, 81] (Fig. 9.2E). PbrC is probably an aspartic peptidase that removes the signal peptide from PbrB before it is transported to the periplasmic space; PbrC is required with PbrB for full resistance [80, 81]. PbrD is an intracellular protein that may bind Pb2+ with a cysteine-rich metal-binding motif but is not essential for Pb2+ resistance [80]. The Pbr system has not yet been identified in Pseudomonas strains. In the P. putida KT2440 genome, of a total of 61 open reading frames with a putative role in metal homeostasis and detoxification, there seems not to be homologues to pbr genes [6].
9.3.2 Mercury Mercuric ions (Hg2+ ) display a rather strong affinity for sulfur-containing ligands, thus its toxicity relates mostly to protein sulfhydryl poisoning and to binding to other relevant thiol compounds. A main ubiquitous resistance mechanism is used by bacterial cells to tolerate mercury: the reduction of Hg2+ to Hg0 . This biotransformation converts highly toxic cationic mercury into the metallic species, an almost innocuous volatile form (Fig. 9.2F). Methylation of mercury will not be included in this chapter as does not seem to represent a resistance mechanism; this microbial modification often yields toxic organomercurial derivatives.
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Mercury resistance has been reported since many years in Pseudomonas species [82, 83], including P. cepacia, P. fluorescens, P. putida, P. putrefaciens, and P. stutzeri [84]. The reduction of Hg2+ to Hg0 is mediated by the mercuric reductase encoded by the merA gene, which is located in the mercury resistance mer operon [85, 86]. The mer operon is often encoded on mobile genetic elements [84]. The simplest and most studied Gram negative mer determinant is that from transposon Tn501, originally identified in P. aeruginosa [82]. The mer operon shows several genetic arrangements depending on its origin. The mer operon of Tn501 consists of merRTPADE genes which encode polypeptides with regulatory, transport and enzymatic functions; other mer genes, such as merB, C, F, E and G, are localized on different mer operons [84, 86]. The mer genes are widely distributed among pseudomonads as shown in Fig. 9.3. Bacteria P. aeruginosa UCBPPPA14 P. aeruginosa PA7 P. aeruginosa plasmid pVS1 , transposon Tn501 P. putida W619 P. stutzeri plasmid pPB P. aeruginosa 07-406 plasmid pMATVIM-7 P. aeruginosa plasmid R1033
Locus 15460 0104 merA
merA
merD merE
2217
1249
0789
0789
Rtn 2200
merA
merD merE
2217 0789 merR merT merP merC
1249 merA
0789 merD merE
2217 0477 0789 merR merT merP merC
1249 merA
2217 0477 0789 merR merT merP
1249 merA
2217 0789 merR merT merP merF
1249 merA
4644 merD merE
2217 0789 merR merT merP merF
1249 merA
0789 merD
merR merT merP
pMATVM7_04 merA 0090
P. aeruginosa 2192
02098
P. putida MU10-2
merR merT merP
2338 merA
P. aeruginosa PA7
Pseudomonas sp. CT14 plasmid pCT14 P. fluorescens plasmid pMER327
mer cluster
pCT14_p48 merA merA2
2217 0789 merR merT merP
0789
2217
0789
TnpA
1249
merA
merG
1249
0790
0789 merB
merD 0789
Fig. 9.3 Schematic representation of the arrangement of the mer genes located in the genomes of pseudomonads. The columns indicate the Pseudomonas strain and the microbial locus for each merA gene. Boxed arrows indicate genes and the direction of transcription. Numbers below genes indicate the COG family to which each gene belongs [87]
The resistance mechanism consists of the initial binding of Hg2+ ions by a pair of cysteine residues on MerP, a mercury binding protein located in the periplasm of Gram-negative bacteria [88]. Mercuric ions are then transferred via a redox exchange mechanism to a pair of cysteines on MerT, an inner membrane transport protein (Fig. 9.2F). MerT is present in most mer operons from Gram negative bacteria. MerT is essential for mercury resistance and is the only Mer protein that interacts directly with MerP. Other Hg2+ membrane transporters are encoded by merC and merF genes. Deletion of merC and merF had no effect on the mercury resistance level [86]. Hg2+ is finally transferred via cysteine residues to the N-terminal domain of MerA, the homodimeric flavoprotein mercuric reductase,
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the key component of the mercury detoxification system. The enzyme catalyzes the two-electron reduction of Hg2+ to volatile elemental mercury (Hg0 ), which is nonenzymatically removed from the growth medium (Fig. 9.2F). The MerA enzyme utilizes NADPH as an electron donor and requires an excess of exogenous thiols for activity. MerA is located in the cytoplasm, where NADPH is abundant [85, 86]. In some bacteria, resistance to organomercurial compounds is also conferred by mer operons encoding the additional organomercurial lyase enzyme MerB (Fig. 9.2G). MerB is a monomeric cytosolic enzyme that cleaves the Hg–C covalent bond of both alkyl and aryl mercurials, releasing Hg2+ , which is then transformed by the MerA mercuric reductase. The activity of the MerB enzyme was first determined in Pseudomonas K-62 [86, 89]. Bacteria possessing MerB are tolerant to both inorganic and organic mercurials (broad-spectrum resistance); in contrast, the narrow-spectrum resistance determinants, where the mer systems lack merB, only confer resistance to Hg2+ [90]. Organomercurials are highly lipid-soluble as to enter the cell efficiently without a specific uptake system. The product of the merG gene lies in the periplasm and probably reduces permeability to organomercurials in soil pseudomonads [86]. Deletion of merG in Pseudomonas K-62 resulted in a decrease in phenylmercury resistance [91]. The regulatory genes for the mer system are merR and merD [86]. MerR belongs to the large MerR family that, as mentioned above, includes transcriptional regulators for Cd2+ , Zn2+ , Cu2+ and Pb2+ . MerR is an activator of the mer cluster and in Gram negative bacteria is divergently transcribed from the major mer promoter [76].
9.4 Toxic Oxyanions Arsenic and chromium are toxic nonessential metalloids that may be present as environmental pollutants. The main tolerance mechanism developed by bacteria for arsenic and chromium oxyanions is their efflux from the cytoplasm by specific orthologous membrane transporters. Oxyanions derived from essential elements that generally lack toxicity (i.e. molybden and tungsten) or those that, while showing toxicity commonly occur at very low levels in the environment (uranium and vanadium) are not described here.
9.4.1 Arsenic Arsenic is a metalloid present in numerous disturbed and natural ecosystems. It can exist in multiple oxidation states, with the most common being arsenite [As(III)] and arsenate [As(V)]. Although some microorganisms can utilize As(V) for anaerobic respiration [92] or oxidize As(III) as a sole energy source, arsenic is generally toxic to most microbes [93]. Arsenate (AsO4 3- ) is a toxic analog of phosphate (PO4 3- ), and most organisms take up arsenate via phosphate transporters [94]. As(V) toxicity is due to the uncoupling of ATP phosphorylation, that would directly impact energy flow, as well as to the inhibition of nucleic acid and phospholipid syntheses
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[95]. In bacteria, the pathway for uptake of trivalent metalloids as As(III) is through the polyol transporter GlpF, which belongs to the family of aquaglyceroporins [96]. As(III) toxicity is predominantly due to its ability to covalently bind protein sulfhydryl groups [93]. Arsenate and arsenite (AsO2 - ) oxyanions are detoxified by an interplay of redox, transport, sequestration, and covalent modification reactions [94] (Fig. 9.4A).
Fig. 9.4 Mechanisms of resistance to toxic oxyanions in pseudomonads. A, arsenic resistance ArsABC and ArsBC systems. B, AsoAB arsenite oxidase complex. C, chromate resistance ChrA transporter. D, ChrR chromate reductase. The functions of the resistance systems are described in the text
Bacteria adapt to arsenic toxicity mainly by the development of resistance mechanisms conferred by chromosomal or plasmid-encoded arsenical resistance (ars) operons [96]. Once the trivalent form of the metalloid accumulates in the cell, resistance is produced by their removal from the cytosol [96]. The ars clusters are widely distributed among pseudomonads, as shown in Fig. 9.5. The mechanism of resistance to arsenic conferred by ars genes has been best characterized from E. coli plasmid R733 [97]. The ars operon consists of genes arsRDABC. The arsA gene encodes the ATPase enzyme subunit of a protein complex composed of an ArsA dimer bound to ArsB, an inner membrane polypeptide
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Bacteria
Locus arsR
arsB
arsC2
40640
1055
0394
arsC1
wrbA
1393
0655
P. syringae pv. syringae B728a P. syringae pv. tomato T1 P. syringae pv. tomato str. DC3000 P. syringae pv. phaseolicola 1448A arsR
arsC2
0640
0394
3702 – 3703 4747 – 4748 1687 – 1686 3723 – 3724
PTPc ArsD
arsA
arsC2
2453
0003
0394
trkA
arsR
2072
0640
Locus
0431
1927 – 1930 2715 – 2718 3034 – 3037 3077 – 3080 2277 – 2280 01002767 – 01002770 01187 – 01190 01380 – 01383 35130, 35110, 35100, 35080 2400 – 2397 2186 – 2183 1504 – 1502
P. putida KT2440 P. putida F1 P. putida GB-1 P. aeruginosa PAO1 P. aeruginosa PACS2 P. aeruginosa C3719 P. aeruginosa 2192 P. aeruginosa UCBPP-PA14 P. fluorescens Pf0-1 P. fluorescens Pf-5 P. syringae pv. syringae B728a
Bacteria
arsH
arsH
ACR3 0798
PTPc
0640
0394
ACR3 0798
P.putida W619
P. entomophila L48
DUF2069
0655
3308
1645 – 1643 4072 – 4074 1247 – 1245 1207 – 1205 0950 – 0948 01001431 – 01001429 05495 – 05493 06125 – 06123 4559 – 4561 51980, 51990, 52000 4456 – 4458 4228 – 4230 2989 – 2991 1354 – 1352 2824, 2826, 2827
P. putida KT2440 P. putida F1 P. putida GB-1 P. putida W619 P. aeruginosa PAO1 P. aeruginosa PACS2 P. aeruginosa C3719 P. aeruginosa 2192 P. aeruginosa PA7 P. aeruginosa UCBPP-PA14 P. fluorescens Pf-5 P. fluorescens Pf0-1 P. mendocina ymp P. entomophila L48 P. stutzeri A1501 ProP
arsB
0477
1055
P. syringae pv. syringae B728a P. syringae pv. tomato str. DC3000 P. syringae pv. tomato T1 P. syringae pv. phaseolicola 1448A
1681 – 1682 3799 – 3798 0009 – 0010 3604 – 3603
trkA
arsC1
0241 – 0234 2095 – 2098 arsC2
wrbA
1393
ProP
DUF619
2453
0431
P. stutzeri A1501 arsR
arsC1
arsH 0431
5146 – 5149
1393
2072
0477
arsR
arsB
arsC2
0640
1055
0394
arsB
arsC2
asoB
asoA
0640
1055
0394
0723
0243
Pseudomonas sp. TS44
arsC1
0431
1393
3165 – 3168 3161 – 3165
P. syringae pv. tomato T1
arsR
3306 – 3308
1247
arsH
aoxA, aoxB
Fig. 9.5 Schematic representation of the local genomic context of ars and aso genes located in the genomes of pseudomonads. All ars and aso genes shown are located in chromosomes. The columns indicate the Pseudomonas strain and the microbial locus for each gene. Preliminary information on aso genes is also included. Boxed arrows indicate genes and direction of transcription. Numbers below genes indicate the COG family to which each gene belongs. [87]
(Fig. 9.4A). As(III) is the substrate of the ArsAB efflux pump, which is an As(III)translocating ATPase [97]. ArsB alone is sufficient for As(III) resistance and proton motive force-dependent As(III) efflux; bacteria lacking ArsA are still resistant to arsenic [96] (Fig. 9.4A). arsC1 belongs to the COG1393 family [87] and encodes an enzyme that reduces As(V) to As(III), which is subsequently extruded from the cell; arsenate reductase activity is required for optimal resistance to As(V) [93] (Fig. 9.4A). In vitro, the reductase activity requires both reduced glutathione (GSH) and any of the three E. coli glutaredoxins, Grx1, Grx2 or Grx3 [96]. The arsD gene is constitutively expressed and encodes a regulatory protein that controls the maximal expression of the ars operon [93]. Finally, arsR encodes a repressor that controls the expression of the ars operon and can be induced by As(III), Sb(III) or even bismuth. ars homologous sequences have been identified in chromosomal DNA from P. aeruginosa [98]. Phylogenetic analysis showed that P. aeruginosa and P. putida possess arsC chromosomal homologs [99]; in the case of P. aeruginosa, its genome contains separate genes for glutaredoxin- and thioredoxin-coupled ArsC reductases [96]. The chromosome of P. fluorescens MSP3 possesses a less complex arsenic operon (arsRBC) which confers resistance to arsenate and arsenite [100]. Analysis of the P. putida KT2440 genome revealed two very similar systems, arsRBCH, for arsenic resistance [6]. The distinct gene arsH is located downstream of arsC and is transcribed in the same direction. The arsH gene product of Yersinia
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enterocolitica was reported to confer resistance to both arsenite and arsenate and was assigned a possible role as a transcriptional regulator [101]. However, P. putida arsH genes, denominated ArsH1 and ArsH2, showed a significant similarity to plant NADH oxidoreductases and to Bacillus subtilis azoreductase [6]. ArsH is widely distributed in bacteria and sparsely in fungi, plants, and archaea [96], but its role in arsenic resistance is still unclear. Bacterial oxidation of As(III) to less toxic As(V) may be considered as a resistance mechanism. Arsenite oxidase (Aso) from Alcaligenes faecalis is the best understood example of this detoxification activity [94]. The enzyme is formed by a molybdopterin-containing subunit and a Fe-S Rieske subunit encoded by the asoA and asoB genes, respectively [102] (Fig. 9.4B). The aso genes form part of the so called “arsenic gene island” encoding proteins related to arsenic resistance and homeostasis; these include putative periplasmic oxyanion binding proteins, probably associated with ABC membrane transporters, as well as an arsenite efflux pump (ArsAB) ATPase. An ortholog of the AsoA Mo-pterin subunit of arsenite oxidase was identified in the genome of Pseudomonas sp. TS44 (Fig. 9.5), suggesting that some pseudomonads also possess the ability to oxidize arsenite [102]. As(III) toxicity via a mechanism involving peroxidation of unsaturated fatty acids was found in P. putida [103, 104]. It was proposed that this process leads to the generation of organic hydroperoxides and oxygen radicals, which induce components of the oxidative stress response such as superoxide dismutase (SOD) and catalase [93]. Catalase activity increased in response to the presence and oxidation of As(III) [104]. These studies also showed that the levels of glutathione reductase (Gor) increased upon exposure of P. putida to As(III). One function of Gor in E. coli is to recycle oxidized glutathione back to reduced glutathione, which is the reductant for the As(V) reductase that converts As(V) to As(III). The latter is then actively removed from the cell by the ArsB effflux pump [93]. Mutants of P. aeruginosa PAO1 affected in the arsB, crc (the catabolite repressor control protein) and gor genes are more sensitive to As(III) than wild-type strain [93]. The crc mutant was more sensitive to H2 O2 in the presence of As(III); the sensitivity to As(III) was assumed to be due to an abnormal regulation of genes under Crc control. Double sodA/sodB mutants also exhibited increased sensitivity to As(III), suggesting that the oxidative stress response is involved in As(III) resistance [93].
9.4.2 Chromium The biological effects of chromium depend on its oxidation state. At the extracellular level, Cr(VI) (usually in the form of chromate, CrO4 2- ) is highly toxic to most bacteria, whereas Cr(III) is relatively innocuous by its inability to traverse cell membranes. In the cytoplasm, chromium toxicity is mainly related to the process of reduction of Cr(VI) to lower oxidation states [i.e. Cr(III) and Cr(V)] in which free radicals may form [105, 106]. Bacterial resistance determinants may be encoded either by chromosomal genes or by plasmids [107, 108]. Usually genes located
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on plasmids encode membrane transporters which directly mediate efflux of chromate ions from the cytoplasm. On the other hand, resistance systems encoded within bacterial chromosomes are generally related to specific or unspecific Cr(VI) reduction, free-radical detoxifying activities, repairing of DNA damage, and processes associated with sulfur or iron homeostasis [106]. The best understood mechanism of resistance to chromate is the efflux of Cr O4 2conferred by the ChrA protein encoded by the P. aeruginosa plasmid pUM505 [109]. ChrA is a membrane protein of 416 amino acid residues which displays a topology of 13 transmembrane segments (TMS) [110]. ChrA functions as a chemiosmotic pump that effluxes CrO4 2- from the cytoplasm using the proton motive force [111, 112] (Fig. 9.4C). Plasmid pMOL28 from C. metallidurans [113], plasmid 1 from Shewanella sp. ANA-3 [114], and transposon TnOtChr from Ochrobactrum tritici 5bv11 [115] all encode ChrA homologs which confer resistance to chromate. The resistance mechanism seems to be the same for these homologs from proteobacteria, as all caused reduced accumulation of CrO4 2- . Structure-function analyses have been conducted with the P. aeruginosa ChrA protein. Random mutagenesis of the chrA gene showed that most essential amino acids are located in the amino terminal end of ChrA [116]. In agreement with this finding, phylogenetic analysis of ChrA homologs revealed that the amino terminal halves are more conserved than the carboxyl halves [117], suggesting that the two halves of ChrA carry out different functions in the transport of CrO4 2- . The ChrA proteins belong to the CHR superfamily of transporters, first described by Nies et al. [107] as a small group of prokaryotic proteins involved in CrO4 2- and sulfate (SO4 2- ) transport. This superfamiliy was classified as TC no. 2.A.51 [118] and includes proteins encoded in chromosomes and plasmids. The protein databases currently contain several dozens of homologs, including proteins from eukaryotes, and has been named as the CHR superfamily of chromate ion transporters [117]. The CHR superfamily contains two families, large proteins (comprising seven LCHR subfamilies) and short proteins (comprising three SCHR subfamilies). All pseudomonads with genomes sequenced have ChrA homologs from subfamilies LCHR1 or LCHR5, as well as other chr related genes (Fig. 9.6). Plasmids pB4 [119], from a Pseudomonas sp. strain, and pUM505, from P. aeruginosa [106], possess chrBAC gene clusters that share high sequence similarity with the resistance determinant from plasmid pMOL28. A function of the ChrB protein in the inducibility of the chrA gene by CrO4 2- had been previously demonstrated in C. metallidurans [113]. The presence of chrC, a gene encoding a probable SOD enzyme, is another variable feature of the chr gene clusters [120] (Fig. 9.6). A second mechanism of resistance to CrO4 2- is the transformation of hexavalent chromium to the trivalent form [105, 108]. Microbial reduction of Cr(VI) to Cr(III) is not a plasmid-associated trait. Chromate reduction has been demonstrated in diverse pseudomonad species, including P. ambigua [121], P. fluorescens [122] and P. putida [123], although only a few enzymes have been characterized to date [105]. Initial studies suggest that chromate reductases may have a different primary role other than chromate reduction; this secondary function for chromate reductases
9
Heavy Metal Resistance in Pseudomonads Bacteria
Locus
Loc.
6202
P
271 Chr cluster chrI
R. metallidurans CH34
chrB
4315
P. putida KT2440 P. putida F1
2556 3159
C
P. putida GB-1
3384
C
4275
atoS GGDEF
2059 chrA5
2202 2199 chrB
2059 chrA5
LCHR5
4275 chrB
P. putida W619
5157
C
P. fluorescens Pf0-1
2147
C
4275 fdhD
moaC moaD moaE
1526
0315
0314
1977
chrB
P. aeruginosa PAO1
chrA
chrB
2247
4275
C 4327
P. stutzeri A1501 P. entomophila L48
2921 3029
chrB
C C
1073 aprE
4275
LCHR1
1404
P. putida W619
3017
C
P. fluorescens Pf-5
3149
C
P. aeruginosa PAO1 P. aeruginosa C3719 P. aeruginosa 2192 P. aeruginosa PA7 P. aeruginosa UCBPPPA14
4289 05218 05773 4856 55740
C
P. mendocina
2228
C
P. stutzeri A1501
2422
2059 chrA5 2059 chrA5
2059 chrA5
chrE chrF
chrC
0605 0607 4275 smtA
0500
rhtB 1280 chrF 4275
glnA 0174
chrC
P 4275
P. mendocina ymp
chrA2
C
2059 chrA5
0605
2059 chrA5
0607 wecD
2059 chrA5
0454
mhpC
2059 chrA5 2059 chrA1
orphan
chrE
wecD
2059 chrA1
0454
2059
0596 pstS
ompA
0226
2885
baeS 0642 baeS 0642
araC
chrA1
tar
2207
2059
0840
zntA
chrA1
padR
2217
1695
DUF1853
2059 chrA1
3782
2059
permease
0730
Fig. 9.6 Schematic representation of the local genomic context of chr genes located in the genomes of pseudomonads. All chrA genes belong to the LCHR1 or LCHR5 subfamilies of the CHR superfamily [117]. The columns indicate the Pseudomonas strain, the microbial loci and the location (Loc.) for each chrA gene (P, plasmid; C, chromosome). Boxed arrows indicate genes and direction of transcription. Identified genes are indicated according to the characterized chr determinant from C. metallidurans plasmid pMOL28. Numbers below genes indicate the COG family to which each gene belongs [87]
may be related to the recent introduction of Cr(VI) to the environment by industrial pollution. The currently best studied chromate reductase is the ChrR enzyme from P. putida, a soluble flavin mononucleotide-binding enzyme which reduces Cr(VI) to Cr(III) [124] (Fig. 9.4D). Purified ChrR revealed that a quinone reductase activity produced a flavin semiquinone during CrO4 2- reduction; this activity transferred >25% of the NADH electrons to reactive oxygen species (ROS) and generated the Cr(V) species transiently. This property of ChrR provides an antioxidant defense mechanism to P. putida by shielding cells against H2 O2 toxicity [125] (Fig. 9.4D). ChrR in one pathway reduces Cr(VI) to Cr(III) generating intermediary Cr(V) and ROS, and, by an additional mechanism, reduces quinones to protect against ROS. ChrR from P. putida belongs to the NADP(H)-dependent FMN reductase (FMN_red) protein family, currently comprising about 250 homologs [106].
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Besides chromate efflux and reduction, several other resistance mechanisms to deal with chromium are displayed by bacteria. Since oxidative stress is responsible for most toxic efects of chromate, protection and detoxification systems against this process is an important part of the defensive barrier. Protection of bacterial cells from DNA damage caused by CrO4 2- is another defensive shield. Cr(VI) has long been known to induce the E. coli SOS repair system that protects DNA from oxidative damage [126]. DNA helicases RecG and RuvB, and the FtsK protein, components of DNA repair and chromosome segregation processes, have ben shown to participate in the response to DNA damage caused by CrO4 2- exposure in P. aeruginosa [127, 128]. Additional protective strategies may be related with sulfur or iron metabolism, but these systems have not been analyzed in pseudomonads.
9.4.3 Tellurium Tellurium is a rare-earth metalloid with a low abundance in the Earth’s crust but which derivatives may be pollutants in industrial waste discharges. The tellurite oxyanion (TeO3 2- ) is highly toxic for most microorganisms, particularly Gramnegative bacteria [129]. Tellurite toxicity in E. coli is several orders of magnitude higher than that of heavy metals such as cobalt, zinc and chromium [23, 130]. Studies on TeO3 2- metabolism and toxicity in E. coli showed that the oxyanion interacts with reduced thiols and that glutathione is the initial target of tellurite reactivity [131]. Tellurite, as chromate, is reduced intracellularly producing toxic intermediates which may cause DNA damage [132]. Despite several genetic determinants related to TeO3 2- resistance have been analyzed to the molecular detail, a general mechanism to explain this phenotype is not available [15, 133]. Instead, a variety of possible biochemical strategies used by bacteria to defend themselves from tellurite toxicity have been reported. Antibiotic-resistant clinical isolates of enterobacteria and P. aeruginosa commonly possess plasmids conferring TeO3 2- resistance [134, 135]. A TeO3 2- resistance determinant from the chromosome of P. syringae pv. pisi encodes a methyl transferase enzyme that may detoxify tellurium by methylation [136]. Similarly, the P. putida genome contains genes that may encode tellurium (and selenium) methylation activities [6]. P. aeruginosa PAO1 mutants affected in the genes encoding DNA helicases RuvB, RecG and the DNA translocase FtsK, that function in DNA repair and chromosome segregation, respectively, showed an increased susceptibility to tellurite [127, 128]. Protection from DNA damage caused by TeO3 2- exposure was proposed as the role of those enzymatic activities in conferring TeO3 2- resistance. The precipitation of TeO3 2- by a siderophore, pyridine-2, 6-bis(thiocarboxylic acid), produced by P. stutzeri KC has been proposed as another mechanism for TeO3 2- detoxification [137]. A detailed metabolomics study of the tellurite hyperresistant Pseudomonas pseudoalcaligenes KF707 strain revealed that the resistance phenotype involves a variety of complex cell modifications, including the induction of the oxidative stress response, resistance to membrane alterations, and a rearrangement of cellular metabolism [138].
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The tehAB operon from the E. coli chromosome encodes TehA, an integral membrane protein, and TehB, a weakly membrane-associated protein [139]. A possible role for this operon in the efflux of TeO3 2- has been discarded [140]. In contrast, the E. coli ArsAB ATPase, which effluxes arsenite, is also able to transport TeO3 2[141]. The TehAB system confers TeO3 2- resistance by a distinct strategy involving thiol redox enzymes, such as glutathione reductase and thioredoxin reductase, as a mechanism of oxidative protection [139]. The TehB protein was found to bind Sadenosyl methionine as a methylation cofactor that detoxifies TeO3 2- , as mentioned above for the P. syringae tellurium resistance determinant [142]. Other E. coli tellurite resistance systems, the kilA and ter determinants, seem to function by protecting glutathione from being reduced by TeO3 2- [131].
9.5 Other Toxic Ions To close the listing of heavy metals displaying deleterious effects on bacteria, a brief outline will be given next for the environmentally important elements silver, tin and selenium, for which resistance mechanisms have been only barely analyzed in pseudomonads. Silver is a highly toxic metal with several biomedical uses, mainly as an antiseptic. Even though numerous examples of silver resistance have been reported in pseudomonads [143–145], no studies on resistance mechanisms have yet appeared. As mentioned in the section of copper, the genomes of P. aeruginosa [3] and P. putida [6] possess homologs for the CusCBA system, a proton-driven RND transporter which may efflux Ag+ from the cytoplasm [24], and for SilP, a P-type ATPase which extrudes Ag+ (see below) [25]. A rather complex Ag+ resistance system, first described from a Salmonella plasmid [25, 146], was later found widely spread in other enterobacterial plasmids [147]. It consists of three operons. silE encodes the Ag+ -inducible periplasmic SilE protein, which binds Ag+ ions and prevents their entry to the cytoplasm. Next, the silCFBAP operon encodes the P-type ATPase SilP, which transports Ag+ from the cytoplasm to the periplasm, the RND complex SilCBA, an efflux pump able to extrude periplasmic Ag+ to the outside, and SilF, a periplasmic chaperone that escorts Ag+ from SilP to the SilCBA pump. A third operon, silRS, encodes a typical two-component regulatory system that controls the expression of the Sil system. Tin is a nonessential metal whose inorganic forms have little toxicity. In contrast, organotins, widely used organometallic compounds, are highly toxic for microorganisms [148]. Organotin-resistant bacteria have been isolated from polluted ecosystems [148, 149]. Several pseudomonad strains resist organotins by breaking Sn-C bonds [150, 151]. A distinct resistance mechanism is displayed by a P. stutzeri strain possessing the tbtABM operon [152]. TbtABM is a RND system which effluxes tributyltin from the cytoplasm. TbtABM also confers resistance to antibiotics and aromatic compounds and shows homology with P. putida multidrug resistance systems. No further details on this organotin resistance mechanism have been reported.
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Selenium, a metalloid related to sulfur and tellurium, is required as a micronutrient by most microorganisms. However, toxic selenium oxyanions selenate (SeO4 2- ) and selenite (SeO3 2- ) may be generated by industrial activities. The reduction of selenite (and less frequently of selenate) to elemental Se0 , considered as a detoxification mechanism, may be carried out by varied bacterial species [153–155], including pseudomonads [156]. Heavy metal-resistant C. metallidurans has the ability to reduce selenite to Se0 , which accumulates as granules in the cytoplasm [155]. The P. putida genome contains genes that may encode selenium methylation activities [6]. No genes or biochemical mechanisms are available for bacterial selenium resistance.
9.6 Concluding Remarks Pseudomonads have evolved diverse resistance mechanisms to cope with heavy metal toxicity. Due to the distinct chemical properties of each toxic metal(loid), and to the different levels to which the microorganisms are exposed, bacteria with varied defense systems have been selected. Strategies involving the exclusion of cytoplasmic ions by membrane efflux pumps seem to be a preferred mechanism. Except for mercury, and probably for tellurite, all the toxic ions treated here may be the substrates of efflux pumps. These transporters belong to a variety of membrane protein families (RND, CDF, MFS, P-type ATPases) frequently widespread among all life domains. A second common resistance mechanism involves the use of redox enzymes. For most chemical elements susceptible to generate different valence species with a lower toxicity (i.e. mercury, arsenic, chromium, selenium), redox detoxification systems are usually deployed by pseudomonads. As may be inferred from the resistance systems described above, understanding the interactions of pseudomonads with toxic heavy metals largely benefited from biochemical and genomic information generated in other bacteria. A conclusion that may be drawn is that the relatively large genomes of pseudomonads are plenty of genetic determinants for heavy metal resistance, which is in tune with the diverse environments that these bacteria use to inhabit. Acknowledgments Research in authors’ laboratories was funded by Coordinación de Investigación Científica (UMSNH), Consejo Nacional de Ciencia y Tecnología (No. 79190), COECYT (Michoacán), and DGAPA-UNAM (IN:208308).
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Index
A AAA+ superfamily, 129 ABC transport system, 75, 250 Acetyl phosphate, 114 Adenylate cyclase, 99–100, 106, 130, 155 Adhesion, 78, 114, 144 AHL-lactonases, 36 Airway epithelial cells, 143, 146, 151 Alg44, 109, 116, 125–128 Alginate, 66, 102, 115–116, 126–127, 130, 151–158, 163, 165, 192 AlgT, 153 Alkaline phosphatase, 78, 100 Allosteric modifications, 97 AmiLEBCRS, 15 Aminoglycoside response regulator (Arr), 103, 119 AmiR, 15 AmrZ, 157 AntABC, 21, 43, 45–46 Anthranilate coenzyme A ligase, 36 Antibiotic resistance, 48, 118, 146, 159, 161, 183, 193 Anti-sigma factor, 59, 66–68, 79, 153–154 Arginine dehydrolase, 178 Arsenate, 256, 266–269 Ars operon, 267–268 Arylsulfatases, 36 AsoA, 268–269 AsoB, 268–269 ATPases, 129, 256–257, 262, 264, 274 ATP-binding cassette, 127, 142 AtpD, 206 Attachment, 114, 119, 121, 150, 158, 186, 188, 207 AttC site, 186 Autoinducer, 29, 67, 90, 155, 160 Autoregulation, 42–43
B Bacterial signal recognition particle (SRP), 12–13 Bacterioferritin, 20 Bacteriophages, 3 BifA, 103, 114–115, 120 Biofilms, 48, 102, 118–121, 146, 151, 153–155, 164–165 Biomass productionm, 189 Biosurfactants, 29, 35 Biovars, 202 Burkholderia, 34, 49–50, 78, 144, 147, 225 Burkholderia cepacia, 144, 147 C cadA, 260–264 Cadmium, 256, 259–264 CadR, 262 CAMP, 97–100, 130, 142 CarA, 8, 206 Carbon tetrachloride, 189 CatA, 178 Cation difusion facilitator (CDF), 256–263, 274 Cationic antimicrobial (CAPs), 48 Cbb3–2 oxidase, 22 C-di-GMP, 97–131 C-di-GMP receptors, 109, 112, 123–131 Cellulose biosynthesis, 108, 188 Ceramide accumulation, 143 CFTR, 142–144, 146, 150–151, 164–165 Cgmp, 97–100 Chaperone(s), 79, 120, 258, 273 Chemotaxis (Che) proteins, 110 CheY, 110 ChiC, 44 Chitinase, 44 Chloride channel, 141–142, 164 ChrA, 267, 270–271
J.L. Ramos, A. Filloux (eds.), Pseudomonas, DOI 10.1007/978-90-481-3909-5, C Springer Science+Business Media B.V. 2010
283
284 Chromate reductases, 270 ChrR, 267, 271 CnrCBA, 257, 259 Cobalt, 256–260, 272 Copper, 256–258, 260, 273 Crc mutant, 269 CsrB, 20 CsrC, 20 CupA fimbriae, 111, 118 CupA-lacZ expression, 121 CupB, 120–122 CupC, 120–122 Cup operons, 120 Cupriavidus metallidurans, 259 CyaA, 99 CyaB, 99 Cyclases Histidine Kinase Associated Sensory Extracellular (CHASE), 105 Cytochromes, 201 Cytotoxicity, 102, 150, 156 CzrSRCBA, 263 D Dendrograms, 180, 224 Denitrification, 22, 178, 187, 191–192 2, 4-diacetylphloroglucinol (DAPG), 18–19 Diamino-butanoic acid, 220 DNA-DNA hybridisation, 178–180, 183 DNA helicases, 272 DNA mobility shift assays, 41–42 D-ornithine, 208 Dyspnea, 142 E EAL proteins, 101, 106, 124 ECF sigma factor fpvI, 67 Ecotypes, 184, 187–194 Efflux pumps, 159, 274 EF-TU proteins, 14 Elastase, 29, 32, 38, 43–44, 47, 49, 157 Electron transport, 201, 256 Endophyte, 187 Endoprotease PrpL, 67 ExbB, 59, 64–65 ExbD, 59, 64–65 Exoenzymes, 152 ExoS, 155–156 ExoT, 155 Exotoxin A, 16, 43, 67, 69, 72, 75, 99, 122–123, 160, 163 ExoU, 155–156 ExoY, 99, 155
Index F FecI, 66, 68–70, 79–81, 83, 85–87 FemI, 69–70, 75, 80–81, 84–87 Ferrichrome uptake, 69, 71 Ferrioxamine uptake, 69, 72 Ffs gene, 13 FimX, 103, 120 FiuI, 69–70, 74, 80–81, 83–84, 87–88 Flagella, 118, 152, 157, 163, 165 Flagella-based vaccine, 163 Flagellar rotation, 110 FleQ, 109, 128–130, 157 FliC, 157 FoxI, 69–70, 74, 82, 84–87 FpvI, 60, 67–70, 73, 79, 82, 85, 87, 89 FpvR, 60, 66–69, 79, 81–82, 85, 87–89 Fur, 20–22, 45, 60, 70, 75–78, 259 G GacS/GacA, 3, 16, 18–20 GDP-mannose 4, 6-dehydratase, 161 Genomovars, 177–183, 191, 194, 202 GGDEF domains, 101, 106, 112, 115, 120, 123–124, 128 GGDEF/EAL containing proteins, 115, 118 Glycoprotein, 142 Gor genes, 269 GyrB, 177–180, 183, 206 H H2 O2 toxicity, 271 Haem uptake, 69, 72, 76 Hairpin-like secondary structure, 10 Heavy metal resistance, 255–274 HHQ, 22, 30–42, 44–50 2-heptyl-4, 1H-quinolone (HHQ), 22, 31, 34 Histidine autokinase activity, 122 Homeostasis, 256–261, 264, 269–270 Homoserine lactone, 16, 18, 29, 155 Horizontal gene transfer, 160–161, 183, 194 HQNO, 30–34, 36–37, 44, 46–49 Hyperbiofilm phenotype, 114 Hypermutable, 161 I IEF profiles, 210, 212, 217–218, 224 IgRs, 4–5 Insertion sequences, 186 Interleukin-2 cytokine release, 48 Iron-regulated genes, 146 Iron-siderophore complex, 59, 62, 73 Iron transport, 45, 73, 90, 201, 210 Isopyoverdines, 207
Index K King’s B medium, 201, 206 KynAU, 47 Kynurenic acid, 36 L Lactoferrin, 164 LadS, 16–17 Las, 29, 39–41, 43, 50 LasI, 16, 29, 39–40 LasR, 16, 29, 32, 39–41, 160 Lipopolysaccharides (LPS), 178 LpxC, 8, 162 Lung infections, 46, 141–165 LysR box, 42 M Major facilitator superfamily, 256 Malonyl-CoA, 37–38 Maltose, 126, 178 Marine environments, 190 Membrane vesicles (MVs), 35 MerA, 265–266 Mercury, 256, 261–262, 264–266, 274 Mer operon, 265–266 MerRTPADE, 265 Metalloids, 255, 266–267 Metallothioneins, 257, 260–261, 264 Metal uptake, 69, 71, 76–77 Methionine auxotrophy, 158 Methyl-accepting Chemotaxis Proteins (MCP proteins), 110–111 Methylesterase activity, 110 MexGHI-opmD, 44 Micronutrient cations, 256–261 Mobile genetic elements, 105, 186, 194, 265 MorA, 103, 118 Motility factors, 152 MucA, 153–155, 159–160 MucB, 153 MucC, 153 MucD, 153 Mucoid, 115, 126, 143–144, 146–148, 152–155, 159–160, 163 MucP, 66 MurC, 162 MurF, 162 MutS, 161 Mutt, 161 N NahA, 191 NahG, 191 NahH, 186, 191
285 NahW, 191 Naphthalene, 181, 186, 191, 194 Natural competence, 184 Natural transformation, 182, 184, 194 N-butanoyl-homoserine lactone, 18 Niche-specific genes, 183–184 Nickel, 256–260 NifH, 187 Nitrate reductases, 192 Nitric oxide, 97, 105, 116, 164 Nitrogen fixation, 187–188 Nonribosomal peptide synthetases, 60 NosZ, 178, 183 N-(3-oxododecanoyl)-homoserine lactone, 16 NreB, 257, 259 Numerical taxonomy, 211, 225 O 3-O lipid A deacylase, 158 Opportunistic bacterial pathogen, 141 OprF, 152, 154 Orotic acid, 36 Outer membrane proteins, 4, 62, 178 P PAC motifs, 105 Pancreatic insufficiency, 141–142 Paralogues of CheW, 110 PbrA, 88–89, 264 PbrABCD, 264 PbrUTR, 264 PchDCBAEGF, 45 P. entomophila, 71–72, 77–78, 81, 89–90, 184, 221, 268, 271 Periplasmic copper-binding proteinsm, 257 PfrI, 79, 85 Phaseolotoxin, 18 Phenazine biosynthesis, 160 PhnAB, 35–38, 42, 46–47 PhoB, 43 PhoPQ, 48 Phoshorylated CheY, 110–111 Phosphorylcholine, 158 PhrS, 7, 10–11, 22 PhuR, 75–76, 80–81, 83–84, 88 PhuS, 75 PhuT, 75 Phylogenetic markers, 225 Phytohormones, 187, 236 Phz operon, 44 PigD, 69–70 Pilus formation, 16–17 PilZ, 109, 116, 125–130 Plasmid pB4, 270
286 P. mendocina, 71–72, 80, 179, 194, 235, 240, 248, 251, 268, 271 pMOL28, 259, 270–271 Polyphasic taxonomical method, 202 Population density, 29, 46 PpGpp, 97–98, 101 PPT23D, 257 PpyR, 43 PQS, 16, 22, 30–50 PqsABCDE, 35–36, 40, 42–45, 47, 49–50 PqsR, 35–36, 39–45, 47–48 Proinflammatory response, 143, 157 PrpL, 60, 67–69, 72 PrrF, 20–22, 45–46 Pseudomonas aeruginosa, 6, 29–51, 97–131, 141–165, 202, 211, 249 Pseudomonas putida, 49, 65, 70, 118, 202, 206, 249, 256 Pseudothioneins, 262–264 P. stutzeri, 80, 177–181, 183–194, 265, 268, 271–273 PtxR, 43, 122 P-type ATPases, 256–257, 262, 274 PUMA3 system, 78 PupA, 65, 79 PupB, 65, 82, 84 PvdS, 43, 45, 60, 67–70, 72–73, 77, 79–82, 85, 88–89, 210, 218, 221, 224 PvrR gene, 119 PvrR (Phenotype variant regulator), 118–119 Pyochelin, 45–46, 60, 77 Pyocyanin, 29, 35–36, 42–45, 47–50 Pyocyanin production, 35, 43–44, 47, 49 Pyoverdine, 45, 60–63, 65–73, 77, 79, 89–90, 201, 205–212, 216–218, 221 Pyoverdine-deficient mutants, 217 Pyridine-2,6-bis (thiocarboxylic acid), 272 Q Quinolone signalling, 29–51 Quorum sensing, 10, 16–18, 21–22, 29, 98–99, 155, 160, 192 R RcnA, 257, 259 Reactive oxygen species, 144, 153–154, 271 RegB, 122 Resistance mechanism, 263–265, 269–270, 274 Resistance-nodulation-cell division, 256 RetS, 16–17, 19, 102, 128, 156 Rhamnolipid, 29, 35, 39, 44, 105, 164 Rhizosphere, 183, 187–188 RhlG, 39
Index RhlR, 16, 29, 40–41, 44 Rhodopseudomonas palustris, 78, 250 RNA-binding protein, 4, 17, 19 RNA polymerase, 4, 10–12, 60, 68, 129 RNase P, 4, 10, 13, 15 RocR, 103, 106, 116, 121–122, 125 Root surface, 187 RpoB, 8, 177–178, 183, 206, 223–224 RpoD, 4, 68, 177–180, 183, 206 RpoN, 4, 156 RpoS, 4, 7, 19–20, 44, 98 RsmA, 11, 16–20, 44 RsmY, 3, 5–6, 11, 16–20 RsmZ, 3, 5, 7, 11, 16–20 S SadC, 103, 114–115 SadR, 120–122 Second messenger purine nucleotides, 97 Sensor histidine kinase, 119 Sensor kinase, 16, 102, 105, 110, 120–121, 156 Siderophore(s), 20, 45–46, 59–65, 67–75, 77, 79–87, 188–190, 201–226 Siderotyping, 60, 201, 203, 205–206, 208–221, 223–225 Siderovars, 203–205, 212–213, 216–218, 221, 223–224 Sigma-54-containing RNA polymerase, 129 Signal transduction, 18, 23, 61–63, 65, 77, 90, 97, 99, 101, 105, 107, 110–111, 118, 130 SilCBA, 273 SilP, 258, 273 Silver, 256, 258, 273 Small colony variant (SCV), 109, 118–119 Small cystein-rich proteins, 264 Small RNAs, 3–23 SodA/sodB mutants, 269 SpoT esterase, 98 6S RNA, 4–5, 9–13 SroG, 16 16S rRNA genes, 149 SsrA gene, 4, 14–15 SsrS, 4, 10–12 Starch, 178–179 Stressors, 49 Succinate dehydrogenase, 20 Superoxide dismutase, 20, 269 Surface adhesion defective (Sad), 114 Swarming motility, 43, 110–111, 115–116 T TbtABM, 273 T-cell proliferation, 48
Index TehAtehB operon, 273 Tellurium, 272–274 Thioredoxin reductase, 273 Thiosulphate, 188, 192 Threo-β-hydroxy-aspartic acid, 208, 220 Tn501, 265 Tobramycin, 102, 119, 146, 155 Toll-like receptors, 144 TonB, 59–66, 68–70, 73–76, 78–80, 82, 84, 86, 88–90 ToxA, 60, 68, 122 Toxic cations, 256, 261–266 ToxR, 103, 109, 122, 125 Transcriptomic analysis, 258 Transition metals, 256, 258 Transmembrane conductance, 142 Transposon, 4, 35, 102, 110, 118–121, 162, 186, 241, 265, 270 Tributyltin, 273 TrpEG, 37–38 Twitching motility, 16, 99, 102, 118, 120, 125, 160 Two-component regulatory systems, 59 Two-component system CzcS-CzcR, 263 Type III secretion system (TTSS), 16–17, 99, 120–122, 155–156
287 Type II secretion system, 78 Type IV pili, 16, 99, 120, 125, 152 V Vfr, 43, 99 Vibrio vulnificus, 102 Viral infections, 144–145 Virulence, 16–18, 29, 36, 43–48, 50, 67–71, 77–78, 90, 98–99, 101, 105, 116, 121, 158, 160–161 Vitamin B–12, 64 W Wringly Spreader Phenotype (Wsp), 110–112 Wrinkled morphology, 129 WspA, 110–111 WspB, 110 WspD, 110 WspR, 102, 110–114, 116, 124 Z Zinc, 36, 70, 256–257, 260–261, 272 Zinc metallo-β-hydrolase, 36 ZntA, 257, 259–260, 262, 264, 271 ZntR, 260, 262
