Production and regulation of fouling inhibitory

compounds by the marine bacterium

Pseudoalteromonas tunicata

Suhelen Egan

A thesis submitted in fulfilment of the requirements for the degree of

Doctor of Philosophy

School of Microbiology and Immunology

Faculty of Life Sciences

The University of New South Wales,

Sydney, Australia

April 2001

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To my family and friends

- Cheers !

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Table of contents

Acknowledgments………………………………………………………………………….9

Abstract…………………………………………………………………………………....11

List of Publications……………………………………………………………………....12

Certificate of Originality……………………………………………………………...…13

List of Figures………………………………………………………………...…………..14

List of Tables……………………………………………………………………………..18

List of Abbreviations…………………………………………………………………….19

1. General introduction and literature review...............................................211.1. Introduction ........................................................................................................21

1.2. Formation of a biofouling community.............................................................22

1.2.1. Molecular fouling..........................................................................................25

1.2.2. Microbial fouling...........................................................................................26

1.2.2.1. The process of bacterial attachment .......................................................26

1.2.2.2. Bacterial biofilm structure......................................................................29

1.2.2.3. Microbial diversity in natural biofilms...................................................30

1.2.3. Macrofouling.................................................................................................32

1.3. Natural inducers and inhibitors of settlement ...............................................33

1.3.1. Influence of bacteria and their exopolymers on the establishment of higher

organisms……...……………………………………………………………………...33

1.3.2. Inducing chemical cues..................................................................................34

1.3.2.1. Neurotransmitters..................................................................................34

1.3.2.2. Induction by free fatty acids ..................................................................37

1.3.2.3. Induction by other compounds..............................................................38

1.3.3. Inhibitory chemical cues................................................................................39

1.3.3.1. Inhibitory cues of eukaryotic origin.......................................................39

1.3.3.2. Inhibitory cues of bacterial origin..........................................................41

1.4. Regulation of bacterial secondary metabolites...............................................42

1.4.1. Two-component signal transduction systems ................................................43

1.4.2. The ToxR regulon .........................................................................................44

1.4.3. Intercellular signalling ...................................................................................48

1.4.3.1. Acyl-HSL signalling systems................................................................48

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1.4.3.2. Non acyl-HSL signalling systems.........................................................51

1.4.3.3. Interference of bacterial signalling.........................................................53

1.5. The genus Pseudoalteromonas..........................................................................54

1.5.1. Biological activities expressed by Pseudoalteromonas sp.............................54

1.5.2. Biological activities expressed by Pseudoalteromonas tunicata ....................57

1.6. Biofouling: the problems and solutions..........................................................59

1.7. Aims of this study ..............................................................................................60

2. Inhibition of algal spore germination by the marine bacterium

Pseudoalteromonas tunicata ......................................................................................622.1. Introduction ........................................................................................................62

2.2. Material and Methods........................................................................................64

2.2.1. Strains and culture conditions........................................................................64

2.2.2. Preparation of mono-culture biofilms............................................................64

2.2.3. Ulva lactuca bioassay....................................................................................64

2.2.4. Preparation of cell-free supernatant ...............................................................65

2.2.5. Dialysis experiment.......................................................................................65

2.2.6. Preparation of crude P. tunicata cell-free supernatant extracts.......................66

2.2.7. Size fractionation of P. tunicata cell-free supernatant....................................66

2.2.8. Assessment of storage conditions on the stability of the anti-algal compound

……………………………………………………………………………...66

2.2.9. Heat treatment of P. tunicata cell free supernatant.........................................66

2.2.10. Protease treatments of P. tunicata cell free supernatant .................................67

2.2.11. Effect of P. tunicata supernatant on the germination of U. lactuca spores post

settlement.......................................................................................................................67

2.2.12. Polysiphonia bioassay...................................................................................67

2.3. Results .................................................................................................................68

2.3.1. Effect of bacterial biofilms on U. lactuca spore germination.........................68

2.3.2. Effect of bacterial supernatant on U. lactuca germination..............................69

2.3.3. Dialysis experiment.......................................................................................70

2.3.4. Effects of crude extracts of P. tunicata cell free supernatant on U. lactuca

spore germination..........................................................................................................72

2.3.5. Fractionation of P. tunicata cell free supernatant...........................................73

2.3.6. Heat treatment of P. tunicata cell free supernatant.........................................74

2.3.7. Enzyme treatments of P. tunicata cell free supernatant..................................75

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2.3.8. Assessment of storage conditions on the stability of the anti-algal compound

……………………………………………………………………………...77

2.3.9. Effect of P. tunicata supernatant on settled spores ........................................77

2.3.10. Activity of P. tunicata cells and cell free supernatant against spores from the

red alga Polysiphonia sp................................................................................................77

2.4. Discussion ...........................................................................................................79

3. Anti-fungal activity of Pseudoalteromonas tunicata................................833.1. Introduction ........................................................................................................83

3.2. Materials and Methods......................................................................................84

3.2.1. Anti-fungal bioassay......................................................................................84

3.2.2. Transposon mutagenesis ...............................................................................85

3.2.3. Phenotypic characterisation of the non anti-fungal transposon mutants.........86

3.2.3.1. Growth curves .......................................................................................86

3.2.3.2. Anti-bacterial activity .............................................................................86

3.2.3.3. Anti-algal activity...................................................................................86

3.2.3.4. Anti-larval activity..................................................................................87

3.2.4. Genotypic characterisation of the non anti-fungal transposon mutants..........87

3.2.4.1. Genomic DNA extractions ....................................................................87

3.2.4.2. Panhandle-PCR method for sequencing within uncloned genomic DNA

………………………………………………………………………...88

3.2.4.3. Preparation of PCR templates and DNA sequencing.............................90

3.2.4.4. Sequence data analysis ..........................................................................92

3.2.5. Preparation of P. tunicata concentrated supernatant ......................................92

3.2.6. Extracts of cells and cell free supernatant of P. tunicata ................................92

3.2.7. Fractionation of the anti-fungal compound from crude cell extracts ..............93

3.3. Results .................................................................................................................93

3.3.1. Activity of P. tunicata against a range of yeast and fungal isolates................93

3.3.2. Transposon mutagenesis ...............................................................................94

3.3.3. Phenotypic characterisation of the non anti-fungal mutants...........................96

3.3.3.1. Growth conditions.................................................................................96

3.3.3.2. Other antifouling properties...................................................................97

3.3.4. Genotypic characterisation of the non anti-fungal mutants ............................98

3.3.4.1. Panhandle-PCR and DNA-sequencing..................................................98

3.3.4.2. DNA sequence analysis.......................................................................100

3.3.5. Identification of the anti-fungal compound produced by P. tunicata ...........112

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3.3.5.1. Anti-fungal activity of P. tunicata supernatant and crude cell extracts .112

3.3.5.2. Fractionation of P. tunicata cell extract................................................112

3.3.5.3. Characterisation of the anti-fungal compound .....................................115

3.3.5.4. Comparison of the active anti-fungal compound with the corresponding

non anti-fungal mutant compound...........................................................................115

3.4. Discussion .........................................................................................................116

4. Generation and analysis of transposon mutants of P. tunicata altered

in normal pigmentation...........................................................................................1224.1. Introduction ......................................................................................................122

4.2. Material and Methods......................................................................................123

4.2.1. Transposon Mutagenesis.............................................................................123

4.2.2. Phenotypic characterisation of pigmented P. tunicata transposon mutants..123

4.2.2.1. Analysis of pigmentation (UV/Visible light spectra)............................123

4.2.2.2. Antifouling activity ..............................................................................123

4.2.2.3. Assessment of bacterial growth ...........................................................124

4.2.3. Genotypic characterisation of pigmented transposon mutants of P. tunicata

…………………………………………………………………………….124

4.2.3.1. Gene sequencing by panhandle-PCR ..................................................124

4.2.4. Analysis of the proteins secreted by wild-type and white mutant 3 (W3) strains

of P. tunicata…...........................................................................................................125

4.2.4.1. Sample preparation and ammonium sulphate precipitation ..................125

4.2.4.2. Protein determination...........................................................................125

4.2.4.3. Sodium dodecyl sulphate - polyacrylamide gel electrophoresis (SDS-

PAGE) ……………………………………………………………………….126

4.2.4.4. Silver staining......................................................................................126

4.3. Results ...............................................................................................................127

4.3.1. Generation of transposon mutants...............................................................127

4.3.2. Phenotypic Characterisation ........................................................................127

4.3.2.1. Analysis of pigmentation (UV/ Visible light spectra)...........................127

4.3.2.2. Antifouling activity ..............................................................................131

4.3.2.3. Assessment of growth .........................................................................135

4.3.3. Genotypic characterisation of transposon mutants.......................................137

4.3.3.1. DNA sequence analysis.......................................................................137

4.3.4. Assessment of secreted protein profiles of wild-type and white mutant 3 (W3)

strains of P. tunicata....................................................................................................166

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4.4. Discussion .........................................................................................................168

5. Identification and characterisation of a putative transcriptional

regulator controlling the expression of extracellular inhibitors in

Pseudoalteromonas tunicata ....................................................................................1745.1. Introduction ......................................................................................................174

5.2. Materials and Methods....................................................................................175

5.2.1. DNA sequencing and analysis.....................................................................175

5.2.2. Two-dimensional gel electrophoresis (2DGE).............................................175

5.2.2.1. Sample preparation..............................................................................175

5.2.2.2. Sample preparation and isoelectric focusing........................................176

5.2.2.3. Second-dimension electrophoresis ......................................................176

5.2.2.4. Staining and analysis...........................................................................176

5.3. Results ...............................................................................................................177

5.3.1. DNA Sequencing analysis...........................................................................177

5.3.2. Global differences in protein expression between wild-type P. tunicata and the

W2 mutant...................................................................................................................189

5.4. Discussion .........................................................................................................194

6. Antifouling activity and phylogenetic relationship of bacteria

isolated from different marine surfaces.............................................................1986.1. Introduction ......................................................................................................198

6.2. Material and Methods......................................................................................199

6.2.1. Bacterial strains ...........................................................................................199

6.2.2. Antifouling activity of the marine isolates....................................................199

6.2.3. Genomic extractions, 16S ribosomal DNA amplification and DNA

sequencing...................................................................................................................200

6.2.4. Phylogenetic analysis ..................................................................................200

6.3. Results ...............................................................................................................201

6.3.1. Settlement of B. amphitrite larvae in the presence of bacterial strains isolated

from different marine surfaces.....................................................................................201

6.3.2. Settlement of B. amphitrite larvae in the presence of dark pigmented bacterial

isolates….....................................................................................................................201

6.3.3. Germination of U. lactuca and Polysiphonia sp. spores in the presence of

biofilms of the U. lactuca isolates ...............................................................................203

6.3.4. Anti-bacterial activity of the U. lactuca isolates ...........................................203

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6.3.5. Anti-fungal activity of the U. lactuca isolates ..............................................205

6.3.6. 16S rDNA sequencing and phylogenetic analysis of the U. lactuca

isolates…….............................................................................................................…205

6.4. Discussion .........................................................................................................207

7. Characterisation of Pseudoalteromonas ulvae, a bacterium with

antifouling activities..................................................................................................2117.1. Introduction ......................................................................................................211

7.2. Materials and Methods....................................................................................212

7.2.1. Source of inoculum and isolation ................................................................212

7.2.2. Phenotypic characterisation .........................................................................212

7.2.3. Negative staining and electron microscopy..................................................213

7.2.4. 16S rDNA amplification, sequencing and phylogenetic analysis.................213

7.2.5. Nucleotide sequence accession numbers .....................................................214

7.2.6. DNA-DNA hybridisation............................................................................215

7.3. Results and Discussion....................................................................................216

7.3.1. Biochemical and phenotypical characterisation of UL12 T and UL13 ..........216

7.3.2. Genotypic characterisation...........................................................................218

7.3.3. Assignment of UL12T and UL13 for a new species.....................................221

7.3.4. Description of Pseudoalteromonas ulvae sp. nov. ......................................223

8. General discussion..............................................................................................2248.1. Antifouling and biocontrol properties of Pseudoalteromonas tunicata ......224

8.2. A model for the synthesis and regulation of pigmentation and fouling

inhibitors in P. tunicata ...............................................................................................228

8.3. Ecological significance of Pseudoalteromonas species..................................232

Appendix I.....................................................................................................................235

Appendix II...................................................................................................................237

Appendix III .................................................................................................................240

References……………………………………………………………………………. 241

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Acknowledgments

Firstly I would like to give a big thanks to my supervisor Staffan Kjelleberg. It has been said

many times before but I am going to do it again, thanks for being such an enthusiastic and

inspiring supervisor and friend. The times when I thought that my project was not going

anywhere your constant positive attitude always had a way of lifting me up...Thanks for your

direction and encouragement.

Thanks to the D2 group, especially to Carola and Sally who have been there from the

beginning and to Sacha and Ashley for going though the last couple of years with me. These

are exciting times for P. tunicata, it is great to see that the project is really taking off now!

Thankyou to all those people in the School of Microbiology and Immunology and the CMBB

for making everyday life in the lab so much fun. Even those days when you are stressed out

and experiments have not been working for weeks there is always someone there that will

make you smile. A special thanks goes to Lyndal, who I have had a great time sharing a lab

and office with, thanks for listening to all my highs and lows on a daily basis...I appreciate the

friendship. Maurice, I know you mean well with all those nasty comments....(C.F. lots)..

thanks!

There are many people who were extremely helpful when it came to getting technical advice in

particular I would like to thank, Geoff Woolcott for his advice when it came to dealing with

Ulva lactuca spores. Sophia for teaching me the Polysiphonia bioassays. Fitri and Julie for

their endless advice with 2D. Carolina for helping me get over my radioactivity fears. A big

thanks to Daniel for the use of the panhandle-PCR technique, which has been the saving grace

for myself and many others in the lab. Finally, thanks to Greg for being a great computer

doctor and for keeping my little beast alive though all of this.

Thanks to Adam, Julie and Kate for patiently dealing with all the administration woes that have

occurred over the years and for keeping me company when waiting to see Staffan.

Thanks to all the friends who I have meet though the University who have been a great source

of support and are a major reason for why I have absolutely enjoyed my PhD. Among them I

would especially like to thank, Carolina for many things especially dragging me to the pool,

Julie for looking after Torsten in the lab and for the Buffy conversations that most people

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walk away from. Di for so often getting us organised, Greg for being a big smiley face,

Deborah for being Deb and Bren for teaching me the words you can’t say in German.

To my bestest friends outside of the University, Alison, Katy, Em, Mike, Russ and Caz. You

guys are so much a part of my past, present and future that I don’t really know where to start,

except to say I love you and thanks for seeing me through this bit.

A big thanks goes to my family, their support for me “still being at uni “ has been amazing.

Thanks Mum and Dad for always being there and encouraging me in everything I chose to

do. Thanks to Becky for being my favourite little sister. Kerrie, Brian, Sarah, Elissa and Gary

thanks for being my extended family and for being so enthusiastic when I blab on about

science. Nan and Pop... I am so grateful to you both that we are such a close family, thanks

for looking after me whenever I needed a bed and a good meal during my studies.

Finally, the person who I owe the most to is Dr T. Thomas!. Thankyou Torsten for giving me

confidence and guidance as a scientist, for being by my side through the good and through the

few bad times. Most importantly thank you for giving me your heart and being my dearest

friend. I know that I would not have done this PhD thing as well without you.

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Abstract

The marine surface-associated bacterium Pseudoaltermonas tunicata, produces a range of

compounds that inhibit fouling organisms, including invertebrate larvae, bacteria, algal spores

and fungi. In addition to these antifouling compounds P. tunicata cells produce both a yellow

and a purple pigment. The aim of this study was to further characterise the antifouling

activities, their regulation and relationship with pigmentation, and the ecological significance of

P. tunicata and related organisms.

It was discovered that the anti-algal compound was extracellular, heat sensitive, polar and

between 3 and 10 kDa in size. The anti-fungal compound was found to be the yellow pigment

and active against a wide range of fungal and yeast isolates. Chemical analysis suggests that

this compound consists of a carbon ring bound to a fatty-acid side chain. Genetic analysis

supports the chemical data for the active compound as a mutant in a gene encoding for a long-

chain fatty-acid CoA ligase was deficient for anti-fungal activity.

To address the regulation of antifouling compounds and their relationship to pigmentation

transposon mutagenesis of P. tunicata was performed. Mutants lacking the yellow pigment

displayed a reduced ability to inhibit fouling organisms. Further analysis of these mutants

identified genes involved with the synthesis and regulation of synthesis of pigment and

antifouling compounds. One of these mutants was disrupted in a gene (wmpR) with similarity

to the transcriptional regulators ToxR from Vibrio cholerae and CadC from Escherichia coli.

Analysis of global protein expression using two-dimensional gel electrophoresis showed that

WmpR is essential for the expression of at least fifteen proteins important for the synthesis of

fouling inhibitors.

The ecological significance of antifouling bacteria was addressed by assessing the antifouling

capabilities of a collection of bacteria isolated from different marine surfaces. Overall, isolates

from living surfaces displayed more antifouling traits then strains isolated from non-living

surfaces. Five dark-pigmented strains originating from the alga Ulva lactuca were further

studied. Phylogenetic and phenotypic analysis revealed that they were all members of the

genus Pseudoalteromonas and were closely related to P. tunicata. Two strains represented a

novel species within the genus and were taxonomically defined as P. ulvae sp. nov.

12

List of Publications

The work presented in this thesis has so far resulted in the following peer-reviewed

publications:

C. Holmström, S. James, S. Egan, S. Kjelleberg (1996) Inhibition of common fouling

organisms by marine bacterial isolates with special reference to the role of pigmented bacteria.

Biofouling, 10: 251-259.

S. Egan, T. Thomas, C. Holmström, S. Kjelleberg (2000) Phylogenetic relationship and

antifouling activity of bacterial epiphytes from the marine alga Ulva lactuca. Environmental

Microbiology, 2: 343-347.

S. Egan, S. James, C. Holmström, S. Kjelleberg (2001) Inhibition of algal spore germination

by the marine bacterium Pseudoalteromonas tunicata. FEMS Microbiology Ecology, 35: 67-

73.

S. Egan, C. Holmström, S. Kjelleberg (2001) Pseudoalteromonas ulvae sp. nov., a bacterium

with antifouling properties from the surface of a marine alga. International Journal of

Systematic and Evolutionary Microbiology, In Press

13

Certificate of originality

I hereby declare that this submission is my own work and to the best of my knowledge it

contains no material previously published or written by another person, nor material which to

a substantial extent has been accepted for the award of any other degree or diploma at UNSW

or any other educational institution, except where due acknowledgment is made in the thesis.

Any contribution made to the research by others, with whom I have worked at UNSW or

elsewhere, is explicitly acknowledged in the thesis.

I also declare that the intellectual content of this thesis is the product of my own work, except

to the extent that assistance from others in the project’s design and conception or in style,

presentation and linguistic expression is acknowledged.

Suhelen Egan

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List of Figures

Figure 1.1: Representation of the marine biofouling process.. ............................................24

Figure 1.2: Model of the activation of the ToxR regulon in response to environmental

stimuli............................................................................................................................47

Figure 1.3: Model for the regulation of bioluminescence by intercellular signalling in V.

fischeri ..........................................................................................................................50

Figure 1.4: Antifouling activities expressed by P. tunicata..................................................59

Figure 2.1: The effect of cell-free supernatant of P. tunicata and the bacterial isolates R60

and JI on the germination of U. lactuca spores. ............................................................70

Figure 2.2: The effect of bacterial cultures on the germination of algal spores....................71

Figure 2.3: The effect of crude extract of P. tunicata cell-free supernatant on the germination

of U. lactuca spores ......................................................................................................72

Figure 2.4: The effect of size fractionated cell-free supernatant of P. tunicata on the

germination of U. lactuca spores...................................................................................73

Figure 2.5: The effect of both unfractionated and the less than 30 kDa fraction (< 30 kDa) of

P. tunicata cell-free supernatant before and after heat treatment (HT) on the germination

of U. lactuca spores. .....................................................................................................74

Figure 2.6: Anti-algal activity of P. tunicata supernatant treated with proteinase K and

carboxypeptidase y . ......................................................................................................76

Figure 2.7: The ability of P. tunicata biofilms and cell free supernatant to inhibit the

germination of spores from the red alga Polysiphonia sp..............................................78

Figure 3.1: Diagrammatic representation of the panhandle-PCR method for sequencing from

uncloned genomic DNA................................................................................................91

Figure 3.2: Anti-fungal activity of P. tunicata wild-type and the mutants FM 1-3...............95

Figure 3.3: Growth of wild-type P. tunicata and the FM1-mutant ......................................96

Figure 3.4: Agarose gel showing the results from a typical panhandle-PCR from FM1

genomic DNA templates................................................................................................99

Figure 3.5: Summary of the sequence strategy for determining the nucleotide sequence of

the region of DNA flanking the transposon insert within the P. tunicata non anti-fungal

mutant genome ............................................................................................................103

Figure 3.6: Nucleotide sequence of the region of genomic DNA surrounding the transposon

within the non-anti-fungal mutant genome...................................................................107

Figure 3.7: Multiple sequence alignment of P. tunicata AfaA with sequences of known long-

chain fatty-acid CoA ligases from three different bacterial species ..............................109

15

Figure 3.8: Multiple sequence alignment of P. tunicata AfaB with Drosophilia Kraken

protein; Pseudomonas putida atropinesterase and Synechocystis sp. esterase. ............111

Figure 3.9: The initial 13 chromatography fractions of the crude cell extract of P. tunicata

....................................................................................................................................113

Figure 3.10: Anti-fungal activity of P. tunicata cell-extract fractions resulting from

separation using solid-phase extraction columns.........................................................114

Figure 3.11: The hypothetical model for the involvement of AfaA and AfaB in the synthesis

of the anti-fungal compound........................................................................................121

Figure 4.1: Transposon mutants of P. tunicata with changes in pigmentation...................128

Figure 4.2: UV/Visible light spectra of cell extracts from P. tunicata wild-type and

pigmented mutants.......................................................................................................130

Figure 4.3: Anti-fungal activity of pigmented mutants of P. tunicata ................................133

Figure 4.4: Growth curves of P. tunicata pigmented mutants compared with wild-type

cultures........................................................................................................................136

Figure 4.5: Summary of the sequencing strategy to determine the nucleotide sequence

flanking the transposon insert within the P. tunicata light purple mutant 2 genome. ...139

Figure 4.6: Nucleotide sequence of the genomic-DNA region surrounding the transposon

within the light purple 2 mutant genome......................................................................142

Figure 4.7: Multiple sequence alignment of the P. tunicata LppA protein with Streptomyces

coelicolor putative oxidase; Synechocystis sp. 3-chlorobenzoate-3,4-dioxygenase;

Comamonas testosteroni toluenesulfonate methyl-monooxygenase oxygenase

component TsaM and Pseudomonas fluorescens aminopyrrolnitrin oxidase PrnD ....144

Figure 4.8: Summary of the sequencing strategy to determine the nucleotide sequence

flanking the transposon insert within the P. tunicata dark purple mutants 3 and 5 and

light purple mutant 3 genomes.....................................................................................147

Figure 4.9: Nucleotide sequence of the genomic-DNA region surrounding the transposon

within the dark purple 3, dark purple 5 and light purple 3 mutant genomes.................151

Figure 4.10: Multiple sequence alignment of the deduced amino acid sequence of P. tunicata

DppB with Bacillus subtilis putative ABC transporter, YvrO and Streptococcus cristatus.

ATP-binding cassette protein.......................................................................................152

Figure 4.11: Hydropathy profile of the inferred amino acid sequence from DppA.. .........153

Figure 4.12: Summary of the sequencing strategy to determine the nucleotide sequence

flanking the transposon insert within the P. tunicata white mutant 3 genome..............156

Figure 4.13: Nucleotide sequence of the genomic-DNA region surrounding the transposon

within the white mutant 3 genome................................................................................160

16

Figure 4.14: Multiple sequence alignment of P. tunicata WmpD with the general secretion

pathway protein, EpsD from Vibrio cholerae; ExeD from Aeromonas salmonicida; the

PulD protein from Klebsiella pneumoniae and the OutD protein from Erwinia

carotovora...................................................................................................................163

Figure 4.15: Multiple sequence alignment sequence of P. tunicata WmpC with ExeC from

Aeromonas hydrophila; EpsC from Vibrio cholerae; ExeC from A. salmonicida and

OutC from Erwinia carotovora...................................................................................165

Figure 4.16: Silver stained SDS-PAGE gel showing supernatant proteins from wild-type

(Wt) and white mutant 3 (W3) strains of P. tunicata during different growth phases…

....................................................................................................................................167

Figure 4.17: Hypothetical model relating antifouling activity and pigment production in P.

tunicata........................................................................................................................172

Figure 5.1: Summary of the sequencing strategy used to determine the nucleotide sequence

flanking the transposon insert within the P. tunicata white mutant 2 genome..............179

Figure 5.2: Nucleotide sequence of the genomic-DNA surrounding the transposon within

the white mutant 2 genome. .........................................................................................183

Figure 5.3: Multiple sequence alignment of P. tunicata WmpR with the transcriptional

activator CadC from Escherichia coli; the ToxR-homologue from Vibrio

parahaemolyticus and ToxR cholera-toxin transcriptional activator from V. cholerae.186

Figure 5.4: Secondary structure prediction of the deduced amino acid sequence of WmpR.

....................................................................................................................................188

Figure 5.5: Two-dimensional gel electrophoresis of the total cell protein from P. tunicata

wild-type in early-logarithmic growth..........................................................................190

Figure 5.6: Two-dimensional gel of the total cell protein from P. tunicata white mutant 2 in

early-logarithmic growth..............................................................................................191

Figure 5.7: Two-dimensional gel of the total cell protein from P. tunicata wild-type in early-

stationary phase growth...............................................................................................192

Figure 5.8: Two-dimensional gel of the total cell protein from P. tunicata white mutant 2 in

early-stationary growth................................................................................................193

Figure 5.9: The differences in the number of proteins expressed by the wild-type (Wt) and

the white mutant 2 (W2) at both early-logarithmic and early-stationary phase of growth.

....................................................................................................................................194

Figure 6.1: Settlement (%) of B. amphitrite larvae on biofilms of dark pigmented marine

bacteria ........................................................................................................................202

Figure 6.2: Distance matrix tree based on a sequence alignment of the 16S ribosomal DNA

of novel isolates with members of the genus Pseudoalteromonas.. .............................206

17

Figure 7.1: Electron micrograph of strain UL12T..............................................................219

Figure 7.2: Distance matrix tree based on a sequence alignment of the 16S rDNA gene of

the isolates UL12T and UL13 (P. ulvae sp. nov.), with members of the genus

Pseudoalteromonas and other closely related bacteria.................................................221

Figure 8.1: Hypothetical model for the regulation of yellow pigment and fouling inhibitors

in P. tunicata ...............................................................................................................231

18

List of Tables

Table 1.1: Examples of the acyl-HSL regulatory systems in bacteria..................................51

Table 1.2: A summary of the biological activities of Pseudoalteromonas sp. .....................57

Table 2.1: Effect of marine surface bacteria on Ulva lactuca spore germination.................69

Table 2.2: Effect of P. tunicata supernatant on the germination of Ulva lactuca spores ...77

Table 2.3: Characteristics of the anti-algal component produced by P. tunicata..................81

Table 3.1: Restriction enzymes used for panhandle-PCR ...................................................89

Table 3.2: Activity of P. tunicata against a range of yeast and fungal species.....................94

Table 3.3: Growth inhibition of bacteria in the presence of P. tunicata wild-type and non

anti-fungal mutant strain FM1.......................................................................................97

Table 3.4: Settlement of marine invertebrate larvae in the presence of biofilms of P. tunicata

wild-type, non anti-fungal mutant strain FM1 and a previously determined non-inhibitory

bacterial isolate. .............................................................................................................97

Table 3.5: Germination of marine algal spores in the presence of biofilms of P. tunicata

wild-type and non anti-fungal transposon mutant strain FM1. ......................................98

Table 3.6: Elution steps and characteristics of the initial 13 fractions from solid phase

chromatography columns. ...........................................................................................113

Table 3.7: Characteristic of the anti-fungal compound produced by P. tunicata cells .......115

Table 4.1: Settlement of marine invertebrate larvae in the presence of biofilms of P. tunicata

wild-type and transposon mutant strains......................................................................131

Table 4.2: Germination of marine algal spores in the presence of biofilms of P. tunicata

wild-type and transposon mutant strains......................................................................132

Table 4.3: Growth inhibition of P. tunicata wild-type by supernatant from P. tunicata wild-

type and transposon mutant strains..............................................................................134

Table 6.1: Anti-larval activity of bacterial strains isolated from different marine surfaces ..

....................................................................................................................................201

Table 6.2: Germination of algal spores in the presence of biofilms of marine bacterial

isolates.........................................................................................................................203

Table 6.3: Anti-bacterial activity of marine isolates against various target bacterial strains

................................................................................................................................... .204

Table 6.4: Anti-fungal activity of marine isolates against various target fungal strains......205

Table 7.1: Phenotypic characterisation of Pseudoalteromonas ulvae UL12T...................217

Table 7.2: Differential characteristics of Pseudoalteromonas species...............................220

19

List of Abbreviations

A: Ampere

AA: Amino Acid(s)

Amp: ampicillin

ANGIS: Australian National Genomic Information Service

acyl-HSL: N-acyl-homoserine lactone

bp: base pair(s)

BLAST: Basic Local Alignment Search Tool

C: Celsius

ci: Curie (= 3.7 x 1010 Becquerel)

Da: Dalton

DCM: dichloromethane

DNA: deoxyribonucleic acid

dNTP: deoxyribonucleotide triphosphate

DP: dark purple mutant

DTT: dithiothreitol

2DGE: two-dimensional gel electrophoresis

EDTA: ethylene diamine tetraacetic acid, trisodium salt

EPS: extracellular polymeric substances

FFAs: free fatty acids

FM: non anti-fungal mutant

g: gram

g: gravitational force

GABA: gamma-aminobutyric acid

GSP: General Secretion Pathway

GSPP: General Secretion Pathway Protein

h: hour(s)

HT: heat treatment

kb: kilobase(s), 1000 bp

kDa: kilodalton(s), 1000 Da

Km: kanamycin

LB: Luria Broth

L-DOPA: L-dihydroxyphenylalanine

l: litre

20

log: logarithmic

LP: light purple mutant

m: milli (10-3)

µ: micro (10-6)

M: Molar (= mole per litre)

MIC: minimal inhibitory concentration

min: minute

MMM: marine minimal media

mol: mole (= 6.022 x 1023)

MOPS: morpholinepropanesulfonic acid

MW: molecular weight

NCBI: National Center for Biotechnology Information

NSS: nine salts solution

OD: optical density

ORF: open reading frame

PAGE: polyacrylamide gel electrophoresis

PCR: polymerase chain reaction

pI: isoelectric point

RBS: ribosome binding site

sp.: species

SDS: sodium dodecyl sulfate

Sm: streptomycin

SmR: streptomycin resistant

SW: seawater

sec: second

TBT: tri-n-butyltin

TSB: tryptone soy broth

v/v: volume per volume

VNSS: V-medium modified from väätänen

w/v: weight per volume

W2: white mutant 2

W3: white mutant 3

Wt: wild-type

21

1. General introduction and literature review

1.1. Introduction

In the marine environment the competition for living space is intense, therefore all surfaces,

living or innate are susceptible to fouling. This process generally begins with the formation of

a biochemical conditioning film onto which bacteria and other microorganisms colonise.

Closely following the microbial fouling is the colonisation of various eukaryotic organisms

including marine invertebrates and algae.

Bacterial colonisation of a surface is influenced by the physico-chemical properties of the

surface (eg. texture, hydrophobicity) and the biological properties of the bacterium, such as

surface motility (eg swarming), surface structures (eg. pili) and the production of adhesive

molecules (eg. extracellular polysaccharides). During the process of adhesion bacterial cells

are reported to alter their gene expression in response to the proximity of the surface and it is

now well accepted that cells in a biofilm differ substantially from their planktonic counterparts

(Costerton et al., 1995; Costerton et al, 1999).

The colonisation of macrofoulers such as sessile invertebrates and algae represents the final

phase of the biofouling process. These organisms seek out new areas to colonise by the

release of free-living larval or spore stages. Similar to the colonisation of bacteria, settlement

of larvae and spores is dependent on physiological, chemical and biological factors.

Communication via chemical signals or cues is important within natural systems, many marine

plants and animals are known to make use of chemical defence strategies as a way of

protecting themselves from becoming fouled (de Nys et al., 1994; Mary et al., 1993;

McCaffrey and Endean, 1985). Alternatively, the same organisms may use separate cues to

signal for the settlement of conspecifics or prey (Pawlik, 1992). The importance of bacterially

derived signals is just becoming realised. Several studies have indicated that it is the primary

colonising bacteria that provide the chemical cues which determine if a higher organism will

settle or not (Berland et al., 1972; Holmström et al., 1992; Maki et al., 1990). For example,

22

the newly established genus Pseudoalteromonas contains numerous marine species that live

in association with higher organisms and produce a range of bio-active molecules (Holmström

and Kjelleberg, 1999). This characteristic may benefit cells of Pseudoalteromonas species in

their competition for nutrients and living space.

The production of secondary metabolites by bacteria can be beneficial, as is the case of a

bacterium attempting to establish itself within the new environment of a host, whether it is as a

pathogen or as a symbiont. However, as they are not essential for growth the synthesis of

secondary metabolites can also be quite costly. As such bacteria have developed complex

ways to regulate the expression of these metabolites which enables them to be produced only

when it will be beneficial for the bacterium.

This review will discuss the role bacteria play in the development and maintenance of a

biofouling community in the marine environment. Complex interactions that take place by

way of specific chemical cues and bacterial secondary metabolites will be discussed. Using

specific examples, some of the ways secondary metabolites are regulated by marine bacteria

will also be addressed. The genus Pseudoalteromonas will be discussed in terms of how it

might benefit from the production of a range of extracellular molecules to compete for

nutrients and living space. Finally, the possibility of using such molecules in commercial

applications will be addressed.

1.2. Formation of a biofouling community

All surfaces in the marine environment are influenced by a variety of biological, physical and

chemical factors that result in the formation of a complex layer of attached microorganisms

and macroorganisms (including sessile plants and animals) referred to as biofouling. The

species composition of the water column is the most important biological factor, bacteria being

especially important as they are often the first to colonise a fresh surface and consequently

alter the physical and chemical properties of the surface. The various physical properties able

to influence biofouling development include surface texture and contour, light availability,

wettability of the surface and rates of heat, gas and nutrient transfer. Chemical properties may

include the presence of various surface-active molecules, the levels of calcium, magnesium or

ion in the water, the availability of specific nutrients and the release of chemical cues from

neighbouring organisms.

23

The sequence of events which lead to the formation of a biofouling community have been

investigated by several authors. Wahl (1989) describes a classical view of the fouling process

where colonisation of a new surface occurs as a succession in three distinct phases. The first

of these events is molecular fouling or the formation of a biochemical conditioning film on the

clean surface, this is followed by microbial fouling (eg: colonisation of bacteria and diatoms)

and macro-fouling (eg: colonisation of macro-algae and invertebrates) events (Figure 1.1).

During the course of these events the nature of the process changes from a physical process

to a predominantly biological process (Clare et al., 1992).

A second model being described as the dynamic model of the fouling process suggests that

the major driving force behind the sequence of events leading to the establishment of a fouling

community is the relative amount of each kind of foulant within the water column. This model

does not rely on a strict successional process and is further complicated by secondary forces

such as physical, molecular and biological interactions between and within each of the

different fouling organisms (Clare et al., 1992).

It should be noted that these models have been intended to describe the biofouling process on

inert surfaces and while much of the basic concepts can be applied directly to living surface

the process is expected to be even more complex. For the purpose of this review the

formation of a biofouling community will be divided into three basic categories, molecular

fouling, microbial fouling and macro-fouling.

24

Molecular fouling

Microbial fouling

Macro-fouling

• colonisation of bacteria, fungi and diatoms

• formation of a biochemical conditioning film

• colonisation of sessile marine invertebrates and algae

Figure 1.1: Representation of the marine biofouling process. The process usually begins

with the formation of a biochemical conditioning film onto which bacteria and other

microorganisms colonise. Microbial fouling is then followed by the establishment of various

sessile eukaryotic organisms, including invertebrates and algae. See text for further details.

25

1.2.1. Molecular fouling

The first event to take place when a new surface is immersed in an aqueous environment is the

adsorption of both organic and inorganic chemical compounds to form what is known as the

conditioning film (Loeb and Neihof, 1975). The composition of the conditioning film appears

to primarily depend on the properties of the water column or site of immersion rather then the

properties of the surface itself. Little (1985) demonstrated that the properties of the immersed

surface influenced the quantity and composition of the adsorbed film during the first hour of

exposure, however after four hours all the surfaces tested had gained an equivalent level and

composition of conditioning film. The chemical compounds, which commonly constitute a

conditioning film, include amino acids, proteins, monosaccharides, polysaccharides, fatty acids

and humic substances. These chemical compounds will change the properties of the surface

such as wettability and surface charge and so may influence the interaction of bacteria with the

substratum (Dexter, 1978). Thus, molecular fouling plays an important role within the

biofouling process as it is the properties of the initial conditioning film which are likely to

have a significant influence over the extent to which subsequent microorganisms will adhere to

the surface. For example, Schneider et al (1994) investigated the effects of borewater

produced conditioning films on the retention times of the non-motile marine Gram-negative

bacterium SW8. The study showed that most films had an effect on the attachment of the

bacterium to both natural and man-made surfaces, with each film resulting in a distinct

adhesion profile for the organism.

In addition to the types of conditioning films described above, surface active compounds

(SACs) produced by microorganisms may interact with the surface-water interface and affect

the adhesion and detachment of bacteria. Bacterial SACs include low molecular weight

biosurfactants and high molecular weight hydrophobic polymers such as lipopolysaccharides

(Neu, 1996). While SACs have generally been studied for their role in the growth of bacteria

on water-insoluble carbon sources, Neu (1996) highlights the importance of these molecules

in the interaction of bacteria with the surface. SACs are important for bacterial motility

phenotypes such as gliding and swarming (Godchaux et al., 1991; Matsuyama et al., 1992).

Cell bound SACs are responsible for the regulation of bacterial cell surface hydrophobicity,

thus allowing the bacterium to adjust to changing environmental conditions. Furthermore,

Neu (1996) suggests that if SACs influence the interactions of other bacteria with the surface

then they may be candidates for chemically mediated bacterial communication.

26

1.2.2. Microbial fouling

The primary colonisers of any fresh surface are predominantly bacteria and diatoms.

Bacterial colonisation occurs via a two step process beginning with reversible attachment to

the substratum followed by non-reversible adhesion (Marshall et al., 1971). Benthic diatoms

attach to the surface via mucus secretions (Cooksey et al., 1984). A number of factors

influence the ability of the primary colonisers to bind to the surface. These can include both

physico-chemical properties of the surface (temperature, texture, hydrophobicity, nutrient

availability) and biological properties of the organisms such as motility, swarming behaviour

and the production of adhesive molecules.

1.2.2.1. The process of bacterial attachment

Marshall et al (1971) describe the colonisation of bacteria to a surface as consisting of an

initial weak attachment of the cells (reversible attachment) followed by a permanent attachment

(non-reversible attachment), which is aided by the production of adhesins or extracellular

polymeric substances (EPS). Cells reversibly attach to the surface are held primarily by

physical forces and can be easily removed by gentle washing (Characklis and Cooksey,

1983). When both the bacterial cell surface and the conditioning film on the substratum are

predominantly negatively charged the opposing forces of electrostatic repulsion and van der

Waals attraction hold the bacterial cell reversibly at distances of 5-20 nm from the surface

(Wahl, 1989). Recent studies have shown that motility and the presence of cell surface

polymers can greatly facilitate bacterial attachment to glass surfaces (Morisaki et al., 1999).

These findings help to explain how the bacterial cell is able to overcome the energy barrier due

to electrostatic repulsion between the cell and the surface long enough to facilitate non-

reversible attachment. Permanent attachment of bacterial cells to the surface is often mediated

via specific mechanisms such as hydrogen bonding, cation bridging, specific receptor ligand

interactions and the production of EPS, some of which will be discussed in more detail below.

During the initial phase of colonisation bacterial cells enhance their ability to attach by the

production of EPS including, glycoproteins, polysaccharides and lipopolysaccharides all

which act as adhesive molecules. In addition to EPS as adhesins bacterial appendages such as

pili and flagella have been found to play important roles in both the initial stages of attachment

to the surface as well as biofilm formation (see below).

It is possible that specific receptor ligand interactions are very important in bacterial

attachment to both living and inanimate surfaces. On the one hand the specificity of such

27

adhesins may limit the choice of habitats available to the bacterium, however it is also possible

that the same adhesin may provide a selective advantage within a specific site (Fletcher, 1996).

Many of the studies in this area focus on the attachment of bacterial pathogens to host cells.

However several specific cases have also been characterised in marine systems. Attachment of

the marine bacterium Vibrio harveyi to chitin is mediated through the expression of at least

two specific chitin-binding proteins (Montgomery and Kirchman, 1993). Other Vibrio

species have also been suggested to carry similar surface proteins able to bind specifically to

chitin (Pruzzo et al., 1996; Yu et al., 1991). In the case of V. harveyi attachment via chitin-

binding proteins is tightly coupled with the expression of chitin degradative enzymes, thus

giving the bacterium a selective advantage for the acquision of nutrients in the marine

environment (Montgomery and Kirchman, 1994). Vibrio shiloi has been demonstrated to be

the causative agent of coral bleaching in Mediterranean coral Oculina patagonica (Kushmaro

et al., 1997). Coral bleaching results from the loss of the photosynthetic microalgae

endosymbionts known as zooxanthellae, which are located in the coral tissues. Studies have

indicated that V. shiloi is able to infect the host coral via the expression of a specific adhesin

that recognises β-D-galactopyranosides on the coral surface. Interestingly, expression of the

adhesin was shown to be temperature sensitive, being produced when the bacterium was

grown at 25 oC but not when grown at 16 oC (Banin et al., 2000; Toren et al., 1998). V. shiloi

cells then penetrate into the coral tissue and produce both a heat-stable extracellular toxin that

inhibits photosynthesis of the zooxanthellae and heat-sensitive toxins believed to cause

bleaching and lysis of the algae (Banin et al., 2000; Ben-Haim et al., 1999). It has generally

been accepted that coral bleaching is a physiological phenomenon of the coral induced by

elevated seawater temperatures (Gates et al., 1992; Glynn, 1991). However, the finding that

the production of the V. shiloi adhesin is temperature regulated suggests an alternative

hypothesis, that is that elevated temperatures causes the coral-bleaching bacterium to become

more virulent (Toren et al., 1998).

Flagella are large complex structures that span the bacterial cell wall and have been well

studied for their role in motility and attachment of bacterial. Swarming motility is a typical

surface induced phenotype found in a wide variety of Gram-negative bacteria. While the

mechanism varies for different bacterial species, swarming cells in general have multiple

flagella, become elongated and move in a coordinated fashion along the surface (Harshey,

1994). Studies involving the role of flagella in the surface colonisation of the marine

bacterium Vibrio parahaemolyticus have suggested an additional role for the flagella as a

mechanosensor (Kawagishi et al., 1996). Attachment to a surface causes the bacterium to

28

alter its morphology from a planktonic or swimmer cell that is approximately 2 µm in length

and possesses a single polar flagellum to a swarmer cell up to 30 µm and possessing many

lateral flagella. This change in morphology to a swarmer cell helps the bacterium efficiently

colonise the surface to which it is attached (McCarter et al., 1992). It was discovered that

slowed rotation of the polar flagella as the swimmer cell approaches the surface provides the

signal to up-regulate surface specific genes such as the lateral flagella (Kawagishi et al.,

1996). Thus the polar flagella are able to function as mechanosensors, which allows the

bacterium to sense the proximity of a surface.

Transposon mutagenesis has been employed to identify genes responsible for both the

initiation and development of bacterial biofilms in a number of organisms including, E. coli

(Pratt and Kolter, 1998), Pseudomonas aeruginosa (O'Toole and Kolter, 1998a), P.

fluorescens (O'Toole and Kolter, 1998b) and Vibrio cholerae (Watnick and Kolter, 1999).

These studies have extended the model for the development of a mature bacterial biofilm into

three distinct phases including, 1) attachment, 2) surface motility, and 3) biofilm formation.

Furthermore, these genetic screens have shown that the initial interaction with the substratum

and subsequent movement along the surface is largely dependent on surface organelles such

as flagella and pili. For example, studies with E. coli have demonstrated that flagella mediated

motility is important for normal biofilm development being required for surface contact as

well as surface spreading (Pratt and Kolter, 1998). In organisms such as P. aeruginosa and

P. fluorescens flagella are believed to have an additional role in directly adhering to inanimate

surfaces (O'Toole and Kolter, 1998a; O'Toole and Kolter, 1998b). Like flagella, pili have

been found to play essential roles in bacterial attachment and biofilm development. The

presence of type I pili is essential for the initial attachment of E. coli cells to inanimate

surfaces such as PVC. More specifically the mannose-specific adhesin FimH, contained

within the type I pili, was found to be critical for attachment and when not bound to mannose

promoted stable adherence of the cell to inanimate surfaces (Pratt and Kolter, 1998). O’Toole

and Kolter (1998a) found that P. aeruginosa strains lacking type IV pili whilst being able to

attach to PVC and form a mono-layer of cells, do not develop into a multi-layered biofilm.

Thus, unlike the E. coli type I pili, the P. aeruginosa type IV pili are not essential for initial

attachment to inanimate surfaces. Rather, it is the twitching motility associated with the type

IV pili that is proposed to be necessary for cells to migrate along the surface and for the

formation of microcolonies within the developing P. aeruginosa biofilm (see below).

29

As indicated above different bacterial species may use different approaches to initiate biofilm

formation, however it is possible that this is also true for individuals in the same bacterial

species. For example, O'Toole and Kolter (1998b) analysed a collection of surface attachment

defective (sad) mutants from P. fluorescens and found that this bacterium was capable of

adopting multiple strategies for initiating biofilm formation and that such strategies were

determined by environmental conditions. Moreover, recent studies with biofilms of E. coli

strains that express cell surface structures termed curli have suggested that the adaptive

programs used by this organism to promote adhesion and biofilm development to surface

within the host are different to those used for non-host surfaces (Prigent-Combaret et al.,

2000). It was observed that E. coli cells adhere to host tissue using flagella and type I pili

while conditions outside of the host enhance the expression of curli, which the bacterial cells

use to attach to inert surfaces (Prigent-Combaret et al., 2000).

It is also possible that bacteria do not need to possess specific mechanisms for sensing and

attaching to surfaces to be able to adapt to the surface environment. As discussed by

McCarter et al (1992), due to the inherent heterogeneity among individual populations of

bacteria, sub-populations suited for attachment to surfaces would be selectively concentrated at

a surface. Variable expression of adhesive traits in bacteria has been observed, for example,

phase variation of extracellular polysaccharide production in the marine bacterium

Pseudoalteromonas atlantica (formally Pseudomonas atlantica) is related to the presence of

a novel insertion sequence within the eps gene (Bartlett et al., 1988). Thus, it is possible that

for some bacterial species the continual generation of genetic diversity together with the

environmental selective pressure is necessary for these bacteria to respond to life in a biofilm.

1.2.2.2. Bacterial biofilm structure

After the initial attachment to a surface bacterial cells are believed to undergo a program of

physiological changes which result in a highly structured, microbial community (Costerton et

al., 1995). These changes may differ according to the environmental conditions at the surface,

for example the colonisation behaviour for the marine bacterium Psychrobacter sp. SW5

varies depending on the hydrophobicity of the substratum. Cells at hydrophobic surfaces

formed tightly packed biofilms consisting of single and paired cells, in contrast, hydrophilic

surfaces were sparsely colonised and were characterised by the formation of long chains of

cells which were more than 100 µm in length (Dalton et al., 1994).

30

While in natural environments biofilms are most likely to consist of a multi-species

community (see below), the studies performed on single species biofilms have provided a

great deal of information with respect to biofilm structure and the molecular mechanisms

behind biofilm formation. The biofilm structure of organisms such as P. aeruginosa, E. coli

and V. cholerae has been extensively studied. For each organism the biofim consists of a

complex three-dimensional structure, with distinct mushroom or pillar like structures of cells

(microcolonies) embedded in a polymer matrix and surrounded by water-filled channels

(Costerton et al., 1995). The water channels are believed to be important for the diffusion of

nutrients and the release of toxic metabolites out of the biofilm (de Beer et al., 1994). Genetic

studies of P. aeruginosa biofilms have highlighted the role of cell to cell communication in

the development of bacterial biofilms. As is discussed later in this chapter (section 1.4.3) cell

to cell communication in a density dependent manner or quorum-sensing is mediated by small

molecules, the most well studied being the N-acyl-homoserine lactones (acyl-HSLs).

Biofilms formed with mutant P. aeruginosa strains that do not produce acyl-HSLs differed

from biofilms formed with wild-type strains. Both strains attached and proliferated on the

surface, however mutant films were thin and the cells densely packed in comparison to the

wild-type which formed the characteristic three-dimensional structure described above (Davies

et al., 1998). In addition, recent studies in our laboratory using transposon mutants of

Serratia liquefaciens defective in acyl-HSL regulation have identified acyl-HSL controlled

genes important for each phase of the colonisation process (i.e. attachment, swarming and

biofilm formation) (Labbate et al., unpubl.).

Quorum-sensing molecules such as acyl-HSLs have also been suggested to be important

within naturally occurring biofilms. Using acyl-HSL responsive reporter strains McLean et al

(1997) were able to detect the presence of naturally occurring acyl-HSL production in

biofilms growing on submerged rocks. While the function of acyl-HSLs in these biofilms is

unclear, it is possible that they are involved in the establishment of a biofilm or facilitating

intercellular communication between different or the same bacterial species within the biofilm

community.

1.2.2.3. Microbial diversity in natural biofilms

In most natural environments biofilms will consist of a mixture of organisms which form

complex communities. Despite this, very few studies have investigated biofilms with a mixed

community, this may be due to the constraints of traditional methods (i.e. cultivation and

microscopy) which are limiting for the study of in situ microbial diversity. However, advances

31

using molecular based techniques, such as amplification of conserved genes for phylogenetic

studies (ie: 16S rRNA and rpoB), fluorescent in situ hybridisation (FISH), denaturing

gradient gel electrophoresis (DGGE) and terminal-restriction fragment length polymorphism

(T-RFLP) analysis, have opened the door for the study of complex microbial communities

like that found in natural biofilms. A number of these studies have been performed on marine

epibiotic communities and the results indicate that the level of microbial diversity observed is

much greater then previously anticipated (Fisher et al., 1998; Gillan et al., 1998; Polz et al.,

1999; Weidner et al., 1996). For example, a DGGE study of the bacterial diversity in

biofilms covering the shells of the bivalve, Montacuta ferruginosa revealed a complex

community containing over 13 different species. Interestingly, while individual biofilms

analysed using the same techniques did not produce identical DGGE profiles, different

biofilms shared common bands suggesting that similar bacteria may be found on different

biofilms and that these bacteria might be important for the biology of the host organism

(Gillan et al., 1998). Other reports highlight the importance of culture independent

experiments when studying interactions between different members of a biofilm community.

Characterisation of the epiphytic community associated with members of the green algae

commonly known as desmids, revealed that many of sequences obtained represented

previously undescribed bacterial species (Fisher et al., 1998). The authors proposed that

because the majority of what is understood about interspecies interactions has been based on

culture studies, the full range of interactions occurring in natural biofilms is not yet fully

appreciated.

The role fungi play in mixed or natural biofilms is often overlooked despite the fact that they

appear to be effective colonisers of surfaces and exhibit much the same adhesion processes as

bacteria (Jones, 1994). In the marine environment fungi consist of an ecological group

occurring in most marine habitats and they play an important role in marine biofilms on both

living and non-living surfaces (Hyde et al., 1998). Marine fungi are the major decomposers

of woody substrates and may also be important in the degradation of dead organisms. In

addition, fungi are common pathogens of marine plants and animals and are also known to

form symbiotic relationships with these organisms (Hyde et al., 1998). A model bacterial and

fungal biofilm community consisting of seven species was developed by Elvers et al (1998) to

investigate the development of a mixed biofilm community on both PVC and later on stainless

steel (Roberts et al., 1999) surfaces. These studies found that some individual species within

the community form more dense biofilms as single cultures rather than as mixed cultures,

however overall mixed populations were often observed to be thicker and more stable than the

single species biofilms (Elvers et al., 1998). These observations may suggest that the

32

individual species are able to influence each other during the attachment process and may also

interact to enhance the stability of the mature biofilm.

Diatoms and protozoa are also important microbial foulers. While the growth of these

unicelluar eukaryotes usually follows that of the development of the bacterial biofilm there

have been observations of diatom colonisation preceding bacterial attachment (Seiburth and

Tootle, 1981). Benthic diatoms attach via the production of mucus secretions and often

densely cover large areas of a surface (Cooksey et al., 1984). Differential feeding by grazing

protozoa can have a significant impact on the bacterial community within a biofilm (Gonzalez

et al., 1993), which can in turn influence subsequent colonisers.

1.2.3. Macrofouling

The settlement of macrofouling organisms such as sessile plants and animals is generally

considered to be the final event in the biofouling process. Marine sessile plants and animals

undergo a complex life cycle. In all cases this involves a free living spore or larval stage that

is responsible for finding new areas in which to settle. Like the colonisation of the

microfoulers, settlement of spores and larvae is influenced by a combination of physical,

chemical and biological factors. Physical factors such as light, hydrophobicity, texture and

orientation of the surface all effect the degree to which macrofoulers will settle. The

hydrophobicity of a surface will influence the types of macrofoulers that will preferentially

settle. Larvae of the barnacle Balanus amphitrite (Rittschof et al., 1989) and the mussel

Mytilus edulis (Crisp, 1984) prefer to settle on hydrophilic surfaces, while larvae of the

bryozoan Bugula neritina (Rittschof et al., 1989) and the tunicate Ciona intestinalis have been

observed to prefer hydrophobic surfaces (Szewzyk et al., 1991). Studies also indicate that the

surface texture greatly influences settlement density of macrofoulers with most species

preferring rough surfaces then smooth (Harlin and Lindberg, 1977; Hills and Thomason,

1998). In addition, a recent study shows that while variations occur between different species,

there is an overall topographical preference for settlement within pits over elevations (Köhler

et al., 1999). Local currents, tides and water flow also influence the settlement of

macrofoulers.

Chemical cues are an important means by which macrofoulers sense and respond to the

surface environment. Detection of specific positive or negative cues may result in a marine

fouling organism being induced to settle on a surface or being repelled from the surface.

Chemical cues affecting the establishment of marine macrofoulers may be derived from adult

33

conspecifics, symbiotic associations, predator-prey associations or even simply other

organisms competing for the same living space or nutrient. In addition, microbial surface

films may inhibit or induce settlement of macrofoulers in a species-specific manner through

the production of specific chemical cues (Pawlik, 1992). The role of natural chemical cues

with respect to the fouling process is discussed in detail in the following sections of this

review.

1.3. Natural inducers and inhibitors of settlement

1.3.1. Influence of bacteria and their exopolymers on the establishment of

higher organisms

The physical, biological and chemical characteristics of a surface play a major role in the

settling behaviour of fouling organisms. Characteristics such as hydrophobicity, surface

texture and light were mentioned above. In addition to these physico-chemical properties,

bacteria (being among the early colonisers of a fresh surface) are likely to influence the

colonisation of subsequent macrobiota. As early as the 1930's ZoBell and Allen (1935)

suggested that bacteria enhance larval settlement by producing extracellular polysaccharides or

glycoproteins that act as an adhesive layer. More recently Szewzyk et al (1991) studied the

relevance of bacterial exopolymers on the attachment of acidian larvae. Pseudomonas sp. S9

(now Pseudoalteromonas sp. S9) was used as the model bacterium and results indicated that

the exopolymers produced by this organism increased the extent by which the larvae attach.

Depending on the amount of exopolymer produced, larvae can either become trapped or they

can actively attach with the aid of sensory organs. Studies by Kirchman et al (1982a)

demonstrated that larvae of the polychaete Janua brasiliensis are induced to settle in response

to both viable and non-viable bacterial surface films suggesting that the essential cue for

settlement was surface associated. Later studies suggest the settlement inducer is lectin

mediated, whereby lectins on the larval surface recognise and bind to glucose molecules within

the exopolymer of the bacterial cell (Kirchman et al., 1982b). While the majority of studies

have involved mono-culture bacterial biofilms, studies with natural mixed microbial

populations have also supported the importance of bacteria as settlement cues for

macroorganisms. Wieczorek et al (1995) observed that the effects of filmed surfaces towards

barnacle larvae settlement changed from inhibitory to stimulatory with increasing age of the

biofilm. They further concluded that the attachment of various larvae to biofilmed surfaces

34

was due to the combined effect of both active selection and passive entrapment of the larvae to

the substratum (Wieczorek et al., 1995; Wieczorek and Todd, 1997).

The settlement and adhesion of algal spores may also be mediated by bacterial exopolymers.

As is discussed in chapter 2 of this thesis, marine algae are known to interact closely with their

bacterial surface films and studies with the common fouling alga Ulva lactuca indicate that

they may even depend upon these bacteria for normal growth (Provasoli and Pinter, 1964).

Adhesion of the marine alga Chlorella vulgaris to glass surfaces depends on the ability of the

cells to secrete an adhesive material largely consisting of proteins and carbohydrates

(Tosteson and Corpe, 1975). Tosteson and Corpe (1975) found that material recovered from

associated marine bacterial cultures was able to further enhance the adhesion of C. vulgaris.

The authors speculated that the material produced by the bacteria may function as an inducer

for the adhesive polymer synthesis, may stimulate its secretion, stabilise the adhesive once it is

secreted or may actually substitute for the adhesive produced by C. vulgaris. The flagellate,

Dunaliella tertiolecta is also stimulated to attach to surfaces covered with a bacterial biofilm

and this is suggested to be due to lectin mediated interactions (Klut et al., 1983; Mitchell,

1984).

1.3.2. Inducing chemical cues

A number of chemical cues have been suggested to induce settlement and metamorphosis of

marine sessile organisms. However with the exception of a few, the molecular structure and

the ecological relevance of these inducers remain a mystery. Different species of invertebrate

larvae respond to different types of chemical signals. For the purpose of this review these

signals have been broadly categorised into the following groups, neurotransmitters, free fatty

acids and other factors.

1.3.2.1. Neurotransmitters

A large proportion of the research concerned with identifying inducers of settlement has dealt

with artificial compounds that directly stimulate the larval nervous system or affect membrane

permeability. Examples of these include L-dihydroxyphenylalanine (L-DOPA) and related

compounds, gamma-aminobutyric acid (GABA) and its analogues, choline, and ions such as

potassium, calcium and sodium. While little evidence is available to suggest that these

compounds occur naturally in the systems being studied, they are thought to act in a manner

similar to compounds that do occur naturally.

35

1.3.2.1.1. Effects of L-DOPA and catecholamines

Several investigators have reported the observations that oyster larvae (Crassostrea gigas) are

capable of being induced to settle and undergo metamorphosis in response to tyrosine derived

neurotransmitters such as L-DOPA and catecholamines; norepinephrine and epinephrine

(Beiras and Widdows, 1995; Bonar et al., 1990; Coon and Bonar, 1985). The current model

being proposed for oyster settlement and metamorphosis involves two pathways. Control of

settlement behaviour appears to act via dopaminergenic receptor-mediated pathway in which

the larvae are likely to respond to exogenous soluble L-DOPA or L-DOPA analogues. Once

settlement has occurred the larvae release irreversible cement that attaches them to the surface.

Metamorphosis, which acts via the adrenergic neural pathway is then triggered by the release

of norepinephrine and epinephrine. Interestingly, in the absence of chemical stimulation,

oyster larvae were found to be able to delay metamorphosis for up to 30 days (Coon et al.,

1990). It should also be noted that L-DOPA and catecholamines can induce settlement in

other larvae including, the polychaete Phargmatipoma lapidosa californica (Pawlik, 1990),

the gastropod Ilyanassa obsoleta (Levantine and Bonar, 1986) and the bivalve Pecten

maximus. However the same molecules failed to induce larvae from species of barnacles

(Rittschof et al., 1986) and mussels (Pawlik, 1990).

Experiments with both Pacific and Atlantic oysters have lead to the hypothesis that bacterial

films on the surface of the juvenile oyster provide the natural source of L-DOPA. Weiner et

al (1985) isolated the bacterial strain Alteromonas colwelliana (now Shewanella colwelliana)

in close association with the oyster. A. colwelliana was found to produce melanin, a pigment

of which L-DOPA is a precursor. While it remains possible that this bacterium or others are

responsible for providing L-DOPA to the larvae, subsequent research failed to support the

view that L-DOPA is important for oyster settlement and metamorphosis under natural

conditions (Fitt and Coon, 1989).

1.3.2.1.2. Effects of GABA or GABA- analogue molecules

Following observations that juvenile Californian red abalone (Haliotis rufscens) settle

preferentially on surfaces covered with crustose red algae, it was found that extracts of the alga

contain inducers for larval settlement and metamorphosis (Morse and Morse, 1984). The

most abundant of these inducers was revealed to be a small water-soluble compound.

36

Although the exact nature of this molecule is unclear, it was found to have structural and

functional resemblance to GABA. This was supported by the fact that GABA itself was able

to mimic the effects of the natural inducer. Furthermore, the natural inducer purified from the

algae was able to bind to GABA receptors from mammalian brain cells (see (Morse, 1990)).

Induction of settlement and metamorphosis by GABA was found to occur in other larvae

including Astraea undosa and Hydroids elegans (Bryan et al., 1997). A marine

dianoflagellate (Hauser et al., 1975) and the chiton Katharina tunicata (Rumrill and Cameron,

1983) have also been shown to display GABA induced behavioural patterns.

The mechanisms involved in triggering the metamorphosis of larvae by GABA and GABA

analogues is similar to the mechanism that occurs in the mammalian nervous system. By

binding to specific membrane receptors that control the chloride ion channels, the original

chemical signal from the environment can be converted to an electrical signal. This signal is

able to activate the nervous system and thereby elicit the behavioural and cellular processes

that result in further development of the larvae (Morse, 1985). In addition to GABA

analogues, abalone larvae were found to respond to the presence of the amino acid lysine. The

larvae do not settle in response to lysine alone but rather lysine is able to increase the larva’s

sensitivity to a morphogenic inducer up to 100 times (Trapido-Rosenthal and Morse, 1986).

Other researchers have indicated that some invertebrate larvae have the capacity to transport

amino acids similar to GABA into their bodies and have suggested that the morphogenic

response is activated by internal stimulation of the nervous system and not by specific

epithelial chemoreceptors (Jaeckle and Manahan, 1989; Pawlik, 1992).

The ecological and biological relevance of GABA and related compounds is questionable.

GABA can be produced and degraded by microorganisms, thus bacteria are likely to have a

strong influence over the settlement patterns of GABA-responsive larvae. To elucidate the

role of GABA as a natural inducer Kasper and Mountfort (1995) studied the dynamics of the

compound on the natural abalone settlement surfaces. They could not detect GABA on

crustose coralline algae surfaces but rather found biofilms containing GABA-degrading

microorganisms, thus they concluded that it was unlikely that this molecule played a

significant role in Haliotis larval settlement within a natural setting. In addition, Johnson and

Sutton (1994) observed that larvae from the crown-of-thorn starfish (Acanthaster planci) was

induced to settle upon contact with crustose coralline algae. However they found that on

removal of the bacterial biofilm from the surface of the algae (through the use of various

antibiotic treatments) settlement was inhibited. While bacterial isolates from the surface of the

algae did not induce larval settlement, it is possible that specific algal metabolites are required

37

for the bacteria to produce the inducing compound or that a member of the non-culturable

bacterial population produces the compound.

1.3.2.1.3. Induction by ions

As highlighted above inducing stimuli are likely to involve a response by the larval nervous

system. The conduction of electrical impulses in nervous tissue relies on the transport of

certain ions across cell membranes within the tissue. Thus it is of interest to mention the

effects of ionic compounds on the settlement behaviour of marine invertebrates. Increases in

potassium ion concentration has been demonstrated to induce settlement and metamorphosis

of the abalone H. rufescens and is believed to trigger these responses in a number of other

species (Baloun and Morse, 1984; Rodriguez et al., 1993). Calcium ions have been indicated

to be important in the control of metamorphosis of polychaete (Pragmatopoma california)

larvae. Excess of external calcium induces metamorphosis in a concentration dependent

manner (Ilan et al., 1993). These results are in contrast to those reported by Baloun and

Morse (1984) who found that increasing the external calcium concentration was inhibitory to

larvae of H. rufescens. This may reflect the different responses by different larvae to the same

stimulus or it is possible that the effects seen by Baloun and Morse were a result of toxicity of

the abalone to calcium ions (Ilan et al., 1993).

1.3.2.2. Induction by free fatty acids

Free fatty acids (FFAs) have been suggested to induce larval settlement and metamorphosis.

Inducers isolated from the red alga Corallina pilulifera were found to be responsible for the

settlement and metamorphosis of larvae from two species of sea urchins (Kitamura et al.,

1993). In addition, FFAs produced from the bacterium Pseudoalteromonas espejiana

(formally Alteromonas espejiana) were found to enhance settlement responses by the hydroid

Hydractinia echinata (Leitz and Wagne, 1993). Pawlik (1986) isolated similar inducers from

the sand tube Phragmaopoma californica and reported that these molecules were the natural

inducers of gregarious settlement and metamorphosis in the larvae. However, subsequent

studies by Jensen et al (1990) using laboratory reared tube-worms, indicates that the presence

of the FFAs in natural tube-worm material is due to contamination of other biological material.

Therefore, Jensen et al (1990) concluded that FFAs do not take part in any natural gregarious

system that induces settlement or metamorphosis in the tube-worm (however, see response by

Pawlik (1992)). Interestingly, fatty acids have also been demonstrated to have an inhibitory

effect against larval settlement (Goto et al., 1992).

38

1.3.2.3. Induction by other compounds

In addition to the molecules described above, other metabolites derived from a number of

different sources have been isolated and characterised for their ability to induce settlement and

or metamorphosis in responsive larvae. Ammonia has been proposed as a settlement cue for

oyster larvae (Coon et al., 1990). Given that most bacteria and marine mammals secrete

ammonia it could act as an indicator of high biological activity. Other weak bases are also

able to induce settlement, suggesting that the mechanism by which ammonia may act is related

to increases in intracellular pH (Coon et al., 1990).

Peptide cues are commonly found to be important for mediating settlement and metamorphic

responses in marine invertebrates such as oysters (Zimmer-Faust and Tamburri, 1994),

barnacles (Tegtmeyer and Rittschof, 1989) and the sand dollar Dendraster excentricus

(Burke, 1984). Decho et al (1998) suggest that peptides represent a logical choice as signal

molecules in the marine environment for several major reasons. They are water-soluble and

easy to synthesise because the structural components, machinery and templates to produce

peptides are readily available in all living organisms. Also, a variety of signals can be

produced depending on the length and sequence of the peptide produced. Finally intra- and

extracellular proteases are able to degrade peptides (at rates depending on length and sequence

of the peptide) thereby terminating the signal.

While there has been extensive screening for compounds that act as potential cues for

invertebrate larval settlement, few studies have identified the chemical inducer or its ecological

relevance. Recently, a metamorphosis-inducing substance identified as lumichrome was

isolated from extracts of the adult ascidian Halcynthia roretiz. The compound was found to

induce metamorphosis of juvenile larvae from the same species but not larvae of other species,

which is suggestive of species specificity (Tsukamoto et al., 1999). Williamson et al (2000)

have studied the recruitment patterns of the sea urchin Holopneustes purpurascens in

response to chemical cues extracted from various red algae. A cue for metamorphosis was

isolated and characterised from the alga Delisea pulchra (a natural host of the urchin) and was

found to be a water-soluble complex of floridoside and isethionic acid. This complex was

also found to trigger settlement responses of the urchin however the effect was less specific.

39

1.3.3. Inhibitory chemical cues

Woodin (1991) highlighted the fact that inhibition of settlement and metamorphosis in sessile

invertebrates is due to the presence of specific cues and not simply to the absence of

stimulatory ones. Woodin also proposed that such inhibitory signals could be more

predominant in the natural environment than stimulatory signals. In natural systems,

inhibitory cues may play a role in the competition of orgainsims for living space or for

protection against predators. The origin of inhibitory cues may be secondary metabolites

produced by the host orgainsm or from associated bacteria.

1.3.3.1. Inhibitory cues of eukaryotic origin

The ability of many sessile marine plants and animals to remain relatively free from fouling

despite the competition for space by epibionts is well documented. These organisms have

developed means of keeping themselves clean by using a combination of physical and

chemical defences. Physical structures such as spines, the production of mucus and the

sloughing of epidermal tissue are some examples of the non-chemical defences commonly

used. With respect to the chemical defences many invertebrates and algae are known to

produce a wide range of secondary metabolites that inhibit the establishment of both

microorganisms and macroorganisms. Inhibitory compounds have been isolated from

sponges (Davis et al., 1991; Tsukamoto et al., 1997), ascidians (Davis and Wright, 1990),

corals (Mizobuchi et al., 1996; Standing et al., 1984), bryozoans (Kon-ya et al., 1994), sea

grass (Jensen et al., 1998) and algae (de Nys et al., 1994; Todd et al., 1993).

The chemical defenses of three Antarctic soft corals (Alcyonium paessleri, Gersemia

antarctica and Clavularia frankliniana) have been studied in some detail. Laboratory

experiments performed by Slattery et al (1997) suggested that both A. paessleri and G.

antarctica, which do not appear fouled in the field, possess waterborne bioactive compounds

with antifouling activity. In contrast, C. frankliniana, which is often fouled, appears to lack

antifouling compounds. Further research indicated that the bioactive molecules produced by

these corals represented very different classes of compounds. The bioactive compounds

released by A. paessleri were primarily sterols including cholesterol and were found to be

responsible for the deterrent effect on common Antarctic echinoderms. G. antarctica released

a diverse array of metabolites into the surrounding water column including homerine,

trigonelline and several other minor metabolites. Homerine was found to be responsible for

the anti-bacterial activity against three Antarctic bacterial isolates. Interestingly, homerine has

also been shown to deter feeding of sea stars (McClintock, 1994) and to act as an antifoulant

40

in gorgonian corals (Leptogorgia sp.) by preventing growth of the diatom Navicula salinicola

(Targett et al., 1983). Additional antifouling agents were isolated from the gorgonian L.

virgulata and were identified as the diterpenoid hydrocarbons, puklide and epoxypukalide.

These compounds inhibited the settlement of larvae from the barnacle Balanus amphitrite

without killing them (Gerhart et al., 1988). Water-soluble extracts of the eelgrass Zostera

marina inhibit the growth of microalga and several marine bacteria (Harrison, 1982). Todd et

al (1993) isolated cinnamic acid, a phenolic sulphate ester from the eelgrass and found it to be

a natural non-toxic inhibitor of attachment of marine bacteria and barnacles to artificial

surfaces. Fatty acids have also been discovered to act as natural antifoulants. Goto et al

(1992) isolated a mixture of fatty acids with antifouling activity from the marine sponge

Phyllospongia papyracea. Interestingly, various mixtures of the fatty acids displayed

stronger activity compared to the same concentration of a single fatty acid suggesting a

synergistic effect of fatty acids on antifouling activity.

Although the phlorotannins and structurally related compounds from brown algae have

primarily been studied for their effect on herbivores (Hay and Fenical, 1988), these water-

soluble molecules have also been studied for their effect on other fouling organisms such as

invertebrate larvae (Lau and Qian, 1997) and algae (Fletcher, 1975). In addition,

phlorotannins have been shown to be inhibitory to microorganisms (Conover and Seiburth,

1964). Lau and Qian (1997) examined the effect of phlorotannins on the settlement of larvae

from the tube-worm Hydroides elegans and suggested that these molecules were acting in two

ways to inhibit settlement. The compounds may target the macrofoulers directly or they may

regulate the growth of microfoulers such as bacteria, which in turn effects larval settlement.

While there is extensive evidence for the production of antifouling metabolites by many

marine sessile organisms and in some cases the active component has been identified little

attention has been given to determine the ecological relevance of these compounds. In the

case of phlorotannins it has been suggested that these water-soluble compounds are unlikely

to be effective against epiphytes within the natural environment. Jennings and Steinberg

(1997) examined the factors that effect the abundance and distribution of epiphytes on the

brown alga Ecklonia radiata. The authors concluded that there was little or no correlation

between the phlorotannin content of the plants and the abundance of epiphytes. In addition,

experiments designed to test the effects of extracted phlorotannins on the spores of Ulva

lactuca indicated that inhibition of germination only occurred at concentrations (> 10 mg/ml)

which were too high relative to the concentrations that a settling propagule would encounter.

Furthermore, while there is little information available relating to the localization of antifouling

41

metabolites in invertebrates or algae, it has been suggested that unless metabolites are

expressed at the surface of these organism they are unlikely to be successful as natural

fouling deterrents (Steinberg et al., 2001).

In contrast to water-soluble metabolites such as phlorotannins, it has been suggested that non-

polar compounds are more likely to be successful as deterrents because they are able to

adsorb to the surface of the producing organisms. An example of this is the halogenated

furanones produced by the red alga Delisea pulchra. These metabolites have broad-range

biological activities including, inhibition of settlement of fouling organisms (de Nys et al.,

1994) and interference of bacterial signal-mediated regulatory systems, such as those involved

in bacterial colonisation phenotypes (Kjelleberg et al., 1997; Maximilien et al., 1998) (also see

section 1.4.3). Methods have been developed which enable the quantification of non-polar

metabolites on the surface of the alga (de Nys et al., 1998). Together with the use of

fluorescence microscopy this has led to the identification of specialised gland cells on the

surface of the plant which are involved in the localisation and release of the halogenated

furanones (Dworjanyn et al., 1999). These studies also found that the concentration of

furanones at the surface of D. pulchra were at a level above that required to inhibit attachment

of bacteria and the settlement of common macro-foulers, thus confirming the role of these

metabolites as natural antifoulants.

1.3.3.2. Inhibitory cues of bacterial origin

A range of marine bacteria have been shown to prevent fouling organisms from developing.

Results from screening studies indicate that inhibitory bacteria may be quite common. Mary

et al (1993) isolated bacterial strains associated with the barnacle B. amphitrite and found that

12 out of 16 isolates inhibited the settlement of B. amphitrite larvae. The strains were

identified as belonging to the genera Vibrio, Aeromonas, Alcaligenes, Flavobacterium and

Pseudomonas (Mary et al., 1993). Similar studies have been performed by others, for

example Maki et al (1988) screened 18 different bacterial strains for their effect on barnacle

settlement and found 7 species to be inhibitory, with the most inhibitory strain being identified

as Deleya marina (now Halomonas marina). Interestingly, aged biofilms of this strain

displayed greater activity then non-aged biofilms which is in contrast to observations with

aged mixed biofilms (Maki et al., 1988). Later studies with D. marina indicated that the same

bacterium adhered to different substrata elicits different attachment responses by the larvae,

suggesting a complex interaction between the substratum, the bacterium and the larva (Maki et

al., 1990).

42

While there is evidence that bacteria inhibit the establishment of higher organisms little

information is known about the nature of this activity. Kon-ya et al (1995) have identified

ubiquinone-8 as the molecule responsible for the anti-larval activity of the marine sponge

isolate Alteromonas sp. KK10304. Cyanobacteria have also been investigated for antifouling

properties, Scytonema hofmanni produces an anti-algal compound identified as cyanobacterin

which is an effective inhibitor of the diatom Nitzschia pulsilla at low concentrations (Abarzua

et al., 1999). One of the extensively studied antifouling bacteria is Pseudoalteromonas

tunicata. This bacterium was isolated from the surface of an adult tunicate (Ciona

intestinalis) located off the west coast of Sweden. It has been shown to inhibit the settlement

and growth of a number of fouling organisms including invertebrate larvae, bacteria, diatoms,

fungi and algal spores. A detailed description of P. tunicata and the specific compounds will

be presented below (section 1.5.2).

Interestingly, in several cases active compounds believed to be produced by the eukaryotic

host have been demonstrated to be the products of associated bacteria. For example,

antifouling and anti-bacterial metabolites, including alkaloids and bryostatins, previously

isolated from bryozoans are now thought to be derived from their surface-associated bacteria

(Anthoni et al., 1990; Davidson and Haygood 1999). In addition, bryozoans have been

observed to display selective activity against different strains of bacteria (Walls et al., 1993).

The ability of the bryozoan to promote growth of one bacterium over another may represent

an efficient way of manipulating the biofilm so that it provides positive cues to some larvae but

negative cues to potential competitors (Walls et al., 1993).

1.4. Regulation of bacterial secondary metabolites

Bacteria are able to sense and adapt to their environment in order to optimise their ability to

survive and grow. This is particularly important for the expression of specific phenotypes

such as colonisation traits, enzymatic pathways for the degradation of certain compounds,

virulence factors, and secondary metabolites such as antibiotics or toxins. The ability to

regulate these phenotypes at the level of gene expression is crucial for the success of bacteria

that are constantly exposed to differing environmental conditions. Furthermore, in a

competitive environment it is important not to be wasteful, for example it would be of no use

for a bacterium to express specific colonisation traits when it is not near a surface. Similarly,

unless the metabolites have a dual function in the bacterial cell, the production of virulence

43

factors or specific secondary metabolites would be a waste of energy if the bacterium is not in

association with the host or symbiotic organism. The study of bacterial gene regulation is not

only of interest for general bacterial physiology but the regulatory systems involved may also

be potential targets for novel biocontrol applications. There are numerous mechanisms by

which prokaryotes regulate gene expression in response to environmental stimuli, however for

the purpose of this review three examples will be discussed, namely those of 1) two-

component signal transduction systems, 2) the V. cholerae ToxR regulon and 3) quorum-

sensing.

1.4.1. Two-component signal transduction systems

A common mechanism which bacteria use to sense and respond to environmental stimuli is

that of two-component signal transduction systems. A wide range of stimuli can be sensed by

two-component systems, including pH, temperature, nutrient limitation or availability, toxins

and specific repellents/ attractants released by host or symbiotic organisms (Moat and Foster,

1995). A typical two-component system is composed of a sensor kinase protein or a histidine

protein kinase (HPK) and a response regulator (RR) protein. The HPK is usually (but not

always) a transmembrane protein and is responsible for detecting a particular environmental

stimulus. Subsequently the HPK transmits a signal to the RR protein located in the

cytoplasm. The process of signal transduction is through a number of phosphorylation and

dephosphorylation reactions. Interaction with a stimulus induces the autophosphorylation of

the HPK, which occurs at a conserved histidine residue located in the C-terminal cytoplasmic

region of the protein. The phosphoryl-group is then transferred to a conserved aspartate

residue in the RR protein causing an alteration in the activity of this protein. Most RR

proteins are DNA-binding proteins and once phosphorylated bind to specific promoter

regions in the target DNA thereby regulating gene expression (Ninfa, 1996; Parkinson and

Kofoid, 1992). Examples of well studied two-component regulatory systems include the Che-

system that regulates chemotaxis, the EnvZ/OmpR-system for osmoregulation in Escherichia

coli and Salmonella typhimurium, the Vir-signal transduction system which regulates

virulence in Agrobacterium tumefaciens and S. typhimurium, and the Spo-system regulation

sporulation in Bacillus subtilis (Stock et al., 1989). Interestingly, there is increasing evidence

that two-component systems play an important role in the regulation of bacterial colonisation

phenotypes. For example, in strains of the soil microorganism Pseudomonas fluorescens the

ColS/ColR system is important for the attachment of the bacterium to root surfaces (Dekkers

et al., 1998). In E. coli two-component systems, EnvZ/OmpR and CpxA/CpxR are important

for attachment and biofilm formation. It was demonstrated that these two-component systems

44

regulate the expression of csgA, which encodes for one of the major components of curli

(Dorel et al., 1999; Vidal et al., 1998). As mentioned in section 1.2.2.1, curli are cell surface

structures necessary for attachment and biofilm formation of E. coli cells to non-host surfaces

(Prigent-Combaret et al., 2000). In a third example two-component systems PilS/PilR and

AlgR/FimS are thought to act together to regulate twiching motility (a surface associated form

of motility mediated by type 4 pili) in strains of Pseudomonas aeruginosa (Whitchurch et al.,

1996).

Perhaps one of the most widely distributed two-component systems among Gram-negative

bacteria that control expression of secondary metabolites consists of the sensor kinase GacS

(formally LemA) and the response regulator GacA. GacA and GacS were first described to

regulate virulence in the plant pathogen Pseudomonas syringae (Hrabak and Willis, 1992)

and to regulate antibiotic and cyanide production in biocontrol strains of P. fluorescens

(Laville et al., 1992). GacA and GacS have recently been demonstrated to regulate the

expression of a variety of other phenotypes including swarming motility in P. syringae

(Kinscherf and Willis, 1999), protease production in P. fluorescens CHAO (Sacherer et al.,

1994), N-acyl-homoserine lactone production in Pseudomonas sp. (Kitten et al., 1998;

Reimmann et al., 1997; Wood and Pierson, 1996), and virulence in S. typhimurium (Johnston

et al., 1996) and in Vibrio cholerae (Wong et al., 1998).

1.4.2. The ToxR regulon

The Gram-negative bacterium Vibrio cholerae is a common inhabitant of aquatic

environments and is the causative agent of the diarrhoeal disease cholera. The success of V.

cholerae as a human pathogen is largely due to the possession of a number of virulence

factors and to its ability to coordinately regulate the expression of these factors in response to

environmental stimuli. The major virulence factors involved in successful pathogenesis of the

host include the cholera toxin (CT), an enterotoxin responsible for the symptoms of the

disease, and the toxin-coregulated pilus (TCP) which is required for colonisation of the

intestinal mucosa (see Kaper et al., 1995). The genes encoding for the TCP are located on a

40 kb stretch of DNA termed the V. cholerae pathogenicity island (VPI) which is unique to

pathogenic strains of V. cholerae (Karaolis et al., 1998). The subunits for the CT are encoded

by the genes ctxA and ctxB and are located on the genome of a lysogenic bacteriophage

(CTXφ) which uses the TCP as a receptor for infection of V. cholerae cells (Waldor and

45

Melkalanos, 1996). Interestingly, recent studies by Karaolis et al (1999) have indicated that

the VPI is also encoded by a bacteriophage termed VPI phage (VPIφ).

Coordinated expression of V. cholerae virulence factors such as CT and TCP occurs via a

cascade of regulatory proteins referred to as the ToxR regulon (Figure 1.2). ToxR is a

transmembrane DNA-binding protein which is stabilised by the interaction with a second

transmembrane protein, ToxS, in a conformation that is optimal for transcriptional activation

(DiRita and Melkalanos, 1991; Miller et al., 1987; Miller et al., 1989). ToxR and ToxS are

encoded in a single operon, which is located within the ancestral genome of V. cholerae (i.e.

not in association with either the CTXφ or the VPIφ). Interestingly, both ToxR and ToxS are

important for the regulation of ancestral genes in addition to those associated with the CTXφ

and the VPIφ (Miller and Mekalanos, 1988). The ToxR regulon is divided into two distinct

branches based on the requirement for the cytoplasmic transcriptional regulator, ToxT. ToxT

is encoded within the VPIφ and its expression is under the control of ToxR/S (Champion et

al., 1997). In the ToxT-independent branch ToxR directly binds to the promoter region and

controls the expression of specific genes. In addition to ToxT, these genes include the outer

membrane proteins OmpU and OmpT (Li et al., 2000; Miller and Mekalanos, 1988;

Sperandio et al., 1995) and cholera toxin genes, ctxA and ctxB (DiRita, 1992). In the ToxT-

dependent branch ToxR/S act together with another pair of membrane regulators, TcpP/H to

activate transcription of toxT (Häse and Mekalanos, 1998; Higgins et al., 1992). ToxT then

directly activates the expression of the majority of V. cholerae virulence factors including the

CT, the TCP (DiRita et al., 1991) and itself via an autoregulatory loop (Yu and DiRita, 1999).

The mechanisms by which the ToxR regulon responds to environmental stimuli is not well

understood, however some observations suggest that different genes in the regulon respond to

different environmental conditions. For example, ToxT-independent genes, including ompU

and ompT are less influenced by stimuli such as pH and temperature than the ToxT-

dependent genes (Miller and Mekalanos, 1988). ToxR itself is thought to act as both the

major sensor and the signal transducer responsible for controlling gene expression within the

regulon. Studies by Wong et al (1998) have identified a homologue of gacA in V. cholerae

(varA) which regulates expression of TCP and CT and is independent of ToxR. As

mentioned in section 1.4.1, GacA is a member of the two-component family of response

regulators involved in the expression of extracellular metabolites in a number of diverse

Gram-negative bacteria. While the corresponding sensor kinase (GacS) is yet to be identified

46

and the environmental signal is not known, it is possible that this system may contribute to

environmental sensing in V. cholerae. A new level of the ToxR virulence cascade has recently

been identified. The transcriptional regulator AphB was found to act together with AphA to

activate expression of the TcpP/H operon in response to environmental stimuli (Kovacikova

and Skorupski, 1999; Skorupski and Taylor, 1999). In addition, it has also been suggested

that environmental stimuli directly regulate ToxT-dependent transcriptional activation of

virulence factors. Schumacher and Klose (1999) demonstrated that ToxT-dependent

transcription of CT and TCP is significantly reduced by an increase in the temperature to

37oC or in the presence of 0.4 % bile (factors which stimulate the expression of the ToxT

protein). The authors hypothesise that this level of regulation may prevent the premature

expression of virulence factors such as TCP, which would attach the bacterium in an

inappropriate location before it penetrates the mucus lining of the intestine. It is also possible

that this allows for a mechanism by which the bacterium is able to exit the host.

Both ToxR and ToxS proteins appear to be widely distributed among the Vibrionaceae.

Homologues have been studied in several different species including, V. parahaemolyticus

(Lin et al., 1993), V. vulnificus (Lee et al., 2000), V. fischeri (Reich and Schoolnik, 1994) and

Photobacterium profundum strain SS9 (Welch and Bartlett, 1998). Interestingly, a number of

these homologues appear to regulate phenotypes involved in virulence and/ or colonisation

traits. For example in V. vulnificus and V. parahaemolyticus the ToxR homologue mediates

the expression of the hemolysin gene, which is an important virulence factor in these

organisms (Lee et al., 2000; Lin et al., 1993). In V. fischeri the ToxR homologue has been

proposed to be important for successful colonisation of the light organ of various fish and

squid species (Reich and Schoolnik, 1994).

Using degenerate PCR techniques a recent study has shown that the ToxR gene is present in

many other species of Vibrio and Photobacterium, suggesting that ToxR itself is an ancestral

gene of the Vibrionaceae (Osorio and Klose, 2000). Therefore, it would appear that after

acquiring the genetic elements that encode the major virulence factors (i.e. CTXφ and VPIφ),

the V. cholerae ToxR/S proteins evolved to become the master regulators controlling the

expression of these virulence factors. Given that V. cholerae ToxR/S regulates OmpU and

OmpT expression in a ToxT-independent fashion and that the ToxR of P. profundum also

controls outer membrane protein expression, it has been suggested that this is likely to be the

original role of the ToxR transcriptional regulator (Osorio and Klose, 2000).

47

TcpP/H TcpA-F ToxT acf genes

ctxA ctxB

OmpU

OmpT

cytoplasmic membrane

periplasm

cytoplasm

Cholera toxinToxT

ToxSToxRTcpH TcpP

AphA

AphB -

+

++++

++

Figure 1.2: Model of the activation of the ToxR regulon in response to environmental

stimuli. Plus or minus symbols indicate either a positive or a negative effect on gene

expression, broken arrows indicate relevant transcripts. See text for detailed description.

48

1.4.3. Intercellular signalling

It is now widely accepted that many bacteria are capable of intercellular communication via the

aid of small diffusible chemical signals. In Gram-negative bacteria the most extensively

studied systems are those which utilise N-acyl homoserine lactones (acyl-HSLs) as the

signalling molecule to monitor their population density. At low population densities the

bacterial cell produces basal levels of acyl-HSLs which diffuse into the surrounding

environment and accumulate as the cell density increases. Once a critical threshold

concentration of acyl-HSL has been reached, the signal molecule binds to a regulatory protein

that in turn results in the induction or repression of specific acyl-HSL regulated genes. Since

this system of gene regulation relies on bacteria monitoring their own population density and

responding by inducing the expression of particular genes only when a sufficient cell

concentration (or a quorum) has been reached, this process has been termed quorum-sensing.

The ability of a bacterium to coordinately control specific phenotypes in response to cell

density provides an obvious competitive advantage. For example, the regulation of virulence

factors by quorum-sensing systems will allow the bacteria to evade the host defenses by

remaining “silent” until sufficient cell numbers are gained therefore increasing the

pathogen’s chance of a successful host infection. Likewise, bioluminescence may be fruitless

if the bacterium is not associated with other luminescent bacteria within the light organ of its

symbiotic host.

In the following sections cell-density dependent gene regulation via both acyl-HSL and non-

acyl-HSL mediated signalling systems will be discussed in more detail. Given that many

acyl-HSL producing bacteria are in association with higher organisms and these system are

often involved in regulating the expression of colonisation traits and virulence factors (see

below) evidence for eukaryotic interference of acyl-HSL systems will also be addressed.

1.4.3.1. Acyl-HSL signalling systems

Acyl-HSL quorum sensing was first described in the control of bioluminescence of the

marine bacterium Vibrio fischeri (Nealson, 1977) and as such has become the paradigm for

acyl-HSL mediated regulation in Gram-negative bacteria (Figure 1.3 and as reviewed in (Swift

et al., 1994)). V. fischeri can live as a free living organism and as a symbiont in the light

organ of some squid species (Ruby and Lee, 1998). During normal growth of V. fischeri, the

acyl-HSL signal molecule (in this case N- (3-oxohexanoyl)-L-homoserine lactone or OHHL)

is synthesised by the LuxI protein and is diffused into the surrounding environment. At high

49

cell densities (for example in the light organ of the squid) a critical concentration of the

OHHL signal is reached allowing OHHL to bind to the receptor protein LuxR. LuxR-OHHL

then binds to the promoter region of the lux operon thereby initiating transcription of the

structural genes required for the bioluminescent phenotype. Since the gene for LuxI is the

first gene in the lux operon, a positive feed back loop or autoinduction takes place leading to a

further increase in OHHL concentration and consequently an increase in bioluminescence. A

second signalling molecule N-Octanoyl-L-homoserine lactone (OHL) and its synthase protein

AinS have also been identified in V. fischeri. OHL is thought to act as a competitive inhibitor

of OHHL and thus may ensure that expression of the lux operon is tightly repressed at low

cell densities (Kuo et al., 1996).

Acyl-homoserine lactone mediated gene regulation is wide spread among a range of Gram-

negative bacteria and controls a diverse number of phenotypes (for a partial list see Table 1.1).

As mentioned earlier (section 1.2.2.2) acyl-HSL regulation has been implicated in various

aspects of bacterial colonisation and biofilm development. A second theme among the acyl-

HSL regulated phenotypes is that of virulence traits. For example, the common plant

pathogens Erwinia carotovora and Agrobacterium tumefaciens control virulence expression

via an acyl-HSL regulatory system (Costa and Loper, 1997; Fuqua and Winans, 1994). In

addition to the growing number of bacteria that mediate gene expression via acyl-HSL

signalling molecules it has become evident that several bacteria utilise more than one quorum-

sensing system. For example the opportunistic human pathogen Pseudomonas aeruginosa

regulates multiple phenotypes through two interlinked quorum-sensing networks. The two

systems, LasI/LasR and RhlI/RhlR participate in the regulation of certain phenotypes such as

elastase production, while other phenotypes are exclusively under the control of just one

regulatory circuit (Pesci and Igleweski, 1997).

50

luxR luxI lux CDABEG

Extracellular environment

Cytoplasm

Bioluminescence

acyl-HSL synthesis

+

LuxI

LuxR

LuxR

Figure 1.3: Model for the regulation of bioluminescence by intercellular signalling in V.

fischeri modified from (Swift et al., 1994).

51

Table 1.1: Examples of the acyl-HSL regulatory systems in bacteria

Bacterial species Phenotype a Regulatoryproteins

Reference

Aeromonashydrophila

Extracellular proteases Ahyl/AhyR (Swift et al., 1997)

Agrobacteriumtumefaciens

Ti plasmid conjugaltransfer

TraI/TraR (Fuqua and Winans,1994)

Burkholderiacepacia

Virulence CepI/CepR (Lewenza et al., 1999)

Chromobacteriumviolaceum

Pigment production CviT/CviR (McClean et al., 1997)

Erwinia carotovorasubsp.betavasculorum

Virulence, antibioticand exoenzymeproduction

EcbI/EcbR (Costa and Loper, 1997)

Pseudomonasaeruginosa

Virulence, exoenzymes LasI/LasR &RhlI/RhlR

(Person et al., 1997;Pesci and Igleweski,1997)

Rhizobiumleguminosarum

Nodulation, starvation RhiI/RhiR (Cray et al., 1996)

Serratia liquefaciens Swarming,biosurfactant andphospholipaseproduction

SwrI/SwrR (Eberl et al., 1996;Lindum et al., 1998)

Vibrio fischeri Bioluminescence LuxI/LuxR &AinS/AinR

(Engebrecht andSilverman, 1984;Nealson, 1977)(Kuo et al., 1996)

Vibrio harveyi Bioluminescence LuxM/LuxN,LuxO, LuxR

(Bassler, 1999)

a Does not represent all phenotypes controlled by acyl-HSL regulatory systems

1.4.3.2. Non acyl-HSL signalling systems

Beside acyl-HSL mediated signalling bacteria employ other systems of cell to cell

communication. In Gram-positive bacteria quorum sensing has been observed to occur via the

active transport of peptide signal molecules and the action of two-component signal

transduction systems (see above). For example, the development of genetic competence in

both Bacillus subtilis and Streptococcus pneumoniae, virulence in Staphylococcus aureus and

the production of antimicrobial peptides such as the lantibiotic nisin by Lactococcus lactis and

subtilin by B. subtilis are all regulated via small peptide signalling molecules (Kleerbezem et

al., 1997).

52

Some Streptomyces species synthesize γ-butyrolactones which structurally resemble acyl-

HSL and are responsible for regulating the production of the antibiotic streptomycin and for

inducing sporulation (Horinouchi and Beppu, 1992). Regulation of exoenzyme production

and virulence in the plant pathogen Xanthomonas campestris is mediated by a small diffusible

extracellular factor, which is believed to be a fatty acid derivative (Barber et al., 1997).

Another plant pathogen, Ralstonia solanacearum has been demonstrated to posess a more

complex hierarchical signalling system. The novel signalling molecule, 3-hydroxypalmitic

acid methyl ester, was identified in this organism and shown to regulate the expression of

virulence factors through the transcriptional activator PhcA. PhcA in turn was found to

control the production of acyl-HSL, thus regulating the quorum-sensing system in R.

solanacearum (Flavier et al., 1997; Flavier et al., 1997). In addition, later studies showed that

the stationary phase sigma factor RpoS was also able to influence acyl-HSL-dependent

autoinduction in this organism (Flavier et al., 1998).

The marine bacterium V. harveyi uses two quorum-sensing systems to control density

dependent phenotypes such as bioluminescence. One of these systems (system 1) requires an

acyl-HSL molecule as the signal and the second system (system 2) uses a yet unidentified

signal molecule referred to as autoinducer 2 (AI-2) (Bassler et al., 1994). System 1 consists

of the signal synthase LuxLM and the sensor protein LuxN. LuxN however differs from the

LuxR protein in V. fischeri as it is membrane-bound and belongs to the two-component signal

transduction system. System 2 is similar to system 1 with the exception that the signal

molecule produced by the synthase LuxS is not an acyl-HSL. LuxPQ, also a member of the

two-component regulatory system acts as the sensor protein for system 2. Information from

both sensors is passed onto the LuxO protein via a phosphorelay system and the integrator

protein LuxU (Freeman and Bassler, 1999; Freeman and Bassler, 1999). In the absence of

the signal molecule at low cell densities, LuxN and LuxPQ phosphorylate LuxO allowing it to

act as a transcriptional repressor of the lux operon. As the cell density increases, interaction of

the autoinducers with LuxN and LuxPQ causes both sensors to change from kinase activity to

phosphatase activity. Dephosphorylation of LuxO inactivates its activity and allows for the

binding of the transcriptional regulator LuxR (not a homologue of LuxR in V. fischeri)

thereby activating transcription of the lux genes (see review (Bassler, 1999)).

Several recent studies have indicated that a wide range of bacteria may use systems

homologous to the AI-2 system of V. harveyi to regulate a diverse range of phenotypes

(Bassler et al., 1997; Jobling and Holmes, 1997; McCarter, 1998; McDougald et al., 2000).

Furthermore, database searches on both complete and partial genome sequences have

53

identified homologues of the V. harveyi luxS gene in a number of bacterial strains including E.

coli and S. typhimurium (Surette et al., 1999). In addition, ongoing studies in our laboratory

have so far indicated that several species of Pseudoalteromonas, including the antifouling

bacterium P. tunicata may also posess the AI-2 signalling system (Franks et al., unpubl.).

1.4.3.3. Interference of bacterial signalling

A common theme among the acyl-HSL regulated phenotypes that have so far been identified

is colonisation or virulence traits (Table 1.1). Taken together with the fact that many acyl-

HSL producing bacteria are associated with higher organisms, it is not surprising that

eukaryotic hosts might evolve mechanisms to interfere with these important regulatory

systems.

As referred to above (section 1.3.3.1), the red alga D. pulchra produces a number of

secondary metabolites known as halogenated furanones that inhibit the colonisation of a

variety of marine fouling organisms. Maximilien et al (1998) discovered that there is a strong

inverse correlation between the bacterial abundance and the furanone content over the surface

of the plant. Fewer bacterial cells are seen at the plant tip where the highest concentration of

furanone is found. More specifically, the authors found that both individual furanones and

crude algal extracts are able to inhibit bacterial attachment in both laboratory and field

experiments without any effect on bacterial growth (Maximilien et al., 1998). Given that the

structure of some halogenated furanones resembles that of a number of bacterial acyl-HSL it

was hypothesised that these molecules act to specifically interfere with acyl-HSL regulatory

systems, thus inhibiting bacterial colonisation phenotypes such as swarming and attachment.

Indeed, studies performed by Givskov et al (1996) show that furanones are able to interfere

with acyl-HSL mediated phenotypes such as swarming in Serratia liquefaciens and

bioluminescence in V. fischeri. Moreover, evidence has recently been obtained to support the

theory that the furanones effect acyl-HSL mediated regulation by binding or altering the

binding site of the LuxR protein thus displacing the native signal molecule (Manefield et al.,

1999). Therefore, D. pulchra appears to have developed a non-toxic chemical defence

mechanism which involves the inhibition of bacterial colonisation by specifically targeting

acyl-HSL regulated gene expression. Since many bacterial phenotypes important for

colonisation are under the control of regulatory systems that are mediated via chemical signals,

it is possible that interference of these systems by host secondary metabolites is a common

occurrence in natural systems.

54

1.5. The genus Pseudoalteromonas

Revision of the genus Alteromonas using phylogenetic studies performed by Gauthier et al

(1995) has led to the division of this genus into the two genera Alteromonas and

Pseudoalteromonas. The newly established genus Pseudoalteromonas currently contains

both pigmented and non-pigmented, Gram-negative, rod-shaped, heterotrophic marine bacteria

which are motile by a single polar flagellum. Members of this genus are frequently isolated

from marine waters around the world and the majority seem to be associated with eukaryotic

hosts (Holmström and Kjelleberg, 1999). In addition, many of the species produce

biologically active secondary metabolites that target a range of organisms. A list of the

different biological activities displayed by members of the Pseudoalteromonas is given in

Table 1.2 and some will be described in more detail in the following sections.

1.5.1. Biological activities expressed by Pseudoalteromonas sp.

The production of biologically active metabolites is a complex process that can be influenced

by a variety of factors. For example, Ivanova et al (1998) demonstrated that the antimicrobial

activity of some Pseudoalteromonas sp. might be due to both proteinaceous and non-

proteinaceous antibiotics, which are produced during different stages of bacterial growth.

Moreover, they were able to provide evidence that the production of these antimicrobial

metabolites was influenced by the degree of hydrophobicity of the surface substratum. The

highest antimicrobial activity was found to occur on hydrophilic surfaces despite the fact that

hydrophobic surface contained more attached bacterial cells (Ivanova et al., 1998). These

findings suggest that the expression of biologically active metabolites by marine bacteria

including Pseudoalteromonas sp. may be regulated in response to different environmental

stimuli.

Pseudoalteromonas species display a broad range of antimicrobial activities that may aid in

the colonisation of surfaces including those of their host organism. P. aurantia (Gauthier and

Breittmayer, 1979), P. luteoviolacea (Gauthier and Flatau, 1976), P. rubra (Gauthier, 1979),

P. citrea (Ivanova et al., 1998) and P. tunicata (James et al., 1996) are among those species

for which anti-bacterial activity has been observed. The anti-bacterial activity of P.

luteoviolacea is thought to be due to the production of two classes of compounds, a large

acidic polysaccharide, which was demonstrated to be associated with proteins (McCarthy et

al., 1994) and a small brominated cell-bound molecule (Andersen et al., 1974; Gauthier and

Flatau, 1976). The macromolecular antibiotic is thought to act by interfering with bacterial

55

respiration since it induces an increased oxygen uptake and the production of peroxidase in

target bacterial cells (Gauthier and Flatau, 1976). This antibiotic is also responsible for the

autoinhibitory activity observed for P. luteoviolacea cells and is closely related to the

polyanionic antibiotics produced by other Pseudoalteromonas sp. including P. citrea and P.

rubra (Gauthier, 1977; Gauthier, 1979). Cells of P. tunicata produce a large anti-bacterial

protein that is able to inhibit the growth of Gram-positive and Gram-negative bacteria from a

variety of habitats (James et al., 1996). Further details concerning the nature of the anti-

bacterial protein will be discussed in section 1.5.2.

In addition to antibiotics, the production of agarases, toxins, bacteriolytic substances and other

extracellular enzymes by many Pseudoalteromonas species may also assist in the competition

for space as well as in the protection against predators. Several Pseudoalteromonas species

have been identified to produce agarolytic substances, including P. espejiana (Uchida et al.,

1997), P. agarolyticus (Vera et al., 1998), P. citrea (Gauthier, 1977), P. atlantica and P.

carageenovora (Akagawa-Matsushita et al., 1992). Agar is a polysaccharide found in the cell

wall of algae. Bacterial degradation of agar occurs through the action of two specific

enzymes, α and β agarase, and the expression of these enzymes may enable the bacteria to

acquire nutrients from the algae. Other bacteria expressing biological activity against marine

algae include strains of P. bacteriolytica which have been isolated from the brown alga

Laminaria japonica and suggested to be the causative agent of red spot disease in this

organism (Sawabe et al., 1998). Strains of P. elyakovii have also been proposed to cause red

spot disease in L. japonica (Sawabe et al., 2000). P. peptidolytica has recently been described

as a bacterium able to cleave the complex protein compounds within the permanent adhesive

of the mussel Mytilus edulis by secreting an unidentified protease/s (Venkateswaran and

Dohmoto, 2000). An extracellular protease is also responsible for the lysis of marine algae by

the Pseudoalteromonas sp. A28. The purified protease, a monomeric protein of 50 kDa,

displays strong killing activity towards the diatom Skeletonema costatum (Lee et al., 2000).

Moreover, the latter authors suggested that the expression of the proteases is regulated by an

acyl-HSL regulatory system (Kato et al., 1999). Algicidal effects have been observed for

other species of Pseudoalteromonas. A yellow pigmented bacterium designated

Pseudoalteromonas sp. strain Y was isolated from estuarine waters in Tasmania, Australia and

was found to display potent algicidal effects against harmful algal-bloom species (Lovejoy et

al., 1998). The toxic compound was released by the bacterium into the surrounding seawater

where it caused rapid cell lysis and death of algal species within the genera Chattonella,

Gymnodinium and Heterosigma. While the minimum bacterial concentration required to kill

the algae was higher than the average concentrations of the isolate under non-bloom

56

conditions, the authors noted that Pseudoalteromonas sp.Y displayed a chemotactic behaviour

that resulted in localised high concentrations of bacterial cells around the target organism

(Lovejoy et al., 1998). Thus, these observations imply that species of Pseudoalteromonas

may play an important role within the natural environment in regulating the onset and

development of harmful algal blooms.

Production of extracellular toxins has been demonstrated for other Pseudoalteromonas

species. These include tetrodotoxin, a neurotoxin produced by P. haloplanktis subsp.

tetraodonis and the causative agent of pufferfish poisoning (Simidu et al., 1990).

Pseudoalteromonas sp. VL-1, closely related to P. haloplanktis subsp. tetraodonis, also

produces tetrodotoxin and has recently been identified as a pathogen causing mortalities in

populations of the sea urchin, Meoma ventricosa (Ritchie et al., 2000). Strains of P.

piscicidia are responsible for releasing a toxin that has been suggested to cause fish mortality

(Bein, 1954; Hansen et al., 1965) and other isolates of this species have been associated with

diseased damselfish eggs (Nelson and Ghiorse, 1999). In addition, P. tunicata has been

demonstrated to be inhibitory towards invertebrate larvae and algal spores (see section 1.5.2).

Several Pseudoalteromonas and Alteromonas species produce exopolysaccharides that have

been demonstrated to benefit the producing strain (by way of facilitating attachment to a

surface) and to aid in the survival of other organisms. For example Alteromonas sp. strain

HYD-1545, isolated from a polychaete (Alvinella pompejana) located in a deep-sea

hydrothermal vent has been demonstrated to produce a specific exopolysaccharide with heavy

metal binding properties (Vincent et al., 1994). Since the polychaete is commonly exposed to

high concentrations of toxic chemicals such as metalic sulfides, it has been proposed that the

heavy-metal binding-exopolysaccharide is important for survival of the polychaete (Vincent et

al., 1994). Further beneficial effects of exopolysaccharide production by bacteria on other

organisms have been reported, for example, Pseudoalteromonas sp. S9 produces an

exopolysaccharide that can induce the settlement and attachment of larvae from the acidian

Ciona intestinalis (Szewzyk et al., 1991). Likewise, the exopolysaccharide of the oyster-

associated bacterium Alteromonas colwelliana (now Shewanella colwelliana) is thought to

influence the settlement and metamorphosis of larvae from the Eastern Oyster (Crassostrea

virginica) (Weiner et al., 1988; Weiner et al., 1985).

57

Table 1.2: A summary of the biological activities of Pseudoalteromonas sp.

Bacterial strain Biological activity Reference

P. agarolyticus Agarolytic (Vera et al., 1998)P. antarctica strain N-1 Agarolytic (Vera et al., 1998)P. atlantica Agarolytic (Akagawa-Matsushita et al.,

1992)P. aurantia Anti-bacterial, anti-fungal (Gauthier and Breittmayer,

1979;Holmström et al., unpubl.)

P. bacteriolytica Causes disease in algae (Sawabe et al., 1998)P. carrageenovora Agarolytic (Akagawa-Matsushita et al.,

1992)P. citrea Anti-bacterial, anti-fungal

and agarolytic(Gauthier, 1977; Ivanova et al., 1998)

P. denitrificans Autotoxic (Enger et al., 1987)P. espejiana Degrades polymers, induces

metamorphosis in hydroidlarvae

(Uchida et al., 1997)

P. haloplanktis strain S5B Extracellular protease,causes fish spoilage

(Odagami et al., 1993)

P. haloplanktis subsp.tetraodonis

Produces tetrodoxin, whichcauses pufferfish poisoning

(Simidu et al., 1990)

P. luteoviolacea Anti-bacterial (Gauthier and Flatau, 1976)P. piscicida Produces a toxin that causes

fish mortality(Bein, 1954; Hansen et al., 1965)

P. rubra Anti-bacterial, anti-fungal (Gauthier, 1979)(Holmström et al., unpubl.)

P. tunicata Anti-fouling against,bacteria, fungi, invertebratelarvae, algal spores anddiatoms

(Holmström et al., 1998)

P. undina Anti-bacterial, anti-viral (Maeda et al., 1997)Pseudoalteromonas sp.strain A28

Algicidal (Lee et al., 2000)

Pseudoalteromonas sp.strain C-1

Agarolytic (Vera et al., 1998)

Pseudoalteromonas sp. F-420

Anti-bacterial (Yoshikawa et al., 1997)

Pseudoalteromonas sp.strain S9

Induces settlement oftunicate larvae

(Szewzyk et al., 1991)

Pseudoalteromonas sp.strain Y

Algicidal (Lovejoy et al., 1998)

1.5.2. Biological activities expressed by Pseudoalteromonas tunicata

One of the most extensively studied species within the genus Pseudoalteromonas is P.

tunicata. P. tunicata strain D2 was isolated from the surface of an adult tunicate (Ciona

58

intestinalis) at a depth of 10 m in coastal waters of Sweden. P. tunicata is dark green

pigmented due to the combined expression of both yellow and purple pigments. In addition,

the isolate has been found to produce at least five extracellular compounds responsible for

inhibiting the establishment of other organisms within a biofouling community (Figure 1.4).

These antifouling compounds inhibit the settlement of invertebrate larvae and algal spores, the

growth of bacteria and fungi, and surface colonisation by diatoms. Larvae of the marine

invertebrates Ciona intestinalis and Balanus amphitrite are inhibited by biofilms and cell free

supernatant of P. tunicata. The anti-larval molecule has been characterised as a heat stable,

polar, stationary phase produced compound. Size fractionation has suggested that the

molecule is less than 500 Da in size. Treatment with metaperiodate resulted in an increase in

activity, indicating that the molecule may be associated with carbohydrate moieties, or is

released by the cell in a form that is bound to or surrounded by carbohydrate containing

molecules (Holmström et al., 1992). The anti-bacterial activity displayed by P. tunicata has

been studied in detail and the component responsible for this activity has been identified as a

large extracellular protein of approximately 190 kDa (James et al., 1996). This protein

inhibits the growth of both Gram-positive and Gram-negative bacteria from a diverse range of

environments including terrestrial, medical and marine isolates. The marine isolates are

among the most sensitive, with minimal inhibitory concentrations (MIC) of the purified

protein being approximately 2-4 µg/ml (James et al., 1996). P. tunicata cells also display

autoinhibition, in which logarithmic phase growing cells are sensitive to this protein.

However, as P. tunicata cells enter into stationary growth phase, which is when the protein is

produced, they become resistant. The anti-bacterial protein has been purified and is known to

contain at least two subunits of 80 kDa and 60 kDa. The N-terminal amino acid sequencing

of both subunits has demonstrated that for the first 27 amino acids these subunits are

identical. Further analyses including Southern hybridisation experiments and DNA-

sequencing from genomic libraries of P. tunicata indicate that the protein is encoded by a

single open reading frame. This suggests that post-translational modification is required to

produce the two different subunits and thus the active protein (Stelzer, 1999). The anti-

bacterial protein produced by P. tunicata appears to be novel since there was no homology to

other proteins. Furthermore, the protein appears to be the only described large anti-bacterial

protein of bacterial origin. The ecological role of this protein is currently being studied,

however, it has been suggested that the production of an anti-bacterial compound by P.

tunicata may aid the efficient colonisation of surfaces in the marine environment. The anti-

algal and the anti-fungal activities of P. tunicata have been investigated in this thesis and

results will be presented in chapters 2 and 3 respectively. The anti-diatom compound has not

yet been characterised.

59

FungiDiatoms

Pseudoalteromonas tunicata

Invertebrate larvaeBacteria

Algal Spores

Figure 1.4: Antifouling activities expressed by P. tunicata

1.6. Biofouling: the problems and solutions

The formation of a biofilm and biofouling community can often be beneficial. For example,

biofilms can play a major role in the biodegradation of natural organic material and of some

artificial materials that have been released into the natural environment. For living organisms,

the formation of a biofilm may also provide protective camouflage from potential predators.

However, biofouling can also have a negative effect resulting in damage of the object to which

the fouling organisms are attached. The natural formation of a biofouling community on

man-made structures exposed to marine conditions poses significant technical, economical

and environmental problems. For example, the build up of a complex community on the

surface of ship hulls can cause corrosion and increased drag leading to a loss in fuel

efficiency (Gitlitz, 1981; Marshall, 1994). On aquaculture structures the build up of

macroalgae and invertebrates can lead to reductions in water transfer and to a decrease in the

flotation capacity of nets and cages. In addition, the growth of fouling organisms on cultured

60

shellfish (i.e. oysters and abalone) greatly reduces their growth due to increased stress and

competition (Evans, 1988; Lewis, 1994).

Traditionally, the control of biofouling has involved costly and labour intensive mechanical

process coupled with the use of toxic antifouling coatings containing metal compounds such

as copper and tri-n-butyltin (TBT). While toxic antifoulants have been widely used in the past

the application of these coatings especially the TBT-based paints has caused a growing

environmental pollution problem. TBT has been shown to be harmful to many forms of

marine life and there is evidence of bioaccumulation of this compound in marine sediments

and throughout the food chain (Clare et al., 1992; Gibbs et al., 1990; Stewart and de Mora,

1990). Due to the increased risk in the application of these toxic coatings governmental

regulations in countries around the world have been established in an effort to limit their use

(Dalley, 1989; Callow, 1999).

The need to develop safe and economically viable alternative antifouling technologies is the

driving force behind the search for biologically active natural products that may influence the

fouling processes. Furthermore, an understanding of the natural process involved in the

formation and maintenance of marine biofilms and biofouling communities will prove

invaluable in the applications of any natural antifouling product. It should also be pointed out

that the application of marine natural products extends beyond the marine environment. Novel

products from a variety of different marine sources are being screened in the pharmaceutical

industry for their activity as anti-viral, anti-cancer and antibiotic agents. In addition, the use of

natural products as biological control agents in the agricultural industry is of growing interest.

1.7. Aims of this study

All surfaces in the marine environment are subject to biofouling and on man-made structures

this causes significant technical and economical problems. Current methods used to prevent

fouling are costly and in the case of toxic metal coatings environmentally hazardous. Thus

there is a need to develop economically viable and environmentally friendly methods for

biofouling control.

As the primary colonisers of a surface bacteria play an important role in the development and

maintenance of a biofouling community and can be potential sources of natural inhibitors of

the biofouling process. The marine surface associated bacterium Pseudoalteromonas

61

tunicata has been extensively studied for its ability to inhibit the settlement and growth of a

number of common fouling organisms including invertebrate larvae and bacteria. While P.

tunicata has also been shown to inhibit algal spore germination and fungal growth little is

known regarding the properties and the mechanism of these activities. In addition regulation

of the expression of antifouling inhibitors by this bacterium has not been studied. It is widely

accepted that epibiotic bacterial such as P. tunicata can inhibit surface colonisation by fouling

organisms, however little information is available regarding the prevalence or diversity of such

bacteria in marine environments.

The overall aim of this thesis was to examine the antifouling and biocontrol properties of

marine bacteria with an emphasis on the bacterium Pseudoalteromonas tunicata. The specific

aims were:

1. To investigate the anti-algal activity of marine bacteria and to characterise the active

component produced by P. tunicata that is responsible for the inhibition of marine algal spore

germination (Chapter 2).

2. To investigate the anti-fungal activity of P. tunicata and to characterise the active compound

(Chapter 3).

3. To study the regulation of the expression of fouling inhibitors produced by P. tunicata

(Chapter 4 and 5).

4. To assess the prevalence and diversity of marine bacteria with antifouling properties similar

to P. tunicata (Chapters 6 and 7).

62

2. Inhibition of algal spore germination by the marine bacterium

Pseudoalteromonas tunicata

2.1. Introduction

Much of the literature relating to the problem of biofouling is concerned with natural

inhibitory compounds that prevent the settlement and growth of invertebrate larvae. However,

where light is sufficient to maintain growth, fouling by macroalgae becomes of great

importance (Evans, 1981). For example, in the aquaculture industry the build-up of

macroalgae can lead to the reduction in water transfer across nets and cages and to decreases

in the flotation capacity of rafts (Evans, 1988; Lewis, 1994). Despite the importance of algae

in biofouling, there are only a few studies involved in this area of research and while natural

inhibitors of invertebrate larvae have been identified (Holmström et al., 1992; Maki et al.,

1988; Rittschof et al., 1986), no such inhibitors have been isolated for algal spores.

The colonisation of a new substratum by free-living algal spores occurs via a specific

sequence of events involving settlement (location of surface and establishing surface contact),

attachment (permanent attachment to the surface), establishment (formation of a cell wall and

cell polarity) and germination (cell division and outgrowth) (Fletcher and Callow, 1992).

Algal spores may be motile or non-motile depending on the species of algae. Non-motile

spores settle predominantly by physical forces such as gravity and water currents. Motile

spores actively reach the surface, however their swimming speed is often slow in comparison

to the speed of local currents and water flow (Lobban and Harrison, 1994). Whether spores

are motile or not the various physical, chemical and biological properties of the surface can

greatly affect the settlement process, for example algal spores tend to prefer to settle on rough

surfaces with a high surface free energy (Lobban and Harrison, 1994).

There have been a few studies detailing the ultrastructure of algal spores during the settlement

process (Callow et al., 1997; Clayton, 1992; Evans and Christie, 1970; Henry and Cole,

1982). Callow et al (1997) used video microscopy to reveal the details of Enteromorpha

spore settlement and adhesion. They observed that contact between the apical portion of the

cell and the surface involves rapid “top-like” spinning of the spore. After some time

(seconds to several minutes) the spore either swims away from the surface or permanently

63

adheres to the surface. Permanent adhesion usually began with the release of cytoplasmic

vesicles containing an adhesive substance. The spore then contracts against the surface and

the flagella are withdrawn into the cell. At this stage the attached spore begins a process of

surface spreading which involves “amoeboid-like” movements against the surface to obtain

maximum contact between the cell and the substratum. Once the spore has attached the

formation of the cell wall begins followed by spore germination.

Marine algae are known to interact with their bacterial surface-films and depend on bacteria

for normal growth and survival. Provasoli et al (1980) have studied in detail the relationship

between the alga Ulva lactuca and its associated microbial flora. Under axenic conditions

strains of U. lactuca rapidly lose normal morphology but aquire their regular morphology

once appropriate bacteria are grown together with the algae. Similar effects have been

described for other algae including Ulva pertusa (Nakanishi et al., 1996), Enteromorpha

linza, E. compressa (Fries, 1975) and Monostroma oxyspermum (Provasoli and Pinter, 1964;

Tatewaki et al., 1983). Along with enhancing the development of algal spores, bacteria also

play a role in the control of algal growth. Berland et al (1972) studied the toxic effects of

bacteria on marine algae and found that some bacteria were able to inhibit the growth of

various species of marine algae. Thomas and Allsopp (1983) found in similar experiments

that some bacterial isolates encouraged while others discouraged the growth of Enteromorpha

sp. Marine bacteria may also regulate populations of different bloom-forming algal species.

For example, a novel marine Pseudoalteromonas isolate demonstrating algicidal activity is

thought to have an important role in controlling the development of harmful dinoflagellate

blooms (Lovejoy et al., 1998).

The genus Pseudoalteromonas contains species that live in association with marine

invertebrates and algae and produce extracellular compounds that inhibit or control adaptive

and behavioural responses in many target organisms (Holmström and Kjelleberg, 1999).

Currently one of the most studied species in this genus is P. tunicata. This surface-

associated bacterium was shown to effect the normal settlement and growth of a variety of

common marine surface fouling organisms, including larvae from invertebrates Ciona

intestinalis and Balanus amphitrite, various bacteria, fungi and spores from the alga, U.

lactuca (Holmström et al., 1998). To date, the activity against marine algae is poorly

understood. This chapter investigates the anti-algal activity of a collection of marine bacteria

and provides a further understanding of the mechanism by which P. tunicata inhibits the

germination of marine algal spores.

64

2.2. Material and Methods

2.2.1. Strains and culture conditions

Marine bacteria were used in the algal spore assay, including a collection of 55 unidentified

isolates from various marine surfaces (Maximilien et al., 1998) and P. tunicata (Holmström

et al., 1998). All strains were grown on the complex marine medium VNSS (Appendix I)

and stored in 30 % (v/v) glycerol at -80 oC.

2.2.2. Preparation of mono-culture biofilms

The effect of bacteria on the germination of marine algal spores was assessed by exposing

spores directly to monoculture biofilms of various surface-associated marine bacteria.

Bacterial isolates were inoculated from overnight pre-cultures into either six wells of a 24-

multiwell culture plate (Sigma) with each well containing one ml of the VNSS medium for U.

lactuca assays or into six 36 mm petri dishes containing 3 ml of VNSS medium for

Polysiphonia assays. Biofilms were developed by incubation at room temperature for 24 h.

Following incubation the growth media was discarded. The wells were washed three times

with sterile filtered (0.22 µm) seawater and fresh sterile seawater was added to each well prior

to the algal bioassay being performed as described below.

2.2.3. Ulva lactuca bioassay

The effect of bacteria on algal spores was assessed using the common marine alga U.

lactuca. During this study spores were assessed for germination which also reflects effects

on settlement, attachment and the initial establishment of spores. Samples of U. lactuca were

collected prior to sporulation, from rock surfaces located on the coast in Sydney, Australia.

Individual algal plants were washed in sterile filtered seawater and dried for 2 h at room

temperature. To induce sporulation, each plant was placed into a separate beaker containing

sterile filtered seawater and the beaker was positioned near a light source (desk lamp). As a

result of phototactic responses, motile spores concentrated in a region closest to the light.

Spores were collected, added to one side of a watch glass (10 cm) containing sterile filtered

seawater and phototactively attracted toward the light source at the opposite end of the watch

glass. After 5 min the spores which reached the other side of the watch glass were collected

and this step was repeated (this process removes other unicellular organisms which may be

associated with the algal spores as well as selecting for spores with a higher chance of

65

survival). The spores were collected and added to sterile filtered seawater. One hundred

microlitres of this spore suspension were added for every millilitre of test sample (see section

2.2.2). After the addition of spores, the plates were placed in the dark for 2 h to allow for an

even settlement of spores. The spores were then incubated at room temperature and under

natural light. After 24 h, one millilitre of fresh sterile filtered seawater was added to each

well. Algal spore germination was assessed after 3 days using an inverted light microscope

(Zeiss), counts of germinated spores were made for 10 fields of view under a 40 x

magnification lens. All samples were tested in duplicate with spores from three separate algal

plants and compared to controls containing only sterile filtered seawater. When no

differences in germination between the three different algal plants were found, then the data

were pooled to increase the number of replicates.

2.2.4. Preparation of cell-free supernatant

Bacterial cultures were grown shaking in one litre of VNSS for 24 h at room temperature.

Cells were harvested by centrifugation at 13200 x g for 30 min at 10 oC. The supernatant

was discarded and the cell pellet was resuspended in sterile filtered seawater. The bacterial

cells were harvested once more and resuspended in sterile filtered seawater at a concentration

of 0.05 g wet cells/ ml (equal to approximately one tenth of the original culture volume). The

concentrated cell suspension was incubated whilst shaking at room temperature for 24 h and

then centrifuged at 20200 x g for 30 min at 10 oC. The supernatant was collected and sterile

filtered (0.22 µm) for use in algal spore bioassays as described in section 2.2.3 and section

2.2.12.

2.2.5. Dialysis experiment

P. tunicata and the non-inhibitory isolate J1 (see section 2.3.1), were inoculated into 10 ml of

VNSS and incubated for 24 h at room temperature, cells were harvested, washed and

resuspended in the same volume of sterile seawater. Approximately 0.5 ml of this

suspension and a 10-2 dilution was placed into pre-washed dialysis tubing (12000-14000 Da)

(Medicell, International Ltd). The tubing containing the cells was then placed into 36 mm

petri dishes containing sterile filtered seawater and U. lactuca spore germination was

assessed as detailed above.

66

2.2.6. Preparation of crude P. tunicata cell-free supernatant extracts

One litre of VNSS medium was inoculated with 1 % (v/v) of an overnight preculture and

incubated for 24 h at room temperature whilst shaking. Cells were harvested by

centrifugation (13200 x g for 30 min) and the supernatant collected. The supernatant was

then sterile filtered and extracted 3 times with 400 ml of dichloromethane (DCM) (EM

Sciences). The non-polar or organic phase was evaporated to dryness under reduced

pressure, redissolved in sterile filtered seawater and tested for effects on U. lactuca spore

settlement at concentrations of 1, 10 and 100 µg dried extract /cm2 of surface area.

2.2.7. Size fractionation of P. tunicata cell-free supernatant

The approximate size of the active anti-algal compound was determined by fractionation of P.

tunicata cell-free supernatant using Macrosep centrifugal concentrators (Amicon) with filter

pore sizes of 1000, 300, 100, 50, 30, 3 and 0.5 kDa. The filtrates were collected, sterile

filtered and thereafter assayed for the ability to inhibit the germination of U. lactuca spores.

Further size fractionation was performed using a combination of the above concentrators and

pressure dialysis (filter sizes 30, 10 and 3 kDa (Amicon)). Preparation of filters, appropriate

centrifugal forces and length of spins varied for the different pore sizes and were carried out

according to the manufacturer’s instructions.

2.2.8. Assessment of storage conditions on the stability of the anti-algal

compound

To assess the stability of the active compound under different storage conditions samples of

the unfractionated and the less than 30 kDa fraction of P. tunicata cell-free supernatant were

either freeze dried or stored at 4 oC. After one week, the freeze-dried samples were dissolved

in milli-Q water and tested with the 4 oC sample for activity against U. lactuca spore

germination.

2.2.9. Heat treatment of P. tunicata cell free supernatant

To determine if the active compound is affected by heat, samples of the unfractionated and the

less than 30 kDa fraction of P. tunicata cell-free supernatant were treated at 80 oC for 10 min,

allowed to cool to room temperature and sterile filtered. The treated and non-treated

supernatants were then tested for the ability to inhibit U. lactuca spore germination.

67

2.2.10. Protease treatments of P. tunicata cell free supernatant

To further characterise the compound effective against algal spores, P. tunicata cell-free

supernatant was treated with the enzymes carboxypeptidase y and proteinase K. The

enzymes were added to active supernatant preparations at a final concentration of 200 µg/ml.

The mixture was incubated at 25 oC for 20 min for carboxypeptidase y and at 37 oC for 1 h

for proteinase K. The sample was then sterile filtered before it was tested for its effect on U.

lactuca spores. Controls for both enzyme treatments included non-treated supernatant,

seawater plus equal concentrations of enzyme and seawater alone.

2.2.11. Effect of P. tunicata supernatant on the germination of U. lactuca spores

post settlement

The effect of P. tunicata supernatant on the survival of algal spores after settlement was

determined. U. lactuca spores were added to test wells containing seawater and allowed to

settle for 10 - 15 h. Thereafter spores were observed using an inverted microscope to ensure

that they had not germinated. The seawater was then discarded (thus removing spores, which

had not settled in this time) and replaced with sterile filtered P. tunicata supernatant. Spore

germination was assessed as outlined above. The percentage of germinated spores was

compared to the percentage of spore germination when exposed to P. tunicata supernatant

before settlement.

2.2.12. Polysiphonia bioassay

The effect of P. tunicata on the settlement and subsequent germination of algae other than U.

lactuca was assessed using spores from a red alga Polysiphonia. Individual plants of the

epiphytic marine red alga Polysiphonia sp. were collected from fouled Sargassum sp.

collected from the rock surfaces of coastal waters in Sydney, Australia. Fertile Polysiphonia

sp. was identified by the presence of cystocarps on female plants. To induce the release of

the spores from the cystocarps the fertile Polysiphonia were removed, placed into glass petri

dishes containing sterile filtered seawater and incubated at room temperature for 1-2 h.

Approximately 50 spores were pipetted into treatment and control (containing sterile seawater

only) petri dishes (36 mm). The dishes were incubated at a constant temperature of 20 oC

under artificial light. After 48 h the number of settled (i.e. attached and germinated) and

unsettled (i.e. not in contact with the surface) spores were counted under a dissecting

microscope and the percentage settlement determined. The treatments included biofilms of P.

tunicata and a non-inhibitory strain (J1) as well as cell-free supernatants of both P. tunicata

68

and J1 prepared as described above. All samples were tested in duplicate and the experiments

repeated three times for each sample. Data were pooled to increase the number of replicates

if no differences in spore settlement between the experiments were found.

2.3. Results

2.3.1. Effect of bacterial biofilms on U. lactuca spore germination

Initial screening for the effect of marine surface bacteria on U. lactuca germination was

performed by exposing spores directly to mono-culture biofilms of 56 isolates. Bacterial

isolates were assessed as non-inhibitory (70-100 % spore germination); slightly inhibitory

(30-70 % spore germination) or strongly inhibitory (0-30 % spore germination). Of the 56

isolates used in this study 13 were found to have an inhibitory effect and the exposure of

spores to three of these isolates, including the bacterium P. tunicata, resulted in strong

inhibition of spore germination (Table 2.1).

69

Table 2.1: Effect of marine surface bacteria on Ulva lactuca spore germination

Strain Characteristic Strain Characteristic Strain Characteristic

P. tunicata + + R62A - R97 -R3 - R62B + R98 -R7 - R65 - R100 -R8 - R66 - R101 -R17 + + R70 - R104 -R23 - R71 - R105 -R25 - R72 - R106 +R27 - R74 - R107 +R29 - R75 - R111 +R30 - R77A - R115 +R31 - R77B - R120 -R34 - R79A - R122 -R39 - R79B - R127 -R43 - R80 - R129 +R48 - R81 - R130 +R49 - R86 - J1 -R51 - R91 - J3 +R60 + + R94 + J7 -R61 + R96B - SW -

- no effect (70-100% germination); + slightly inhibitory (30-70% germination); ++ totally inhibitory (0-30% germination)

2.3.2. Effect of bacterial supernatant on U. lactuca germination

Three bacterial strains P. tunicata, R60 and J1, were selected from the original screen for

further analysis. P. tunicata and R60 were both selected due to their inhibitory effects on

algal spore germination. Isolate J1 was chosen because it did not affect spore germination.

In order to determine if inhibitory effects are due to cell-bound components or components

released into the surrounding medium in these strains, the effects of bacterial stationary-phase

supernatants were assayed. Stationary phase supernatant was used because in the marine

environment most bacteria spend long periods of time in stationary phase and commonly

produce active secondary metabolites in this stage (Moriarty and Bell, 1993). Moreover, the

anti-larval molecule and the anti-bacterial protein produced by P. tunicata are both reported to

be stationary phase products (Holmström et al., 1992; James et al., 1996). Cell-free

stationary phase supernatants from P. tunicata were shown to have inhibitory effects upon

the germination of U. lactuca spores. As is shown in Figure 2.1, few spores germinated after

exposure to P. tunicata supernatant. The supernatant from bacterial strain R60 was also

inhibitory toward algal spore germination, approximately 30 % of algal spores survived

exposure to the supernatant from this strain (Figure 2.1).

70

SW J1 R60 P. tunicata0

20

40

60

80

100

120

Type of bacterial supernatant

% G

erm

inat

ed s

pore

s

Figure 2.1: The effect of cell-free supernatant of P. tunicata and the bacterial isolates R60

and JI on the germination of U. lactuca spores. The positive control contains seawater only

(SW) and is representative of no inhibition (i.e. 100 % germination). Error bars indicate the

standard deviations for 6 replicates.

2.3.3. Dialysis experiment

Although the cell-free supernatant of P. tunicata was found to be active against spore

germination, when compared to the results for the effects of P. tunicata biofilms, the

supernatants were relatively less active. There are two possible explanations for this

observation: 1) P. tunicata cells produce two active compounds, one which is surface

associated while the other is extracellular. Thus when exposed to biofilms the combined

effect of both components on algal spores would be greater then the extracellular component

alone. 2) The activity is the result of an extracellular product that is unstable and is required

to be continually produced by the cells for inhibition of spore germination.

71

To test if the effect of P. tunicata on algal spore germination is the result of an extracellular

compound alone or in combination with a cell-surface associated compound, bacterial cultures

were placed into dialysis bags within the test wells. The system was successful in keeping the

bacteria in the bags, yet still allowing the transfer of secondary metabolites. As Figure 2.2

shows, the number of spores which settled in test wells containing P. tunicata was

significantly reduced in both the undiluted and the 1:99 dilution of the culture, compared to

the control and to the wells with cultures of a non-inhibitory isolate (J1). These results favour

the hypothesis of an extracellular anti-algal compound that is released by P. tunicata cells.

SW A A* B B*0

20

40

60

80

100

120

Type of bacterial culture

% G

erm

inat

ed s

pore

s

Figure 2.2: The effect of bacterial cultures on the germination of algal spores. Contact

between the bacteria and the spore was avoided by the use of dialysis tubing (see text).

Undiluted 24 h grown cultures of P. tunicata (A) and a 1:99 dilution (A*). Undiluted 24 h

grown cultures of strain J1 (B) and a 1:99 dilution (B*). The positive control for spore

germination contains seawater (SW) only and is representative of no inhibition (i.e. 100 %

germination). Error bars indicate the standard deviations for three replicates.

72

2.3.4. Effects of crude extracts of P. tunicata cell free supernatant on U. lactuca

spore germination

To characterise the anti-algal component from P. tunicata supernatant and to determine if it is

a polar compound an active supernatant was extracted with a 1:1 volume of dichloromethane

(DCM). The dried residue of the DCM phase was resuspended in sterile filtered seawater

and assayed for activity in the U. lactuca spore assay. Exposure of the supernatant extract to

spores had no effect on spore germination at any of the concentrations (i.e. 1, 10 and 100

µg/cm2) tested compared with the media controls (Figure 2.3).

SW

NS

S 1

NS

S 1

0

NS

S 1

00

Supe

rnat

ant 1

Supe

rnat

ant 1

0

Supe

rnat

ant 1

00

0

20

40

60

80

100

120

Type and concentration of extract

% G

erm

inat

ed s

pore

s

Figure 2.3: The effect of crude extract of P. tunicata cell-free supernatant on the germination

of U. lactuca spores. Spores were exposed to 1, 10 and 100 µg/cm2 of supernatant extract

and the equivalent concentrations of an extract of the medium NSS as a control. The positive

control for spore germination contains seawater (SW) only and is representative of no

inhibition (i.e. 100 % germination). Error bars indicate the standard deviations for three

replicates.

73

2.3.5. Fractionation of P. tunicata cell free supernatant

The approximate size of the anti-algal component was determined by size fractionation of the

cell-free supernatant from P. tunicata. Supernatant was filtered through filters with different

cut-off sizes ranging from 1000 kDa to 500 Da and each fraction tested separately in the U.

lactuca germination assay. The P. tunicata supernatant that passed through a cut-off filter of

3 kDa or less did not inhibit germination of algal spores as compared to the control

containing seawater alone (Figure 2.4). The fraction of the supernatant that remained above

the 10 kDa filter also lost the ability to inhibit spore germination. These results suggest that

the active compound is between 3 and 10 kDa in size.

unfractionated <30 kDa >10 kDa <10 kDa >3 kDa <3 kDa <500 Da SW0

20

40

60

80

100

120

140

Size fraction of cell free supernatant

% G

erm

inat

ed s

pore

s

Figure 2.4: The effect of size fractionated cell-free supernatant of P. tunicata on the

germination of U. lactuca spores. The positive control for spore germination contains

seawater (SW) only and is representative of no inhibition (i.e. 100 % germination). Error

bars indicate the standard deviations for three replicates.

74

2.3.6. Heat treatment of P. tunicata cell free supernatant

To further characterise the nature of the inhibitory compound produced by P. tunicata,

supernatant fractions were heat treated at 80 oC for 10 min. The results obtained after

exposure of the algal spores to heat-treated supernatant, indicate that the active compound is

sensitive to heat (Figure 2.5). In the unfractionated sample, 75 % of spores were able to

germinate compared to 10 % germination when exposed to the supernatant prior to treatment.

A similar reduction in activity after heat treatment occurred for the fraction of supernatant less

than 30 kDa.

SW Unfractionated Unfractionated HT <30 kDa <30 kDa HT0

20

40

60

80

100

120

Type of cell free supernatant

% G

erm

inat

ed s

pore

s

Figure 2.5: The effect of both unfractionated and the less than 30 kDa fraction (< 30 kDa) of

P. tunicata cell-free supernatant before and after heat treatment (HT) on the germination of U.

lactuca spores. The positive control for spore germination contains seawater (SW) only and

is representative of no inhibition (i.e. 100 % germination). Error bars indicate the standard

deviations for three replicates.

75

2.3.7. Enzyme treatments of P. tunicata cell free supernatant

In addition to heat treatment, enzymes that target specific molecules can be used to determine

the nature of the component responsible for algal spore inhibition. Inhibitory fractions of P.

tunicata supernatant were treated with broad range proteases to determine if the inhibitory

component is a protein or peptide, as suggested by its heat sensitivity. Results (Figure 2.6)

show that whilst proteinase K did not significantly alter the activity of inhibitory fractions, a

slight reduction in anti-algal activity was evident after treatment with carboxypeptidase y.

76

A

SW Treated Untreated0

20

40

60

80

100

120

Type of cell free supernatant

% G

erm

inat

ed s

pore

s

B

SW Treated Untreated0

20

40

60

80

100

120

Type of cell free supernatant

% G

erm

inat

ed s

pore

s

Figure 2.6: Anti-algal activity of P. tunicata supernatant treated with proteinase K (A) and

carboxypeptidase y (B). The smaller than 30 kDa inhibitory fraction of P. tunicata cell free

supernatant was treated with the enzyme and thereafter assayed for the ability to inhibit U.

lactuca spore germination and compared with untreated samples. The positive control for

spore germination contains seawater (SW) and enzyme and is representative of no inhibition

(i.e. 100 % germination). Error bars indicate the standard deviations for three replicates.

77

2.3.8. Assessment of storage conditions on the stability of the anti-algal

compound

The effect on the stability of the inhibitory fraction of P. tunicata supernatant upon different

storage conditions was assessed. It was found that once partly purified (i.e. the less than 30

kDa fraction) the anti-algal activity remained after one week of storage at 4 oC and the freeze

dried samples retained partial activity as seen in Table 2.2. However, the unfractionated

supernatant stored at 4 oC or freeze dried lost activity.

Table 2.2: Effect of P. tunicata supernatant on the germination of Ulva lactuca spores

Germinated spores (%)

Storage Unfractionated <30 kDa fraction

Freeze dried 76.3 26.24 oC 60.9 7.8Fresh 0.3 4.7

Seawater 100.0 100.0

2.3.9. Effect of P. tunicata supernatant on settled spores

To begin to study the mode of action of the anti-algal component, comparisons were made

between the effect of P. tunicata supernatant before and after settlement of the algal spores.

The results showed that 58.5 % of the spores germinated when exposed to P. tunicata

supernatant after settlement compared with 0.8 % germination of spores that were exposed

prior to settlement. This demonstrates a reduction in the effectiveness of the anti-algal

compound against settled spores.

2.3.9.1. Activity of P. tunicata cells and cell free supernatant against

spores from the red alga Polysiphonia sp.

The ability of the P. tunicata to effect the settlement and subsequent germination of spores

from other groups of marine algae was assessed using spores from the red alga Polysiphonia

sp. The effects of surface biofilms and cell free supernatant of P. tunicata on the germination

of spores from Polysiphonia are shown in Figures 2.7A and 2.7B, respectively. Both

biofilms and the cell-free supernatant were found to prevent the germination of the spores

from the red alga Polysiphonia as compared to that of a non-inhibitory isolate (J1) and

seawater alone.

78

A

SW P. tunicata J10

20

40

60

80

100

Type of bacterial biofilm

% G

erm

inat

ed s

pore

s

B

SW P. tunicata J10

20

40

60

80

100

Type of bacterial supernatant

% G

erm

inat

ed s

pore

s

Figure 2.7: The ability of P. tunicata biofilms (A) and cell free supernatant (B) to inhibit the

germination of spores from the red alga Polysiphonia sp. compared with that of a non-

inhibitory bacterial isolate (J1) and seawater (SW) as a positive control for spore germination.

Error bars indicate the standard deviations for six replicates.

79

2.4. Discussion

This chapter demonstrates that a high proportion of the marine surface-associated bacteria

that were tested inhibited the germination of marine algal spores. Of the 56 isolates used in

this study 13 were found to have an inhibitory effect and the exposure of spores to 3 of these

isolates, including the bacterium P. tunicata, resulted in total inhibition of spore germination

(Table 2.1). The numbers of inhibitory bacteria found in this study is comparable to similar

screens for the ability of bacteria to influence settlement in invertebrate larvae. Holmström et

al (1992) found that 5 out of 40 marine bacterial isolates tested were strongly inhibitory to

the settlement of Balanus amphitrite larvae. Maki et al (1988) also studied the effect of

bacterial surface films on larvae and demonstrated that attachment of B. amphitrite larvae can

vary with the species of bacteria. Such observations are suggestive of strain specificity with

respect to the production of stimulatory or inhibitory cues in the marine surface environment.

Thus, the presence or absence of a certain species of bacteria can potentially influence the

normal growth and survival of other marine surface organisms such as invertebrates and

algae.

While several reports are available demonstrating the effects of bacterial assemblages on the

settlement or subsequent germination of algal spores (Berland et al., 1972; Dillon et al.,

1989; Provasoli and Pintner, 1980; Thomas and Allsopp, 1983), few attempts have been made

to determine the nature of such interactions. Dillion et al (1989) suggested that the high level

of free energy found on surfaces containing microbial biofilms contributed to an increase in

adhesion by Enteromorpha spores, however the possibility of a specific biochemical

interaction was not ruled out. To further investigate the nature of the algal inhibitory activity

observed in this work three of the bacterial strains, two inhibitory (P. tunicata and R60) and

one non-inhibitory (J1) strain, were selected and the cell-free supernatant from each strain

was tested for its effect on algal spores. The cell-free supernatant from strain J1 had no

effect, however supernatant from strains P. tunicata and R60 prevented the spore germination,

with the supernatant of P. tunicata being the more effective of the two (Figure 2.1). These

data indicate that the anti-algal activities of the bacterial strains are due to the release of

extracellular inhibitors. These finding are further supported by the results of the dialysis

experiments (Figure 2.2) which show that contact between cells of P. tunicata and U. lactuca

spores is not necessary for inhibition. From this study it appears that P. tunicata is

particularly effective in its ability to regulate the settlement and/ or germination of algal spores

in the marine environment. Interestingly, other species of Pseudoalteromonas have been

reported to express algicidal activity. The bacterium Pseudoalteromonas sp. strain Y is

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suggested to play an important role in the control of harmful algal-blooms by the production

of an extracellular inhibitor that causes rapid lysis of algal species (Lovejoy et al., 1998). In

addition, the bacterium, Pseudoalteromonas sp. A28 was demonstrated to lyse marine algae

via the production of extracellular proteases (Lee et al., 2000).

P. tunicata secretes a range of biologically active metabolites, these include a polar, heat

stable, anti-larval molecule of less than 500 Da (Holmström et al., 1992), a 190 kDa anti-

bacterial protein (James et al., 1996) and a pigment that inhibits the growth of fungi (chapter

3). To determine if one of the above metabolites was also responsible for the effect on algal

spore germination P. tunicata was analysed in more detail with respect to the nature of the

anti-algal factor. The basic characteristics of the anti-algal compound are summarised in

Table 2.3. Extraction of active supernatant with the organic solvent DCM and subsequent

testing of the non-polar phase indicated that the molecule is polar and thus likely to be

efficiently dispersed in the marine environment. In addition, these data suggest that the

activity is not related to the anti-fungal pigment produced by P. tunicata cells, which is a non-

polar molecule (chapter 3). Size fractionation of the supernatant using pressure filtration

indicates that the anti-algal compound is between 3 and 10 kDa (Figure 2.4). Anti-algal

activity was not observed in the fraction less than 500 Da or within the range of 190 kDa,

clearly showing that neither the anti-larval nor the anti-bacterial protein are responsible for the

activity against algal spore germination. Heat treatment of the active supernatant indicates that

the anti-algal factor is heat sensitive. Seventy-five percent of spores are able to germinate

when exposed to heat treated unfractionated supernatant compared with 10 % germination

when exposed to untreated supernatant, a similar reduction occurred for the less that 30 kDa

fraction (Figure 2.5). To further characterise the anti-algal component, inhibitory fractions

were treated with the broad range proteolytic enzyme, proteinase K and carboxypeptidase y.

Treatment with these enzymes had little effect on the anti-algal activity of the inhibitory

fractions of P. tunicata supernatant (Figure 2.6). While loss in activity due to the exposure

of these enzymes would give a strong indication that the inhibitory compound is

proteinaceous, no effect does not prove that it is not. It is feasible that low molecular weight

proteins and peptides are less affected by heat and more resistant to the effects of proteolytic

enzymes. Examples of such peptides are the heat stable enterotoxins produced by

Escherichia coli (Robins-Browne, 1994). In addition, the anti-algal component was found to

be unstable in the unfractionated supernatant when stored at 4 oC or freeze dried as indicated

by a loss in activity over a 7 days period. In contrast, when partially purified the compound

appeared less sensitive to degradation (Table 2.2).

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Table 2.3: Characteristics of the anti-algal compound produced by P. tunicata

Conditions or treatments of bacterial supernatant Anti-algal activity

Biofilms active

Cell free supernatant active

DCM extraction (non-polar phase) inactive

Size fraction active within 3 - 10 kDa range

Heat treatment inactive

Proteinase K treatment active

Carboxypeptidase y treatment active

The need to replace current methods for antifouling has opened up a new field of research

involved in understanding the interactions that occur in marine fouling communities. This

study has raised several new questions and suggestions for future research. For example,

studies concerning the mode of action of the anti-algal component produced by P. tunicata

would not only be of ecological interest but also important for commercial application.

Preliminary evidence indicates that the anti-algal compound has a reduced effectiveness once

the spores have settled (section 2.3.9). This observation may be explained by a biocidal

mode of action for the anti-algal compound that targets components within the plasma

membrane of the spore. Prior to settlement, the plasma membrane of the spore is exposed,

however, upon settling spores rapidly begin to form a protective cell wall (Braten, 1971). A

second proposed mechanism to explain the reduced effect of P. tunicata supernatant on

settled spores is that the inhibitory component serves as a negative cue for settlement, thus

simply preventing settlement of the spores until death ensues.

The ability of the P. tunicata to effect the settlement and subsequent germination of spores

from other groups of marine algae was assessed using spores from the red alga Polysiphonia

sp. Polysiphonia is a filamentous marine alga with world-wide distribution. Both biofilms

and cell-free supernatant of P. tunicata were shown to significantly inhibit the germination of

Polysiphonia sp which indicates a more general inhibitory effect against marine algal spores

(Figure 2.7). This observation is similar to that found for the anti-larval molecule produced

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by P. tunicata which is effective against a range of marine invertebrate larvae (Holmström et

al., 1992). Berland et al (1972) have also demonstrated a broad range algicidal activity for a

number of bacterial isolates. In contrast, the chemically mediated attack on unicellular algae

by marine bacteria such as Flavobacterium sp. strain C49 and Pseudoalteromonas sp. strain

Y appears to be specific for certain algal species (Lovejoy et al., 1998; Yoshinaga et al.,

1997).

In summary, marine surface organisms are under intense competition for living space. It is

therefore not surprising that complex interactions occur between not only individuals of the

same or closely related species, but also between vastly different organisms. The aim of

chapter 2 was to study the inhibitory effect of marine surface bacteria on the germination of

marine algal spores. It was shown that a number of marine surface bacteria, including the

previously described antifouling bacterium, P. tunicata, possess anti-algal activities. Further

characterisation of the anti-algal activity of P. tunicata was performed and results indicate that

this bacterium produces an extracellular component with specific activity toward algal spores

that is heat sensitive, polar and between 3-10 kDa in size. This biologically active compound

was also found to prevent germination of spores from the red alga Polysiphonia sp.

suggesting that it may be effective against a variety of marine algae.

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3. Anti-fungal activity of Pseudoalteromonas tunicata

3.1. Introduction

In the marine environment fungi represent an ecological group which occurs in most marine

habitats and which plays an important role in biofilms on both living and non-living surfaces.

Fungi are often essential in both marine and non-marine environments as decomposers and

commonly form important symbiotic relationships with plants and animals (Hyde et al.,

1998). In addition fungi also cause disease in humans, are responsible for food spoilage and

are a major cause of plant diseases in many economical valuable crops (Prescott et al., 1990).

Methods of fungal control used today include traditional practices such as crop rotation,

eradication and quarantine, together with the use of either chemical fungicides or biological

control agents. The characteristics of chemical fungicides mean that they are often non-

specific and can also effect many beneficial organisms. Therefore, safety towards humans

and the environment as well as the development of resistance are some of the major concerns

with the use of chemical fungicides. Due to these increasing concerns the search for new and

safe methods for efficient fungal control has turned toward the use of natural products and

biological control methods.

Within their natural habitat many animals, plants and microbes have developed efficient

methods to protect themselves against fungal over-growth by the production of anti-fungal

compounds. In some cases animals and plants are able to use the anti-fungal properties of

symbiotic bacteria to provide a defence against potentially pathogenic fungi. For example, the

surface of embryos of the American lobster, Homarus americanus, are inhabited by a single

microbe which produces the anti-fungal substance 4-hydroxyphenethyl alcohol (tyrosol),

thus providing protection for the embryos against fungal disease (Gil-Turnes and Fenical,

1992). In a similar interaction, embryos of the shrimp Palameom macrodactylus are

protected by an anti-fungal compound (2-3-Indoline-dione) produced by Alteromonas sp.

(Gil-Turnes et al., 1989).

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The use of bacteria and their metabolites as biocontrol agents is of great interest. For

example, in the agriculture industry strains of Pseudomonas and Bacillus have been studied

for their effectiveness as biological control agents of fungal plant diseases such as take-all

and damping-off (Ryder and Rovira 1993). The biocontrol capabilities of the Pseudomonads

largely result from their ability to produce a variety of anti-fungal metabolites each with a

unique spectrum of activity. Some of the specific metabolites that have been characterised

from Pseudomonas sp. include phenazine carboxylic acid (Thomashow and Weller, 1988),

pyrrolnitrin (Arima et al., 1964), pyoluteorin (Howell and Stipanovic, 1980), 2, 4-

diacetylphloroglucinol (Keel et al., 1990), and hydrogen cyanide (Voisard et al., 1989).

Interestingly, studies have shown that expression of these metabolites is under the coordinate

regulation of the GacS/GacA two-component regulatory system (Gaffney et al., 1994; Laville

et al., 1992). Strains of Bacillus subtilis have potential as biocontrol agents of important

plant diseases due to the production of the novel anti-fungal metabolite, fengycin

(Vanittanakom et al., 1986). Fengycin is a lipopolypeptide that inhibits filamentous fungus

but is ineffective against yeast and bacteria. Studies using transposon mutagenesis identified

the genes involved in the production of the fungicide and suggest that it is synthesised non-

ribosomally by the multi-enzyme thiotemplate mechanism (Chen et al., 1995; Tosato et al.,

1997).

As discussed in chapters 1 and 2, Pseudoalteromonas tunicata is an effective inhibitor of

many common marine biofouling organisms including larvae, algae, diatoms and bacteria.

Previous work has established that cells of P. tunicata are also able to inhibit the growth of

fungi (James, 1998). This chapter describes the identification, using chemical analysis

complemented with genetic analysis, of a compound with specific growth inhibitory activity

towards a number of fungal isolates.

3.2. Materials and Methods

3.2.1. Anti-fungal bioassay

Fungal isolates were cultured on VNSS (Appendix I) agar plates (stock cultures stored on

VNSS plates at room temperature) and incubated at 30 oC for 48 h prior to being used in the

bioassay. Fungal suspensions were prepared by inoculating a loop full of the target fungus

into 500 µl of VNSS medium. Two-hundered mircolitres of the fungal suspension were

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plated onto VNSS agar plates and allowed to air dry. Thereafter Pseudoalteromonas

tunicata wild-type and mutants were inoculated in small circles from a fresh agar plate onto

the plate containing the target fungal suspension. Plates were incubated for 48 h or until the

fungi had created an even lawn of growth. Zones of inhibition were visible at this time

surrounding the bacterial inoculations.

Testing of bacterial extracts for anti-fungal activity was performed using sterile filter paper

discs (5 mm diameter Whatman paper). Dried extracts were resuspended into the appropriate

solvent at equal concentrations according to total dry weight of extract, spotted onto filter

paper discs and thereafter allowed to dry at room temperature. The filter paper discs

containing the extracts and solvent controls were then placed onto the surface of a VNSS agar

plate containing a lawn of the target fungus (Penicillium sp). Plates were dried and incubated

for 48 h at 30 oC then assessed for zones of fungal growth inhibition.

3.2.2. Transposon mutagenesis

The transposon mutagenesis protocol established by James (1998) was used to generate a

specific non anti-fungal mutant of P. tunicata. Fifty millilitre overnight cultures of both

donor E. coli Sm10 (containing pLOF mini-Tn10 system) and the streptomycin resistant

recipient P. tunicata (Sm R) were prepared. The E. coli strain was grown shaking at 37 oC in

LB10 medium (Appendix I) containing 85 µg/ml kanamycin (Km) and 100 µg/ml ampicillin

(Amp). P. tunicata (Sm R) isolates were grown shaking at room temperature in VNSS with

200 µg/ml streptomycin added.

Donor and recipient cultures were mixed at a volume ratio of 1:3 (50 µl E. coli + 150 µl P.

tunicata) in 5 ml of wash solution (50% NSS: 50% 10 mM MgSO4) and gently mixed by

inversion. The mixture of donor and recipient was filtered through a 0.22 µm filter (2.5 cm

diameter) and washed with another 5 ml of wash solution. The filters were then placed cell

side up onto LB15 plates (Appendix I) containing 3 mM isopropyl-β-D-thiogalactoside

(IPTG) and incubated for 4 h at 30 oC. After incubation the filters were placed into

Eppendorf tubes with 1 ml of NSS (Appendix I) and vortexed.

Screening for the loss of anti-fungal activity was performed by mixing fungal spores with the

conjugation mix just after their suspension from filters. The mixture of fungal spores and

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bacterial cells was then plated together onto VNSS with Km (85 µg/ml) and Sm (200 µg/ml)

to select for recipient P. tunicata strains carrying the mini-Tn10 transposon. Agar plates

were then incubated for 48 h at 30 oC. Mutant isolates without inhibition zones and

remaining dark green in pigmentation were selected and retested for the loss of anti-fungal

activity as described above.

3.2.3. Phenotypic characterisation of the non anti-fungal transposon mutants

3.2.3.1. Growth curves

A comparison of the growth rates for the non anti-fungal transposon mutants and wild-type

P. tunicata was performed. The strains were grown in 500 ml flasks containing 200 ml

VNSS medium for wild-type and VNSS medium with the antibiotics Km (85 µg/ml) and Sm

(200 µg/ml) for the non anti-fungal transposon mutants, FM1, FM2, FM3 (see section 3.3.2).

One percent (v/v) of an overnight culture was inoculated into appropriate flask and incubated

shaking at 23 oC. Growth was monitored by absorbence readings (610 nm) over a 24 h

period. This experiment was carried out in duplicates.

3.2.3.2. Anti-bacterial activity

Activity against the growth of bacteria was performed on VNSS agar plates (Appendix I).

Fresh colonies of each of the test strains including FM1, FM2, FM3 and wild-type P.

tunicata were spot inoculated and air dried. In order to obtain sufficient growth of test

bacteria to provide measurable inhibition zones of the target strains, the plates were incubated

for 7 days at 23 oC. However, it should be noted that growth inhibition could also be detected

from 2 days old bacterial colonies (James et al., 1996). The test strains were thereafter

overlaid with agar containing 0.4 ml of an overnight culture of a target bacterial strain per 3

ml agar. Anti-bacterial activity was determined after a further 24 h incubation by a zone of

inhibition surrounding the target strains.

3.2.3.3. Anti-algal activity

Activity against the germination of algal spores was determined for both Ulva lactuca spores

and Polysiphonia sp. spores. Overnight cultures of the strains FM1, FM2, FM3, wild-type

P. tunicata and the previously determined non-inhibitory marine isolate J1 (see section 2.3.1)

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were prepared. The culture were used to inoculate 24 well culture plates or petri dishes (36

mm) containing VNSS medium for wild-type P. tunicata and the marine isolate J1. VNSS

containing the antibiotics Km (85 µg/ml) and Sm (200 µg/ml) was used for the non anti-

fungal mutant strains. Dishes were incubated for 24 h to form biofilms and washed twice

with sterile filtered seawater prior to assays being performed as outlined in chapter 2 (section

2.2.3 and 2.2.12).

3.2.3.4. Anti-larval activity

Invertebrate larval settlement assays were performed with the help of Dr Carola Holmström

and Sophia McCloy from the Centre for Marine Biofouling and Bio-Innovation, UNSW.

The assays were performed using standard settlement assays against larvae of the tube worm

Hydroides elegans and cyprid larvae of the barnacle Balanus amphitrite (de Nys et al., 1994;

Holmström et al., 1992). Briefly, overnight cultures of the strains FM1, FM2, FM3, wild-

type P. tunicata and a non-inhibitory marine isolate (Holmström et al., unpubl.) were

prepared. The cultures were used to inoculate petri dishes (36 mm) containing VNSS

medium for wild-type P. tunicata and the non-inhibitory marine isolate and VNSS containing

the antibiotics Km (85 µg/ml) and Sm (200 µg/ml) for the non anti-fungal mutant strains.

Dishes were incubated for 24 h to form biofilms, washed twice with sterile filtered seawater

and invertebrate larvae were added. The number of settling larvae was determined

microscopically after 3 days incubation at 25 oC and compared to controls containing sterile

filtered seawater.

3.2.4. Genotypic characterisation of the non anti-fungal transposon mutants

3.2.4.1. Genomic DNA extractions

Genomic DNA was extracted from bacterial cultures using the XS-buffer protocol outlined

below (Tillett and Neilan, 2000). An XS-buffer solution was prepared from the following

reagents: 0.5 g potassium ethyl xanthogenate (Fluka); 10 ml 4M ammonium acetate; 5 ml 1M

Tris-HCl pH 7.4; 2 ml 0.45M EDTA; 2.5 ml 20% SDS (w/v) and water up to 50 ml. Two

millilitres of an overnight culture were pelleted in an Eppendorf tube, the supernatant was

removed, cells resuspended into 1 ml of XS buffer and incubated at 70 oC for 60 min. After

incubation the tube was vortexed for 10 sec and incubated on ice for 30 min. The tube was

then centrifuged at 21000 x g for 10 min at 4 oC and the supernatant carefully removed into a

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fresh 2 ml Eppendorf tube. To precipitate the nucleic acids one volume of isopropanol was

added and the tube gently shaken until a stringy white precipitate was visible. The precipitate

was removed by spooling with a glass rod. Thereafter the nucleic acids were washed with

70% (v/v) ethanol, air dried and resuspended into 50-100 µl of sterile milli-Q water. When

little or no white precipitate was visible the nucleic acids were pelleted by centrifugation

(21000 x g for 5 min at 4 oC) and the supernatant removed. The pellet was then washed once

with 70 % (v/v) ethanol, lyophilised in a vacuum centrifuge and resuspended in 30-50 µl of

milli-Q water.

Following extraction, DNA was visualised on a 1% (w/v) agarose gel along side a molecular

weight standard (λ-DNA digested with EcoRI/ HindIII) to assess integrity and concentration.

If required DNA samples were treated with RNase, then extracted with phenol / chloroform /

isoamylalcohol and ethanol precipitated (see Appendix III).

3.2.4.2. Panhandle-PCR method for sequencing within uncloned genomic

DNA

Since the DNA sequence of the mini-Tn10 transposon is known it is possible to use the

panhandle polymerase chain reaction (panhandle-PCR) method to obtain sequence

information of the genes disrupted by the transposon. The panhandle-PCR method (adapted

by D. Tillett from (Siebert et al., 1995) and as summarised in Figure 3.1) relies on the

concept of “suppression PCR”. Due to the nature of the adaptor molecule that contains

inverted terminal repeats, PCR products with adaptors at both ends will form so called

“panhandle” structures following every denaturation step. Since these structures are more

stable than a primer / template hybrid, further amplification is suppressed. In contrast, PCR

products which are formed by a specific primer (in this case, one designed from the

transposon sequence) and an adaptor primer have the adaptor sequence at one end only and

will not form the panhandle structure, allowing PCR amplification to continue.

3.2.4.2.1. Preparation of adaptor ligated DNA

Genomic DNA extracted from non anti-fungal mutants was used for restriction digest and

ligation of adaptor molecules in a one step process. One microgram of genomic DNA, 1µl

(10 pmol/µl stock) of adaptor 1 (Appendix II), 1µl (10 pmol/µl stock) adaptor 2 (Appendix

II), 40 mM ATP, 2.5 units T4 ligase (Boehringer Mannheim), 10 units of blunt-end

89

restriction enzyme (various, see Table 3.1), 10 x One-Phor-All buffer PLUS (Pharmacia) (see

Table 3.1) and milli-Q water to give a final reaction volume of 20 µl were incubated at 20 oC

for 16 h. After incubation the reaction was deactivated at 68 oC for 10 min. DNA was then

precipitated using ethanol (see Appendix III) the resulting pellet washed with 70 % (v/v)

ethanol and resuspended in 50 µl of sterile milli-Q water. This solution served as the

template DNA for the PCR reactions described below.

3.2.4.2.2. Panhandle-PCR

PCR were performed in a 20 µl volume, containing 1 µl of template DNA (as described

above), 2 µl Taq 10 x reaction buffer containing 25 mM MgCl2 (Boehringer Mannheim), 1 µl

of a 10 mM dNTP mix (Boehringer Mannheim), 10 pmol of adaptor primer 1 (Appendix II)

and 10 pmol of the gene specific primer (Tn10C or Tn10D see Appendix II). One unit of a

thermostable DNA polymerase blend of Taq (Boehringer Mannheim) and PFU (Stratagene)

in a unit ratio of 10:1 was added to each reaction after a hot start (95 oC for 3 min). The cycle

parameters were as follows: denaturing step at 95 oC for 30 sec and annealing/ extension at

68 oC for 7 min, the number of cycles varied between 25 and 30 depending on the template.

Table 3.1: Restriction enzymes used for panhandle-PCR

Restriction enzyme 1 Recognition sequence Final concentration of One-Phor-All buffer PLUS

Dra I TTT↓AAA 1xEcoRV GAT↓ATC 2xHincII GT (T,C)↓(A,G)AC 1xHpaI GTT↓AAC 1xPvuII CAG↓CTG 1xRsaI GT↓AC 1xScaI ACT↓ACT 2xSspI AAT↓ATT 2xXmnI GAANN↓NNTTC 1x

1 All enzymes were purchased from either Boehringer Mannheim or Pharamacia.

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3.2.4.3. Preparation of PCR templates and DNA sequencing

PCR products were visualised on a 1 % (w/v) agarose gel using a molecular weight standard

to estimate size and concentration of product. Single band products were purified by either

ethanol precipitation (see Appendix III) or using Prep-a gene DNA purification kit (BioRad)

according to the manufacture’s instructions. When non-specific products were present the

sample was run on a preparative agarose gel and the band of interest excised. DNA was

extracted from the gel slice using the prep-a-gene DNA purification Kit (BioRad).

A standard DNA sequencing reaction consisted of 50-100 ng of double stranded template

DNA, 15-20 pmol of sequencing primer (various, see Appendix II), 4 µl of CSA buffer

(Applied Biosystems) and 4 µl of BigDye terminator cycle sequencing reaction mix

(Applied Biosystems) in a final volume of 20 µl. Amplifications were conducted using the

following thermoprofile: an initial denaturation step at 94 oC for 1 min, followed by 25 cycles

of 94 oC for 10 sec, 50 oC for 5 sec and 60 oC for 4 min. After cycling 16 µl of milli-Q water

and 64 µl of 95% (v/v) ethanol were added and vortexed briefly. The reaction mix was then

incubated for 1 h at room temperature and thereafter centrifuged at maximum speed for 20

min. The pellet was washed in 70 % ethanol and lyophilised in a vacuum centrifuge.

Separation of sequencing products was performed on a ABI 377 DNA sequencing system at

the Automated Sequencing Facility, UNSW.

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Genomic DNA

Restriction digest and ligation of adaptors

Panhandle PCR using specific primers and adaptor primers

(b) No panhandle structure formed, direct sequencing of PCR fragments

(a) Panhandle structure formed, no amplification

Construction of specific primers based on known mini-Tn10 sequence information

Tn10 D

Tn10 C

AP1 AP1

AP1

AP1

Tn10 D

Tn10 C

Figure 3.1: Diagrammatic representation of the panhandle-PCR method for sequencing from

uncloned genomic DNA. n = adaptor molecule; = mini-Tn10 transposon. PCR

fragments generated only from adaptor primers will form panhandle structures and

amplification will be suppressed (see text) (a). Fragments generated by both an adaptor

primer and a gene specific primer will not form a panhandle structure and amplification can

continue generating a PCR fragment that can be sequenced directly by primer walking (b).

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3.2.4.4. Sequence data analysis

The DNA-sequence-electropherograms provided by the UNSW sequencing facility were

analysed with ABI-PRISM software (available through the UNSW sequencing facility).

Results were processed and multiple sequence alignment was performed using the

AutoAssembler program within the INHERIT package (Applied Biosystems). The

assemblages were manually edited to erase ambiguous positions and frame-shifts. The

completed DNA-sequence was compared to known sequences in the GenBank-database

using the BLAST-search algorithm (Altschul et al., 1990) and open reading frames (ORF)

were defined using the ORF finder program both made available through the National Center

for Biotechnology Information (NCBI) web site (http://www.ncbi.nlm.nih.gov). Further

analysis (molecular weight, pI, hydrophobic index, secondary structure etc.) was performed

using the appropriate programs in the GCG-software package provided by the Australian

National Genomic Information Service (ANGIS) web site

(http://www.angis.org.au/WebANGIS/). Where appropriate the molecular biology analysis

tools available through the ExPASy site (http://expasy.proteome.org.au/index.html) were also

used.

3.2.5. Preparation of P. tunicata concentrated supernatant

Concentrated supernatant from P. tunicata was prepared as previously described (section

2.2.4). The cells from a 24 h culture of P. tunicata in VNSS were harvested by

centrifugation and resuspended into NSS and incubated for a further 24 h. The cells were

removed by centrifugation and filtration (0.22 µm). The cell-free supernatant was then tested

for anti-fungal activity by a drop plate method whereby aliquots of supernatant were dropped

onto a VNSS plate containing a lawn of the target fungus (Penicillium sp.). Plates were dried

and incubated for 48 h at 30 oC then assessed for zones of fungal growth inhibition.

3.2.6. Extracts of cells and cell free supernatant of P. tunicata

The cells from a 24 h culture of P. tunicata in VNSS were harvested by centrifugation

(13200 x g for 30 min) and were subsequently extracted three times with re-distilled

methanol (EM Sciences) at an approximate concentration of 40 ml of methanol per gram of

wet cells. Cell debris was removed by filtration through a 150 mm Whatman filter paper and

the remaining liquid methanol phase dried under reduced pressure. The dried methanol

extract was then resuspended into a 1:1 dichloromethane (DCM) (EM Sciences) and milli-Q

93

water mix. The water phase was extracted a further two times with equal volume of DCM.

This was followed by three extractions of the remaining water phase with equal volumes of

isobutanol (EM Sciences). After each extraction both the DCM phase and isobutanol phase

were collected using a separating funnel, evaporated separately under reduced pressure and

thereafter tested for anti-fungal activity as detailed above. The cell-free concentrated

supernatant of P. tunicata (section 3.2.5) was similarly extracted three times with DCM. All

fractions were evaporated under reduced pressure and tested for anti-fungal activity (see

section 3.2.1).

3.2.7. Fractionation of the anti-fungal compound from crude cell extracts

Thin Layer Chromatography (TLC) on silica gel plates (MERCK) was used to identify the

chromatographic procedures and the solvent systems that give the best separation of

compounds within the crude extract. Further purification was performed using solid phase

extraction columns (Alltech), which fractionate samples based on polarity with more polar

compounds being retained on the column. The active DCM cell extract (see section 3.3.5.1)

was loaded onto the column using diethylether (EM Sciences). Based on the results from

TLC assays, fractions were eluted from the column using the following steps hexane, 10%

(v/v) ethylacetate / hexane, chloroform, 4% (v/v) isopropanol / chloroform, 20% (v/v)

isopropanol / chloroform and methanol. After elution from the column all fractions were

dried under reduced pressure and tested for anti-fungal activity using the filter disc method as

described above. Sub-fractionation of active fractions were performed as above with the

exception that samples were loaded onto the column with DCM and eluted using 2% (v/v)

isopropanol / chloroform.

3.3. Results

3.3.1. Activity of P. tunicata against a range of yeast and fungal isolates

To investigate the effectivness of the anti-fungal activity of P. tunicata a sensitivity screen

with a range of yeast and fungal isolates was performed. Results in Table 3.2 show that P.

tunicata cells are capable of affecting the growth of a wide range of yeast and fungal species.

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Table 3.2: Activity of P. tunicata against a range of yeast and fungal species

Target organism Growth inhibition (mm)

Alternaria alternata 3.5

Aspergillus niger 3

Aureobasidium pullulans 6

Candida albicans 4

Cladosporium cladosponoides 6

Penicillium digitatum 5

Penicillium expansium 2.5

Saccharomyces cerevisiae 3.5

Rhizopus nigricans 5

Rhodotorula rubra 3.5

3.3.2. Transposon mutagenesis

A transposon mutant of P. tunicata specifically altered in its ability to inhibit fungal and yeast

growth was generated for two main reasons. Firstly, to gain information regarding genes

essential for anti-fungal activity and secondly, for comparative studies with the wild-type

strain during chemical identification and analysis of the active compound. The transposon

mutagenesis protocol was successful in generating 3 mutants (designated FM1, FM2, FM3)

of P. tunicata which are not able to inhibit fungal growth (Figure 3.2). The mutants FM1

and FM2 were generated by Sally James and mutant FM3 was generated in collaboration

with Ashley Franks. FM3 was obtained from a screen of 45000 transposon mutants and was

the only one that specifically lost the anti-fungal activity.

95

Figure 3.2: Anti-fungal activity of P. tunicata wild-type (wt) and mutants (FM1, FM2 and

FM3) were inoculated in small circles from a fresh agar plate containing the target fungus

.

96

3.3.3. Phenotypic characterisation of the non anti-fungal mutants

3.3.3.1. Growth conditions

The P. tunicata non anti-fungal transposon mutants displayed similar phenotypic

characteristics as the wild-type strain. They all remained dark green in pigmentation and were

able to grow under the same conditions as wild-type strain. Figure 3.2 is a typical growth

curve over a 24 h period of P. tunicata wild-type and one of the non anti-fungal mutants. The

results show that there is no difference in the general growth pattern or rate. This result was

also seen for the other non anti-fungal mutants.

3 02 01 000.0

0.2

0.4

0.6

0.8

1.0

Wild-type (A)Wild-type (B)non anti-fungal (A)non anti-fungal (B)

Time

OD

610

nm

Figure 3.3: Growth of wild-type P. tunicata and the FM1-mutant. Duplicate cultures (A

and B) were inoculated as indicated in the text (3.2.3.2) and growth was monitored by

absorbence readings at 610 nm over a 24 h period.

97

3.3.3.2. Other antifouling properties

The ability of the P. tunicata non anti-fungal mutants to inhibit other fouling organisms was

assessed. The non anti-fungal mutants (FM1-FM3) were found to display the same pattern

of antifouling properties as the wild-type P. tunicata strain. Table 3.3, Table 3.4 and Table

3.5 summarise representative results of these assays for the FM1-mutant.

Table 3.3: Growth inhibition of bacteria in the presence of P. tunicata wild-type and non

anti-fungal mutant strain FM1.

Growth inhibition (mm)

Target bacterium Wild-type Non anti-fungal mutant

P. tunicata 3.5 3

Bacillus subtilis 3 3

Table 3.4: Settlement of marine invertebrate larvae in the presence of biofilms of P. tunicata

wild-type, non anti-fungal mutant strain FM1 and a non-inhibitory marine isolate.

Percentage settlementa

Target organism Wild-type Non anti-

fungal mutant

Non-inhibitory

isolate bNo biofilm

control

Balanus amphitrite

larvae1 ± 0.6 6 ± 3.3 80 ± 6.5 95 ± 1.6

Hydroides elegans

larvae

0 0 49 ± 6.8 45 ± 4.6

a All values are means ± standard deviations (n=3)

b Previously determined non-inhibitory marine isolate (Holmström et al., unpubl.)

98

Table 3.5: Germination of marine algal spores in the presence of biofilms of P. tunicata

wild-type, non anti-fungal transposon mutant strain FM1 and a non-inhibitory marine isolate

Percentage germinationa

Target organism Wild-type Non anti-

fungal mutant

Non-inhibitory

isolate bNo biofilm

control

Ulva lactuca

spores

0 0 91± 3 100

Polysiphonia sp.

spores

0 4 ± 1.4 87 ± 5.6 82 ± 2.1

a All values are means ± standard deviations (n=3)

b Previously determined non-inhibitory marine isolate, see section 2.3.1.

3.3.4. Genotypic characterisation of the non anti-fungal mutants

3.3.4.1. Panhandle-PCR and DNA-sequencing

To identify the genes, which have been disrupted by the transposon insertion in the P.

tunicata genome, panhandle-PCR was employed. Since the DNA sequence of the

transposon was known specific primers could be designed to amplify the genomic DNA of

regions flanking the transposon. The primers were designated Tn10C and Tn10D (sequence

is given in Appendix II). Panhandle-PCR products were generated from the non anti-fungal

mutants. Figure 3.4 shows the results of a typical panhandle-PCR. Single bands were

amplified from different genomic digests, which would indicate that only one copy of the

transposon was present. This supports previous Southern-blot experiments (James, 1998)

showing that the mini-Tn10 transposon consistently inserts only once within the P. tunicata

genome. The PCR products were purified and then sequenced by primer walking.

99

Figure 3.4: Agarose gel showing the results from a typical panhandle-PCR from FM1

genomic DNA templates. Lane 1 and 8: 250 ng and 125 ng of molecular weight marker (λ

EcoRI/HindIII digest) respectively; Lane 2: Xmn1 digest and Tn10C primer; Lane 3: Dra1

digest and Tn10C primer; Lane 4: HincII digest and Tn10D primer; Lane 5: Xmn1 digest and

Tn10D primer; Lane 6: PvuII digest and Tn10D primer; Lane 7: Dra1 digest and Tn10D

primer.

100

3.3.4.2. DNA sequence analysis

Initial sequence analysis of the regions directly flanking the transposon in each of the three

non anti-fungal mutants indicated that they were all disrupted in the same DNA region. The

region flanking the Tn10C side of the transposon in FM1 is identical to that of the Tn10D

side of FM2 and the Tn10C side of FM3. Based on this fact sequencing was only continued

with the FM1 strain to obtain a total of 3736 bp of sequence which flanked both sides of the

transposon. The sequencing strategy employed using specific primers and primer walking is

shown schematically in Figure 3.5. Additional panhandle-PCR primers (FMpan1 and

Fmpan2) were designed and PCR products obtained to continue sequencing further along the

genomic DNA. Sequences for each of the specific primers used are given in Appendix II.

After sequence assembly, a consensus sequence was obtained and the nucleotide sequence

was submitted to the programs ORF-finder and BLAST from NCBI. Figure 3.6 shows the

primary sequence data highlighting ORFs and their predicted AA sequence. The results of

the analysis indicate that the transposon had disrupted a 1662 bp ORF, designated afaA (anti-

fungal activity A), with 63 % identity and 78 % similarity (over 539 amino acid residues) to

the E. coli long-chain-fatty-acid CoA ligase gene (fadD).

Further sequence analysis of AfaA revealed a putative ribosome binding site (RBS)

5'AGGAGCT 3' located 4 bp upstream of the alternative GTG start codon. Following the

translational stop of afaA is a GC-rich inverted repeat followed by a series of six thymidine

residues (nucleotide 3336 to 3360) which may act as a ρ-independent terminator of

transcription (Mathews and van Holde, 1990). A potential transcription start point was

identified at base-pair position 1651. The region upstream contains sequences, as highlighted

in Figure 3.6, that are within reasonable agreement with potential -10 and -35 sequences for

E. coli σ70-responsive promoters.

Analysis of the deduced amino acid sequence of the AfaA gene indicates that this protein has

a molecular weight (MW) of 61081.6 Da and a predicted isoelectric point (pI) of 5.84. The

hydropathy profile shows that the protein is primarily hydrophilic with an average

hydrophobicity of -0.028. No predominate hydrophobic regions were predicted thus

implying that the protein is not membrane integrated. In addition, no secretion signal

sequences were identified indicating that the protein is likely to be located in the cytoplasm.

101

Figure 3.7 shows the multiple sequence alignment of the deduced amino acid sequence AfaA

with the sequences of the genes with high sequence similarity. A putative AMP-binding

domain signature motif (LQYTGGTTGVAK) was detected between amino acids 214 and

226. This domain is shared between a number of prokaryotic and eukaryotic enzymes which

act via an ATP-dependent covalent binding of AMP to their substrate, including insect

luciferase (Ye et al., 1997), Gramicidin S synthetase I and II (Turgay et al., 1992) and long-

chain-fatty-acid Co-A ligase (Black et al., 1992). A second motif was located between

positions 437 and 461. This region known as the fatty acyl-CoA synthetase signature motif

(FACS signature motif) (DGWLHTGDIGXWXPXGXLKIIDRKK) has been identified to

be highly conserved for fatty acyl-CoA synthetases. The FACS motif is believed to function

in part to promote fatty acid chain length specificity and may compose part of the fatty-acid

binding site (Black et al., 1997).

3.3.4.2.1. Analysis of flanking ORFs

Directly upstream of afaA is a 852 bp ORF (ORF 2) designated here as afaB. A potential

RBS was located 5 bp from the alternative start codon GTG at positions 680 - 686. The

predicted transcriptional start point is at nucleotide 639 and potential -10 and -35 sequences

of prokaryotic promoters, are indicated in Figure 3.6. Although no direct experimental

evidence was obtained regarding operon structure (i.e. primer extension or Northern-blot

analysis), the close proximity of ORF 2 with afaA and the lack of an obvious terminator of

transcription downstream of the termination codon, suggests that afaA and afaB are

potentially co-transcribed.

Analysis of the deduced amino acid sequence of the afaB gene indicates that this protein has

a MW of 32047.7 and a predicted pI of 6.76. The hydropathy profile shows that the protein

is primarily hydrophilic with an average hydrophobicity of -0.188. No predominate

hydrophobic regions were predicted thus implying that the protein is not integrated into the

membrane.

AfaB has sequence similarity (46 % over 252 amino acid residues) to a group of enzymes

known as serine hydrolases whose common feature is the hydrolysis of substrates with a

carbonyl-containing group (Chan et al., 1998). Figure 3.8 shows the multiple sequence

alignment of the deduced amino acid sequence of AfaB with the sequences of proteins with

high sequence similarity. A conserved motif corresponding to the serine active site of

triglyceride lipases was detected between amino acid positions 131 and 141. Triglyceride

102

lipases are lipolytic enzymes that hydrolyse the ester bonds of triglycerides and can be of

animal, plant or prokaryote origin. Within the prokaryotes many of the lipases are

extracellular and encode a N-terminal signal sequence that is cleaved from the mature

polypeptide (Upton and Buckley, 1995). However, analysis of this region within the

predicted ORF of afaB did not indicate the presence of a conserved N-terminal signal

sequence. This suggests that P. tunicata AfaB is potentially intracellular or that it is released

into the extracellular environment by another mechanism.

Downstream of the afaA gene at position 3465 bp is the beginning of another ORF (ORF3)

with similarity (69 % over 59 amino acid residues) to E. coli Ribonuclease D (D90825). A

predicted RBS is located 12 bp upstream of the ATG start codon. Ribonuclease D functions

as a putative tRNA processing enzyme and is also located directly down stream of the fadD

gene within the E. coli genome (Zhang and Deutscher, 1988).

Located upstream of the afaB gene and encoded in the opposite direction is ORF 4. The

initial sequence of this gene indicates that it is similar (65 % over 135 amino acid residues) to

an alkaline phosphatase from Enterococcus faecalis (AF154110).

103

0 1000 2000 3000

FMpan2-S3

FMpan2-S2

FMpan2-S5

FMpan2

FMpan2-S4

FMTnD-S6

FMTnD-S5

FMTnD-S3

Tn10D-S1

FMTnD-S7

FMTnC-S8

Ap2

Tn10C-S1

FMTnC-S2

FMTnC-S4

FMpan1-S3

FMpan1-S4

FMpan1

FMpan1-S2

Figure 3.5: Summary of the sequence strategy for determining the nucleotide sequence of

the region of DNA flanking the transposon insert within the P. tunicata non anti-fungal

mutant. Arrows indicate length and direction of sequence products. Blue arrows represent

transposon or adaptor specific primers and black arrows represent sequence specific primers.

All primers used are listed in Appendix II. The nucleotide sequence is shown in base pairs

along the top of the diagram. Open reading frames are indicated by the bold coloured lines;

red = ORF 1 (afa A); blue = ORF 2 (afa B); purple = ORF 3; green = ORF 4.

104

1 CTGTACTGATAACCTGACTGGATGAATTCAGCCACTAAATCACGATCGTTACGTTTGAAA Y Y Q Y G S Q I F E A V L D R D N R K

61 TAGTCAGTACCACCGCCTAACAGTAAATCAACAGGTAGGCGACCATTAATTTTGTTATCT F Y D T G G G L L L D V P L R G N I K N

121 ATGTAGTCATTTGCGATTTCGTTGTAGTTTTTACGATGAACATTGTGTGCAGCAAAGCTT D I Y D N A I E N Y N K R H V N H A A F

181 GCAGGTGTGGCATGGTTAATTTGAGATGTTGCAACAAGAGCTGTCAACATGCCGCGTTTT S A P T A N I Q S T A V L A T L M G R K

241 TTTGCGATTTCTAGCATGGTTTCA AGCGGT TTTTTTGCCGTATCGACAGCAATAGCACCA K A I E L M (RBS)

301 TTGTAACTTTTATGGCCTGTACTTAGGGCTGTCGCACCTGCAGCGCTATCGGT AACATA A (-10) 361 GTATGATCATCAGGAAAGTACGTGCCATACC GGTCAA AATAGAATCGAATACGGTTGTCT (-35) 421 CGACGTCTTTTGTTGTTAGGTCATCGGAGTAATAGCGATAAGCAGTGGTATAGGCTGGGC

481 CCATACCGTCACCAATCATGTAAATAATATTTTTTGGCGCACTGGCATAAACGGATGATG

541 AAAGTAAACCTAATGCGGTTAGGGTAAATATTTTTTTCATAGTATGTTCTTA TTGTCT GG (-35) 601 GAAGCTTTATTTTACGTGCATCATTG TATAAT AAAAACAAATTTATGTGCGTAACCGTTA (-10) 661 GATAACAGTAACCAATTGT AAGTGAG AAAACTGTGACGTTCGATTCAATAATTGTAAAAA (RBS) V T F D S I I V K S

721 GCAATGATTTAACTTTACGCGGTATTAAGCATGGAGACAAAACTCAGCAAACAATTTTAG N D L T L R G I K H G D K T Q Q T I L A

781 CACTGCATGGCTGGCAAGATAATTGCCATAGTTTTATTCCTTTATTTAATTTTTTAACTG L H G W Q D N C H S F I P L F N F L T E

841 AATATCAATGTTATGCTTTTGATTTTCCTGGGCATGGACTCTCTGACTGGCGCCATTCGT Y Q C Y A F D F P G H G L S D W R H S S

901 CAGCACATTATTATTTAACTGAATATGTTGATGATGTGCTAAACATGATCAAAAACGAAA A H Y Y L T E Y V D D V L N M I K N E I

961 TTAAAGAACCTATTCATTTGGTCGGGCATTCAATGGGGGCAATGGTCGCGACATTATTTA K E P I H L V G H S M G A M V A T L F T

1021 CGGCATGCTTTCCTGAAAAAGTGAAAAGCCTAACGTTGATCGACGGTATTGGATTTGTGA A C F P E K V K S L T L I D G I G F V T

1081 CCACCGCGGCAAATAAATCATCACAACAGCTACGCCAAGCGCTCGAAAATCGTTCTCGAC T A A N K S S Q Q L R Q A L E N R S R L

105

1141 TTCATAATAAACCAGCTAAAATTTTTCAAGATTTAGAATCATTAATTCTGATGAGAATGC H N K P A K I F Q D L E S L I L M R M Q

1201 AGGTTTCTGATTTAAATAAAGAAAACAGTGAATTAATTATGCAGAGAAATTGTATTCCTA V S D L N K E N S E L I M Q R N C I P I

1261 TTAATAATGGCGTAAAACTATCCATTGACCCAAAACTGAAACTTGCATCTGCATTTCGTT N N G V K L S I D P K L K L A S A F R F

1321 TTTGTGATGAGCAAGCCCATGAAATTTGTAAGAGTATTCCACATAATGTGCATGTAGTAC C D E Q A H E I C K S I P H N V H V V L

1381 TTGCCAGCTCCAATAGTGCTGGTTTTAGTGAAAAATATGCAGAGTATGTGAAGGATTTTA A S S N S A G F S E K Y A E Y V K D F N

1441 ACGCAATTACCCGCTATGATTTAGACGGTTGCCATCATTGTCATATGGAGCAACCACAAC A I T R Y D L D G C H H C H M E Q P Q R

1501 GGCTTGCCGCCATTTTGCGTCAGATTGTCGCTTTGGCAAGTTGAGGGAATTGTATGTTAA L A A I L R Q I V A L A S *

1561 GAATAAATGTGTGATTATATGCGGCCATTTAGAGTAATTACTCAA TTGTTA AATCGCCTT (-35) 1621 AGTTAAATTAAGGTAAC TATAAT GAATAAAAATAAG AGGAGCT AACGGTGGATAAAATCT (-10) (RBS) V D K I W

1681 GGCTAAACAGGTTTCCAGAAGGCATGCCTGAAGAAATAGATCCAAGACACTACAACTCCC L N R F P E G M P E E I D P R H Y N S L

1741 TGCTAGACCTATTTGAAATAAGTTTTGCGGAGTATGCCCAATTACCAGCATTTTCTAACA L D L F E I S F A E Y A Q L P A F S N M

1801 TGGGTAGGGCGTTGAGCTATCAAGAGCTAGACGTTGCAACTAAAAAGTTTGCTGCGTATT G R A L S Y Q E L D V A T K K F A A Y L

1861 TACAACATGATTTAGGTCTTAAAAAAGGTGATAAAGTGGCTGTTATGATGCCTAATCTAT Q H D L G L K K G D K V A V M M P N L L

1921 TGCAAACTCCAATTGCAATTTTAGGTATTTTACGAGCCGGCTGTACGGTTGTGAACGTCA Q T P I A I L G I L R A G C T V V N V N

1981 ATCCTCTGTACACAGCGCGTGAGCTTGAACATCAGCTTAATGACTCAGAAACTACCGCTA P L Y T A R E L E H Q L N D S E T T A I

2041 TTGTTATTTTAGCCAATTTTGCCCATACACTTGAAGAAGTGCTTGGCAAAACGGGTGTGA V I L A N F A H T L E E V L G K T G V K

2101 AACACATAATCCTAAGTGAAATTGGTGATATGTGTGGCGGGCTGAAAAAACACCTAGTGA H I I L S E I G D M C G G L K K H L V N

2161 ATTTTGTTGTTAAACATATTAAAAAAATGGTACCTGCTTTTTCATTACCAAATGTAATTC F V V K H I K K M V P A F S L P N V I P

106

t 2221 CATATGCACGTTTGATGGCTGATGCTGATGCGAAGCATTATTCACGACCAGAGCTTACTC Y A R L M A D A D A K H Y S R P E L T H

2281 ATTTAGATTTAGCCTTTTTACAATATACCGGTGGTACAACGGGTGTTTCTAAAGGCGCAA L D L A F L Q Y T G G T T G V S K G A M

2341 TGTTAAGTCATGGCAATATGGTTGCTAATCTTGAACAAGTATCAGGCTGTTTAGATACTG L S H G N M V A N L E Q V S G C L D T V

2401 TACTTGATCGCGGTAAAGAAATTGTAGTCACTGCGTTACCGCTTTATCATATTTTTGCGT L D R G K E I V V T A L P L Y H I F A L

2461 TAACGGCTAACTGCTTAACCTTCATGAAATATGGTGGTTTAAATTTATTAATCACAAATC T A N C L T F M K Y G G L N L L I T N P

2521 CTCGTGATATGAAAGGCTTTGTAAAAGAGCTAAGTAATAACCGTTTTACTGCAATCACAG R D M K G F V K E L S N N R F T A I T G

2581 GTGTGAATACGCTTTTTAACGGATTGCTTAATACTCCAGGTTTTGATGAACTTGATTTTT V N T L F N G L L N T P G F D E L D F S

2641 CGAACCTAAAGTTATCTTTAGGTGGTGGTATGGCTGTGCAGCGTCCTGTTGCTGAGCGCT N L K L S L G G G M A V Q R P V A E R W

2701 GGCAAGAAGTAACTAAAACACGTTTAGTTGAAGGCTACGGTTTAACCGAATGTGCGCCAC Q E V T K T R L V E G Y G L T E C A P L

2761 TTGTTACTATCAGCCCGTACGATTTAGCTGGTTATAATGGCTCAATTGGTTTACCAGCAC V T I S P Y D L A G Y N G S I G L P A P

2821 CTAGCACTGATATTAAAATTATGGGTGAAGATGGCCAAGAAGTTGCAAAAGGTGAAGCGG S T D I K I M G E D G Q E V A K G E A G

2881 GTGAGCTTTGGGTTAAAGGCCCACAAGTAATGCTTGGTTATTACAAACGACCAGAAGCGA E L W V K G P Q V M L G Y Y K R P E A T

2941 CAGCTGAATGTATGCATGATGGTTGGTTTGCAACTGGCGACATTGCGACCTACGATGATG A E C M H D G W F A T G D I A T Y D D E

3001 AAGGGTTCTTTTATATCGTCGATCGTAAAAAAGATATGATCATTGTCTCTGGTTTTAATG G F F Y I V D R K K D M I I V S G F N V

3061 TATTTCCAAATGAAATCGAAGAAGTATGCATGATGAATTCAGGCGTGCTTGAAGTGGCAG F P N E I E E V C M M N S G V L E V A A

3121 CAATTGGTGTACCTCATGAGGTCAGTGGTGAACAAGTTAAGATTTTTGTTGTTAAAAAAG I G V P H E V S G E Q V K I F V V K K D

3181 ACCCTTCTTTAACAGAAAAAGATATAATAGCTCATTGTCGTAAGAATCTAACTAACTACA P S L T E K D I I A H C R K N L T N Y K

3241 AGGTCCCTAAATTCGTTGAATTTAGAGAGGAACTTCCTAAGACTAATGTAGGTAAAATTT V P K F V E F R E E L P K T N V G K I L

107

3301 TAAGGCGCGCCTTAAAAGAAGCAAGTTAAGAAAA GCCGGCTCAGCCGGCTTTTTTAT TAA R R A L K E A S

3361 AGGGGCTCACACTTGCACTATCAATATATCCAAGAACAATCACAACTTGATGAATTTTTA

3421 CAAGCCATTTCGACTCAATCAGTATT AGCAATT GATACTGAGTTTATGCGCAGACGTACA (RBS) M R R R T

3481 CTTTATCCTGAAATAGCCCTTATCCAAGTGTTTGATGGTCAACACTTAGGTTTAATCGAT L Y P E I A L I Q V F D G Q H L G L I D

3541 CCATGTTGCGATCTCGATTTAAGCCGTTTTTGGCAGATTATGTCTGATGCTGCAATTCTG P C C D L D L S R F W Q I M S D A A I L

3601 AAAGTACTACATTCTCCCTCTGAAGATGTTGAAGTATTTTT 3641 K V L H S P S E D V E V F

Figure 3.6: Nucleotide sequence of the region of genomic DNA surrounding the

transposon within the non anti-fungal mutant genome. The nucleotide sequence is shown

along with the translated amino acid sequence in one-letter code. The inverted solid triangle

(t) indicates where the mini-Tn10 transposon insertion occurred. Specific open reading

frames (ORF) as indicated in the text are highlighted as follows: afaA (ORF 1) is shown in

red, afaB (ORF 2) is shown in blue, ORF 3 is shown in purple and ORF 4 is shown in green.

Potential promoter regions are underlined as are predicted ribosome binding sites (RBS).

For ORF 3 the reverse complement of the actual sequence is given. A probable

transcriptional terminator following afaA is indicated in italics and underlined.

108

1 50P. tunicata ..vdkiwlnr fpegmpeeid prhynslldl feisfaeyaq lpafsnmgraE. coli ..mkkvwlnr ypadvptein pdryqslvdm feqsvaryad qpafvnmgevB. subtilis mqsqkpwlae ypndiphel. plpnktlqsi ltdsaarfpd ktaisfygkkH. influenzae ..mekiwfqn ypkgsekfld tskyesildm fdkavrehpd rpayinmgqv •••••• • • • ••• • • •••• •• • •••• •••••••

51 100P. tunicata lsyqeldvat kkfaaylqhd lglkkgdkva vmmpnllqtp iailgilragE. coli mtfrkleers rafaaylqqg lglkkgdrva lmmpnllqyp valfgilragB. subtilis ltfhdiltda lklaaflqcn .glqkgdrva vmlpncpqtv isyygvlfagH. influenzae ltfrkleers rafaaylqne fklqrgdrva lmmpnllqyp ialfgilrag • • ••••••• ••••••• •• •••••••••• •• ••••••

101 150P. tunicata ctvvnvnply tarelehqln dsettaivil anfahtleev lgktgvkhiiE. coli mivvnvnply tprelehqln dsgasaiviv snfahtlekv vdktavqhviB. subtilis givvqtnply teheleyqlr daqvsviitl dllfpkaikm ktlsivdqilH. influenzae liavnvnply tprelelqlq dsgavaivvv snfastlekv vfntnvkhvi •••••••• • •••••••• •• ••••• •••••••• • •• •••••

151 200P. tunicata lseigdmcgg lkkhlvnfvv khikkmvpaf slpnvipyar lmadadakhyE. coli ltrmgdqlst akgtvvnfvv kyikrlvpky hlpdaisfrs alhngyrmqyB. subtilis itsvkdylpf pknilypltq kq.kvhidfd ktanihtfas cmkqektellH. influenzae ltrmgdqlsf gkrtlvnfvv kyvkklvpky klphavtfre vlsigkyrqy • •• • •••••• • ••• •• ••• • • • •

201 250P. tunicata srpelthl.d laflqytggt tgvskgamls hgnmvanleq vsgcldtvl.E. coli vkpelvpe.d laflqytggt tgvakgamlt hrnmlanleq vnatygpll.B. subtilis tipkidpehd iavlqytggt tgapkgvmlt hqnilantem ...caawmydH. influenzae vrpeisre.d laflqytggt tgvakgamlt hgniitnvfq akwiaepfig •••• • •••••••••• ••• ••••• •••• ••••• • • •

251 300P. tunicata .drgkeivvt alplyhifal tancltfmky gglnllitnp rdmkgfvkelE. coli .hpgkelvvt alplyhifal tincllfiel ggqnllitnp rdipglvkelB. subtilis vkegaekvlg ivpffhvygl tavmnysikl gfemillpkf dpletl.kiiH. influenzae dhsrtrsail alplyhvfal tvncllflel gvtailitnp rdiegfvkel ••• ••• •••••••••• ••••• • • •• ••••••• •• ••••••

301 350P. tunicata snnrftaitg vntlfnglln tpgfdeldfs nlklslgggm avqrpvaerwE. coli akypftaitg vntlfnalln nkefqqldfs slhlsagggm pvqqvvaerwB. subtilis dkhkptlfpg aptiyigllh hpelqhydls siksclsgsa alpvevkqkfH. influenzae kkyrfeaitg vntlfnalln nenfkevdfs alklsvgggm aiqqsvatrw ••••••• •••••••••• • • ••••• •••••••••• ••• •••••

109

351 400P. tunicata qevtktrlve gygltecapl vtispydlag yngsiglpap stdikimgedE. coli vkltgqylle gygltecapl vsvnpydidy hsgsiglpvp steaklvdddB. subtilis ekvtggklve gyglseaspv thanfiwgkn kpgsigcpwp stdaaiyseeH. influenzae heltgcniie gygmtecspl iaacpinvvk hngtigvpvp ntdikiikdd ••• ••• •••••••••• • ••• ••••••• • •••••• ••

401 450P. tunicata gqevakg.ea gelwvkgpqv mlgyykrpea taecmhdgwf atgdiatyddE. coli dnevppg.qp gelcvkgpqv mlgywqrpda tdeiikngwl htgdiavmdeB. subtilis tgelaapyeh geiivkgpqv mkgywnkpee taavlrdgwl ftgdmgymdeH. influenzae gsdakig.ea gelwvkgdqv mrgywqrpea tsevlkdgwm atgdivimde • ••• • •• •••••••••• •••• •••• ••• ••• •••••• •

451 500P. tunicata egffyivdrk kdmiivsgfn vfpneieevc mmnsgvleva aigvphevsgE. coli egflrivdrk kdmilvsgfn vypneiedvv mqhpgvqeva avgvpsgssgB. subtilis egffyiadrk kdiiiaggyn iypreveeal yeheaiqeiv vagvpdsyrgH. influenzae syslrivdrk kdiilvsgfn vypneiedvv mlnykvseav aigvphavsg •••••••••• •••••••••• • ••••••• • • •• ••• •••••• •••

501 550P. tunicata eqvkifvv.k kdpsltekdi iahcrknltn ykvpkfvefr eelpktnvgkE. coli eavkifvv.k kdpslteesl vtfcrrqltg ykvpklvefr delpksnvgkB. subtilis etvkafvvlk kgakadteel dafarsrlap ykvpkayefr kelpktavgkH. influenzae etikifvv.k kddsltrdel rnhcrqyltg ykvpkeiefr delpktnvgk • •••••• • ••••••• •••• •• ••••• •••• •••••••••

551 568P. tunicata ilrralkeas ........E. coli ilrrelrdea rgkvdnkaB. subtilis ilrrrlleee tenhhik.H. influenzae ilrrvlrdee iakrpkh. •••• • •

Figure 3.7: Multiple sequence alignment of the deduced amino acid sequence of P. tunicata

AfaA with sequences of known long-chain fatty-acid CoA ligases from three different

bacterial species. Sequences were from the following accession numbers P29212 (E. coli);

P94547 (B. subtilis) and P46450 (H. influenzae). The sequence corresponding to the AMP-

binding domain motif is indicated (red) as is the FACS signature motif (blue). Residues

identical between P. tunicata and one other protein are indicated by black dots (•); residues

identical in two other proteins are indicated with blue dots (•) and residues which are identical

between all proteins are indicated by green dots (•). Small dots (.) denote gaps.

110

1 50 AfaB .......... .......... .......... ......vtfd siivksndlt AJ000516 mgqtrvaatt aaqspaaels petngqteep lqllgedswe efsiavpwgt P07383 .......... .......... .....eiipv pdqaawnask ksiqindaik S75226 .......... .......... .......... .......mpt ldllgfphhy • •

51 100 AfaB lrgikhgdkt qqtilalhgw qdnchsfipl fnfl.teyqc yafdfpghgl AJ000516 veakwwgske rqpiialhgw qdncgsfdrl cpllpadtsi laidlpghgk P07383 mryvewgnps gdpvlllhgy tdtsrafssl apflskdkry laldlrghgg S75226 qqsgsdrqga apslifvhgw llshhywlpl mellsgqysc vsydlrgfga • • • • ••••••• ••••••• •• •• • • • • ••••

101 150 AfaB .sdwrhssah yylteyvddv lnmikneike pihlvghsmg amvatlftac AJ000516 sshypmgmqy fifwdgicli rrivrkynwk nvtllghslg galtfmyaas P07383 ts...ipkcc yyvsdfaedv sdfidkmglh nttvighsmg smtagvlasi S75226 sqslghprse ydleaygqdl idlleklnie qawlvghslg gsvaiwaahl • • ••• • •• • • ••••••• ••• •

151 200 AfaB fpekvksltl idgigfvtta ankssqqlrq alenrsrlhn kpakifqdle AJ000516 fpteveklin idiagptvrg tqrmaegtgr aldkfldyet lpeskqpcys P07383 hpdkvsrlvl istalktgpv lewvydtvlq kdfplddpse fakewvaapg S75226 cpervkgvvc vnagggi... ..ylkeefek frsageklld frppwlgrlp •••••• • • •• • • •• • •

201 250 AfaB slilmrmqvs dlnkense.. ....limqrn cipinngvkl sidpklklas AJ000516 ydemiklvld aydgsvdeps vrvlmnrgmr hnpskngylf ardlrlkvsl P07383 khdngmaknl kteelavpkh vwlsaargfs iinwtaasky ltaktlilwg S75226 lldlafsrmm vekplarkwg rqrlldflra dqqaargsll estteaavhl • • • • •• •• •• • ••• •

251 300 AfaB afrfcdeqah eicksiphnv hvv.....la ssnsagfsek yaeyvkdfna AJ000516 lgmftaeqtl ayarqircrv lnirgipgmk fetpqvyadv iatlrenaak P07383 nqnqpmtesm qndiraalpk akfiqyngfg hsmfwedpem vakdlneflk S75226 lpklvaelpq pmyflagqnd rvmelqyvky lasfhglfaq lgtnvveien • •• • •• • • • • • • •

111

301 332 AfaB itrydldgch hchmeqpqrl aailrqival as AJ000516 vvyvevpgth hlhlvtpdrv aphiirflke a. P07383 .......... .......... .......... .. S75226 cghfamleql pvvanklqqi latdh..... .. • • • • ••• •• •

Figure 3.8: Multiple sequence alignment of the deduced amino acid sequence of P. tunicata

AfaB with Drosophilia Kraken protein (AJ000516); Pseudomonas putida atropinesterase

(P07383) and Synechocystis sp. esterase (S75226). The putative lipase serine active site is

indicated in red. Residues identical between P. tunicata and one other protein are indicated

by black dots (•); residues identical in two other proteins are indicated with blue dots (•) and

residues which are identical between all proteins are indicated by green dots (•). Small dots

(.) denote gaps. Numbers shown in parentheses are the GenBank accession numbers.

112

3.3.5. Identification of the anti-fungal compound produced by P. tunicata

3.3.5.1. Anti-fungal activity of P. tunicata supernatant and crude cell extracts

To determine if the anti-fungal activity is due to cell bound compounds or to compounds that

are released into the cell free supernatant, the effect of bacterial stationary-phase supernatants

on fungal growth was assayed. Anti-fungal activity was not found to be present in cell-free

supernatant preparations or in concentrated crude extracts of the supernatant. However,

whole cell preparations extracted with methanol were shown to have anti-fungal activity

indicating that the anti-fungal activity is cell associated. DCM was used to separate the more

polar compounds from the initial methanol extract and was followed by an extraction with

isobutanol to separate the more non-polar compounds. The resulting DCM fraction was

yellow and had anti-fungal activity. In contrast the isobutanol fraction was dark purple in

colour and did not inhibit the growth of the target fungal isolate. The minimal inhibitory

concentration (MIC) of the methanol crude cell extract indicate that the compound may be

active at low concentrations, as less than 1 µg/ml crude cell extract inhibited fungal growth.

3.3.5.2. Fractionation of P. tunicata cell extract

As the initial step for the purification of the anti-fungal compound, fractionation of the crude

cell extract was performed using solid-phase chromatography columns, which separate

compounds based on their polarity. Fractionation using these columns and the solvent

system outlined in section 3.2.7, resulted in the separation of the extract into 13 fractions

(Table 3.6, Figure 3.9). These fractions were then tested in the anti-fungal bioassay for

activity. Results of the test showed that the initial fractionation was successful in separating

the active compound into a single fraction. The active fraction, fraction 6, also contained the

majority of the yellow pigment (Figure 3.10) and was further fractionated using the same

solid phase chromatography columns into 4 sub-fractions of which only one, sub-fraction 2,

remained both yellow in colour and active.

113

Table 3.6: Elution steps and characteristics of the initial 13 fractions from solid phase

chromatography columns.

Fraction Elution step Colour Anti-fungal activity

1 100% Hexane clear negative

2 10% ethylacetate/ hexane very light yellow negative

3 100% chloroform clear negative

4 100% chloroform clear negative

5 100% chloroform light yellow negative

6 100% chloroform yellow positive

7 100% chloroform light yellow negative

8 4% isopropanol/ chloroform clear negative

9 4% isopropanol/ chloroform purple negative

10 4% isopropanol/ chloroform purple negative

11 20% isopropanol/ chloroform very dark purple negative

12 20% isopropanol/ chloroform dark purple negative

13 100% methanol purple negative

Figure 3.9: The initial 13 chromatography fractions of the crude cell extract of P. tunicata.

The vials are numbered 1 to 13 and correspond to the fractions listed in Table 3.6 (see text

for details).

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Figure 3.10: Anti-fungal activity of P. tunicata cell extract fractions resulting from

separation using solid phase extraction columns (see text). Filter discs were soaked in equal

amounts of each fraction and placed onto agar plates containing a suspension of target fungal

strain (Penicillium sp.). Zone of fungal growth inhibition is visible surrounding the filter disc

with fraction number 6.

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3.3.5.3. Characterisation of the anti-fungal compound

Final purification and identification of the anti-fungal compound was performed in

collaboration with Dr Naresh Kumar from the School of Chemistry, UNSW and Ashley

Franks from the School of Microbiology and Immunology, UNSW. A scale up of the initial

purification to approximately 200 g (40 L culture) of wet cell weight was required to obtain

sufficient material for characterisation of the active compound. The basic characteristics of

the compound as determined by nuclear magnetic resonance (NMR), gas chromatography -

mass spectrometery (GCMS) and UV/light spectrometery are given in Table 3.6. The

purified compound was show to have retained anti-fungal activity.

Table 3.7: Characteristic of the anti-fungal compound produced by P. tunicata cells

Characteristic Result

Molecular weight (MW) 354

Absorbance (nm) 241 and 421

Predicted molecular formula C23H20O3

Predicted molecular structure Ring and aliphatic chain

3.3.5.4. Comparison of the active anti-fungal compound with the

corresponding non anti-fungal mutant compound

Cells of the transposon generated non anti-fungal mutant were extracted and the extract

fractionated in parallel with the wild-type cells. The fractions of the non anti-fungal mutant

extract which corresponded to the active fractions in the wild-type extract whilst remaining

yellow in colour did not affect the growth of the target fungus. The mutant fraction

corresponding to the final wild-type active fraction was analysed by NMR and GC-MS to

assess the differences with respect to structure that may result in loss of activity. Data

obtained thus far indicate that the two compounds have similar characteristic and only differ

slightly in molecular weight. The purified wild-type compound has a molecular weight of

354 compared to 356 for the corresponding mutant compound.

116

3.4. Discussion

The marine antifouling bacterium P. tunicata has been previously shown to inhibit the growth

of Helminthosporium sp. (James, 1998). The effectiveness of the anti-fungal activity is

highlighted in the current study by its ability to inhibit the growth of a variety of fungi and

yeasts (Table 3.2). Among the fungal and yeast strains tested were several of ecological and

medical importance. For example, strains common to food spoilage such as Aspergillius

niger (Doster and Michailides, 1995), Penicillium expansium (Janisiewicz, 1988) and

Rhizopus nigricans (Battilani et al., 1996) were found to be sensitive to the anti-fungal

activity of P. tunicata cells. Candida albicans commonly causes opportunistic infections in

both humans and animals (Prescott et al., 1990) and represents one of the medically

important yeast strains shown to be growth inhibited by P. tunicata cells. The feature of

broad range inhibition of yeast and fungal growth by P. tunicata not only opens the way for

many applications but is also of great ecological interest. Fungi are present in many marine

ecosystems as colonisers of living and inanimate surfaces (Hyde et al., 1998) and may even

be in direct competition with bacteria for nutrients and living space. Therefore it is not

surprising that surface colonising bacteria such as P. tunicata have developed methods to

control fungal growth.

To further investigate the anti-fungal activity of P. tunicata genetic analysis was performed

using transposon mutagenesis. Due to their ability to produce single, stable and random

insertions into the target genome, transposons are useful tools for genetic manipulation.

High efficiency of mutagenesis allows a large number of colonies to be produced that contain

random insertions within a specific gene, thus resulting in the loss of function of that gene.

The use of transposon technology has previously been restricted to E. coli, however methods

have been successfully developed for manipulations of other Gram-negative bacteria (de

Lorenzo et al., 1990). One such system is the modified version of the transposon Tn10

known as mini-Tn10 (Herrero et al., 1990). This transposon carries a kanamycin-resistance

marker that allows for easy selection of mutants. In addition, the transposase gene is outside

of the mobile element, which allows for a stable insertion because of the loss of the

transposase gene during the transfer. Using mini-Tn10 and the suicide vector pLOF for

delivery (Way et al., 1984), transposon mutants defective in the ability to inhibit fungal

growth were successfully generated in P. tunicata. The non anti-fungal mutants remained

both dark green in pigmentation and inhibitory to other target organisms including,

invertebrate larvae, algal spores and bacteria, showing that the compound responsible for

inhibiting fungal growth is target specific.

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Using an alternative method to cloning, known as the panhandle-PCR method (described in

section 3.2.4.2), it was possible to sequence regions of genomic DNA that surround the

transposon insertion site. This provided information about the gene/s that have been

disrupted to cause the observed phenotype. Sequence analysis of the three non anti-fungal

mutants indicated that they had all been disrupted in the same DNA region. The transposon

had inserted into identical positions and orientation within the genome of mutants FM1 and

FM3, whereas in mutant FM2 the transposon had inserted in the same position but in the

reverse orientation. Since each of these mutants were selected from separate experiments it is

not possible that they are simply clones of the same mutant. The low frequency of specific

non anti-fungal mutants generated by this method should also be noted. In one case

approximately 45000 transconjugants were screened resulting in only one non anti-fungal

mutant (FM3). Such low numbers of transposon mutants with a specific phenotype may be

due to several reasons. Firstly, it is possible that during mutagenesis a saturation point is

reached whereby all genes in the pathway have been mutated. This would suggest that the

pathway for the production of the anti-fungal compound is quite short, consisting of only one

gene. A second possibility is that the production of the compound is linked to essential

genes or cellular metabolites, in which case a mutation in any other gene would result in lethal

phenotype. A third possibility is the insertion of the transposon into so called “hot-spots”.

Hots-pots are specific DNA sites where a transposon will preferentially insert. Because

mini-Tn10 was found inserted not only in the same gene but also specifically within the same

location of that gene it is possible that the mini-Tn10 has such hot-spots in the P. tunicata

genome. The transposon Tn10 is known to insert preferentially into specific sites based on

the presence of a 9 bp GC-rich target sequence (5’NGCTNAGCN3’). While improvements

have been made to reduce target site specificity for the mini-Tn10 transposon (Kleckner et al.,

1991), as shown in this and other studies within our laboratory (Srinivasan, 2000), such

improvements have not completely prevented the transposon from having preferred insertion

sites. Interestingly, in the case of mutants FM1-3 the insertion site for mini-Tn10

(5’AGCATTATT3’) does not exactly match that of the consensus sequence, indicating that

other factors are involved in creating hot-spots for mini-Tn10. Since the presence of hot-

spots could interfere with the random insertion of the transposon it is not surprising that a

limited number of mutants were generated which lack anti-fungal activity. Despite the

potential influence of hot-spots, it is possible that the theory of pathway saturation may also

still apply. Wild-type P. tunicata is dark green in colour and a loss of pigmentation results in

a general loss of antifouling activity (see chapter 4). Therefore in this screen for a non anti-

fungal mutant only transposon mutants which remained pigmented were selected. Given that

118

the yellow pigment has been identified as the active anti-fungal molecule (see below) it is

possible that mutations before the last step in the production of the active compound may

have resulted in altered pigmentation and thus were not selected for in this study.

DNA-sequence analysis of the mutants show that the transposon had disrupted a gene

(designated anti-fungal activity gene A or afaA) with high sequence similarity to genes from

various organisms encoding for a long-chain-fatty-acid CoA synthetase (fatty acid:CoA

ligase, AMP-forming; EC 6.2.1.3). This enzyme plays a central role in cellular metabolism

by catalysing the formation of fatty acyl-CoA from fatty acid, ATP and CoA. Activated fatty

acids (fatty acyl-CoAs) are bioactive compounds involved in a variety of processes such as,

intracellular protein transport (Glick and Rothman, 1987), cell signalling (Barber et al., 1997;

Downard and Toal, 1995; Korchak et al., 1994), transcriptional control as well as beta-

oxidation and phospholipid biosynthesis (Black and DiRusso, 1994).

The P. tunicata afaA gene product was found to be most similar to the E. coli acyl-CoA

synthetase known as FadD. This inner-membrane associated enzyme has been extensively

studied for its role in the activation of exogenous long-chain fatty-acids into metabolically

active CoA thioesters concomitant with their transport across the inner membrane. The

activated fatty acids then serve as substrates for beta-oxidation or are incorporated into

cellular phospholipids (Black et al., 1992). Interestingly, Barber et al (1997) have also

identified a gene (rpfB) with high similarity to FadD. RpfB is predicted to be involved in the

synthesis of a small diffusible signal molecule (DSF) which regulates pathogenicity in the

plant pathogen Xanthomonas campestris (Barber et al., 1997). Fatty-acid derivatives have

been shown to function also as signal molecules in other organisms including Myxococcus

xanthus (Downard and Toal, 1995), Ralstonia solanacearum (Flavier et al., 1997) and

Salmonella dublin (El-Gedaily et al., 1997).

Downstream of the fadD gene in E. coli is the rnd gene that encodes ribonuclease D, a

putative tRNA processing enzyme. A partial sequence with similarity to the E. coli rnd gene

was also identified downstream of afaA in the P. tunicata genome. However, the organisation

of genes upstream of the afaA gene in P. tunicata differs from the region upstream of the

fadD gene in E. coli. Directly upstream of the afaA gene is a gene (afaB) encoding for a

putative protein with similarity to a group of enzymes known as the serine hydrolases. The

common feature of this group of enzymes is that they all catalyse the breakage of substrates

with a carbonyl-containing group or ester (Upton and Buckley, 1995). In addition, a

conserved motif corresponding to the serine active site of triglyceride lipases was detected.

119

This may suggest that the substrate for the putative esterase is a triglyceride. The proximity

of the genes afaB and afaA and the absence of a terminator sequence downstream of afaB

suggest that they may be transcribed together within same operon. Following the

translational stop of afaA is a sequence which resembles that of a ρ-independent terminator of

transcription, indicating that any downstream ORFs are unlikely to be encoded on the same

operon and thus will not be directly affected by the insertion of a transposon upstream.

Chemical characterisation of the anti-fungal compound was performed to determine its

structure. The agar-plate based bioassay (see section 3.2.1) suggests that the active

compound is secreted into the extracellular environment. However, testing of cell-free

supernatant and extracts of the supernatant and the cell pellets indicate that the compound is

relatively non-polar and cell-associated. As an initial step in the purification process the

crude cell-extract was separated into 13 fractions using solid-phase chromatography which

separates compounds based on their polarity. The anti-fungal activity was concentrated in

one fraction, which also contained the majority of the yellow pigment of P. tunicata. Further

purification was carried out using a combination of chromatographic procedures and the

purified compound was further characterised using a number of analytical tools such as GC-

MS and NMR. While the exact chemical structure of the active compound has not yet been

determined it is believed to consist of a carbon ring bound to a aliphatic or fatty-acid side

chain.

The role the putative enzymes AfaA and AfaB play in the production of the anti-fungal

compound is yet not clearly defined. However it seems certain that fatty-acids are involved as

supported by the nature of the active molecule and by the strong homology of the AfaA and

AfaB proteins to fatty-acid synthetases and lipases, respectively. The current working model

is illustrated in Figure 3.11. AfaB may function as an extracellular lipase that acts to degrade

certain lipids resulting in the release of specific fatty-acids. AfaA is then required to activate

(by the addition of co-enzyme A) and transport these fatty-acids from the environment, the

activated fatty-acids then form the fatty-acid side chain of the anti-fungal compound (yellow

pigment). In this model a second (yet unidentified) enzyme is required to ligate the activated

fatty-acid to the carbon ring structure of the yellow pigment. Several variations on this model

are possible. Firstly, AfaB may be intracellular, in which case it is the lipids already in the

cell which are degraded and subsequently activated by AfaA. Secondly, it is possible that

AfaA acts to ligate the fatty-acid directly to the carbon ring structure of the yellow pigment

rather then ligating a co-enzyme A.

120

In the case of the non anti-fungal mutant, which remains dark green in colour, preliminary

chemical analysis of its yellow pigment indicates that it differs from the wild-type pigment by

a slight increase in its molecular weight. It is possible that without the uptake of exogenous

fatty-acids, a different side chain (possibly larger) is added to the carbon ring, which gives

this molecule its yellow colour but not the anti-fungal activity.

This chapter has highlighted the advantages gained from using a multi-directional approach

toward the identification of natural products. The use of specific bioassays, genetics and

chemistry makes it possible not only to identify individual components but to gain a more

comprehensive understanding of the role, biosynthesis and mode of action of important

biologically active molecules. This information will be invaluable for the future application of

natural products as successful biological control agents.

121

A

AfaB

AfaA

CoA

afaB afaA

+

?

LipidsFatty-acids

Cytoplasm

Extracellular environment

Yellow pigment with anti-fungal activity

B

AfaB

afaB afaA +

?

LipidsFatty-acids

Cytoplasm

Extracellular environment

Yellow pigment without anti-fungal activity

Figure 3.11: The hypothetical model for the involvement of AfaA and AfaB in the synthesis

of the anti-fungal compound. A) wild-type cells, B) non anti-fungal mutant cells (see text for

details).

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4. Generation and analysis of transposon mutants of P. tunicata

altered in normal pigmentation

4.1. Introduction

The antifouling bacterium P. tunicata produces both a yellow pigment and a purple pigment

which when combined give the bacterium a dark green appearance. Chapter 3 investigated the

anti-fungal activity of P. tunicata and it was shown that the yellow pigment was responsible

for the ability of P. tunicata cells to inhibit the growth of a wide range of yeast and fungal

isolates.

Microorganisms produce a variety of pigments that are often important for the general

physiology and survival of the producing organism in its natural habitat. Photosynthetic

bacteria utilise chlorophyll pigments to capture light for ATP synthesis. The biological role of

pigment production in heterotrophic bacteria is varied. The brown/ black melanin pigments

produced by a number of bacteria have been suggested to protect the cells from desiccation

and UV irradiation (Margalith, 1992). Carotenoid pigments (yellow to red) have a protective

role against photo-oxidation or damage caused by visible light irradiation. Carotenoids have

also been suggested to substitute for sterols as an important structural component of microbial

membranes (Margalith, 1992). Several bacterial pigments function as antagonists against

other organisms. For example, violacein, the purple pigment produced by Chromobacterium

violaceum is known to have anti-bacterial activity (Lichstein and van de Sand, 1945).

Phenazine pigments commonly produced by members of the genus Pseudomonas and the

genus Streptomyces also display anti-bacterial activity (Thomashow and Weller, 1988;

McDonald et al 1999). Bacterial quinones vary in colour from yellow, orange to red and

display a number of biological properties including anti-fungal, anti-bacterial and insecticidal

activity (Margalith, 1992).

In addition to the yellow pigment being responsible for the anti-fungal activity a second

correlation between pigment and antifouling activity has been observed in P. tunicata. When

grown in rich medium such as Luria broth or TSB, P. tunicata loses its pigmentation and

along with this the inhibitory activity against each of the different target organisms

(Holmström et al., unpubl.). These observations have led to the hypothesis that the synthesis

123

of all inhibitory compounds by P. tunicata is linked to the production of pigment in this

organism. To address the relationship between pigmentation and antifouling compounds,

transposon mutagenesis of P. tunicata was employed. Mutants altered in wild-type

pigmentation were selected and characterised on a phenotypic and genotypic level. The data

obtained from the analysis supports the link between pigmentation and antifouling properties

in P. tunicata and has lead to the identification of genes potentially involved in synthesis and

regulation of pigment and antifouling compounds.

4.2. Material and Methods

4.2.1. Transposon Mutagenesis

The basic transposon mutagenesis protocol is as outlined in section 3.2.2 with the exception

that fungal spores were not used in the screening process. The conjugation mix was plated

onto VNSS media containing 200 µg/ml of streptomycin (Sm) and 85 µg/ml kanamycin (Km)

and incubated for 48 h. Mutant colonies altered in normal pigmentation were then selected for

further analysis.

4.2.2. Phenotypic characterisation of pigmented P. tunicata transposon

mutants

4.2.2.1. Analysis of pigmentation (UV/Visible light spectra)

The ultra-violet and visible light spectra of pigments produced by each of the transposon

mutants were determined. Pigments were extracted by adding re-distilled methanol (40 ml/g

wet cells) to the cells and stirring the solution over gentle heat for approximately 10 min. The

solution was filtered (Whatman filter paper 5 mm) to remove cell debris and the UV/Visible

spectra determined with a Beckman DU 640 spectrophotometer.

4.2.2.2. Antifouling activity

Each of the different pigmented transposon mutants were assessed for activity against

different target organisms. The settlement of invertebrate larvae using the barnacle Balanus

amphitrite and the hydroid Hydroides elegans was determined as described in section 3.2.3.4.

124

Activity against the germination of spores from the algae Ulva lactuca and Polysiphonia was

determined as described in sections 2.2.3 and 2.2.12, respectively. Anti-fungal activity of the

pigmented mutants was assessed using the bioassay outlined in section 3.2.1. Anti-bacterial

activity was assayed on agar plates using the overlay method as described in section 3.2.3.2.

Anti-bacterial activity of supernatant samples was assessed by the drop plate method (James et

al., 1996). A bacterial lawn of target P. tunicata cells was spread onto a VNSS agar plate and

allowed to air dry. Thereafter, 20 µl drops of test sample was added to the plate and after

incubation (24 h at 23 oC) the bacterial lawn was assessed for zones of growth inhibition

surrounding the drop. Supernatant from high cell density cultures of both P. tunicata wild-

type and pigmented mutant strains were prepared as outlined in section 2.2.4 with the

exception that cells were resuspended into sterile NSS and not seawater. The total protein

concentration was determined using the BCA method described in section 4.2.4.2.

4.2.2.3. Assessment of bacterial growth

A comparison of the growth rates of the pigmented mutants and the wild-type strain of P.

tunicata was performed in 500 ml flasks. Each flask containing 200 ml of VNSS medium for

the wild-type and VNSS medium containing the selective antibiotics Km (85 µg/ml) and Sm

(200 µg/ml) for the transposon mutants. One percent (v/v) of an overnight culture was

inoculated into appropriate flasks and incubated shaking at 23 oC. Growth was monitored by

absorbancy readings (610 nm) over a 24 h period. These experiments were carried out in

duplicates for each strain.

4.2.3. Genotypic characterisation of pigmented transposon mutants of P.

tunicata

4.2.3.1. Gene sequencing by panhandle-PCR

Genomic DNA was extracted from each of the pigmented transposon mutants of interest

using the XS-buffer method detailed in section 3.2.4.1. DNA flanking the transposon

inserted into the genome of each mutant was amplified using the panhandle-PCR method.

Panhandle-PCR, DNA-sequencing and sequence analysis was performed as described in

sections 3.2.4.2, 3.2.4.3 and 3.2.4.4, respectively.

125

4.2.4. Analysis of the proteins secreted by the wild-type and the white

mutant 3 (W3) strains of P. tunicata

4.2.4.1. Sample preparation and ammonium sulphate precipitation

Overnight precultures of both wild-type and white mutant 3 (W3) (see section 4.3.1) were

used to inoculate 200 ml of VNSS (Appendix I). The growth of these cultures was monitored

and 30 ml samples removed at early-stationary growth phase (12 h) and late-stationary growth

phase (24 h). The cells were gently pelleted (2000 x g for 10 min) the supernatant removed

and sterile filtered (0.22 µm).

To concentrate the proteins from each sample ammonium sulphate precipitation was used.

The supernatant was transferred to a pre-chilled beaker and 85 % (w/v) of ammonium

sulphate was added slowly whilst stirring on ice. After the salt had fully dissolved, the

solution was left stirring gently on ice for 30 min. The precipitated proteins were collected by

centrifugation at 75 600 x g for 30 min at 4 oC and resuspended in approximately 500 µl

milli-Q water. For long term storage the samples were placed in 50 % (v/v) glycerol at –20oC.

4.2.4.2. Protein determination

The total protein concentration in a sample was estimated by the bicinchoninic acid (BCA)

method using a microtitre plate based assay. This system combines the reaction of proteins

with Cu 2+ ions to yield cuprous ions (Cu 1+ ) which in the presence of BCA leads to a colour

reaction. Ten microlitres of each protein sample (diluted in NSS when necessary) were added

in triplicate to microtitre wells. To each well 200 µl of freshly prepared working reagent,

consisting of 50 parts bicinchoninic acid solution (Sigma) and 1 part 4 % (w/v) CuSO4

solution (Sigma), was added. The plate was incubated at 37 oC for 30 min. The colour

reaction was measured by absorbence at 595 nm using a model 3550 microplate reader

(BioRad). Samples were compared to a standard curve consisting of known concentrations of

bovine serum albumin (BSA) (Sigma) ranging from 0.2 to 1.2 mg/ml. A blank consisting of

only the diluent and the working reagents was used as a base line reaction.

126

4.2.4.3. Sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-

PAGE)

Electrophoretic analysis of supernatant samples were performed using a discontinuous buffer

system based on an established protocol (Laemmli, 1970). The stacking gels consisted of 5.5

% (v/v) acrylamide / bis acrylamide solution (15:1 ratio) (BioRad), 0.125 M Tris-HCl (pH

6.8), 0.3 % (w/v) sodium dodecyl sulphate (SDS), 0.1 % (w/v) ammonium persulphate (APS),

0.2 % (v/v) tetramethylethylenediamine (TEMED). The separating gels contained 0.375 M

Tris-HCl (pH 8.8), 0.1 % (w/v) SDS, 0.06 % (w/v) APS, 0.06 % (v/v) TEMED and 12 %

(v/v) acrylamide / bis acrylamide solution. Gels were cast in a BioRad Mini-protean II

electrophoresis unit. Equal concentrations of total protein from each sample were mixed with

sample buffer containing 0.125 M Tris-HCl (pH 6.8), 4 % (w/v) SDS, 20 % (v/v) glycerol, 10

% (v/v) 2-mercaptoethanol and a twentieth volume of 10 % (v/v) bromophenol blue solution.

Broad range molecular weight markers (BioRad) were prepared by adding sample buffer.

Both samples and markers were boiled at 100 oC for 90 seconds. The samples were loaded

on the gel (15 µl) and electrophoresis was performed at a constant current of 30 mA until the

loading dye had migrated to the opposite end of the gel. The tank buffer consisted of 0.025

M Tris-HCl pH 8.3, 0.192 M glycine, 0.1 % (w/v) SDS.

4.2.4.4. Silver staining

For sensitive detection of proteins, silver staining was performed. Gels were fixed overnight

in a solution of 40 % (v/v) methanol and 10 % (v/v) acetic acid and thereafter soaked in a

second solution containing 5 % (v/v) ethanol and 1 % (v/v) acetic acid for a period of 30 min.

The solution was discarded and the gels washed in milli-Q water for 2 min. This step was

repeated twice. After washing, the gels were soaked in a 0.02 % (w/v) sodium thiosulphate

solution for 1 min and thereafter washed a further 3 times with milli-Q water for 2 min each

wash. Gels were then stained shaking gently in a solution of 0.2 % (w/v) silver nitrate and

0.075 % (v/v) formaldehyde for 20 min. After this the gels were rinsed briefly in milli-Q

water then developed in a pre-chilled solution of 6 % (w/v) sodium carbonate, 0.004 % (w/v)

sodium thiosulphate and 0.05 % (v/v) formaldehyde. After the desired intensity of staining

had been achieved the development reaction was stopped by placing the gels in a 5 % (v/v)

acetic acid solution for a minimum of 5 min and gels were thereafter stored in milli-Q water.

127

4.3. Results

4.3.1. Generation of transposon mutants

To investigate the link between the production of pigment and antifouling components,

transposon mutants of the antifouling bacterium P. tunicata were generated. The mini-Tn10

system used for transposon mutagenesis was successful in generating several mutant strains

of P. tunicata with altered pigmentation. Approximately 10 % of the transposon mutants were

changed in colour, including purple, yellow and white mutants (Figure 4.1). The frequency

with which each pigmented mutant was obtained varied, however in general the most abundant

of the pigmented mutants were dark purple followed by yellow, light purple and white

phenotypes. Each of different mutant phenotypes were screened for their antifouling ability

and the results of phenotypic and genotypic analysis of these mutants are presented below.

4.3.2. Phenotypic Characterisation

4.3.2.1. Analysis of pigmentation (UV/ Visible light spectra)

To obtain a qualitative guide to the pigment/s produced by each of the different transposon

mutants, UV/Visible light scans were performed on cell extracts. Figure 4.2 shows the results

of these scans. As can be seen, the dark green colour of P. tunicata is a combination of a

purple pigment (575 nm) and a yellow pigment (425 nm). Both the dark and light purple

mutants have lost the yellow pigment as indicated by the loss of a peak at 425 nm. Similarly,

the yellow mutant has lost the peak corresponding to the purple pigment. White mutants have

lost the pigments corresponding to both peaks, indicating a loss of both the purple and yellow

pigments.

128

Figure 4.1: Transposon mutants of P. tunicata with changes in pigmentation. Mutants were

generated using the mini-Tn10 mutagenesis system and those with an altered pigmentation

were selected.

129

A

B

C

130

D

E

Figure 4.2: UV/Visible light spectra of cell extracts from P. tunicata wild-type and

pigmented mutants. A) wild-type; B) yellow mutant; C) dark purple mutant; D) light purple

mutant and E) white mutant.

131

4.3.2.2. Antifouling activity

4.3.2.2.1. Anti-larval activity

Activity against both the cyprid larvae of the barnacle B. amphitrite and larvae from the tube

worm H. elegans was assessed. Results from representative assays in Table 4.1 show that the

yellow mutant retained wild-type activity and was capable of preventing the settlement of B.

amphitrite larvae and of H. elegans larvae as compared to controls containing seawater alone.

However, the light purple, dark purple and white mutants were shown to have strongly reduced

or no ability to inhibit larval settlement.

Table 4.1: Settlement of marine invertebrate larvae in the presence of biofilms of P. tunicata

wild-type and transposon mutant strains

Percentage settlement a

Target

organism

Wild-

type

Yellow Dark

purple

Light

purple

White No

biofilm

Balanus

amphitrite

larvae

0 0 77 ± 6.5 86 ± 4.7 79 ± 6.5 95 ± 1.6

Hydroides

elegans

larvae

0 0 88 ± 4 83 ± 8 57 ± 7 60 ± 5.5

a All values are means ± standard deviations (n=3)

4.3.2.2.2. Anti-algal activity

The ability of the pigmented mutants to inhibit the germination of spores from two types of

common macroalgae was determined and the results of representative assays are given in

Table 4.2. From these data it can be seen that the yellow mutant is able to maintain the ability

to inhibit the germination of both U. lactuca spores and that of Polysiphonia sp. whilst the

132

anti-algal activities of the white, light purple and dark purple mutants are greatly reduced or

lost.

Table 4.2: Germination of marine algal spores in the presence of biofilms of P. tunicata

wild-type and transposon mutant strains

Percentage germination a

Target

organism

Wild-

type

Yellow Dark

purple

Light

purple

White No biofilm

Ulva lactuca

spores

0 0 71 ± 23 122 ± 19 91 ± 3 100

Polysiphonia

sp. spores

0 0 60 ± 4 79 ± 19 87 ± 6 81.5 ± 2

a All values are means ± standard deviations (n=3)

4.3.2.2.3. Anti-fungal activity

To determine if the pigmented mutants were able to inhibit fungal growth each mutant was

tested against a range of fungal isolates. Figure 4.3 shows the results of a typical anti-fungal

bioassay. It can be seen that all pigmented mutants with the exception of the yellow mutants

were defective in their ability to inhibit the growth of fungi.

133

Figure 4.3: Anti-fungal activity of pigmented mutants of P. tunicata. The target fungus

(Penicillium sp.) was spread plated onto VNSS agar plates and air dried. P. tunicata

pigmented mutants were then inoculated in small circles onto the agar plate containing the

target fungus and incubated at 30 oC for 48 h. Wt = wild-type P. tunicata, W = white mutant,

Lp = light purple mutant, Dp = dark purple mutant, Y = yellow mutant.

134

4.3.2.2.4. Anti-bacterial activityThe pigmented mutants were assessed for anti-bacterial activity against a range of bacterial

isolates including wild-type P. tunicata. Experiments using the overlay method (section

3.2.3.2) showed that with the exception of the white mutant variety all of the mutants retained

their ability to inhibit bacterial growth. However, it was observed that in comparison to the

other strains the light purple mutants whilst remaining inhibitory had a reduced anti-bacterial

activity (data not shown). The anti-bacterial compound produced by P. tunicata cells has been

identified as a large extracellular protein (James et al., 1996). To clarify the observations

concerning the light purple mutant a second assay was performed in which supernatant from

each of the mutant and wild-type strains were collected and the concentration of total protein

determined. Equal concentrations (with respect to the total protein) of the supernatant sample

were assayed in the drop test method (James et al., 1996) for anti-bacterial activity against a

lawn of wild-type P. tunicata cells. The results of this assay are summarised in Table 4.3.

At concentrations of 6 mg/ml the supernatant from yellow, dark purple and light purple mutant

strains displayed a similar degree of inhibition as the wild-type. However, whilst the yellow

and dark purple mutants remain comparable to the wild-type over the full range of

concentrations tested, the light purple strains appear to have reduced effect at lower

concentrations of total supernatant protein.

Table 4.3: Growth inhibition of P. tunicata wild-type by supernatant from P. tunicata wild-

type and transposon mutant strains

Growth inhibition (cm) a

Concentration of

total protein (mg/ml)

Wild-type Yellow

mutant

Dark purple

mutant

Light purple

mutant

White

mutant

6 1.3 1.2 1.3 1 0

3 1.1 1 1.2 0.4 0

1.5 0.8 0.7 1 0 0

0.6 0 0 0 0 0

0 0 0 0 0 0

a The values are of the diameter, in cm, of growth inhibition zones of the target strain

135

4.3.2.3. Assessment of growth

To ensure that the differences seen in the antifouling bioassays were not due to differences in

the growth of the transposon mutants, growth curves were determined. No differences in the

growth pattern between wild-type P. tunicata and the light purple or dark purple transposon

mutants were observed. However, the white mutants tended to reach maximum growth

slightly earlier than the wild-type. Wild-type P. tunicata appears to enter into a bi-phasic

growth pattern at approximately mid to late logarithmic growth (Figure 4.4). Interestingly,

this coincides with the appearance of pigment in the wild-type strain.

136

3 02 01 000.0

0.2

0.4

0.6

0.8

1.0

Wild-type (A)Wild-type (B)White mutant 3 (A)

White mutant 3 (B)

Time (hours)

OD

610

nm

3 02 01 000.0

0.2

0.4

0.6

0.8

1.0

Wild-type (A)

Wild-type (B)

White mutant 2 (A)

White mutant 2 (B)

Time (hours)

OD

610

nm

A

B

Figure 4.4: Growth curves of P. tunicata pigmented mutants compared with wild-type

cultures A) wild-type and white mutant 2 B) wild-type and white mutant 3.

137

4.3.3. Genotypic characterisation of transposon mutants

4.3.3.1. DNA sequence analysis

4.3.3.1.1. Sequence analysis of the light purple mutants

Eight mutants with a light purple phenotype, designated LP 1 to LP 8, were further analysed.

The mutants were selected from different sets of experiments in order to eliminate the

possibility of analysing clones. Initial sequence analysis of the regions directly flanking the

transposon in each of the 8 light purple mutants indicated that they had been disrupted in the

in two different DNA-regions. Sequencing from both sides of the transposon in mutants LP

2 and LP 3 was continued using the primer walking strategy in Figure 4.5 and Figure 4.8.

After sequence assembly, a consensus sequence was obtained and the nucleotide sequence

was submitted to the programs ORF-finder and BLAST X from NCBI. The primary

sequence data for LP 2 is shown in Figure 4.6.

The results of the analysis indicate that the transposon had disrupted a 1110 bp open reading

frame (ORF 1) with 33 % identity and 52 % similarity (over 353 amino acid residues) to a

putative oxidase (GenBank accession number AL021529) from Streptomyces coelicolor.

Further sequence analysis of this region revealed a putative ribosome binding site (RBS)

5'AGGAGGT3' located 6 bp upstream of the ATG start codon. A potential transcriptional

start point was identified 45 bp upstream of the start of the ORF. Upstream of this region

contains sequences as highlighted in Figure 4.6 that are in agreement with -10 and -35

sequences for E. coli σ70 -responsive promoters.

Analysis of the deduced AA sequence of ORF 1 indicates that this protein has a molecular

weight of 43201 Da and a predicted isoelectric point (pI) of 7.56. The protein is primarily

hydrophilic and was not found to contain any transmembrane regions.

Figure 4.7 shows the multiple sequence alignment of the deduced amino acid sequence of

ORF 1, designated LppA (Light purple phenotype A) with the sequences of other genes with

high sequence similarity. These include 3-chlorobenzoate-3,4-dioxygenase (GenBank

accession number D90912) from Synechocystis sp., toluenesulfonate methyl-monooxygenase

TsaM (GenBank accession number U32622) from Comamonas testosteroni, and

aminopyrrolnitrin oxidase (GenBank accession number U74493) from Pseudomonas

fluorescens. The region between amino acid residues 56 and 106 shows homology to a

138

Rieske iron-sulfur [2Fe-2S] cluster. This provides further evidence that LppA has an oxidase

function.

Directly downstream of lppA is the start of a second open reading frame. This ORF has

similarity to enzymes with transferase activity, including 56 % similarity (over 286 amino acid

residues) to a probable transferase from Streptomyces coelicolor, 57 % similarity (over 120

amino acid residues) to 8-amino-7-oxononanoate synthase (BioF) from Bacillus sphaericus

and 57 % similar (over 120 amino acid residues) to glycine acetyl-transferase (2-amino-3-

ketobutyrate) from B. subtilis. The proximity of this open reading frame to lppA and the lack

of an obvious terminator of transcription downstream of the lppA termination codon, suggests

that both of these open reading frames may be in the same operon and thus the genes are co-

transcribed. This putative gene was therefore designated lppB.

Sequence analysis of the DNA flanking the transposon in the mutant strain LP 3 revealed that

the sequence aligned with a region of DNA downstream of the disrupted genes in mutant

strains DP 3 and DP 5 (Figure 4.8). Therefore, this sequence analysis is presented together

in section 4.3.3.2.2.

139

0

Lp2TnD-S5

Lp2TnD-S3

500 1000 1500 2000

Lp2TnD-S14

Lp2TnD-S11

Lp2TnD-S4

Lp2TnC-S13 Lp2TnC-S10

Lp2TnC-S8

Lp2TnC-S12

Lp2TnC-S7

Lp2TnC-S9

Ap2Ap2

Ap2 Tn10C-S1Tn10D-S1

Figure 4.5: Summary of the sequencing strategy to determine the nucleotide sequence

flanking the transposon insert in the P. tunicata light purple mutant 2 (LP 2) genome. Arrows

indicate length and direction of sequence primer products. Blue arrows represent transposon

or adaptor specific primers and black arrows represent sequence specific primers. All primers

used are listed in Appendix II. The nucleotide sequence is shown in base pairs along the top

of the diagram. Open reading frames are indicated by the bold coloured lines; red = lppA

(ORF1); blue = lppB (ORF2).

140

1 TCATGTTAGATTCTACGTTTGAGAACGTTTCAACCAAATTGTGCCAGTTTATCGCACAGA

61 GTTATTTGGATGATGAGGAACAAATTTTACCCAATACTCCAATCATCAAACTGAACATTC

121 TTGATTCAGCATCCATTTTTGATGTGGTTGAGTTTATTCATCGCGAATTTTCAGTGCGCC

181 TTCCGGTGC TTGAAA TTCACCCAGATAATT TTAATT CAGTCCAAGTACTGAGTGAGCTGG (-35) (-10) 241 TTTACCGTCAATTTAGCA AGGAGGT GGAACAATGATCCCTAATCAATGGTATGCGATTTT (RBS) M I P N Q W Y A I L

301 ACGCACCCAAGACGTTAAGAAAAAACCTGTAGGTATTAAACGATTAGAAAAGCTGTTGGT R T Q D V K K K P V G I K R L E K L L V

361 ACTTTATCGTGATACAGCGGGTGAATTAGTTTGTCTTGATGATCGTTGCCCTCACAAAGG L Y R D T A G E L V C L D D R C P H K G

421 CGTTAAATTAAGTTTGGGTCAACAACATGGTGATCTGATTGCCTGTCCTTATCATGGTTT V K L S L G Q Q H G D L I A C P Y H G F

481 TCAATACGATCAGCAAGGTGATTGTGTTCATATGCCAGTGCTAGGTCAAAAAGGTAATGT Q Y D Q Q G D C V H M P V L G Q K G N V

541 GCCAAAAGGCATGTGCGTTAAAAGCTACAAGGTTAAAGAAGAGTTAGGCCTGATTTGGTT P K G M C V K S Y K V K E E L G L I W F

601 TTGGTTTGGAGACGTGCGTCAAGAGCATGAATATCCAGATATCCCAATGTTTAAACAACT W F G D V R Q E H E Y P D I P M F K Q L

661 TAAGGAATATAAAGGCCACTATTCCTATTATGGCTGGGATGCTCCAATCAATTACACCCG K E Y K G H Y S Y Y G W D A P I N Y T R

721 CTATGTGGAAAGTGTGTGTGAGATTTATCATATTCCATTTGTCCATAAAGGTTCAGCCAT Y V E S V C E I Y H I P F V H K G S A I

781 CAATATTTGGGATCCAAAAGGAGGTCGAGTTGATAATTTCGAGTGTAAGGTCGAAGACAC N I W D P K G G R V D N F E C K V E D T

841 GCTGATCACCAGTGACTTTATTTTACGTCCAGATGATGACCGCACCGCTGAGGAAACCTT L I T S D F I L R P D D D R T A E E T L

901 AGTTGCTCGTAAACCTTGGACTCGCGGCTGGCGTTTTGGCATTGACGTGCAAATGCCGAA V A R K P W T R G W R F G I D V Q M P N

961 TTTAATCCTTATTCGCAGTGATGTGTTTGATGTGATTTTTATCCCTACACCTATCGATGA L I L I R S D V F D V I F I P T P I D E t 1021 AAATACCTGCTGGGTATGCGTTTGTTATCAAGAGCCTAAACGTGACTTGCTTTTACCACT N T C W V C V C Y Q E P K R D L L L P L

1081 CATTAAACCGTTGCCAATTCCATTTTGGGGGCCTGTGCGACCTTGGTATATGTGCCGAAT I K P L P I P F W G P V R P W Y M C R I

141

1141 TGAGCGTTTTGTGCAGCAAGCAAAAGATATGGCAGTCATTGCCCATCAAGAGCCTAAAGT E R F V Q Q A K D M A V I A H Q E P K V

1201 GTCTCATCCAAAAGCAAACCGTTTAATCCCACTTGATAAAGTTAATGCTCATTATATTCG S H P K A N R L I P L D K V N A H Y I R

1261 CTTTCGTGAAAAGCTAATTCGTGAAGCAAAAGAGCAACAACAAGTGAAAGAGCTTGCCAA F R E K L I R E A K E Q Q Q V K E L A K

1321 GTCTAATCCAGAATTTGCAGCGCCGACACTTGAACAGATTATCGGCATTGAAGAAGTACA S N P E F A A P T L E Q I I G I E E V H

1381 TTAAAGCAA ACGGAAT TTGATATGACAGACAATAAAAACACCGCTATTGAGCAAATACAC * (RBS) M T D N K N T A I E Q I H

1441 GCCCTAGTTATCGACGTAGTAACAGAGCAAACGTGTTATGCCGAGTCGGATTTAATTTTA A L V I D V V T E Q T C Y A E S D L I L

1501 GATGCGCCGATGGAAGAAGGCTTGGGGATAGACTCCATTATCCTTGCTTCTATCGTCAGT D A P M E E G L G I D S I I L A S I V S

1561 GAAATTCAAAAATTGTTTATGTTTGAGACCCGTCTCAATACTGGCAGTTTTAATACCATT E I Q K L F M F E T R L N T G S F N T I

1621 CAAGCATTACTCGACATTTGTCACAATGCGATGCTATCAGACGCAGGAGTGCAAAAACTG Q A L L D I C H N A M L S D A G V Q K L

1681 GCACAATTAGGACTTGCAGCAGCACCACAAGCTGTTTGTGTAAGTTCGCAGCCAGAGCCT A Q L G L A A A P Q A V C V S S Q P E P

1741 GAACAGCGTTCAACTCAGGCACAAACAATGCGAGATTTTGTCGCAGATGGTAGCCCTGAC E Q R S T Q A Q T M R D F V A D G S P D

1801 TTATTTAGTAAAGTGCGTAAGTTTGACCAGTTTTATAAAAATCAGGCTGAGCAAGGTAAC L F S K V R K F D Q F Y K N Q A E Q G N

1861 TTTTGGTACGGCATGCCACTTAGCTCCAGATGTGAAAATCGAGCGACTATTTATGATGGC F W Y G M P L S S R C E N R A T I Y D G

1921 TATCAGAAAAAAGAACGTGAATTCTTAATGTTTGCCTCGAATAATTATTTGGGGTTAGCT Y Q K K E R E F L M F A S N N Y L G L A

1981 AACGACCCGCGGGTCATCAAAGCAATCTGTGATGCTACGCAAAAATACGGTGCAACAAAT N D P R V I K A I C D A T Q K Y G A T N

2041 ACAGGTTGTCGTCTGATTGGTGGCACTAATCATTTGCACCTTGAACTGGAAGCACGTTTG T G C R L I G G T N H L H L E L E A R L

2101 GCAGCGTTTAAAGGTCGCGAAGCCTGTATTGTTTTCCCCTCTGGTTATTCGGCTAACCTT A A F K G R E A C I V F P S G Y S A N L

2161 GGTACGATTTCTGCGTTAACTGGTCCAAAAGACACTGTGATTTCAGATGTTTATAATCAC G T I S A L T G P K D T V I S D V Y N H

142

2221 ATGAGTATTCAAGATGGTTGTAAGTTATCAGGTGCAAAACGCCGTATTTACAAACATAAC M S I Q D G C K L S G A K R R I Y K H N

2281 GATATGGATTCGTTAG D M D S L

Figure 4.6: Nucleotide sequence of the genomic-DNA region surrounding the transposon

within the light purple 2 mutant (LP 2) genome. The nucleotide sequence is shown along with

the translated amino acid sequence in one-letter code. The inverted solid triangle (t)

indicates where the mini-Tn10 transposon insertion occurred. Specific open reading frames

(ORF) as indicated in the text are highlighted as follows: ORF 1 (LppA) is shown in red,

ORF 2 (LppB) is shown in blue. Potential promoter regions are underlined as are predicted

ribosome binding sites (RBS).

143

1 50 LppA .......... .......... ..mipnqwya ilrtqdv.kk kpvgikrlek AL021529 .......... .......... ..mipnqwyp iveaqevgnd kplgvrrmgq D90912 .....msals shsqvlsmlr ttpinfnhwy vvaqagelgq gplgivlwek U32622 .......... .......... .mfirncwyv aawdteipae glfhrtllne U74493 mndiqldqas vkkrpsgayd attrlaaswy vamrsnelkd kpteltlfgr ••••••• • • • • •• •• ••••

51 100 LppA llvlyrdtag elvclddrcp hkgvklslgq qhgdliacpy hgfqydqqgd AL021529 dlvlwrdidg nlvcqgarcp hkganlgdgr mkgntiecpy hgfrygadga D90912 aiaiyrdqdg qvravedrcp hrqvklsegk vlgnnlecay hgwqfdaqgh U32622 pvllyrdtqg rvvalenrcc hrsaplhigr qegdcvrcly hglkfnpsga U74493 pcvawrgatg ravvmdrhcs hlganladgr ikdgciqcpf hhwrydeqgq ••••••• • ••••••••• ••••••• • • •• • ••• •••••• ••

101 150 LppA cvhmpvlgqk gnvpkgmcvk sykvkeelgl iwfwfgdvrq eheypdipmf AL021529 crvipamgse aripgslrvp typvreqfgl vwmwwgderp tadlppvaap D90912 cakipyfsed qklppcrlrt ypvqekdgfi wlypgdldhl ashgpeplai U32622 cveipgqeqi ppktciksyp vvernrlvwi wmgdparanp ddivdyfwhd U74493 cvhipghnqa vrqlepvprg arqptlvtae rygyvwvwyg splplhplpe ••• • •• • • • • • •• • • • •

151 200 LppA kqlkeykghy syygwdapin ytryvesvce iyhipfvhkg sainiwdpkg AL021529 aevtdnrkly atkrwtrpvh ytryieslle fyhvtyvhrd hwfnyidyll D90912 pewhhlnhig sfaafdcpgh fsylienlmd myhghlhdny qawasaslre U32622 spewrmkpgy ihyqanykli vdnlldfthl awvhpttlgt dsaaslkpvi U74493 isaadvdngd fmhlhfafet ttavlriven fydaqhatpv halpisafel • • • • •• • •••• ••• • •• • •• • •• ••

201 250 LppA grvdnfeckv edtlitsdfi lrpdddrtae etlvarkpwt rgwrfgidvq AL021529 lygtpskfgl dgrerylaat ritnhrvete aegqtirysf dhcqeddptn D90912 ietngdqvvv dynaqsyyki dkiwsisqlf fptlrrlhpe nlrvsyiyph U32622 erdttgtgkl titrwylndd msnlhkgvak fegkadrwqi yqwsppallr U74493 klfddwrqwp eveslalaga wfgagidftv dryfgplgml sralglnmsq • • •• • • • •• •• • • •

251 300 LppA mpnlilirsd vfdvifiptp identcwvcv cyqepkrdll lplikplpip AL021529 tthyvitftf pcmvhvqteq fettswlvpi ddqntehilr wyeyeqvkpv D90912 wsstlgadfk iyclfcpisa nktkaylvhf tslea..... fpklhklpva U32622 mdtgsaptgt gapegrrvpe avqfrhtsiq tpetettshy wfcqarnfdl U74493 mnlhfdgypg gcvmtvaldg dvkykllqcv tpvsegknvm hmlisikkvg • • ••• •• ••• ••

144

301 350 LppA fwgpvrpwym cr....ierf vqqakdmavi ahqepkvshp kanrlipldk AL021529 lrfeplrrll pwaslymekw vqdpqdvrim ehqepkisag gvnkfipvde D90912 frsflknwls gtarpllegl idqdirmisq eqaafeqnpd rqnvevnpal U32622 ddealtekiy qgvvvafeed rtmieaheki lsqvpdrpmv piaadaglnq U74493 gilrratdfv lfglqtrqaa gydvkiwngm kpdgggaysk ydklvlkyra • • ••• ••• • ••••• • • ••••

351 400 LppA vnahyirfre klireakeqq qvkelaksnp efaaptleqi igieevh... AL021529 mnakyismra kliadasaap ssparaaepe peaagrggsa aratgngrga D90912 akvqqlirqq alasena... .......... .......... .......... U32622 grwlldrllk aenggtap.. .......... .......... .......... U74493 fyrgwvdrva ser....... .......... .......... .......... •• ••••• ••• •• • ••

401 417 LppA .......... ....... AL021529 aggrrgtkpk edaaarp D90912 .......... ....... U32622 .......... ....... U74493 .......... .......

Figure 4.7: Multiple sequence alignment of the deduced amino acid sequence of the P.

tunicata LppA protein with Streptomyces coelicolor putative oxidase (AL021529);

Synechocystis sp. 3-chlorobenzoate-3,4-dioxygenase (D90912); Comamonas testosteroni

toluenesulfonate methyl-monooxygenase oxygenase component TsaM (U32622) and

Pseudomonas fluorescens aminopyrrolnitrin oxidase PrnD (U74493). A Rieske iron-sulfur

cluster region is highlighted in blue. Conserved cysteine residues at positions 69 and 88 and

conserved histidine residues at positions 71 and 91 are the predicted iron-sulfur binding sites.

Residues identical between the P. tunicata LppA protein and one other protein are indicated

by black dots (•); residues identical in two other proteins are indicated with blue dots (•);

residues which are identical between three proteins are indicated by green dots (•) and

residues which are identical in all 5 proteins are indicted by red dots (•). Small dots (.) denote

gaps. Numbers shown in parentheses are the GenBank accession numbers.

145

4.3.3.1.2. Sequence analysis of the dark purple mutants

The genomic DNA flanking the transposon in two transposon mutants with dark purple

phenotype (DP 3 and DP 5) was sequenced and was found to align directly upstream of the

DNA sequence surrounding the transposon in mutant LP 3 (see section 4.3.3.1.1). The

combined DNA sequence of DP 3, DP 5 and LP 3 resulted in a total of 3913 bp of DNA.

The sequencing strategy using PCR and primer walking is shown schematically in Figure 4.8.

An additional panhandle-PCR primer (Lp3pan2) was designed and PCR products obtained to

continue sequencing further along this region of the genomic DNA.

Analysis of the sequencing data indicates that the transposon mutants DP 3 and DP 5 were

disrupted in different regions of a 1257 bp ORF (Figure 4.9). Sequence analysis of the

region surrounding this ORF designated (dark purple phenotype A) dppA, revealed a putative

RBS (5'AAGGAAT3') located 9 bp upstream of the ATG start codon. A potential

transcriptional start point was identified and upstream of this region contains sequences, as

highlighted in Figure 4.9 that are in agreement with -10 and -35 sequences for E. coli σ70 -

responsive promoters.

Analysis of the deduced amino acid sequence of DppA shows that the protein has a molecular

weight of 45129.6 Da and a theoretical pI of 6.46. The hydrophobicity profile as predicted by

the method of Kyte and Doolittle (1982) and secondary structure prediction by the SOSUI

method (available through the ExPASy web site) indicated that the protein has 4 possible

transmembrane regions (Figure 4.11), thus suggesting that this protein is membrane bound.

DppA is similar to conserved hypothetical integral membrane proteins including; 34 % similar

(over 344 amino acid residues) to protein RP699 from Rickettsia prowazekii (GenBank

accession number AJ235272); 42 % similar (over 304 amino acid residues) to permease

HI1548 from Haemophilus influenzae (GenBank accession number P44250) and 40 %

similar (over 225 amino acid residues) to a putative integral membrane protein from Neisseria

meningitidis (GenBank accession number CAB84643).

Approximately 10 bp downstream from dppA is a 702 bp ORF termed dppB. Analysis of this

region identified the putative RBS (5'AGGAAAT3') located 3 bp upstream of the start codon.

The deduced amino acid sequence indicates that the dppB gene encodes a 25937.8 Da protein

with a theoretical pI of 6.21. Hydrophobicity profile indicates that this protein is soluble and

without any transmembrane regions.

146

DppB shows high similarity to ABC-transporter proteins including 45 % identity and 60 %

similarity (over 226 amino acid residues) to the Bacillus subtilis putative ABC transporter,

YvrO (GenBank accession number AJ223978) and 42 % identity and 61 % similarity (over

219 amino acid residues) to the Streptococcus cristatus ATP-binding cassette protein, TptC

(GenBank accession number AAB97961) (Figure 4.10). A putative ATP/GTP-binding site

motif (GPSGSGKS) was detected between amino acids 38 and 45 of DppB. This glycine-

rich region typically forms a flexible loop within the protein, which interacts with one of the

phosphate groups of the nucleotide. This motif is also known as the 'A' consensus sequence

(Walker et al., 1982) or the 'P-loop' motif (Saraste et al., 1990) and can be detected in most

proteins which bind ATP or GTP. In addition, a signature sequence for the ABC-transporter

protein family was detected (LSGGQQQRVAVRAI) between amino acids 142 and 156.

Analysis of the sequence surrounding the transposon of the LP 3 mutant indicated that this

strain was disrupted in a 1179 bp ORF located downstream of dppB. This new ORF was

designated dppC. A predicted RBS (5'CTGAAGT3') was located 5 bp upstream of the ATG

start codon for dppC. From the deduced amino acid sequence the protein has a molecular

weight of 44758.9 Da and a theoretical pI of 4.92. The hydrophobicity profile indicated that

the protein is soluble and is unlikely to contain any transmembrane regions. No similarity

was found between DppC and other proteins currently in the Swissprot or EMBL databases.

The first 431 bp of a fourth ORF (dppD) was sequenced downstream from dppC. Initial

sequence analysis of this region indicated that dppD is 36 % identical and 55 % similar (over

142 amino acid residues) to a putative methyl transferase from Streptomyces coelicolor

(GenBank accession number CAA16186). Due to the proximity of the genes encoding each

of these proteins and to the lack of identified transcription terminators following each of the

ORFs, it is predicted that dppA through to dppD are encoded in the same operon. This

operon may also encode other genes further downstream of dppD.

147

1000 2000 30000

Dp3TnD-S2

Ap2

Dp3TnD-S7

Dp3TnC-S3

Dp3TnC-S5

Tn10D-S1

Tn10D-S1

Tn10C-S1

Dp3TnC-S6 Dp3TnD-S4

Ap2

Tn10C-S1

Lp3TnC-S2Ap2 Tn10D-S1

Lp3TnD-S3

Tn10C-S1 Ap2

Lp3TnD-S2

Lp3pan2-S3

Lp3pan2

Lp3pan2-S2

Lp3pan2-S4

Ap2

Figure 4.8: Summary of the sequencing strategy to determine the nucleotide sequence

flanking the transposon insert in the P. tunicata dark purple mutants 3 and 5 (DP 3, DP 5)

and light purple mutant 3 (LP 3) genomes. Arrows indicate length and direction of sequence

primer products. Blue arrows represent transposon or adaptor specific primers and black

arrows represent sequence specific primers. All primers used are listed in Appendix II. The

nucleotide sequence is shown in base pairs along the top of the diagram. Open reading

frames are indicated by the bold coloured lines; red = dppA; blue = dppB; green = dppC and

purple = dppD.

148

1 AAAAGGTGGCAATGTTGTTTGGCTCGTGACAGTGGAAACGTTATTACTGACTTTTGTTGC

61 GTCTTTGGCTGGCATCGCGATGGGACTTATTTTGGGCAGCTATTTGCAGCAAAATGGTTG

121 GGACATCAGTCAGTTTGGTGAATTTAGCCTGGCAGGTGTTGGGATGACCAGTGCCCTAAA

181 AGCAAAACTAACCG TTGAGA ATGTGATCACCCCAGTTGTGGTGATGTT TATTAT TGCCAT (-35) (-10) 241 ACTTGCTGCGCTTTATCCGGCCTTCTCTGCAGCACGTTTGGTGCCAGCAC AAGGAAT GAG (RBS) 301 AGCCACATGATACATAAGTTAGCTTTACGCAATTTATTGCGCAATAAACGCCGTTCAATA M I H K L A L R N L L R N K R R S I

361 CTGACCTCTGTGATTATTATTTTTGCCTTTAGCATGATGATCTTGTTTATGGGGTTGTCT L T S V I I I F A F S M M I L F M G L S

421 GATGGAGGCCATAAAGCCATGGTCGACATAGGTGTAAAAATGGGGCTTGGCCATGTTGTG D G G H K A M V D I G V K M G L G H V V

481 GTGCAGCACCCACAATATCGGGATGATCCTGCGTTAGCGCATTTGATTCGAACACCAGAA V Q H P Q Y R D D P A L A H L I R T P E

541 GAAGTTAAGCAAACCATTTTGAGCCAGCAGCCACAACTGCAAGTTGTTGCACGCTTGCGG E V K Q T I L S Q Q P Q L Q V V A R L R t 601 GCTGATGCGTTAATTCAAGCGGGTCGACATGGTATTGCGCTGAGTATTTCAGGGGTTGAG A D A L I Q A G R H G I A L S I S G V E

661 CCTGAGCTTGAGCGTCAGGTATCAGCAATTGCCGATGACAAGGCTATTGAGCAAGGGGAA P E L E R Q V S A I A D D K A I E Q G E

721 ACCTTAGCTGCATTTACTCAGTCACATCCCCATGGCAATTTGGCTGGCATCGTACTGGGT T L A A F T Q S H P H G N L A G I V L G

781 GCAACCCTTGCCACCAATCTTGAAGTGCGAATTGGAGACACGGTTACGCTTACGGTAAAA A T L A T N L E V R I G D T V T L T V K

841 CCTGCCAGTGGGGGCGATTTAGCTCGCTCCGCATTTCAAGTTGCTGGGATATTTAAAACC P A S G G D L A R S A F Q V A G I F K T

901 GGATTACATGAACTTGATACCTTCTGGGCTGAAGTGCCGATTACTGCGCTGCAAAGGCTA G L H E L D T F W A E V P I T A L Q R L

961 CTTGAAGTCGACGGTCAAGTCAGTGAATTGGCATTATATTCGCCAAGTGGCAGTGATGGT L E V D G Q V S E L A L Y S P S G S D G

1021 GTGGGCCTGTTGACAGCATCTATTCGTGCACAGTTGCCAGAGTATGGCGTACAACCTTGG V G L L T A S I R A Q L P E Y G V Q P W

1081 CAAACGGCTGCCCCTGAGCTGTATTCGGCAGTGACCTTGGATGCTGCGGGCATGTACTTA Q T A A P E L Y S A V T L D A A G M Y L

149

1141 TTGATGCTGATTGTCTATGTCGTGGTCGCAGTTGGGATCTTAAATACGGTATTGATGTCG L M L I V Y V V V A V G I L N T V L M S

1201 ACTTTTCGCCGCCAAAAAGAATTTTCGATGATGATAGCCGTTGGTGCAAGGGCAAGTACG T F R R Q K E F S M M I A V G A R A S T

1261 GTGACTAAAGTGGTGCTTTTAGAAGCGGTTTACCTCAGTGCTTTTTCACTCTTACTTGGC V T K V V L L E A V Y L S A F S L L L G

1321 CTCGGATTTGGGCTGTGGGGGCATTACTATTTTGCCACCGAAGGACTGAACTTTAAGGAA L G F G L W G H Y Y F A T E G L N F K E

1381 GTGTTTGGCACCGCAATGGAAGCCGGTGGCGTGCTCCTTCCTGAGAAATTTTATTCTACG V F G T A M E A G G V L L P E K F Y S T t 1441 TTGTACACCGATAAATTACTGCTCAGTGTGCTGTTTATTTTTGTTATCACTATCGTCGTT L Y T D K L L L S V L F I F V I T I V V

1501 ACGTTGGTTCCAGCTATTCGCGCTGGTCACCGCTCACCAGTAGCAGCAAGTCACGAATCG T L V P A I R A G H R S P V A A S H E S

1561 TA AAGGAAT GACCATGATTTCACTCACCAAAATCAATAAAATATTTTCTGACAAACATCA * (RBS) M I S L T K I N K I F S D K H Q

1621 ATCATTTCATTGTTTGAAAGACATAGATCTCACCATAGATAAGGGCGAGTTTACTGTGAT S F H C L K D I D L T I D K G E F T V I

1681 TGCCGGACCTTCAGGCTCTGGTAAATCAACGTTGTTAAACATTATAGGCTTATTAGATAA A G P S G S G K S T L L N I I G L L D K

1741 AGCGACCTCAGGAACTTATTTATTTGATGATCTGGATGTATCAACCATGACAAATAATGC A T S G T Y L F D D L D V S T M T N N A

1801 GCTGGCAGATATTCGTCGAGAAAAAATAGGCTTTGTATTTCAAGCATATAATTTAATGCC L A D I R R E K I G F V F Q A Y N L M P

1861 GGTGTTAACCGCATTAGAAAATACTGAAATGATAATGGAGTTTTGTGGTTTGGATAAAAA V L T A L E N T E M I M E F C G L D K K

1921 GCTGCGTCGCCAACGCGCGATGGAGACGTTGACATCGGTTGGACTTGCGGATTTAAAAGA L R R Q R A M E T L T S V G L A D L K D

1981 CCGTTTTCCAGCTCAATTGAGTGGTGGGCAACAGCAGCGAGTCGCAGTTGCTCGTGCGAT R F P A Q L S G G Q Q Q R V A V A R A I

2041 AGCGGCGCAACCTTTGTTAGTTGTTGCTGATGAACCGACCGCAAACTTGGATTCTCACTC A A Q P L L V V A D E P T A N L D S H S

2101 TGCGGAAAATTTGCTTAATTTGATGGTAAAACTCAATCACGATTTAGGCATTACTTTCTT A E N L L N L M V K L N H D L G I T F L

2161 GTTTAGTTCGCACGATCAGCGCGTTATTCAACGAGCGCAGCGAGTACTGCAATTACAAGA F S S H D Q R V I Q R A Q R V L Q L Q D

150

2221 TGGCCAGATAGTGAGCGATGAACGTAAAGATCAGCAACCAAAGCTGGTGGCTCTGTGAAT G Q I V S D E R K D Q Q P K L V A L *

2281 CTGAAGT TTAGCATGCTGTTACTGGCGGGGGCTTTGCACCATGCCAGTAGTTTTGCTCAA (RBS) M L L L A G A L H H A S S F A Q t 2341 GAGGATGATTGGTCTGCGTTATTAGATGATGCTACAGCTCCGGTGTCAAAATGGTCTGGT E D D W S A L L D D A T A P V S K W S G

2401 GTGGTTGAAGGTAATGCCACCGCGATTGATGCTGCTAGCGACAACAGCCTTGGCAGTAAT V V E G N A T A I D A A S D N S L G S N

2461 TGGTTTGCTCGATTGGAATATAAATACACTCAACACGCTTCGCAATGGGTGATACATGCA W F A R L E Y K Y T Q H A S Q W V I H A

2521 CAGCTTGATTATGACTATCTTGATCAAGCATGGGCACTTCCTTTGTTTGCTAATCGTGAT Q L D Y D Y L D Q A W A L P L F A N R D

2581 GTTACCGATCGGCTGGTCGATTTAGAAAAGGACAGTGATAGCTCAAAGCAGTTATGGTAC V T D R L V D L E K D S D S S K Q L W Y

2641 GGACAGATTGACTGGGCTTATTGGCAACATGATTGGCAACAAGGCCGTTTAACGTTAGGT G Q I D W A Y W Q H D W Q Q G R L T L G

2701 CGTCAACCAGTTACAGTCGGATTAGGGCGGATTTTTTCACCTGTCGATCCATTGGGGGCA R Q P V T V G L G R I F S P V D P L G A

2761 TTTAGTGTTTTTGACTTAGACCGTTTATATAAACGTGGGGTTGATGCAATTCGTTATGAC F S V F D L D R L Y K R G V D A I R Y D

2821 TACTTTGCGAGCAATGATTTACTGACCCAAACCGTGGTAACAGGTAACCAAAATGATAAG Y F A S N D L L T Q T V V T G N Q N D K

2881 CTAAATCTGTTACAGGTGGTTAAAGGCACTTTGGAGCAAGGTGTGTGGCTGTTAACCGCA L N L L Q V V K G T L E Q G V W L L T A

2941 GCAATTCGTGAAGAGCAGCGTTACCTTTCGGCCAGTTTACAGCAGTATGTCAGCTGGCTT A I R E E Q R Y L S A S L Q Q Y V S W L

3001 GATGCCGATGTATATGGTGAGGTGCTCAGTGCTCAGTTACGGGGTACTGAGCGTCTGTTA D A D V Y G E V L S A Q L R G T E R L L

3061 GCCCATCAAAATAAGCAGCAAAGATACCTCGTGGGGCTCAGCACTAAAGTTGGCAAAAAT A H Q N K Q Q R Y L V G L S T K V G K N

3121 GGCACTTTCACTTTGGAGTGGCAACAACAATCTTTAGCTGCAAGCACTACCTCAGACTAC G T F T L E W Q Q Q S L A A S T T S D Y

3181 GATTTTTGGCAGCAGCAACAAGCCCGCAGCCAAATCATGCCAATGGGAGTGGCAAAGCGC D F W Q Q Q Q A R S Q I M P M G V A K R

3241 TATCTCGCTGTGTCATACAGTGATGAATGGCGTGATCTAACCCGCTACGAAATCTTGGCA Y L A V S Y S D E W R D L T R Y E I L A

151

3301 ATGCAAAACTTAATCGACAATCATCCCACGTTGTCATTGGCTGTAACTCATTCTGCTAGT M Q N L I D N H P T L S L A V T H S A S

3361 GACAACTTGCAGTTAAGGTTGGCCTTGGCGTGGACACCAGAACAAGGAGCGCGTGATAGC D N L Q L R L A L A W T P E Q G A R D S

3421 GAATTTGAGCAATTTGCCAACGTTCTGCAGCTCGGTTTTCGCTATTACTTCT AGGTAGA A E F E Q F A N V L Q L G F R Y Y F * (RBS)

3481 TCATGCAACTGACTACATTAAAAAAACTCATCTTCGACAAAAATATTTTCGCTTTTTTGA M Q L T T L K K L I F D K N I F A F L K

3541 AGCTTGGCAAACATGTGGATACCATGTATCGCACTAGTTTTGTGACCGCCGCAAACTCCA L G K H V D T M Y R T S F V T A A N S S

3601 GTGGAGTATTAGATTTTTTAAATCAAGGTGGCAAAACCTTAGAGCAATTACAACAATTAC G V L D F L N Q G G K T L E Q L Q Q L L

3661 TGACCGTAGACGAAAGCAAAAAAGGGGCGCTCAGCGCGTGGCTAAATTGCGGTGTAACCC T V D E S K K G A L S A W L N C G V T L

3721 TAAAAGAGTTGAGTTTGCGTGATGGTAAGTACCAATTAAAAGGCAAACTTAGCAAGCACT K E L S L R D G K Y Q L K G K L S K H L

3781 TAGCTAAAGATGACAATCAAATTGCCGCCGCTCTTTTTGAAGAAGCAATCCGTTACCACT A K D D N Q I A A A L F E E A I R Y H Y

3841 ATGACGCCTTACTGAGTGCACCTCAGCGTTTTTGTGGCGGGCAAGCTTATCAGCTAGCAG D A L L S A P Q R F C G G Q A Y Q L A D

3901 ATCAAGATGGTCG Q D G

Figure 4.9: Nucleotide sequence of the genomic-DNA region surrounding the transposon

within the dark purple 3, dark purple 5 and light purple 3 mutant genomes. The nucleotide

sequence is shown along with the translated amino acid sequence in one-letter code. The

inverted solid triangle (t) indicates where mini-Tn10 transposon insertions occurred.

Specific open reading frames (ORF) as indicated in the text are highlighted as follows: ORF 1

(dppA), ORF 2 (dppB), ORF 3 (dppC) and ORF 4 (dppD) are shown in red, blue, green and

purple, respectively. Potential promoter regions are underlined as are predicted ribosome

binding sites (RBS).

152

1 50DppB misltkinki fsdkhqsfhc lkdidltidk geftviagpsgsgkstllniAJ223978 mltlnnisks yklgkeevpi lkhinltvqa geflaimgpsgsgkstlmniAAB97961 mlklknihks yqqgsqefpi lkgidlhvke gdflaimgpsgsgkstlmni • • • • • • •• •••• ••• • ••• •••••••••

51 100DppB iglldkatsg tylfddldvs tmtnnaladi rrekigfvfq aynlmpvltaAJ223978 igcldrptsg tytldqidil kgkdgalaei rnesigfvfq tfhllprltaAAB97961 igcldkasag syhiegtdvs dlsdnqlsdl rnqkigfvfq nfnlmpklta •• ••••••• •• • ••• •••••• • •••••••• •••• •••

101 150DppB lentemimef cgldkklrrq rametltsvg ladlkdrfpaqlsggqqqrvAJ223978 lqnvelpmiy nkvkkkerrq rayealekvg lkdrvsykppklsggqkqrvAAB97961 cqnvelplty mkvpkkerre ralemlrlvg leersdfkpmelsggqkqrv • • • • •• ••• •• • • •• • • • ••••••••

151 200DppB avaraiaaqp llvvadepta nldshsaenl lnlmvklnhd lgitflfsshAJ223978 aiaralvnqp rfiladeptg aldtksseqi lalfselhre .gktiimithAAB97961 aiaralvtnp sfilgdeptg aldtktsvqi melfkqfneq .gktiviith • ••• •• ••••• •• • • • • •• • • •

201 234DppB dqrviqraqr vlqlqdgqiv sderkdqqpk lvalAJ223978 dpdvakkadr tvfirdgelv ldergdisha ....AAB97961 epevaqlckq tvvlrdgnie tralg..... .... • • • • • •• •• ••• •

Figure 4.10: Multiple sequence alignment of the deduced amino acid sequence of P. tunicata

DppB with Bacillus subtilis putative ABC transporter YvrO (AJ223978) and Streptococcus

cristatus ATP-binding cassette protein (AAB97961). The conserved ATP/GTP-binding site

motif A is indicated in blue and the ABC transporter family signature motif is highlighted in

red. Residues identical between P. tunicata DppB and one other protein are indicated by

black dots (•); residues identical in all three proteins are indicated with blue dots (•). Small

dots (.) denote gaps. Numbers given in parentheses are GenBank accession numbers.

153

Figure 4.11: Hydropathy profile of the inferred amino acid sequence from DppA. At least

four predominate hydrophobic regions are evident between amino acids 17 and 39; 273 and

295; 314 and 336; 382 and 404 which represent potential membrane spanning regions of the

protein.

154

4.3.3.1.3. Sequence analysis of the white mutants

Two transposon mutants showing a white phenotype (W2 and W3) were analysed and the

genomic-DNA region flanking the transposon was sequenced for both. Sequence analysis

indicated that the W2 mutant had been disrupted in a gene (wmpR) encoding for a putative

transcriptional regulator and in chapter 5 a detailed analysis of this gene and its encoded

protein is discussed.

A total of 3517 bp of DNA surrounding the transposon in the W3 mutant was sequenced

using the primer walking strategy outlined in Figure 4.12. The primary sequence data for W3

is shown in Figure 4.13. The consensus-sequence obtained was submitted to the programs

ORF-finder and BLAST X from NCBI. Analysis of the region flanking the transposon

indicated that a 2181 bp ORF had been disrupted. This ORF was designated wmpD (white

mutant phenotype D). A predicted RBS was located 9 bp from the ATG start codon.

Analysis of the deduced amino acid sequence of WmpD has indicated that the protein has a

molecular weight of 75105.3 Da and a theoretical pI of 5.21.

WmpD shows similarity to a group of General Secretion Pathway Proteins (GSPP) including,

51 % identity and 70 % similarity (over 676 amino acid residues) to the cholera toxin

secretion protein, EpsD from Vibrio cholerae (GenBank accession number P45779); 51 %

identity and 69 % similarity (over 643 amino acid residues) to the ExeD protein from

Aeromonas salmonicida (GenBank accession number P45778); 48 % identity and 66 %

similarity (over 688 amino acid residues) to ExeD from Aeromonas hydrophila (GenBank

accession number P31780); 50 % identical and 66 % similar (over 631 amino acid residues)

to the pullulanase secretion envelope protein, PulD from Klebsiella pneumoniae (GenBank

accession number P15644) and 47 % identity and 64 % similarity (over 634 amino acid

residues) to the pectic enzymes secretion protein, OutD from Erwinia carotovora (P31701).

A multiple sequence alignment of these proteins with the deduced amino acid sequence for

WmpD is given in Figure 4.14. A putative signal sequence was detected (using the

SIGCLEAVE program available through ANGIS) at the N-terminal of WmpD, suggesting

that the protein is exported from the cytoplasm. This is consistent with the fact that GSPPs

with sequence similarity to WmpD also begin with a signal sequence (Figure 4.14), and are

thought to be localised in the outer membrane (Pugsley, 1993).

Directly upstream of wmpD is a 936 bp ORF designated wmpC. A possible RBS

(5'AGCAGCT3') was located 2 bp from the ATG start of wmpC. Further analysis of this

155

region predicts a transcriptional start point and the upstream region contains potential -10 and

-35 sequences as highlighted in Figure 4.13. Analysis of the deduced amino acid sequence of

WmpC shows that the protein has a molecular weight of 34402.2 Da and a theoretical pI of

4.86. Secondary structure prediction (using PredictProtein programs (Rost, 1996) available

through ExPASy) of WmpC suggests that the protein contains a single transmembrane region

located at the N-terminus between amino acids 22-39 (Figure 4.15).

WmpC is similar to a second group of GSPPs, including 36 % identity and 56 % similarity

(over 294 amino acid residues) to EpsC from V. cholerae (GenBank accession number

P45777); 35 % identity and 52 % similarity (over 309 amino acid residues) to ExeC from A.

hydrophila (GenBank accession number X66504) and 30 % identity and 46 % similarity

(over 285 amino acid residues) to OutC from E. carotovora (GenBank accession number

P31699). Figure 4.15 shows the multiple sequence alignment of these proteins with WmpC.

A signature motif for bacterial type II secretion system protein C was detected (using the

MOTIFS program available through ANGIS) providing further evidence that WmpC

functions as a member of the GSP.

The genes encoding for the GSP in other bacteria are transcribed in one operon

(Wandersman, 1996). The proximity of the P. tunicata wmpC and wmpD genes to each other

and their high similarity to known GSPP genes indicates that wmpC and wmpD may also be

co-transcribed. Preliminary DNA-sequence information downstream of the wmpD gene has

indicates that this region contains an additional ORF similar to a third component of the

general secretion pathway in a number of Gram-negative organisms, namely ExeE from A.

hydrophila, OutE from E. carotovora and EspE from V. cholerae.

156

0 1000 2000 3000

Ap2

W3TnD-S4 W3TnD-S7

W3TnD-S6

W3TnD-S3

Tn10D-S1

Tn10C-S1

W3TnC-S2

W3TnC-S7

W3TnD-S2

W3TnC-S3

W3TnC-S5

W3TnD-S5

W3TnC-S4

Ap2

W3pan2

Figure 4.12: Summary of the sequencing strategy to determine the nucleotide sequence

flanking the transposon insert in the P. tunicata white mutant 3 (W3) genome. Arrows

indicate length and direction of sequence primer products. Blue arrows represent transposon

or adaptor specific primers and black arrows represent sequence specific primers. All primers

used are listed in Appendix II. The nucleotide sequence is shown in base pairs along the top

of the diagram. Open reading frames are indicated by the bold coloured lines; red = wmpC

(ORF 1); blue = wmpD (ORF 2).

157

1 AGCCACTATGGTTGAATATTACCTAGCTGAAATTGGCAAAAATGTAGCATAAATTGCGTG

61 GTAAGAACAGTTGCTACATTTATTTACAATCATCGCTGCGCTAGATGATTTATACAACAA

121 ATGGCTGCTTTTACCTGAAATA TTGACA AACAAGGGCAGAATTCACA TTACTG CGCTTGT (-35) (-10) 181 TGAATGAAAAATGAAAGGGTACTATTGGGCTGGATTTCAGCATTTAAATTC AGCAGGT CA (RBS) M

241 TGCAAGTAAAACTAGAACAATTACAAAAATTAATTGCAAAATTACCAGAAAAAAAAATAA Q V K L E Q L Q K L I A K L P E K K I S

301 GTTACGGCCTTTTTATTTTAATTTTGGTTTACCTTGCTTTCTTAGCTGCGCAGATGGTTT Y G L F I L I L V Y L A F L A A Q M V W

361 GGCTGTTAATGCCTGTGCCAAAATCTGATGCGGTTACCTTTCCTTTAAATAGTGTTCGCA L L M P V P K S D A V T F P L N S V R S

421 GCCAAACGTCACATGGTTTTAATAGCCGCACTCTGACTGACCTCAATATGTTTGGTTCAG Q T S H G F N S R T L T D L N M F G S V

481 TATCGCTGGCACCAAAAACTGCGCCAGTTGAAGCACCCAAAGTGATTAATAGCGCACCCG S L A P K T A P V E A P K V I N S A P E

541 AAACACGCTTAAGTATTACTTTAACGGGTGTAGTTGCAATTAATGGTGATGAAACTGCTG T R L S I T L T G V V A I N G D E T A G

601 GTTCGGCAATTATTGAAAGTCAAAATAGCCAAGAAACATATCAAGCTGAAGATGTTATCA S A I I E S Q N S Q E T Y Q A E D V I K

661 AAGGCACTCGGGCACAATTAAAACAAATTTTTTCAGATCGCGTTATTTTGCAAGTTAATG G T R A Q L K Q I F S D R V I L Q V N G

721 GTGGCTTTGAAACCTTAATGCTCGATGGGTTTGAATTTAGTAAAACGTTTAGTGCGGCAA G F E T L M L D G F E F S K T F S A A S

781 GCCCTACTAATGATGACAATCGCGGCAGATTAGTTGCCAATGATCATCATGCAGTGTTGA P T N D D N R G R L V A N D H H A V L K

841 AACCAACGGATGATCCTGAGGTTCAGTCCGATTTAACTGAAACACGTGACGAAATTTTAC P T D D P E V Q S D L T E T R D E I L Q

901 AAGAGCCAGGTAAGTTATTTGAATACATTCAGGTTTCTCCGGAGCGTCAAGATGGTGAGC E P G K L F E Y I Q V S P E R Q D G E L

961 TAGTTGGTTACCGTCTTCGACCTGGCAAAGACCCTGAGTTATTTAATCGAATGGGTTTAC V G Y R L R P G K D P E L F N R M G L Q

1021 AAAACAATGATTTAGCGATTTCAATTAACGGTTATCCGCTAAACGATATGAAACAAGCTA N N D L A I S I N G Y P L N D M K Q A M

158

1081 TGAGTGCAATTAATGAATTGCGCACTGCAACATCAGCAAACATCACCATTGAGCGTGATG S A I N E L R T A T S A N I T I E R D G

1141 GTGAGCAAATTGATGTGCAATTTAGCCTTGAATAATACACATAAGATT CGGAGT TTATTA E Q I D V Q F S L E * (RBS)

1201 GGAATGAATCACTTGCTACACCTAACAAAAATTAAAAAGGGGTTAGCAAAATACGCTACG M N H L L H L T K I K K G L A K Y A T

1261 TTATTGCTTGCGACAGGCCTTTCTTGTTCAGCTATGGCAGTACAGTATTCTGCCAATTTT L L L A T G L S C S A M A V Q Y S A N F

1321 AAAGGTACAGATATTAATGAGTTTATTAATATCGTGGGTCAAAATTTAAATAAAACCATC K G T D I N E F I N I V G Q N L N K T I

1381 ATTATCGATCCTAATGTGCGCGGTAAAATCAATGTCCGCAGTCCAGAGCTTATGGATGAA I I D P N V R G K I N V R S P E L M D E

1441 GAGCTGTATTACCAGTTCTTTTTAAATGTACTAGAGGTATATGGTGTTGCTGTTGTTGAG E L Y Y Q F F L N V L E V Y G V A V V E

1501 ATGGACAATGGTATTTTAAAAGTTAAAAAAAGTTCTGATGCTAAAAAATCAAATGTTCCG M D N G I L K V K K S S D A K K S N V P

1561 GTATTGGGCGATGATTTTGACGTGCAGGGTGACATGCTAGTAACTCGTGTTGTACGAGTG V L G D D F D V Q G D M L V T R V V R V

1621 AAAAATGTCAGTGTGCAAGAGCTTGGGCCAATTATTCGTCAATTTAGCGACCAAAAAGAT K N V S V Q E L G P I I R Q F S D Q K D t 1681 GGGGGTCATGTAACAAATTATAACCCATCAAACGTATTGATGATGACGGGCCATGCGTCT G G H V T N Y N P S N V L M M T G H A S

1741 TCAGTAAATCGTTTGGTTGAAATCATTCGCTTAGTTGACCAAGCGGGCGATCAACAAGTT S V N R L V E I I R L V D Q A G D Q Q V

1801 GATATTGTAAAATTACGATATGCCACCTCAGCTGATGTGGTCTCTGTGGTTGATAACATT D I V K L R Y A T S A D V V S V V D N I

1861 TATAAGCCAGCCTCGGGTAAATCTGATATCCCAGCCTTTTTAATTCCTAAAGTGGTCGCT Y K P A S G K S D I P A F L I P K V V A

1921 GATGAGCGTACCAATAGCGTTATTGTCAGTGGTGAAGCGCAAGCACGTGAGCGTGCAATT D E R T N S V I V S G E A Q A R E R A I

1981 ACCTTAATTAAACGTCTTGATGATGAGTTAGAAACTCAAGGTAATACAAAGGTGTTTTAT T L I K R L D D E L E T Q G N T K V F Y

2041 ATCAATTACGCCAAAGCTGAAGATTTAGTTAAAGTATTGCAAGGGGTCAGCAAAACTATT I N Y A K A E D L V K V L Q G V S K T I

2101 GCTGAAGAACAAAAGCAAGGCGCTAAAACTAGCTCCCGTGGTCGTAATGATATTAGTATT A E E Q K Q G A K T S S R G R N D I S I

159

2161 GAGGCGCATCCTAATTCGAACTCGCTCGTGATTACTGCTCAACCTGACATAATGCGTTCA E A H P N S N S L V I T A Q P D I M R S

2221 TTAGAAGGTGTTATTGCTAAGCTTGATGTGCGCCGTGCTCAAGTGCTGGTCGAAGCGATT L E G V I A K L D V R R A Q V L V E A I

2281 ATTGTTGAAGTGTTTGAAGGTGATGGTGTTAATTTAGGTTTTCAATGGATTAACAAACAA I V E V F E G D G V N L G F Q W I N K Q

2341 GGCGGCATGTTGCAGTTTAACAATGGTACTACCGTACCGGTTGGTAGTTTAGGTGTTGCT G G M L Q F N N G T T V P V G S L G V A

2401 GGTGAATTAGCTCGCGATAAAACAATCAAAAAAACGGTACTTGGAACAAACGAGGGTTCG G E L A R D K T I K K T V L G T N E G S

2461 GCTAATCAATACGAAGAAACTAAAGAAGGTGACTTAACGGCCCTTGCTAGTTTGCTTGGT A N Q Y E E T K E G D L T A L A S L L G

2521 GGCGTAAATGGTTTAGCGCTTGGTTTTGCCCGGGGTGATTGGGGTGCAATTTTACAAGCG G V N G L A L G F A R G D W G A I L Q A

2581 GTTTCAACCGATACAAATTCTAATATCCTAGCAACACCGTCGGTGACGACTATGGATAAC V S T D T N S N I L A T P S V T T M D N

2641 GAAGAAGCTTCGATGATTGTCGGTCAAGAAGTACCCATTATTACCGGTTCGCAAACGGGT E E A S M I V G Q E V P I I T G S Q T G

2701 AATAATAATACCAATCCATTCCAAACAGTTGAACGTCAAGAAGTGGGTATCAAACTAAAA N N N T N P F Q T V E R Q E V G I K L K

2761 GTCACACCACAAATCAACGATGGCAGTGCAGTACAGCTGACAATTGAGCAAGAAGTATCA V T P Q I N D G S A V Q L T I E Q E V S

2821 AGCGTTAGTGGTGCAACAGCGGTTGATATCACCATTAATAAACGTGAAGTCACTACCACA S V S G A T A V D I T I N K R E V T T T

2881 GTGCTGGCAGATGATGGCGCTATGGTAGTACTTGGTGGCTTAATTGATGAAGATGTGCAA V L A D D G A M V V L G G L I D E D V Q

2941 GAAAGTGTCTCTAAAGTGCCACTATTGGGTGACTTACCGATTATTGGTCACTTATTTAAA E S V S K V P L L G D L P I I G H L F K

3001 TCAACTAGCACCAACCGTCGTAAACGTAACTTATTGATTTTTATTCGCCCAACCATTATT S T S T N R R K R N L L I F I R P T I I

3061 CGTGACAGTGCGACTATGAACCAGCTAAGTAATAGCAAATATAATTACATTCGCACTGAG R D S A T M N Q L S N S K Y N Y I R T E

3121 CAGCAAAAACAAAAAGACGACGGTGTTGATTTAATGCCGACAATCGATACACCTATGTTG Q Q K Q K D D G V D L M P T I D T P M L

3181 CCAGCTTGGAACGATGCCTTGGTTTTACCACCAACGTATGAGCAATATTTACACTCGCAA P A W N D A L V L P P T Y E Q Y L H S Q

160

3241 AATGTAAAAGAACAAGAGCAAAAAAATGACTGAACAAGTAGGTCTAATTAAGCGCTTACC N V K E Q E Q K N D *

3301 TTTTGCATTTGCTAAACGTTTTGGCGTATTGCTTTCGCAGCAACCAACTGGCTATACCTT

3361 ATATTGCCATGGACAAATAAACCCCGAAACGTTGCTTGAAGTTCGCCGCGTTGCGGGAGC

3421 TGAATTTATTGTCGAGCCTTTAAGTGATGAAAAATTTGAGTTGTTGCTTGAATCTGTTTA

3481 TCAACGCGATAGCTCTGAAACCCAGCAAATAATGGAAGA

Figure 4.13: Nucleotide sequence of the genomic-DNA region surrounding the transposon

in the white mutant 3 (W3) genome. The nucleotide sequence is shown along with the

translated amino acid sequence in one-letter code. The inverted solid triangle (t) indicates

where the mini-Tn10 transposon insertion occurred. Specific open reading frames (ORF) as

indicated in the text are highlighted as follows: ORF 1 (wmpC) is shown in red, ORF 2

(wmpD) is shown in blue. Potential promoter regions are underlined as are predicted

ribosome binding sites (RBS).

161

1 50 WmpD mnhllhltki kkglakyatl llatglscs. .amavqysan fkgtdinefiVcEspD .......... mkywlkkssw llagsllstp lamanefsas fkgtdiqefiAsExeD .........m inkgkswrla tvaaalmmag sawateysas fknadieefiKpPulD .......mii anvirsfslt llifaallfr paaaeefsas fkgtdiqefiEcOutD .......... ......mlll sgsvllmass lawsaefsas fkgtdiqefi • • • • ••• • •••• ••• •••••• •••

51 100 WmpD nivgqnlnkt iiidpnvrgk invrspelmd eelyyqffln vlevygvavvVcEspD nivgrnlekt iivdpsvrgk vdvrsfdtln eeqyysffls vlevygfaavAsExeD ntvgknlskt iiiepsvrgk invrsydlln eeqyyqffls vldvygfavvKpPulD ntvsknlnkt viidpsvrgt itvrsydmln eeqyyqffls vldvygfaviEcOutD ntvsknlnkt viidpsvsgt itvrsydmmn eeqyyqffls vldvygftvi •••• ••••• ••••• •••• ••••• •• •• •••••• •••••• •••

101 150 WmpD emdngilkvk kssdakksnv pvlgddfdvq gdmlvtrvvr vknvsvqelgVcEspD emdngvlkvi kskdaktsai pvlsgeeran gdevitqvva vknvsvrelsAsExeD pmdngvlkvv rskdaktsai pvvdetnpgi gdemvtrvvp vrnvsvrelaKpPulD nmnngvlkvv rskdaktaav pvasdaapgi gdevvtrvvp ltnvaardlaEcOutD pmdnnvlkii rskdakstsm platdrqpgi gdevvtrvvp vnnvaardfg ••••• ••• •• ••• • • ••• • •• ••••• •••••• •••

151 200 WmpD piirqfsdqk dgghvtnynp snvlmmtgha ssvnrlveii rlvdqagdqqVcEspD pllpqlidna gagnvvhydp aniilitgra avvnrlaeii rrvdqagdkeAsExeD pllrqlndna gggnvvhydp snvllitgra avvnrlvevv rrvdkagdqeKpPulD pllrqlndna gvgsvvhyep snvllmtgra avikrlltiv ervdnagdrsEcOutD rssrelndna wrgtcgdyep anvvvmtgra gvihavmtiv ervdqtgdrn • •• • •• • • • •••• ••• • •••••••• • •••••••

201 250 WmpD vdivklryat sadvvsvvdn iykpasgksd ipafli.pkv vadertnsviVcEspD ievvelnnas aaemvrivea lnkttdaq.n tpeflk.pkf vadertnsilAsExeD vdiiklryas agemvrlvtn lnkdgntqgg ntslllapkv vadertnsvvKpPulD vvtvplswas aadvvklvte lnkdtsksal pgsmv..anv vadertnavlEcOutD vttiplsyas stevvkmvne lnkmdeksal pgmlt..anv vadertnsaa ••••••••• ••••• • • • • • •• ••• •••••••••

251 300 WmpD vsgeaqarer aitlikrldd eletqgntkv fyinyakaed lvkvlqgvskVcEspD isgdpkvrer lkrlikqldv emaakgnnrv vylkyakaed lvevlkgvseAsExeD vsgepkarar iiqmvrqldr dlqsqgntrv fylkygkakd mvevlkgvstKpPulD vsgepnsrqr iiamikqldr qqatqgntkv iylkyakasd lvevltgissEcOutD gfgepnsrqr vidmvkqldr qqavqgntkv iylkyakaad lvevltgvgd •••• •••• • ••• •• •• ••••••• •• •••••• •• •• •••

162

301 350 WmpD tiaeeqkqga ktssrgr... ..ndisieah pnsnslvita qpdimrslegVcEspD nlqaekgtgq pttsk.r... ..nevmiaah adtnslvlta pqdimnamleAsExeD sieadkkggg ttagggnasi gggklaisad ettnalvita qpdvmaeleqKpPulD tmqsekqaak pvaaldk... ...niiikah gqtnalivta apdvmndlerEcOutD siqtdqqna. .lpalrk... ...disikah eqtnslivna apdimrdleq •• ••• • • • •• •••• •• ••••••• •••••• ••

351 400 WmpD viakldvrra qvlveaiive vfegdgvnlg fqwinkqggm lqfnngttvpVcEspD vigqldirra qvliealive maegdginlg vqwgslesgs viqygntgasAsExeD vvakldirra qvlveaiive iadgdglnlg vqwantnggg tqftdtnlpiKpPulD viaqldirrp qvlveaiiae vqdadglnlg iqwanknagm tqftnsglpiEcOutD viaqldirrp qvlveaiiae vqdadgmnlg vqwanknagv tqftntglpi •••••• ••• •••••••••• • •••• ••• •• •• ••• •• • •

401 450 WmpD vgslgvagel ardktikktv lgtnegsanq yeetkegdlt alasllggvnVcEspD ignvmiglee akdttqtkav ydtnnnflrn ettttkgdyt klasalssiqAsExeD gsvaiaakdy nengt.t... .......... .........t gladlakgfnKpPulD staiaganqy nkdgtvs... .......... .........s slasalssfnEcOutD ttmmagadqf rrdgtlg... .......... .........t aattalggfn • • • ••• • • • •• •• • •••••••• •

451 500 WmpD glalgfargd wgailqavst dtnsnilatp svttmdneea smivgqevpiVcEspD gaavsiamgd wtalinavsn dsssnilssp sitvmdngea sfivgeevpvAsExeD gmaagfyhgn waalvtalst stksdilstp sivtmdnkea sfnvgqevpvKpPulD giaagfyqgn wamlltalss stkndilatp sivtldnmea tfnvgqevpvEcOutD giaagfyqgn wgmlmtalss nskndilatp sivtldnmea tfnvgqevpv • • ••• •• ••• • •••• •• ••••••• • ••••• •• • •••••••

501 550 WmpD itgsqtgnnn tnpfqtverq evgiklkvtp qindgsavql tieqevssvsVcEspD itgstagsnn dnpfqtvdrk evgiklkvvp qinegnsvql nieqevsnvlAsExeD qsgsqsstts dqvfntierk tvgtkltvtp qinegdsvll nieqevssvaKpPulD ltgsq.ttsg dnifntverk tvgiklkvkp qinegdsvll eieqevssvaEcOutD lagsq.ttsg dnvfqtverk tvgiklkvkp qinegdsvll eieqevssva ••••• • •• •••••••• •••••••••• ••• • ••• ••••••••

551 600 WmpD ga.....tav ditinkrevt ttvladdgam vvlgglided vqesvskvplVcEspD ga....ngav dvrfakrqln tsvmvqdgqm lvlgglider aleseskvplAsExeD qkqatgtadl gptfdtrtik navlvksget vvlgglmdeq tqekvskvplKpPulD daasstssdl gatfntrtvn navlvgsget vvvgglldks vsdtadkvplEcOutD daasssstnl gatfntrtvn navlvssgdt vvvgglldks tnesankvpl •• ••• • • ••• • • •• •• • ••••••••• ••••••••••

163

601 650 WmpD lgdlpiighl fkststnrrk rnllifirpt iirdsatmnq lsnskynyirVcEspD lgdipllgql frstssqvek knlmvfikpt iirdgvtadg itqrkynyirAsExeD lgdipvlgyl frstnnttsk rnlmvfirpt ilrdahvysg issnkytmfrKpPulD lgdipvigal frstskkvsk rnlmlfirpt virdrdeyrq assgqytafnEcOutD lgdipvlgyl frsnstetkk rnlmlfirps iirdrsqfqs asaskyhsfs ••• • •• • • •••• • ••• ••••• •••• • • • ••••••

651 700 WmpD teqqkqkddg vdlmptidtp mlpawndalv lpptyeqylh sqnvkeqeqkVcEspD aeqlfraekg lrllddasvp vlpkfgddrr hspeiqafie qmeakq....AsExeD aeqldaaaqe syltspkrqv lpeygqdvaq spevqkqiel mkarqqatadKpPulD daqskqrgke nnd.amlnqd lleiyprqdt aafrqvsaai dafnlggnl.EcOutD aeenkqrnvs ngegglldnd llrlpeggna ytfrqvqssi vafypaggk. •• •• • • • • •• • •• • •

701 WmpD nd........VcEspD ..........AsExeD gaqpfvqgnkKpPulD ..........EcOutD ..........

Figure 4.14: Multiple sequence alignment of the deduced amino acid sequence of P. tunicata

WmpD with the general secretion pathway protein, EpsD from Vibrio cholerae (VcEspD,

GenBank accession number P45779); ExeD from Aeromonas salmonicida (AsExeD,

GenBank accession number P45778); the PulD protein from Klebsiella pneumoniae

(KpPulD, GenBank accession number P15644) and the OutD protein from Erwinia

carotovora (EcOutD, GenBank accession number P31701). Putative N-terminal signal

sequences are highlighted in blue. Residues identical between P. tunicata WmpD and one

other protein are indicated by black dots (•); residues identical in two other proteins are

indicated with blue dots (•); residues which are identical between three proteins are indicated

by green dots (•) and residues which are identical in all 5 proteins are indicted by red dots (•).

Small dots (.) denote gaps.

164

1 50 WmpC .......mqv kleqlqklia klpekkisyg lfililvyla flaaqmvwllAhExeC ..mtlpfrnd llssllarck tvplsrfsqp lfwlllllla hqcagltwrlVcEpsC mefkqlppla awprllsqnt lrwqkpiseg ltllllvasa wtlakmvwvvAsExeC ..mtlpfrdd llssllarck tvplsrfsqp lfwlllllla hqcagltwrlEcOutC . .marlqafk dpsfhslvat frslplirrf vlglilllic qqlavltwrf • • • • •• • •• •••• •• • ••• •

51 100 WmpC mpvpksdavt fplnsvrsqt shgfnsrtlt dlnmfgsvsl apktapveapAhExeC ldlgsqqasq pwqpamvasq gqgsarldls gisrlslfg. kakqqaqaadVcEpsC saeqtpvptw sptlsglkae rqpldisvlq kgelfgvft. epkeapvveqAsExeC ldlgsqqssq pwqpavagsq gqgkarldlg gvsrlalfg. kakqqaraadEcOutC llpedsrivg vsvtpaqake kpatp....g dftlfghap. dadastvnda •• • •• • • • •• •• ••• •

101 150 WmpC kvinsapetr lsitltgvva ingdetagsa iiesqnsqet yqaedvikgtAhExeC avaadapktq lnaqlngvla .ssdpaksia iiahngvqns ygigdfidgtVcEpsC pvvvdapktr lslvlsgvva .sndaqksla vianrgvqat ygineviegtAsExeC avaadapktq lnaqlngvla .ssdpaksia iiahsgvqns ygigdfidgtEcOutC alsgdiplts lnisltgvla .sgdakrsia iiakdsqqys rnvgdaipgy • •• •• ••• • •••• •• • •• • • • ••• ••

151 200 WmpC raqlkqifsd rvilqvnggf etlmldgfef sktfsaaspt nddnrgrlvaAhExeC qakirqvfad rviierdgrd etlmldgeey gkplpkpgnq ..........VcEpsC qaklkavmpd rviisnsgrd etlmlegldy tapatasvsn pprprpn...AsExeC qakvrqvfad rviierdard etlmldgeey gkplpkqgnp ..........EcOutC eakivtisad rvvlqyqgry ealhlyqeee atgapsssga .......... • ••••• • ••••• • ••••••• • • • • •

201 250 WmpC ndhhavlkpt ddpevqsdlt etrdeilqep gklfeyiqvs perqdgelvgAhExeC .......... .....ddkls svrsellgnp gkitdylnis pvrvdgrmvgVcEpsC .......qpn avpqfedkvd aireaiarnp qeifqyvrls qvkrddkvlgAsExeC .......... .....dekls svrsellgnp gkitdylnis pvrvdgrmvgEcOutC .......... ........fn qvkdeiqkdp fsaqdyltis pvteeevlkg • ••••• • •• • • • • • •• ••

251 300 WmpC yrlrpgkdpe lfnrmglqnn dlaisingyp lndmkqamsa inelrtatsaAhExeC yrlnpgsnpe lfnqlglvan dmavsingld lrdnaqamqa mqqvagatemVcEpsC yrvspgkdpv lfesiglqdg dmavalngld ltdpnvmntl fqsmnemtemAsExeC yrlnpgsnpe lfnqlglvan dmavsingld lrdnaqamqa mqqvagatemEcOutC yqlnpgknpd lfyraglqdn dlavslngmd lrdadqaqqa maqlagmskf ••• •••••• •••• ••• • ••• •••• • • ••• • • ••

165

301 321 WmpC nitierdgeq idvqfsle.. .AhExeC tvtverqgql ydvyvglse. .VcEpsC sltverdgqq hdvyiqf... .AsExeC tvtverqgql ydvyvglse. .EcOutC nltverdgqq qdiylaldgd h • • •••• • •• • •

Figure 4.15: Multiple sequence alignment of the deduced amino acid sequence of P. tunicata

WmpC with ExeC from Aeromonas hydrophila (AhExeC, GenBank accession number

P45790); EpsC from Vibrio cholerae (VcEpsC, Genbank accession number P45777); ExeC

from A. salmonicida (AsExeC, GenBank accession number P45772) and OutC from Erwinia

carotovora (EcOutC, GenBank accession number P31699). Potential transmembrane region

is highlighted in blue. The bacterial type II secretion system protein C signature motif is

indicated in red. Residues identical between P. tunicata WmpC and one other protein are

indicated by black dots (•); residues identical in two other proteins are indicated with blue

dots (•); residues which are identical between three proteins are indicated by green dots (•)

and residues which are identical in all five proteins are indicted by red dots (•). Small dots (.)

denote gaps.

166

4.3.4. Assessment of secreted protein profiles of wild-type and white mutant

3 (W3) strains of P. tunicata

Genotypic characterisation of the W3 transposon mutant suggested a role for the general

secretion pathway in the expression of pigment and antifouling compounds. The general

secretory pathway is required by a number of Gram-negative bacteria for secretion of a variety

of structurally unrelated extracellular enzymes and toxins (Pugsley, 1993). To further assess

the function of the putative secretion proteins WmpD and WmpC, the extracellular proteins of

both wild-type P. tunicata and the W3 mutant were analysed by SDS-PAGE and silver

staining (Figure 4.16).

It was discovered that the protein profiles of W3 differed from wild-type P. tunicata

supernatant profiles. As highlighted in Figure 4.16, four of the major protein bands in both

the early-stationary and late-stationary phase wild-type samples were absent from both early-

stationary and late-stationary phase mutant samples. In addition, an approximately 60 kDa

band present in both samples appears to be over expressed in the mutant late-stationary phase

sample.

167

Figure 4.16: : Silver stained SDS-PAGE gel showing supernatant proteins from the wild-

type (Wt) and the white mutant 3 (W3) strains of P. tunicata during different growth phases.

Lane 1: Broad-range molecular weight marker; Lane 2: Wt early stationary; Lane 3: W3 early

stationary; Lane 4: Wt late stationary; Lane 5: W3 late stationary. The major proteins missing

in W3 but present in Wt are indicated by red arrow. The band indicated by the blue arrow is

over expressed in W3 during late stationary phase of growth.

.

168

4.4. Discussion

Previous observations have indicated that pigmentation is possibly linked to the expression of

biologically active secondary metabolites in P. tunicata (Chapter 3; Holmström et al.,

unpubl.). In this chapter transposon mutagenesis was used as a tool to further study the

relationship between pigmentation and the production of fouling inhibitors by P. tunicata.

Using the mini-Tn10 system as described in chapter 3, transposon mutants altered in normal

pigmentation were generated (Figure 4.1). Wild-type P. tunicata produces a yellow and a

purple pigment, which when combined give the bacterial colony a dark green appearance.

Four different categories of pigmentation mutants were isolated including, yellow, dark purple,

light purple and white phenotypes. The UV/Visible light scans (Figure 4.2) showed that the

yellow phenotype is due to the loss of purple pigmentation. Similarly, the purple phenotypes

are due to the loss of yellow pigmentation and white mutants lack both pigments. Analysis of

the antifouling properties of the four different pigmentation phenotypes revealed that the

purple and white mutants differed from the wild-type and yellow mutants in their ability to

inhibit each of the target organisms. Yellow mutants retained full activity toward invertebrate

larvae (Table 4.1), algal spores (Table 4.2), bacteria (Table 4.3) and fungal growth (Figure

4.3). This suggests that the purple pigment is unlikely to be involved in the antifouling

activity of P. tunicata. In contrast, the white and light purple had lost some or all of their

ability to inhibit each of the target fouling organisms, and the dark purple mutants had lost

their ability to inhibit fungal growth, larval settlement and algal spore germination. These

observations indicate that the yellow pigment is important, either directly or indirectly, for

antifouling activity in P. tunicata. Quantitative analysis for the presence of the anti-bacterial

protein in each of these mutant strains has not yet been performed. However, preliminary

observations using SDS-PAGE indicate that protein bands corresponding to both subunits

(60 kDa and 80 kDa) of the active anti-bacterial protein are present but vary in intensity for

each of the mutant samples (data not shown). These observations appear to correspond well

with the levels of activity observed for each of the mutants (Table 4.3).

The data describing the antifouling properties of the transposon mutants suggest that the

production of fouling inhibitors is linked to the synthesis of yellow pigment or that fouling

inhibitors and pigment are jointly regulated in P. tunicata. Therefore, genes disrupted to

cause a change in pigmentation will also provide information regarding the identity and/ or

regulation of antifouling components in this organism. The panhandle-PCR method

described in section 3.2.4.6 was used to sequence the regions of genomic DNA where the

transposon had inserted. Initial sequence analysis revealed that of the eight mutants with a

169

light purple phenotype, the transposon had inserted into only two different sites. The

possibility of mini-Tn10 having hot-spots was discussed in chapter 3. Sequence information

resulting from the study of the pigmented mutants gives further evidence for the existence of

hotspots for mini-Tn10 insertion in the P. tunicata genome.

Sequence information obtained from the DNA surrounding the transposon in light purple

mutant 2 indicates that a protein with a putative oxidase function has been disrupted. The

ORF designated lppA shows sequence similarity to both known and putative oxidase enzymes

such as 3-chlorobenzoate-3, 4-dioxygenase (CbaB) (Kaneko et al., 1996), toluenesulfonate

methyl-monooxygenase (TsaM) (Junker et al., 1997) and aminopyrrolnitrin oxidase (PrnD)

(Hammer et al., 1997). Oxygenases and oxidases are enzymes that catalyse the oxidation of a

substrate by molecular oxygen (O2). Oxygenases incorporate oxygen molecules from O2 into

the substrate, whereas oxidases do not incorporate oxygen into the substrate (Mathews and

van Holde, 1990). Interestingly, PrnD is involved in the synthesis of pyrrolnitrin, an anti-

fungal compound derived from tryptophan and produced by strains of Pseudomonas

fluorescens (Vincent et al., 1991). An important feature of these oxygenases and oxidases is

that they each contain a conserved Rieske iron-sulfur cluster (Figure 4.7). Iron-sulfur clusters

play a critical role in a wide range of oxidation/ reduction reactions in biological systems and

proteins containing these clusters are known as iron-sulfur proteins (Mathews and van Holde,

1990). The possibility that LppA is an iron-sulfur protein provides further evidence for its

role as either an oxidase or oxygenase. Downstream of lppA (and most likely in the same

operon), is a second ORF (lppB) with similarity to enzymes having transferase activity (i.e.

catalyses the transfer of specific molecular groups from one molecule to another). Including

8-amino-7-oxonanoate synthase (BioF) which is involved in the synthesis of biotin (Bower et

al., 1996).

The genomic DNA flanking the transposon in the dark purple mutants (DP 3 and DP 5) was

found to align with the sequence obtained from the second light purple mutant (LP 3).

Together this region of DNA encodes four ORFs (dppA, dppB, dppC, and dppD), which are

potentially co-transcribed. DppA was found to have homology to integral membrane proteins.

A secondary structure prediction of the deduced amino acid sequence also supported the

membrane location of this protein.

Downstream of dppA is dppB. This ORF encodes a protein with high homology to ABC

transporter proteins. This family of proteins derives its name due to the existence of a

conserved ATP-binding cassette (ABC). ABC-transporters use the energy derived from the

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hydrolysis of bound ATP to transport (import or export) substrates across the cell membrane.

While a range of organisms use ABC-transporters for a variety of substrates, individual ABC

transporters are specific for a given substrate in a given direction (i.e. import or export).

Eukaryotic ABC-proteins include members of the multidrug resistance transporter (MDR)

which can pump hydrophobic drugs out of the cell. In Gram-positive bacteria they are

involved in drug efflux and lantibiotic secretion. The ABC transporters of Gram-negative

bacteria are implicated in the uptake of various substrates such as maltose, histidine,

oligopeptides and iron siderophore complexes. In addition, ABC transporters are involved in

both protein secretion and the export of non-protein molecules such as antibiotics and

carbohydrates (Fath and Kolter, 1993; Higgins, 1992; Reizer et al., 1992; Wandersman,

1996).

Typically ABC-transporters require the function of multiple protein domains, including the

ATP-binding domain and a highly hydrophobic domain consisting of several membrane

spanning segments. The membrane spanning domains are believed to form a pathway

through which the substrate crosses the membrane and to be responsible for the substrate

specificity of the transporter. The individual domains of an ABC transporter may be

expressed within a single protein or as separate polypeptides (Higgins, 1992). Analysis of the

deduced amino acid sequence from DppB indicates that this protein does not contain a

transmembrane domain, suggesting the involvement of an additional polypeptide/s. As

mentioned above, DppA (encoded directly upstream of DppB) is a hypothetical integral

membrane protein consisting of four putative transmembrane segments. Therefore it is likely

that DppA functions together with DppB as part of a putative ABC-transporter apparatus.

The deduced gene product of dppC, a 45 kDa protein, was not found to share similarity with

proteins currently available in the Swissprot or EMBL databases. Initial sequence analysis of

a region further downstream of DppC indicates the presence of a fourth ORF (DppD) in the

putative operon structure. This gene has similarity to a predicted methyl-transferase, which is

likely to be involved in the transfer of a methyl group from a donor molecule to a recipient

molecule.

The similarity of the ORFs identified from the study of the purple transposon mutants of P.

tunicata to enzymes with oxidase and transferase activity suggests that these genes are

involved in the synthesis of the yellow pigment. In addition, the putative operon consisting of

DppA through to DppD, which includes the ABC protein, resembles that of other ABC

transporters where the genes for transport are linked to structural genes of the exported

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molecule (Wandersman, 1996). As such the putative ABC transporter may act as an exporter

of the yellow pigment. Alternatively, it may be important for the uptake of a precursor to the

synthesis of the yellow pigment. Sequence analysis of the regions downstream of DppD is

likely to reveal other enzymes involved in the pathway for yellow pigment production in this

organism. As new enzymes are identified it will be important to confirm their function. This

may be possible using site directed mutagenesis to disrupt specific ORFs and can be followed

by complementation of the gene expressed on a plasmid vector to restore the function.

Molecular biology tools are currently being developed in our laboratory, which will enable

such experiments to be performed on P. tunicata.

The current working model being proposed in relation to the production of antifouling

compounds and pigmentation is outlined in the figure below (Figure 4.17). Precursors of the

pigment could be directly involved in the inhibitory effects or it may be that they lead to

additional side branching pathways, each one resulting in a different antifouling component.

The anti-bacterial protein is unlikely to be a precursor of the yellow pigment. Therefore, the

reduced levels of anti-bacterial activity in the light purple mutants may be due to the pigment

and the anti-bacterial protein being jointly regulated (see chapter 8).

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yellowpigment

Light purple mutants-lost yellow pigment + inhibitors

Dark purple mutants-lost yellow pigment + inhibitors

A B C ED

anti-larval anti-algal anti-fungal

Figure 4.17: Hypothetical model relating antifouling activity and pigment production in P.

tunicata.

White mutants that lack the expression of both the yellow and purple pigments were also

studied. It was discovered that one of these mutants, W3, had been disrupted in a gene

encoding for a General Secretion Pathway Protein (GSPP). Extracellular proteins secreted by

the General Secretion Pathway (GSP) contain an N-terminal signal sequence (or signal

peptide) which allow them to cross the inner membrane via the sec-dependent pathway

(Pugsley, 1993). Crossing of the outer membrane can occur via several ways, the most

common one is the GSP and requires a number of specific helper-proteins that span the cell

envelope; these are commonly known as the GSPPs (Wandersman, 1996). This secretory

apparatus (also known as the type II secretion system) has been identified in a number of

Gram-negative bacteria including Vibrio cholerae (Overbye et al., 1993), Xanthomonas

campestris (Dums et al., 1991), Erwinia carotovora (Reeves et al., 1993), Klebsiella oxytoca

(Pugsley, 1993) and Pseudomonas aeruginosa (Bally et al., 1992). Most of these bacteria

secrete a number of structurally diverse polypeptides yet possess only one secretion

machinery which is common for all. The genes encoding for the GSPPs are clustered and

organised in one or two operons. It would appear that a similar structural organisation exists

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for P. tunicata as analysis of surrounding ORFs revealed other components of the secretory

apparatus including GSPP C and GSPP E. DNA-sequence analysis of the GSPP from a

diverse number of bacteria has revealed that they have homologous proteins with sequence

identities ranging from 30 to 60 % (Pugsley, 1993). Despite their sequence homology the

secretion machinery from one bacterial species is usually unable to secrete an exopolypeptide

from another closely related species (Wandersman, 1996).

The genotypic and phenotypic characterisation of white mutant 3 would suggest that the gene

products of wmpD and wmpC are involved in the secretion of yellow pigment, purple pigment

and each of the specific antifouling molecules. This direct link is unlikely as the pigments and

antifouling molecules are extremely different and would require different transport systems.

In addition, the GSP is specific for proteins or peptides which is not the case of a number of

these molecules (eg. the anti-fungal and anti-larval components). It is possible that the GSP

described here is required to secrete extracellular enzymes or surface-structures needed by the

bacterium to sense environmental cues or to obtain specific metabolites required as precursors

for pigment/ antifouling production. In this case analysis of the extracellular proteins

differing between W3 mutant and wild-type strains will provide information regarding the

requirements for the production of fouling inhibitors by P. tunicata. Supernatant samples

were collected and the differences in the proteins secreted during two different growth phases

were assessed. Production of fouling inhibitors and pigmentation begins during early-

stationary and continues into late-stationary phase of growth, at this time several differences

can be seen between the proteins released by the wild-type and W3 mutant. These include the

loss or reduced expression of a number of proteins and the accumulation of a major protein as

indicated in Figure 4.16. Future studies will aim to identify these proteins using methods

such as protein mass-spectrometry finger-printing together with N-terminal amino acid

sequencing.

In conclusion, this chapter has investigated the relationship between pigmentation and the

synthesis of fouling inhibitors in the marine bacterium P. tunicata. Using transposon

mutagenesis genes involved in the synthesis of the yellow pigment have been identified and

these are also likely to play a role in the production of the other antifouling molecules. Given

that the same or related pathways are likely to be involved in producing pigment and inhibitors

it seems feasible that their expression may also be coordinated. In this regard, phenotypic

characterisation of a second white mutant (W2) was carried out in this chapter and initial

sequence analysis revealed that it had been disrupted in a gene with a potential transcriptional

regulatory role. A detailed analysis of this mutant is presented in the following chapter.

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5. Identification and characterisation of a putative transcriptional

regulator controlling the expression of extracellular inhibitors

in Pseudoalteromonas tunicata

5.1. Introduction

In order to optimise their ability to survive and grow bacteria must be able to sense and

respond to changes in their immediate environment. This is especially important for the

expression of non-essential phenotypes such as colonisation traits, virulence factors and

secondary metabolites including antimicrobials and toxins. The study of bacterial gene

regulation is of interest to both general bacterial physiology and for the purpose of application

of novel bacterial metabolites. The regulatory systems of pathogens and pests may be

potential targets for novel biocontrol agents. Conversely, understanding how bacteria such as

P. tunicata regulate expression of genes involved in the synthesis of biologically active

metabolites is likely to be essential for the identification of environmental conditions that lead

to an improved production of these metabolites.

Transposon mutagenesis was used in chapter 4 to study the correlation between pigment

production and antifouling activity in P. tunicata. A variety of mutants altered in the wild-type

pigmentation were isolated and characterised. The study of the transposon mutants lead to the

identification of several genes potentially involved in the syntheses of pigment and specific

fouling inhibitors. One of these mutants displayed a white phenotype and was found to have

lost the wild-types ability to inhibit each of the target organisms. Preliminary sequence data

indicated that this mutant, designated W2 might have been disrupted in a gene involved in the

regulation of pigment production and the expression of fouling inhibitors.

This chapter demonstrates that the putative regulatory protein (WmpR) is most similar to

transcriptional activators CadC from Escherichia coli and ToxR from Vibrio cholerae. Both

CadC and ToxR are required to sense the environment and respond by increasing

transcription of specific genes. For example, ToxR regulates the expression of colonisation

and virulence traits in response to changes in pH and temperature, which signal that the

bacterium is within the host (DiRita et al., 1991). This chapter also presents a detailed

175

analysis of wmpR and molecular evidence is provided to suggest that WmpR functions as a

regulator of antifouling activity and pigmentation in the marine bacterium P. tunicata.

Through the use of the putative WmpR protein P. tunicata may be able to sense

environmental signals and respond to them by increasing the expression of fouling inhibitors.

This may be an important means by which P. tunicata cells out-compete other surface-

associated organism in a biofouling community and will be discussed in this chapter and the

remaining chapters of this thesis.

5.2. Materials and Methods

5.2.1. DNA sequencing and analysis

The DNA region flanking the transposon within the W2 mutant was amplified using the

panhandle-PCR method as described in section 3.2.4.2. DNA sequencing and data analysis

of this region was performed as outlined in sections 3.2.4.3 and 3.2.4.4, respectively.

5.2.2. Two-dimensional gel electrophoresis (2DGE)

5.2.2.1. Sample preparation

Total cellular-protein samples were prepared from early-logarithmic and early-stationary

growth phase cells of both the wild-type strain and the W2 mutant strain of P. tunicata. Two

hundred microlitres of an overnight culture were inoculated into 50 ml of VNSS medium for

the wild-type strain and 50 ml of VNSS with the addition of the antibiotics Km (85 µg/ml)

and Sm (200 µg/ml) for the W2 mutant strain. Growth was monitored by optical density

(OD) at 610 nm and 5 ml samples removed at an OD of 0.3 for the early-logarithmic sample

and 0.6 for early-stationary phase sample. These points represent prior to and the onset of

pigmentation and antifouling activity in wild-type P. tunicata, respectively. Cells were

collected by centrifugation (2000 x g for 10 min) and washed once in 0.2 mM sucrose

solution, followed by a second centrifugation step (2000 x g for 10 min). The cell pellet was

resuspended into 200- 500 µl of sterile milli-Q water and stored in small aliquots at -80 oC.

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5.2.2.2. Sample preparation and isoelectric focusing

Total protein concentration of each preparation was determined using the BCA method as

described in section 4.2.4.2. For each sample 50 µg of total cell protein was resuspended in

450 µl rehydration buffer [8 M urea, 0.1 M dithiothreitol, 40 mM Tris-HCl (pH 8.8), 1.2 %

(v/v) Pharmolytes (pH 4-8, Amersham Pharmacia), 4 % (w/v) CHAPS; (freshly prepared)].

Samples were sonicated on ice three times for 1 min with a Branson Sonifier on a 30 % duty

cycle and a setting of 3.5. To remove nucleic acids, 10 µl of nuclease buffer (1 mg/ml

DNAse, 0.25 mg/ml of RNAse A, 24 mM Tris-Base, 476 mM Tris-HCl, 50 mM MgCl2) was

added to the sample and incubated on ice for 20 min. Samples were centrifuged (21000 x g

for 30 min at 4 oC) and the supernatant loaded onto 18 cm Immobiline DryStrips (pH 4-7,

linear, Amershan Pharmacia). Strips were kept for a minimum of 6 h for rehydration at room

temperature with the gel side facing upwards. Isoelectric focusing (separation in the first

dimension) of the strips was performed using a Multiphor II system (Pharmacia) according to

the manufacturer’s instructions at 15 oC programmed for 0.5 h sequential intervals of 300

Volt, 1 kVolt and 2.5 kVolt, followed by 17 h at 3.5 kVolt, for a total of 61400 Volt-h. After

isoelectric focusing the strips were stored at -80 oC until ready to run the second dimension.

5.2.2.3. Second-dimension electrophoresis

Prior to second dimension electrophoresis, the strips were equilibrated by washing them in

freshly prepared equilibration buffer (6 M urea, 2 % (w/v) SDS, 20 % (v/v) glycerol, 0.375 M

Tris-Base; pH 8.8) supplemented with 2 % (w/v) dithiothreitol for 20 min whilst gently

shaking. After the wash, the solution was discarded and replaced for 10 min with

equilibration buffer supplemented with 2.5 % (w/v) acrylamide. Separation in the second

dimensions was performed with 11.5 % SDS-PAGE gels made with Duracryl (0.8 % bis-

acrylamide; Genomic Solutions) and using a Protean II system (BioRad). Gels were run for

approximately 5 h at a constant current of 45 mA. Following the completion of the run, the

gels were fixed overnight in fixing solution consisting of 40 % (v/v) methanol and 10 % (v/v)

acetic acid.

5.2.2.4. Staining and analysis

All gels were stained using the silver staining protocol outlined in section 4.2.4.4, with the

exception that the times of each wash was extended as follows: water washes for 10 mins,

sodium thiosulphate wash for 2 mins. After a brief wash in milli-Q water, the silver stained

gels were scanned using a BioRad GS-700 Imaging Densitometer. Images were saved as

177

TIFF files and the data analysed using BioRad Melanie II software. Duplicates of each

condition were performed and compared. Only spots that appear in both duplicates of one

condition and not in any of the duplicates of the other condition were considered to be

significant changes.

5.3. Results

5.3.1. DNA sequencing analysis

The genomic DNA flanking the transposon in the white mutant 2 (W2) strain was sequenced

as described in section 4.2.3. A total of 3674 bp of DNA sequence was obtained using the

primer-walking strategy outlined in Figure 5.1. After sequence assembly, the consensus

sequence was submitted to programs ORF-finder and BLAST X. The primary sequence data

for W2 is shown in Figure 5.2.

Analysis of the region flanking the transposon indicated that a 2088 bp ORF encoding for a

putative transcriptional regulator had been disrupted. The ORF was therefore designated

wmpR (White mutant phenotype regulator). Further sequence analysis of this region revealed

a putative RBS (5' AAGAAG 3') located 2 bp upstream of the ATG start codon. Promoter

prediction analysis resulted in the identification of a potential transcriptional start point at

nucleotide position 553. This region contains putative -10 and -35 sequences as highlighted

in Figure 5.2. Following the translational stop of wmpR is a GC-rich inverted repeat, which is

followed by a series of thymidine residues (nucleotides 3394 to 3441). This region may act

as an ρ-independent terminator of transcription (Mathews and van Holde, 1990).

The deduced amino-acid sequence of WmpR was found to be similar in the N- terminus to a

sub-group of the OmpR-like transcriptional activators. These transcriptional regulators have

their DNA-binding domain located at the N-terminus rather than the C-terminus of the

protein. More specifically, WmpR was shown to be 38 % identical and 66 % similar (over 85

amino acid residues) to E. coli CadC transcriptional regulator protein (P23890); 38 %

identical and 64 % similar (over 71 amino acid residues) to the ToxR homologue from Vibrio

parahaemolyticus (Q05938) and 39 % identical and 62 % similar (over 71 amino acid

residues) to the Vibrio cholerae transcriptional activator ToxR (P15795). A multiple

178

sequence alignment of these proteins with the deduced amino acid sequence of WmpR is

given in Figure 5.3.

Further analysis of WmpR shows that the protein has a predicted molecular weight of

79202.9 Da and a theoretical pI of 5.43. The hydrophobicity profile as predicted by the

method of Kyte and Doolittle (1982) and the secondary structure predicted by the SOSUI

method (available through the ExPASy web site) indicated that the protein has a

transmembrane region between the amino acid residues 153 and 175. This provides evidence

that the protein is membrane integrated. A more extensive presentation of the secondary

structure of the protein as predicted using the PredictProtein program (Rost, 1996) (available

through ExPASy) is given in Figure 5.4. The secondary structure prediction suggests that the

N-terminus of the protein is located inside the cytoplasm. Based on sequence similarity with

other bacterial transcriptional regulators this region also contains the DNA-binding domain,

thus further supporting the cytoplasmic location. Interestingly, a very large proportion of the

protein is exposed to the periplasm of the bacterial cell and may be involved in environmental

sensing as has been suggested for similar transcriptional regulatory proteins such as CadC

(Watson et al., 1992) and ToxR (Miller et al., 1987).

179

1000 2000 30000

Ap2W2pan3-S5

W2pan3-S2

W2pan3-S3

W2pan1-S3

Ap2

W2pan1

Ap2

Tn10C-S1

Ap2

Ap2

W2TnC-S4

W2pan4

W2TnC-S3 W2pan1-S2 W2pan3 W2pan3-S4

W2pan4-S2

Figure 5.1: Summary of the sequencing strategy used to determine the nucleotide sequence

flanking the transposon insert within the P. tunicata white mutant 2 (W2) genome. Arrows

indicate length and direction of sequencing primer products. Blue arrows represent

transposon or adaptor specific primers and black arrows represent sequence specific primers.

All primers used are listed in Appendix II. The nucleotide sequence is shown in base pairs

along the top of the diagram. The bold red line indicates the wmpR open reading frame.

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1 TGTCTGCCGACATCTTGATTTGATTATTTTGTTCAGTGATGTAAGTTTTTATAGTCAAAA

61 ATACTAAAGCAAACAGTAATATCATAGGTACCGCAAATAAAACTCTTCTTACTTTCATCA

121 TTTGGTCCATGTTTTGGTCCATCTGTTCAGACGAATATTGCGAACTATGTGCAGATCTTT

181 GCGGCATATTGGTGACAGACATAATATTAACCTTCCTATTTATTGATACTTAACTCGTCC

241 GTTTAAGGGCTCAATAAGTTGTTATCGAATGCTCAAGTAAAATAAAAAAGGCGTAATCTA

301 AAAAAGATAAATGTACGCCAAACTTAGGGAAATCTATGAAATTAAGGAGTTGTCATGCCT

361 CAATAGCCTTATATCGTACTTTTAATGCCGCTTAACCTAATATGATTGTTTTTGTTGTCT

421 TTTAAACTCAAAACTTTCACTTTACGTTTGTCAGCCATAAAAGTTATCTTAAGGTAACTC

481 AGTATTGAATTTGAACAATAATGACTGTGTAAT TTGTCT GTAATTAAAGTGTAACTTC TA (-35) 541 TATT TTTCATTGATTTAGATGTTGTTTGGCCATGTAGGATGTTGTTAGTAAATGAAAGGC (-10) 601 GGTTGAGTTACTATATAAAGCTGTTCTTAATTGCTCGACAAAATAATAAAAAGGGTAGTC

661 AGTGATTCAAATTGGTCGCTATCAATTAGATG AAGAAG AGATGGTGCTCAGCTGTGACGA (RBS) M V L S C D D

721 CCAGCGTGTTCTACTTGAACCTAAAGTATTTGATGTGCTTACATACTTTTGCCAGCATCA Q R V L L E P K V F D V L T Y F C Q H H

781 TAACCGTTATATCTCAATGACTGAGTTACACGAAAATATTTGGCAAGGTCGGTGTGTCTC N R Y I S M T E L H E N I W Q G R C V S

841 TGATGCAGCGGTCCGTCGTATCATTAGTAAAATTCGCATCTTAATGAACGACGATCATAA D A A V R R I I S K I R I L M N D D H K

901 AAACCCAACGTATATTCAATCTTTACCTAAGCGAGGCTATAAGCTGATCTGCCCGGTTGA N P T Y I Q S L P K R G Y K L I C P V E

961 ATATGATATTGAAGATGCTGCTGAATCATCCTCTGTAGAGAGTACAGCAGTAGCTATAAC Y D I E D A A E S S S V E S T A V A I T t 1021 GGATTTGTCTGATCATAATGAAGCCAATAATTATAATGAACCTACAGAGCCTGAAGAACA D L S D H N E A N N Y N E P T E P E E H

1081 TCAAGATTTAGCTGAAGAGTTGGCTGGTAATTTTGTTCACGTTGTTAAAAAGCCAAAAAA Q D L A E E L A G N F V H V V K K P K K

1141 ATTCAAGTATACCTTTTTGTCCCTTTTAATGTTATGTATATGTGTATTTGGTTACCTAGC F K Y T F L S L L M L C I C V F G Y L A

1201 TAAATCGTGGTTTTTCCCTGCTATAGTTCAAACTCAAGTGGTCAATACTTTACCCGGCGA K S W F F P A I V Q T Q V V N T L P G D

181

1261 TAAAATAGCTGTAACTCAATCAGCGGACGGTAGTTACCTCGCGTTTTCAGGTCAAGTGCA K I A V T Q S A D G S Y L A F S G Q V H

1321 TGATGAGTCGGGCTTTCAAATTTATGTGAAACACCAATCTGATTTTGATTTTAGGCCGAT D E S G F Q I Y V K H Q S D F D F R P I

1381 AACGCACCATGCACATCTACCAAGCTCAATAGCTTTTTCATTCGATAATAAGAGTTTATA T H H A H L P S S I A F S F D N K S L Y

1441 TTTTTCTGATACTTCAAAAATTAATTCATCATTAAATCAAATTAAGTTAGACGGAGAGAA F S D T S K I N S S L N Q I K L D G E N

1501 TCGGGAGATTGAAATATTAGTCGATAACTATTTTTTGATATCCGATGTATTTACTGCCAG R E I E I L V D N Y F L I S D V F T A R

1561 AACATCGAACAATGTCTTTTTTGCTGCGAAAAAGTCCACCGAGGGACCTTTTTTGATTTA T S N N V F F A A K K S T E G P F L I Y

1621 TGAGTATGATGTTGTTAATAAAGCTGTAACTGCAATTACTGCATCCTCTCAAGCTGAAAG E Y D V V N K A V T A I T A S S Q A E S

1681 TTTAGACATTAAAGGTGATGTATCGTTTGATGGCTCAAAATTAGCGGTATTAAGGACGAA L D I K G D V S F D G S K L A V L R T N

1741 TCGCTTAAGTCATAGTGATGAGATTCGCGTTATTGATTTAAAAACTAAAGAGGTAGTGAT R L S H S D E I R V I D L K T K E V V I

1801 ACGCAGGCAACATCCGGCTAGAGTTTATGATGTTGCCTGGGGTGATAATAATAATTTGCT R R Q H P A R V Y D V A W G D N N N L L

1861 GATCTTGAGTCGAGGCCAGCTACTGAAAATAAATATTGCAACAAGTGAAGAAACTCTCCA I L S R G Q L L K I N I A T S E E T L Q

1921 ATTCGCCAATGGGGTTAAACTTGCCAGCCTTGATTCAATTAAAAACAGAATAGTTTCTAT F A N G V K L A S L D S I K N R I V S I

1981 TAATTTAGGATTGAAAGAAAAGCTATTTATAGAAAAAAAACTGCCTTTCGGTGAGTTAGA N L G L K E K L F I E K K L P F G E L E

2041 AACGAAGCGAGTCCTCAAAAAAGATATTTATCAAATGAACTATTTTGGAGATAAAATTTT T K R V L K K D I Y Q M N Y F G D K I L

2101 AGCTTTGTTAAAAAATCATGACGTAACACAATTAGGTTTCTTGGATTTGGAAGCTGATCG A L L K N H D V T Q L G F L D L E A D R

2161 TTTTGATTCTGTGATTGCAACTGAGTACAATTTGGCTGTCTTAGATGTTGCTCCATTGCA F D S V I A T E Y N L A V L D V A P L Q

2221 AGGAAAAATTTTAGTTAGAATAAACAGAAGAATTGCATTATTAGATCCCAGTAACATAGA G K I L V R I N R R I A L L D P S N I D

2281 TCTTCAATATATCTCCTCCGGTGATGATTTAATTGGTGACGCAACTTTTTCAGCTGATAA L Q Y I S S G D D L I G D A T F S A D N

182

2341 TCTAAGTATTTTATTTTCTACTCAGAATTATGAGCAATGGGATGTAAATATTTTTAATAT L S I L F S T Q N Y E Q W D V N I F N I

2401 TGCTAAGAAAACTACTGAGCCGTTCTTGAGGGATATACGTTATATTCGACCGTACGGTGA A K K T T E P F L R D I R Y I R P Y G E

2461 GAGCTTTATAATTGGAGATTCTAAAGGTGAGTTATCTTTTTTTAGTCCATCAATAAATAA S F I I G D S K G E L S F F S P S I N K

2521 GAAGATAGCGTTGAATCACGCATTATCAAAAGAGCCAAATACACAATGGTTAGTTCGAGG K I A L N H A L S K E P N T Q W L V R G

2581 GGATTATATTTATTGGAGCTCACATGATTTAGTAAATACAACTTTTCATCAATTAAATAT D Y I Y W S S H D L V N T T F H Q L N I

2641 AAGTAATCTAAATCAGCCTGAGTTGGAAGTGCAGCAATTTATTTACAATGAGGTTAAGCC S N L N Q P E L E V Q Q F I Y N E V K P

2701 AGAATTTGCAATTGATCTTAATAATTTAAATTTTCTTATGTCGAAATCTGAGAGTGTAAC E F A I D L N N L N F L M S K S E S V T

2761 TTCAGAAATAGTAGAAATTCCATTCAGGTGAATCAGTGTTTTTGCCTTCAAATTGATAAA S E I V E I P F R *

2821 ATAATCGATAAGTTTGATGTTAGTTACGCGTTAGTTATTGTTAAAAGCTTTGTTTGCCAA

2881 TAAATTGTTCATAGGAATTAATTTTTACATTACTTTTTACATTACTTTTTACAACACTTT

2941 AAAAGAAGGCAAATCAATATGCTACATCAACTTGTTTTTACATTACTTCTTCCTATTTTA

3001 TCAACTTTATCAACAGTAACTGGTGACCCTGTAGTTGTGACAGATCCTCAGCCGCAAGAA

3061 TTATGTTACTTACTACCAGCTCGTTGTGATAAAGATCTTGAAGCTAACTCAGCAAATAGT

3121 AACTAATTATTCAGTATCGGCTGCATAGCAATATGTTGCCTTGTCAAAGCTGGGGTCATT

3181 TAATTAGTCGCACGCATTACATGTTTTGCTAATTAAGTTACAACGGCTTTGGCTAAAACG

3241 ACTAACGTGATTTTTTAACTTATTGGTTATAACAATGTCAGCTAGCTTTTTGTCAATGCC

3301 CATAGCAAAAAGCACTATTAGTTTGAATGCTTGTAATCGATATGTCGTACTTAAATTTCA

3361 GTATTAGTCATTATATACATCCATCCTACAATG ATGTTTCAAACATAAAGCGCATTTATT

3421 GCGCTTTTTTTTGTAAGCTT GCTTGCCATTTCATTAGTTTCATTTAAACTGTCACATCTT

3481 TCGAATCGAAACTTAATCTTTCACTGTCAGTCACTAATCCTGTTTGCAGCTGATGGATGT

3541 TATTATTGTTTTATTTCTCTTACTTTTATCGCTCTAATTATGATGATTTTTATATTTTTA

3601 TGACAAAACGAAATACCCAACAGCGTCGCCATACTATTTTAAGTCGTGTAAATGAACACG

3661 GTGAGGTGAGTGTT

183

Figure 5.2: Nucleotide sequence of the genomic-DNA surrounding the transposon within

the white mutant 2 (W2) genome. The nucleotide sequence is shown along with the translated

amino acid sequence of WmpR in one-letter code. The inverted solid triangle (t) indicates

the site where the mini-Tn10 transposon insertion occurred. The specific open reading frames

(ORF) as indicated in the text is highlighted in red. Potential promoter regions are underlined

as are predicted ribosome binding sites (RBS). A putative transcriptional terminator following

the ORF is underlined and in italics.

184

1 50WmpR .......... .......... .......... .....mvlsc ddqrvllepkEcCadC .......... ........mq qpvvrvgewl vtpsinqisr ngrqltleprVpToxR .......... ..mtnigtkf llaqrftfdp nsnsladqqs gnevvrlgsnVcToxR mfglghnske ismshigtkf ilaekftfdp lsntlidked seeiirlgsn • • •••

51 100WmpR vfdvltyfcq hhnryismte lheniw..qg rcvsdaavrr iiskirilmnEcCadC lidllvffaq hsgevlsrde lidnvwkrs. .ivtnhvvtq siselrkslkVpToxR esrillmlae rpnevltrne lhefvwreqg fevddssltq aistlrkmlkVcToxR esrilwllaq rpnevisrnd lhdfvwreqg fevddssltq aistlrkmlk • • • • • • •• • •••• • •• • • • •• •

101 150WmpR d.dhknptyi qslpkrgykl icpveydied aaesssvest avaitdlsdhEcCadC dndedspvyi atvpkrgykl mvpviwysee egeeimlssp ppipeavpatVpToxR d.stkspefv ktvpkrgyql ictverlspl ssdsssieve epasdnndasVcToxR d.stkspqyv ktvpkrgyql iarve..... .....tveee mareneaahd • • • • •• ••••••• ••••• • ••••••• •

151 200WmpR neannynept epeehqdlae elagnfvhvv kkpkkfkytf lsllmlcicvEcCadC dspshslniq ntatppeqsp vkskrfttfw vwfffllslg icvalvafssVpToxR anevetivep slatssdaiv epeapvvpek ahvasavnpw iprvilflalVcToxR isqpesvney aesssvpssa tvvntpqpan vvanksapnl gnrlfiliav • • •• • • • • •

201 250WmpR fgylakswff paivqtqvvn tlpgdkiav. .tqsadgsyl afsgqvhdesEcCadC ldtrlpmsks rillnprdid inmvnkscns wsspyqlsya igvgdlvatsVpToxR llpic.vllf tnpaesqfrq igeyqnvpv. .mtpvnhpqi nnwlpsieqcVcToxR llpla.vlll tnpsqssfkp ltvvdgvav. .nmpnnhpdl snwlpsielc •• • • • •• •• ••• • •

251 300WmpR gfqiyvkhqs dfdfrpithh ahlpssiafs fdnkslyfsd tskinsslnqEcCadC lntfstfmvh dkinynidep sssgktlsia fvnqrqyraq qcfmsiklvdVpToxR ieryvkhhae dslpveviat ggqnnqliln yihdsnhsye nvtlrifagqVcToxR vkkynekhtg glkpieviat ggqnnqltln yihspevsge nitlrivanp •• • • • • • • • •

185

301 350WmpR ikldgenrei eilvdnyfli sdvftartsn nvffaakkst egpfliyeydEcCadC nadgstmldk ryvitngnql aiqndllesl skalnqpwpq rmqetlqkilVpToxR ndptdick.. .......... .......... .......... ..........VcToxR ndaikvce.. .......... .......... .......... .......... • •

351 400WmpR vvnkavtait assqaesldi kgdvsfdgsk lavlrtnrls hsdeirvidlEcCadC phrgalltnf yqahdyllhg ddkslnrase llgeivqssp eftyaraekaVpToxR .......... .......... .......... .......... ..........VcToxR .......... .......... .......... .......... .......... • • • • •

401 450WmpR ktkevvirrq hparvydvaw gdnnnllils rgqllkinia tseetlqfanEcCadC lvdivrhsqh pldekqlaal nteidnivtl pelnnlsiiy qikavsalvkVpToxR .......... .......... .......... .......... ..........VcToxR .......... .......... .......... .......... .......... • • •

451 500WmpR gvklasldsi knrivsinlg lkeklfiekk lpfgeletkr vlkkdiyqmnEcCadC gktdesyqai ntgidlemsw lnyvllgkvy emkgmnreaa dayltafnlrVpToxR .......... .......... .......... .......... ..........VcToxR .......... .......... .......... .......... .......... • • • • • • •

501 550WmpR yfgdkilall knhdvtqlgf ldleadrfds viateynlav ldvaplqgkiEcCadC pgantlywie ngifqtsvpy vvpyldkfla se........ ..........VpToxR .......... .......... .......... .......... ..........VpToxR .......... .......... .......... .......... .......... • • •

551 600WmpR lvrinrrial ldpsnidlqy issgddligd atfsadnlsi lfstqnyeqwEcCadC .......... .......... .......... .......... ..........VpToxR .......... .......... .......... .......... ..........VcToxR .......... .......... .......... .......... ..........

186

601 650WmpR dvnifniakk ttepflrdir yirpygesfi igdskgelsf fspsinkkiaEcCadC .......... .......... .......... .......... ..........VpToxR .......... .......... .......... .......... ..........VcToxR .......... .......... .......... .......... ..........

651 700WmpR lnhalskepn tqwlvrgdyi ywsshdlvnt tfhqlnisnl nqpelevqqfEcCadC .......... .......... .......... .......... ..........VpToxR .......... .......... .......... .......... ..........VcToxR .......... .......... .......... .......... ..........

701 736WmpR iynevkpefa idlnnlnflm sksesvtsei veipfrEcCadC .......... .......... .......... ......VpToxR .......... .......... .......... ......VcToxR .......... .......... .......... ......

Figure 5.3: Multiple sequence alignment of the deduced amino acid sequence of P. tunicata

WmpR with the transcriptional activator CadC from Escherichia coli (EcCadC, GenBank

accession number: P23890); the ToxR-homologue from Vibrio parahaemolyticus (VpToxR,

GenBank accession number: Q05938) and ToxR cholera-toxin transcriptional activator from

V. cholerae (VcToxR, GenBank accession number: P15795). Residues identical between P.

tunicata and one other protein are indicated by black dots (•); residues identical in two other

proteins are indicated with blue dots (•) and residues which are identical between all proteins

are indicated by red dots (•). Small dots (.) denote gaps.

187

.........10........20........30........40........50.......60AA MVLSCDDQRVLLEPKVFDVLTYFCQHHNRYISMTELHENIWQGRCVSDAAVRRIISKIRISec_pred EE E HHHHHHHHH HHHHHHHHH HHHHHHHHHHHRel_sec * ** * ********* *** ******** ** **********Mem_top iiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiii

.........70........80........90........100.......110......120AA LMNDDHKNPTYIQSLPKRGYKLICPVEYDIEDAAESSSVESTAVAITDLSDHNEANNYNESec_pred HHH EHHHH EEEE EE HHHHH HHHEEEERel_sec ** ***** * * ** ** **** * *******Mem_top iiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiii

.........130.......140.......150.......160.......170......180AA PTEPEEHQDLAEELAGNFVHVVKKPKKFKYTFLSLLMLCICVFGYLAKSWFFPAIVQTQVSec_pred HHHHHHHHHHH EEEEEE HHHHHHHHHHHHHHHHHHHH EEEEEERel_sec **** ********** * ***** *** ******************* ****Mem_top iiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiMMMMMMMMMMMMMMMMMMooooooooo

.........190.......200.......210.......220.......230......240AA VNTLPGDKIAVTQSADGSYLAFSGQVHDESGFQIYVKHQSDFDFRPITHHAHLPSSIAFSSec_pred E EEEEE EEEEEEEEE EEEEEE EE EEEERel_sec **** ***** *** ****** **** ***** ***** * **** ***Mem_top oooooooooooooooooooooooooooooooooooooooooooooooooooooooooooo

.........250.......260.......270.......280.......290......300AA FDNKSLYFSDTSKINSSLNQIKLDGENREIEILVDNYFLISDVFTARTSNNVFFAAKKSTSec_pred E EEEEE EE EEEEE EEEEEEEEEE HHHHHHHHHRel_sec *** ** * **** *** * ******* **** *Mem_top oooooooooooooooooooooooooooooooooooooooooooooooooooooooooooo

.........310.......320.......330.......340.......350......360AA EGPFLIYEYDVVNKAVTAITASSQAESLDIKGDVSFDGSKLAVLRTNRLSHSDEIRVIDLSec_pred EEEEEE HHHHHHHHHHH EEE EEEEEE EEEEERel_sec *** **** ********* ** *** **** ***********Mem_top oooooooooooooooooooooooooooooooooooooooooooooooooooooooooooo

.........370.......380.......390.......400.......410......420AA KTKEVVIRRQHPARVYDVAWGDNNNLLILSRGQLLKINIATSEETLQFANGVKLASLDSISec_pred EEEEEE EEEE EEE EEE HHHHHHH EEEERel_sec ** **** *** * **** * *** ***** *** *Mem_top oooooooooooooooooooooooooooooooooooooooooooooooooooooooooooo

.........430.......440.......450.......460.......470......480AA KNRIVSINLGLKEKLFIEKKLPFGELETKRVLKKDIYQMNYFGDKILALLKNHDVTQLGFSec_pred EEEEE EEEEE HHHHHHHHHHHHHHHHHHHHHHHHH EEERel_sec * ******* ***** ************ ********* ***Mem_top oooooooooooooooooooooooooooooooooooooooooooooooooooooooooooo

188

.........490.......500.......510.......520.......530......540AA LDLEADRFDSVIATEYNLAVLDVAPLQGKILVRINRRIALLDPSNIDLQYISSGDDLIGDSec_pred EEEE EEE EEEE EE EEEEERel_sec ** ** ***** *** ***Mem_top oooooooooooooooooooooooooooooooooooooooooooooooooooooooooooo

.........550.......560.......570.......580.......590......600AA ATFSADNLSILFSTQNYEQWDVNIFNIAKKTTEPFLRDIRYIRPYGESFIIGDSKGELSFSec_pred EEEEEE HHHHHHHHHH HHHHHHHHHEE EEEEE EEERel_sec ** ** ** *** *** **** *** *Mem_top oooooooooooooooooooooooooooooooooooooooooooooooooooooooooooo

.........610.......620.......630.......640.......650......660AA FSPSINKKIALNHALSKEPNTQWLVRGDYIYWSSHDLVNTTFHQLNISNLNQPELEVQQFSec_pred HHHHHHHHHHH EEEE EEEEE EEE HHHHHHHHRel_sec ** ******* ***** * **** * ***** *******Mem_top oooooooooooooooooooooooooooooooooooooooooooooooooooooooooooo

.........690.......700.......710....AA IYNEVKPEFAIDLNNLNFLMSKSESVTSEIVEIPFRSec_Pred HHHHH EEE HHHH EEEERel_sec **** * ***** * ** **Mem_top oooooooooooooooooooooooooooooooooooo

Figure 5.4: Secondary structure prediction of the deduced amino acid sequence of WmpR as

predicted by PredictProtein (Rost, 1996). AA = amino acid sequence. Sec_pred = Predicted

secondary structures, H = helix, E = extended (sheet), blank = other (loop). Rel_sec

represents the reliability index, strong predictions are indicated by *. Mem_top = predicted

membrane topology, M = helical transmembrane region, i = inside membrane, o = outside.

189

5.3.2. Global differences in protein expression between wild-type P. tunicata

and the W2 mutant

Based on the phenotypic and genotypic study of the P. tunicata W2 mutant it would appear

that the putative transcriptional regulator, WmpR, is responsible for the expression of genes/

proteins required for the syntheses of pigments and fouling inhibitors. To further investigate

the role of WmpR and to identify specific proteins that are influenced by this regulator,

differences in the expression of proteins at a global level were examined using two-

dimensional gel electrophoresis (2DGE). Proteins expressed by both the wild-type and the

W2 mutant strains were compared at two growth phases. The first samples were taken during

early-logarithmic growth (i.e. before the expression of pigments and inhibitors). The second

samples were taken during early-stationary phase of growth, when pigments and antifouling

inhibitors are beginning to be expressed by wild-type cells. Approximately 950 spots were

detected on the gels and analysed (Figure 5.5, Figure 5.6, Figure 5.7 and Figure 5.8).

Specific differences in protein expression were found between the wild-type and mutant

samples and a summary of this analysis is given in Figure 5.9. A total of 39 proteins were

up-regulated and 9 down-regulated when the early-logarithmic wild-type sample was

compared with the early-stationary phase wild-type sample. Similarly, 24 protein spots were

up-regulated and 9 down-regulated in the mutant when comparing the early-logarithmic

sample with early-stationary phase. A comparison of the mutant with the wild-type at the

early-stationary growth phase found 15 protein spots to be missing or down-regulated in the

mutant. These proteins consisted of a sub-population of those that were found to be up-

regulated in wild-type cells in early-stationary phase of growth. In contrast, no difference was

found between the wild-type and the mutant strain for the early-logarithmic growth phase

samples.

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pH 4 pH 7

Figure 5.5: : Two-dimensional gel electrophoresis of the total cell protein from P. tunicata

wild-type (Wt) in early-logarithmic growth. Fifty micrograms of protein were separated and

the gel silver stained. Protein spots circled in blue indicate proteins found in P. tunicata Wt

during early-logarithmic growth, but not in Wt during early-stationary phase growth (Figure

5.7).

191

pH 4 pH 7

Figure 5.6: Two-dimensional gel of the total cell protein from P. tunicata white mutant 2

(W2) in early-logarithmic growth. Fifty micrograms of protein were separated and the gel

silver stained. Protein spots circled in blue indicate proteins found in P. tunicata W2 strain

during early-logarithmic growth, but not in P. tunicata W2 strain during early-stationary

phase growth (Figure 5.8).

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pH 4 pH 7

Figure 5.7: Two-dimensional gel of the total cell protein from P. tunicata wild-type (Wt) in

early-stationary phase growth. Fifty micrograms of protein were separated and the gel silver

stained. Protein spots circled in either blue or red indicate proteins found in P. tunicata Wt

during early-stationary growth, but not in Wt during early-logarithmic growth (Figure 5.5).

Protein spots circled in red represent proteins present in P. tunicata Wt during early-

stationary phase, but not in the white mutant 2 P. tunicata strain during early-stationary phase

(Figure 5.8).

.

193

pH 4 pH 7

Figure 5.8: Two-dimensional gel of the total cell protein from P. tunicata white mutant 2

(W2) in early-stationary growth. Fifty micrograms of protein were separated and the gel

silver stained. Protein spots circled in blue indicate proteins found in P. tunicata W2 during

early-stationary growth, but not in P. tunicata W2 during early-logarithmic growth (Figure

5.6).

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15 down-regulated

39 up-regulated

24 up-regulated

0

Wt early logarithmic

Wt early stationary

W2 early logarithmic

W2 early stationary

9 down-regulated

9 down-regulated

Figure 5.9: The differences in the number of proteins expressed by P. tunicata wild-type

(Wt) and the white mutant 2 (W2) at both early-logarithmic and early-stationary phase of

growth as detected by two-dimensional gel electrophoresis.

5.4. Discussion

As described in the previous chapter transposon mutants of P. tunicata deficient in yellow

pigment (i.e. purple and white mutants) are unable to inhibit the target fouling organisms such

as invertebrate larvae, algal spores, fungi and bacteria. The phenotypic characterisation of

these mutants suggests that the pigments and inhibitors share a common biosynthetic pathway

or are coordinately regulated. Sequence analysis of the DNA region where the transposon

had inserted in the white mutant 2 (W2) strain revealed that a gene with similarity to

transcriptional regulators was disrupted. The deduced gene product of the ORF, designated

WmpR, was found to be similar in the amino-terminus to a sub-group of the OmpR-like

transcriptional activators, including CadC from E. coli and ToxR from V. cholerae. This class

of regulators differ from other members of the OmpR group as they are membrane bound and

are able to both sense and respond to changes in the external environment. In addition, these

regulators have their DNA-binding domain located at the amino-terminus rather than the

carboxy-terminus of the protein (Miller et al 1987; Watson et al 1992).

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ToxR coordinately regulates the expression of a number of virulence genes such as cholera

toxin (CT) and the toxin co-regulated pilus (TCP) in V. cholerae, the causative agent of the

diarrhoeal disease cholera. The ToxR protein is a transmembrane DNA-binding protein

which also functions as a sensor and whose activity is enhanced by the presence of ToxS.

The genes encoding for ToxR and ToxS are expressed as a single operon separate from the

structural genes they regulate. A detailed description of the complete ToxR regulon is

provided in section 1.4.2. Homologues to ToxR have been characterised in a number of other

species in the Vibrionaceae, including V. parahaemolyticus (Lin et al., 1993), V. fischeri

(Reich and Schoolnik, 1994), V. vulnificus (Lee et al., 2000) and Photobacterium profundum

(Welch and Bartlett, 1998). The common theme among this group of organisms is the need

to respond to relatively extreme changes in environmental conditions. For example from a

free-living state in seawater to a form closely associated with their host, as is the case of V.

cholerae and V. fischeri, or in response to high pressure as for the deep-sea bacterium P.

profundum.

The cadC gene from E. coli also encodes a membrane-bound transcriptional activator that is

capable of sensing and responding to changes in environmental conditions. The CadC protein

activates the transcription of cadBA under conditions of low external pH and in the presence

of lysine. The cadBA operon encodes proteins involved in the decarboxylation of lysine to

cadaverine (CadA) and for the lysine/ cadaverine transport (CadB) (Watson et al, 1992; Dell

et al., 1994). The production and excretion of cadaverine leads to an increase in the external

pH and has been suggested to provide a selective advantage for the bacterium under acidic

growth conditions (Neely and Olson, 1996).

Unlike toxR, cadC is located in close proximity to the structural genes for which it regulates

and it does not appear to be associated with a toxS homologue. Sequence analysis of the

DNA flanking the wmpR gene of P. tunicata has not yet identified a toxS homologue nor any

other ORF, which may suggest that WmpR functions at on a regulatory level in a similar way

to CadC. Both CadC and WmpR are predicted to consist of a much larger periplasmic

domain than the ToxR protein. Since ToxS functions to enhance activity of ToxR, the larger

periplasmic domain of CadC and WmpR may compensate for the lack of a ToxS homologue.

Two-dimensional gel electrophoresis (2DGE) was used to investigate the role of WmpR with

respect to global changes in protein expression. Total cellular proteins were compared

between wild-type and the W2 mutant at two different growth phases, before (early-

logarithmic) and during (early-stationary) the expression of pigment and inhibitors in P.

196

tunicata wild-type. Analysis of the 2DGE patterns between the wild-type and the mutant

found no differences in proteins expressed by the two strains during early-logarithmic phase

of growth. However a comparison during early-stationary growth phase found 15 protein

spots missing in the mutant. The missing proteins consisted of a sub-population

(approximately 40 %) of those found to be up-regulated in the wild-type when the cells

entered early-stationary phase of growth (Figure 5.9).

The results of the 2DGE analysis show a difference in protein expression, which coincides

with pigment/ inhibitor production in P. tunicata. The W2 mutant expresses fewer proteins

than the wild-type, which correlates well with the loss of pigment/ inhibitor production in this

strain. Furthermore, these results provide the first evidence that WmpR may function as a

regulator of protein expression in P. tunicata cells. Moreover, the up-regulation of 15

proteins in the wild-type and not in the W2 mutant indicates that WmpR is an activator of

protein expression rather than a represser.

Global protein expression studies of other bacteria entering stationary phase of growth have

demonstrated both an up-regulation and a down-regulation of specific proteins (Nyström,

1993). This pattern has also been observed for P. tunicata during the transition from

logarithmic growth to stationary growth phase (see Figure 5.9). Due to limitations in protein

detection using silver staining, it is possible that not all the proteins were detected in this

study. Thus several other proteins regulated by WmpR may be identified in future studies.

Radioactively labelling proteins during cell growth may improve the detection limit. In

addition, this method would allow for the distinction between constitutively expressed proteins

and those that are regulated in response to growth conditions, because only the proteins that

are synthesised at or after the addition of the radiolabel are visualised. Nevertheless, it is clear

that several proteins are under the control of the putative WmpR regulator.

The nature of the signal/s needed for WmpR mediated expression of the pigments and the

inhibitors is unclear. The large periplasmic C-terminal domain of the protein would suggest

that it is responding to external environmental conditions such as the presence or absence of

specific nutrients, toxins or signal molecules. Observations of pigment/ inhibitor expression

under different culture conditions indicate that the response is mediated by various

environmental signals. For example, in nutrient rich media a reduced pigment expression

occurs and during growth at 35 oC only the yellow pigment is expressed (data not shown). In

a natural setting WmpR may be required by P. tunicata to sense and respond to signals

generated by the host organism (eg. tunicates) which in turn will lead to the expression of

197

antifouling phenotypes useful for the symbiotic host. To identify environmental signals

involved in WmpR mediated regulation, future studies in which genes of interest (eg: wmpR,

afaA, lppA etc. See chapter 4) are fused to reporter genes, such as green fluorescent protein

(GFP) will be conducted. Expression of these genes can then be monitored under different

environmental conditions to identify the conditions needed to stimulate productions of

antifouling inhibitors.

Finally, it has also been observed that the knock-out mutation in wmpR can be complemented

during extended periods of incubation, i.e. reversion to wild-type phenotype (data not shown).

PCR and sequence analysis suggests that the return to a green phenotype is not the result of

an unstable transposon insertion but rather to some other effect. One explanation could be

that the putative transcriptional regulator is part of a more complex regulatory cascade. For

example, many bacteria are capable of sensing and responding to the environment via the use

of small diffusible signal molecules. This form of gene regulation is also known as quorum-

sensing and can be linked to complex regulatory cascades within the cell (see section 1.4.3).

The possibility that quorum-sensing is involved in pigment and fouling inhibitor production

has been addressed and a putative homologue to the luxS gene involved in the AI-2 regulatory

system of V. harveyi has been identified in P. tunicata (Franks et al, unpubl.).

In conclusion, this chapter has identified the first regulator involved in controlling the

expression of both pigment and fouling inhibitors in the marine antifouling bacterium P.

tunicata. Two-dimensional gel electrophoresis was used as a powerful tool to determine the

proteins under the control of WmpR. Further work will involve the identification of WmpR-

regulated proteins with techniques such as N-terminal amino acid sequencing and peptide

mass-spectrometry fingerprinting. It is possible that the study of a CadC/ ToxR homologue

in P. tunicata will aid in the understanding of this organism ability to coordinate the

expression of pigments and the specific extracellular inhibitory compounds. In addition, these

studies will provide a greatly improved understanding of stationary phase biology in this

organism as well as add to the general understanding of bacterial regulatory systems.

198

6. Antifouling activity and phylogenetic relationship of bacteria

isolated from different marine surfaces

6.1. Introduction

As discussed in the previous chapters, biological interactions between different marine surface

associated organisms play a major role in the development and maintenance of a biofouling

community. Many sessile marine plants and animals have evolved defence mechanisms

against fouling by producing metabolites which can influence the settlement, growth and

survival of other competing organisms (de Nys et al., 1994; Mary et al., 1993; McCaffrey and

Endean, 1985). For example, natural products from the leaves of the eelgrass Zostra marina

have been found to prevent attachment of marine bacteria and barnacles to artificial abiotic

surfaces (Harrison, 1982; Todd et al., 1993). Another well studied example is the red alga

Delisea pulchra, which produces secondary metabolites known as halogenated furanones.

These compounds are able to interfere with bacterial colonisation traits and in addition prevent

settlement of invertebrate larvae and spores of common algae (de Nys et al, 1994; Kjelleberg

et al., 1997; Maximilien et al., 1998). Plants and animals may also rely on the secondary

metabolites produced by bacteria as their defence against other surface colonising organisms.

The sponge isolate Alteromonas sp. KK10304, has been shown to produce components

responsible for preventing the settlement of marine invertebrate larvae (Kon-ya et al., 1995).

The antifouling bacterium P. tunicata isolated from the surface of tunicates, is an effective

inhibitor of many fouling organisms. The anti-algal activity and anti-fungal activity were

discussed in chapters 2 and 3, respectively. In both cases the components released by this

bacterium were found to be target-specific and appeared unique in their effectiveness towards

the target organism.

While it is now widely accepted that bacteria can inhibit the colonisation of surfaces by

fouling organisms, little information is available regarding the diversity and properties of these

antifouling bacteria (Holmström et al., 1992; Maki et al., 1988; Mary et al., 1993). The aims

of this chapter are (i) to assess the frequency with which bacterial strains isolated from living

and inanimate surfaces in the marine environment show inhibitory activity against the

199

settlement or growth of fouling organisms and (ii) to determine the identity and the

phylogenetic relationship of inhibitory isolates.

6.2. Material and Methods

6.2.1. Bacterial strains

A total of 164 bacterial strains were isolated from various surfaces (including rocks, algae and

sessile animals) in the marine waters around Sydney, N.S.W., Australia. Isolation of the

bacterial strains was performed in collaboration with Dr Carola Holmström from the Centre

for Marine Biofouling and Bio-Innovation. The object of interest was resuspended into sterile

NSS (Appendix I) and surface bacteria were removed by vortexing. Aliquots of the samples

were then spread on VNSS agar (Appendix I) and incubated at 23 oC for 48 h.

Morphologically distinct bacterial colonies were selected. Bacteria were stored at -70 oC in 30

% (v/v) glycerol. Ninety-three strains were obtained from various rock surfaces, twenty-three

were obtained from a variety of seaweeds and twenty-three strains were isolated from the

surface of sessile animals such as tunicates, barnacles and nematodes. An additional twenty-

five dark pigmented strains were isolated from various seaweeds.

6.2.2. Antifouling activity of the marine isolates

The effect of the various marine bacterial isolates on the settlement of barnacle larvae B.

amphitrite was assessed as described in section 3.2.3.4. Activity against the germination of

spores from the algae Ulva lactuca and Polysiphonia was determined as described in sections

2.2.3 and 2.2.12, respectively. Activity against the growth of a wide range of bacterial strains

was performed on agar plates using the overlay method as described in section 3.2.3.2.

Among the bacterial strains tested were a collection of eight unidentified marine sponge

isolates (Longford, 1999). Anti-fungal activity of the bacterial isolates against three different

target fungi was determined using the bioassay outlined in section 3.2.1.

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6.2.3. Genomic extractions, 16S ribosomal DNA amplification and DNA

sequencing

To identify bacterial isolates of interest and determine their phylogenetic relationship, genomic

DNA was extracted using the XS-buffer method as described in section 3.2.4.5. Following

genomic DNA extraction the 16S ribosomal DNA was PCR-amplified using primers

corresponding to positions 27 in the forward direction (F27) and 1492 in the reverse direction

(R1492) of the Escherichia coli 16S rRNA gene sequence (see Appendix II). The

thermoprofile consisted of 25 cycles of denaturation at 95 oC for 30 sec, annealing at 54 oC

for 30 sec and extension at 72 oC for 2 min. The PCR products were visualised on a 1 %

(w/v) agarose gel using a molecular-weight standard to estimate the size and concentration of

products. Single band products were excised from the agarose gel and purified using a Prep-

a-Gene DNA purification kit (BioRad). Approximately 100 ng of the template DNA was then

sequenced in a thermocycling reaction with BigDyeTM terminator cycle sequencing mix

(Applied Biosystems) as outlined in section 3.2.3.7. The 16S rRNA gene was sequenced in

both directions by primer walking using primers directed to the conserved regions of the gene.

The specific sequence of each primer is given in Appendix II.

6.2.4. Phylogenetic analysis

Phylogenetic analysis was performed with the assistance of Torsten Thomas (School of

Microbiology and Immunology, UNSW). The DNA sequences of the 16S rRNA gene from

isolates of interest were aligned using the programs PILEUP and CLUSTAL W (GCG-

software package) (Wisconsin, 1992). The aligned sequences were applied to genetic distance

and maximum parsimony methods for phylogenetic inference. Gaps and ambiguous

positions were manually deleted and distances were calculated using the formula of Jukes and

Cantor (1969), Kimura (1980) and maximum likelihood (Felsenstein, 1981). Phylogenetic

inference protocols, EDNAML, EDNADIST, NEIGHBOR, EDNAPARS, CONSENSE and

SEQBOOT were supplied by the PHYLIP packages (version 3.57c) (Felsenstein, 1989). All

sequence manipulation and phylogeny programs were made available through the Australian

National Genome Information Service (ANGIS, Sydney, Australia).

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6.3. Results

6.3.1. Settlement of B. amphitrite larvae in the presence of bacterial strains

isolated from different marine surfaces

The frequency of bacterial strains isolated from living and inanimate surfaces that display

inhibitory effects towards the settlement of invertebrate larvae was assessed. The results of

these assays showed that bacterial strains isolated from different surfaces displayed varying

degrees of anti-larval activity. Ten percent of bacterial isolates from rock surfaces were found

to be inhibitory to the settlement of larvae compared with 30 % of isolates from marine

animals and 74 % of bacteria isolated from marine algae (Table 6.1).

Table 6.1: Anti-larval activity of bacterial strains isolated from different marine surfaces

Source of marine bacterial

isolates

Number of isolates tested Number of inhibitory isolates

Rock surfaces 93 9

Animal surface 23 7

Algal surface 23 17

6.3.2. Settlement of B. amphitrite larvae in the presence of dark pigmented

bacterial isolates

The previous chapters of this thesis demonstrated that pigmentation correlates with the

expression of antifouling activities in P. tunicata. To determine if other dark pigmented

marine bacteria also display antifouling activity twenty-five dark pigmented isolates taken

from the surface of various seaweeds were tested against invertebrate larval settlement. With

the exception of one (isolate 14), all of the pigmented isolates tested were able to reduce the

number of settled larvae compared to the control containing seawater alone (Figure 6.1).

Among the pigmented strains having the strongest activity, five isolated from the alga Ulva

lactuca (designated UL1, UL12, UL13, UL14 and UL15) were selected for further

investigation.

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SW

UL

1

UL

12

UL

13

UL

14

UL

15 1 2 3 4 5 6 7 8 9

10 11 12 13 14 15 16 17 18 19 21

0

20

40

60

80

Bacterial isolate

% S

ettl

emen

t

Figure 6.1: Settlement (%) of B. amphitrite larvae on biofilms of dark-pigmented marine

bacteria. The values represent means ± standard deviations (n=3).

203

6.3.3. Germination of U. lactuca and Polysiphonia sp. spores in the presence of

biofilms of the U. lactuca bacterial isolates

To assess if the U. lactuca isolates also display anti-algal activity, biofilms of each isolate

were prepared and germination assays performed with spores from the green alga U. lactuca

and the red alga Polysiphonia sp. The data presented in Table 6.2 show the percentage of

algal spores that germinated when exposed to biofilms of the U. lactuca bacterial isolates.

Isolates UL1, UL14 and UL15 inhibited the germination of algal spores compared to controls

containing seawater alone and biofilms of the non-inhibitory marine strain J1. Isolates UL12

and UL13 were inhibitory to the germination of U. lactuca spores, however compared to

isolates UL1, UL14 and UL15 they showed a reduced activity against the germination of

spores from Polysiphonia sp. In addition, spores of U. lactuca were more sensitive towards

the bacterial isolates compared with the spores of Polysiphonia.

Table 6.2: Germination of algal spores in the presence of biofilms of marine bacterial

isolates

Percentage germination a

Target

organism

UL1b UL12 b UL13 b UL14 b UL15 b J1c No

Biofilm

Ulva lactuca

spores

1.5 ± 2 2.5 ± 2 6 ± 2.2 0 0 83 ± 9 100

Polysiphonia

sp. spores

1 ± 1.7 34 ± 5.3 23 ± 13 0 3 ± 2.4 87 ± 5.6 88 ± 6.1

a All values are means ± standard deviations (n=3); b Bacteria from the surface of the marine alga U. lactuca ;

c Non-inhibitory marine bacterium (see section 2.3.1)

6.3.4. Anti-bacterial activity of the U. lactuca isolates

Anti-bacterial activity of each of the U. lactuca isolates was assessed against a collection of

twenty different bacterial strains. Isolates UL1, UL14 and UL15 inhibited 90 % of the

bacteria tested, including a variety of Gram-positive, Gram-negative, marine, terrestrial and

204

pathogenic bacteria (Table 6.3). These three isolates were also strongly inhibitory against

their own growth and against the growth of other isolates from U. lactuca. While isolates

UL12 and UL13 did not display an overall anti-bacterial effect they were found to slightly

inhibit the growth of some of the unidentified marine sponge isolates (Table 6.3).

Table 6.3: Anti-bacterial activity of marine isolates against various target bacterial strains

Growth inhibition (mm) a

Target strain UL1b UL12b UL13b UL14b UL15b

Pseudoalteromonas tunicata c 6.5 0 0 9 7

Pseudomonas aeruginosa c 3 0 0 2.5 2.5

Vibrio harveyi c 0.5 0 0 0.5 0.5

Escherichia coli c 0 0 0 0 0

Staphylococcus aureus c 1.5 0.5 0.5 1.5 1

Serratia marcescens c 0 0 0 0 0

Bacillus megaterium c 2.5 0 0 2.5 3

Marine isolate SI 1 (Gram +ve) d 3 0.5 0.5 3 3

Marine isolate CA 3 (Gram +ve) d 3 0.5 0.5 2.5 3

Marine isolate UL 164 (Gram +ve) d 2.5 1 1 2 2

Marine isolate AC 24 (Gram +ve) d 2 1.5 1.5 1.5 2

Marine isolate SI 9 (Gram -ve) d 2.5 0 0 3 3

Marine isolate CBb 16 (Gram -ve) d 3 1 1 2.5 2.5

Marine isolate S 15 (Gram -ve) d 3.5 2 1.5 3.5 2.5

Marine isolate CA 24 (Gram -ve) d 3 0 0 3.5 3.5

UL1 b 7 0 0 7.5 7

UL12 b 7 0 0 6 7

UL13 b 6 0 0 6.5 6.5

UL14 b 7.5 0 0 8.5 7

UL15 b 7.5 0 0 6 8

a All values are representative of the radius, in mm, of inhibition zones of the target strain; b Bacteria from the

surface of the marine alga U. lactuca ; c Bacterial strains obtained from the School of Microbiology and

Immunology culture collection, University of New South Wales, Sydney; d Unidentified marine isolates

(Longford, 1999).

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6.3.5. Anti-fungal activity of the U. lactuca isolates

Inhibition of fungal growth by the U. lactuca isolates was assessed against three different

fungal strains. The results of these assays as presented in Table 6.4, indicate that isolates

UL12 and UL13 had no effect on fungal growth, while isolates UL1, UL14 and UL15

displayed strong growth inhibition against all three fungi.

Table 6.4: Anti-fungal activity of marine isolates against various target fungal strains

Growth inhibition (mm) a

Target strain UL1b UL12b UL13b UL14b UL15b

Aureobasidium pullulans 5.5 0 0 6 7

Cladosporium cladosponoides 6.5 0 0 6 6

Penicillium digitatum 5 0 0 4.5 5

a All values are representative of the radius, in mm, of inhibition zones of the target strain; b Bacteria from the

surface of the marine alga U. lactuca.

6.3.6. 16S rDNA sequencing and phylogenetic analysis of the U. lactuca

isolates

In order to identify the U. lactuca isolates and determine their phylogenetic relationship,

phylogenetic analysis based on the 16S rRNA gene was performed. The DNA sequencing

strategy using primers (Appendix II) directed to conserved regions within the 16S rRNA gene

was successful in generating 1386, 1355, 1403, 1365 and 1384 base pairs of sequence for

isolates UL1, UL12, UL13, UL14 and UL15, respectively. Based on the analysis of the

sequence data all isolates belong to the genus Pseudoalteromonas and are most closely

related to the species P. tunicata. In addition, the phylogenetic tree represented in Figure 6.2

shows that the isolates UL12 and UL13 form a distinct phylogenetic group separate from

isolates UL1, UL14 and UL15. Nucleotide sequences have been deposited in the

DDBJ/EMBL/GenBank database under the accession numbers AF172987 through

AF172991.

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0.01Vibrio anguillarum

Vibrio fischeri

Pseudoalteromonas bacteriolyticaPseudoalteromonas strain UL12Pseudoalteromonas strain UL13

Pseudoalteromonas tunicataPseudoalteromonas strain UL1Pseudoalteromonas strain UL14Pseudoalteromonas strain UL15

Pseudoalteromonas denitrificansPseudoalteromonas aurantia

Pseudoalteromonas citreaPseudoalteromonas sp. ANG.RP2

Pseudoalteromonas prydzensis

Pseudoalteromonas sp. S9Pseudoalteromonas luteoviolacea

Pseudoalteromonas rubraPseudoalteromonas piscicida

Pseudoalteromonas sp. YPseudoalteromonas peptidolytica

Pseudoalteromonas sp. MB8-02

Pseudoalteromonas undinaPseudoalteromonas nigrifaciens

Pseudoalteromonas sp. SWO8Pseudoalteromonas antarctica

Pseudoalteromonas haloplanktis

Pseudoalteromonas gracilisPseudoalteromonas sp. MB6-05

Pseudoalteromonas sp. IC006Pseudoalteromonas carrageenovora

Pseudoalteromonas tetradonisPseudoalteromonas espejianaPseudoalteromonas atlantica

Pseudoalteromonas distinctaPseudoalteromonas elyakovii

Pseudoalteromonas sp. MB6-03Pseudoalteromonas sp. IC013

Figure 6.2: Distance matrix tree based on a 1111 bp sequence alignment of the 16S

ribosomal DNA of novel isolates with members of the genus Pseudoalteromonas. Distances

were calculated according to the algorithm of Jukes and Cantor (1969) and trees calculated

according to Saitou and Nei (1987). Outgrouping was performed with Vibrio anguillarum.

Bar indicates 1 substitution per 100 nucleotide positions.

207

6.4. Discussion

Many marine bacteria both free-living and attached, have been shown to influence the

settlement of invertebrate larvae. Metabolites derived from bacteria can provide larvae with a

negative cue, which would cause them to search else where for a suitable substratum for

settlement. Likewise, the degree to which marine bacteria respond to metabolites from marine

algae and invertebrates can have a profound effect on the distribution of bacteria on a living

surface. Due to this complex interaction between different surface-associated organisms,

living marine surfaces (in general) do not show the relatively regular pattern of biofilm

formation that can be observed on inanimate surfaces (Wahl. et al., 1994). For example, the

numbers of bacterial epiphytes found on the red alga D. pulchra is inversely related to the

concentration of the algal derived secondary metabolites over its surface (Maximilien et al.,

1998). The importance of bacterial settlement cues for sessile organisms is well documented

however little is known about the diversity and distribution of the bacteria on the surface of

these organisms. This chapter has demonstrated that bacterial isolates from different surfaces

in the marine environment vary with respect to their ability to inhibit the settlement of

invertebrate larvae. Overall, isolates from living surfaces such as marine algae and sessile

animals were more active in preventing settlement as compared to isolates from rock surfaces

(Table 6.1). These data suggest that a high frequency of bacteria on living surfaces are able to

regulate biofouling and indicate that the marine surface environment is niche-specific with

respect to bacterial strains that colonise specific surfaces.

A correlation between the expression of pigmentation and the ability for bacteria to inhibit

settlement of invertebrate larvae was also observed. High proportions of dark-pigmented

bacteria were found to have a negative effect on larval settlement (Figure 6.1). This finding

nicely correlates with the results presented in chapter 4, which demonstrate that transposon

mutants of P. tunicata defective in pigment production are also defective in antifouling traits.

Of the pigmented strains having strong activity against B. amphitrite, five isolated from the

green alga U. lactuca (designated UL1, UL12, UL13, UL14 and UL15) were selected at

random for further study. The extent to which these isolates were able to inhibit a wider

variety of common fouling organisms was assessed using previously established bioassays.

The data presented in Table 6.2 show the percentage germination of algal spores when

exposed to biofilms of the different bacterial isolates. Isolates UL1, UL14 and UL15

inhibited the germination of algal spores as compared to controls containing seawater alone

and biofilms of the non-inhibitory marine strain J1. Isolates UL12 and UL13 were inhibitory

208

to the germination of U. lactuca spores, however compared to isolates UL1, UL14 and UL15

they showed a reduced inhibitory activity against the germination of spores from Polysiphonia

sp. In addition, spores of U. lactuca are more sensitive towards the inhibitory activity of these

bacterial isolates compared with the spores of Polysiphonia. Further variation in the pattern of

antifouling properties can be seen with respect to the anti-fungal activity. As presented in

Table 6.3, isolates UL12 and UL13 showed no effect on the growth of three different fungi,

while the other three isolates displayed strong growth inhibition against all three fungi. These

variations in the pattern of inhibition might be due to differences in the quantity of compound

produced or to the presence of different or modified antifouling compounds produced by the

isolates.

Table 6.4 summarises the effect of each isolate upon the growth of twenty different bacterial

strains. Isolates UL1, UL14 and UL15 inhibited 90 % of the bacteria tested, including a

variety of Gram-positive, Gram-negative, marine, terrestrial and pathogenic organisms. These

isolates are also strongly inhibitory against their own growth and against the growth of other

isolates from U. lactuca. Autoinhibitory activity has been observed for P. tunicata. In this

case, as with the U. lactuca isolates, it is not yet known how the bacteria survive despite the

production of an autoinhibitory factor. However, it has been demonstrated that as the P.

tunicata cells progress into stationary growth phase they become increasingly resistant

towards the effects of an anti-bacterial protein which is also produced during stationary phase

of growth (James et al., 1996). As the U. lactuca isolates are clearly able to survive at high

cell densities within colonies on an agar plate it is likely that a similar growth-phase dependent

mechanism of resistance occurs for these bacteria.

From an ecological perspective the inhibition of other marine bacterial epiphytes competing

for the same surface (i.e. U. lactuca) would give a selective advantage during colonisation.

Furthermore, the inhibition of closely related species in a growth-phase dependent fashion

prevents a pre-existing bacterial population from being out-competed by faster growing cells.

The less pronounced inhibitory effect against other non-marine and marine bacteria (see Table

6.3), can therefore be seen in the context of not competing for the same niche. However, other

isolates (UL12 and UL13) from U. lactuca displayed in general weak or no activity against

the majority of the target bacterial strains including themselves, indicating that more complex

interactions and mechanisms are involved in the development of a bacterial surface

community. In a study by Maki et al (1990) it was observed that cells of the same bacterial

strain attached to different surfaces caused altered settlement responses by barnacle larvae.

Therefore, it is also possible that changes in the physiology of the U. lactuca isolates when

209

growing on the algal surface may have a major influence upon the extent by which the bacteria

display antifouling activity.

To identify the U. lactuca isolates and to determine their phylogenetic relationship,

phylogenetic analysis based on 16S ribosomal DNA was carried out. Results of the analysis

revealed a close affiliation of all isolates with members of the genus Pseudoalteromonas

(Figure 6.2). The isolates were most closely related to the species P. tunicata, with UL12 and

UL13 representing a distinct phylogenetic group from the group containing the isolates UL1,

UL14 and UL15. This classification correlates with the phenotypic characterisation showing

that these two groups have different properties with respect to their antifouling activities (see

above).

As discussed in chapter 1 the genus Pseudoalteromonas contains many species that produce

biologically active molecules and have been found to live in association with higher organisms

(Holmström and Kjelleberg, 1999). P. tunicata in particular has been studied for its ability to

influence the behaviour of higher organisms. This strain was originally isolated from the

surface of a tunicate (Ciona intestinalis) in Sweden (Holmström et al., 1998). The data

presented in this chapter show that close relatives of P. tunicata species can also be isolated

from the surface of the common green alga U. lactuca in Australia, indicating that this group

of organisms may be widely distributed in a range of marine environments. Both the tunicate

C. intestinalis and the alga U. lactuca, unlike many other sessile marine plants and animals,

have not been reported to produce secondary metabolites for their protection against fouling

processes. It is possible that the success of these organisms in remaining free from fouling is

due to the colonisation of surface-associated antifouling bacteria such as P. tunicata and the

isolates used in this study.

Interestingly, other Pseudoalteromonas species have been described to posses a range of

biological activities including anti-bacterial (Gauthier, 1979; Gauthier and Breittmayer, 1979;

McCarthy et al., 1994), agarolytic (Akagawa-Matsushita et al., 1992; Vera et al., 1998) and

algicidal (Imai et al., 1995; Lovejoy et al., 1998) traits. This study demonstrates a link

between the phylogenetic assignment and the diversity of biological activities for bacterial

epiphytes with pronounced antifouling properties. The bacterial isolates collected from the

green alga U. lactuca were found to be members of potentially new species of

Pseudoalteromonas most closely related to P. tunicata. It would appear that P. tunicata and

closely related strains exist in association with different eukaryotic hosts and in different

210

geographical waters, suggesting that members of this genus may be present in marine

environments as successful and beneficial colonisers of living surfaces.

The following chapter of this thesis details the phenotypic and genotypic characterisation of

isolates UL12 and UL13 for the purpose of taxonomic assignment as a new species of

Pseudoalteromonas with antifouling properties.

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7. Characterisation of Pseudoalteromonas ulvae, a bacterium with

antifouling activities

7.1. Introduction

Bacteria frequently cultured from marine environments are Gram-negative, heterotrophic and

motile by the use of flagella. These bacteria have traditionally been divided into two groups

based on their ability to ferment sugars. The fermentative group include members of the

genera Vibrio, Photobacterium, Aeromonas, Listonella and Colwellia and among the non-

fermentative group are members of the genera Alteromonas, Pseudomonas, Alcaligenes,

Halomonas, Deleya, Halomonas, Marinomonas, Shewanella and Flavobacterium (Kita-

Tsukamota et al., 1993).

With the increasing use of molecular techniques such as 16S and 23S rDNA sequencing and

DNA-DNA hybridisation to determine the relatedness of bacteria, many of the traditional

groups have been reclassified. The genus Alteromonas is a recent example of these changes.

Originally the genus Alteromonas contained four species, A. macleodii. A. vaga, A. communis

and A. haloplanktis but was later used for any marine, Gram-negative, heterotrophic bacterium

which differed from Pseudomonas species by having a lower G+C content (38-50%

compared with 55-64%). As a result before the reclassification there were 14 species

assigned to this highly heterogeneous group, including a former member of the genus

Pseudomonas, P. piscicida (Gauthier et al., 1995). Based initially on rRNA-DNA

hybridisation studies (Van Landschoot and De Ley, 1983) and then followed with 16S rDNA

phylogenetic analysis (Gauthier et al., 1995), most members of the group Alteromonas have

been reclassified within the new genus Pseudoalteromonas, leaving A. macleodii as the only

species of Alteromonas.

The genus Pseudoalteromonas currently includes both pigmented and non-pigmented, Gram-

negative, rod-shaped, heterotrophic marine bacteria that are motile by means of polar flagella.

Species of the genus Pseudoalteromonas are frequently isolated from marine waters around

the world, the majority of which appear to be associated with eukaryotic hosts (Holmström

and Kjelleberg, 1999). Species have been isolated from various animals such as tunicates

212

(Holmström et al., 1998), mussels (Ivanova et al., 1998; Ivanova et al., 1996), pufferfish

(Simidu et al., 1990), sponges (Ivanova et al., 1998) and from a range of marine algae

(Akagawa-Matsushita et al., 1992; Yoshikawa et al., 1997). The two bacterial strains (UL12

and UL13) characterised in this study were isolated from the surface of the marine alga Ulva

lactuca. Both strains have been shown to inhibit the settlement of larvae from the marine

invertebrate Balanus amphitrite and the germination of spores from the green alga U. lactuca

and spores from a species of the red alga Polysiphonia (Chapter 6). The aim of this chapter

was to describe by phenotypic and genetic characterisation the isolates designated UL12T (T =

type strain) and ULl3 for the purpose of taxonomic assignment.

7.2. Materials and Methods

7.2.1. Source of inoculum and isolation

Isolation of the bacterial strains UL12T and UL13 was performed as described in section

6.2.1. Specifically, UL12T and UL13 were isolated from the surface of the common marine

alga U. lactuca, which was collected from the rocky intertidal zone near Sydney, on the east

coast of Australia. The algal thallus was suspended into sterile NSS (Appendix I) and surface

bacteria removed by vortexing. Aliquots of the samples were then spread on the complex

marine medium VNSS agar (Appendix I) and incubated at 23 oC for 48 h. The type strain

UL12T has been deposited in the Culture Collection of the University of New South Wales as

UNSW 095600 T and in the National Collection of Industrial and Marine Bacteria, Aberdeen,

Scotland as NCIMB 13762 T.

7.2.2. Phenotypic characterisation

The morphological and biochemical properties of isolates UL12T and UL13 were determined

using the tests described below and a fresh bacterial inoculum. The bacterial strains were

routinely prepared by inoculating cells in VNSS medium (Appendix I) and incubating at 23oC

overnight.

Oxidative or fermentative utilisation of glucose was determined by the method of Hugh and

Leifson (Hugh and Leifson, 1953). Catalase activity was determined by the method of

Skerman (Skerman, 1967) and oxidase activity tested using the Kovacs method (Kovacs,

1956).

213

The optimal growth condition of isolates UL12T and UL13 with respect to salt concentration

was tested using the medium VNSS with NaCl concentrations ranging from 0 to 10 % (w/v).

Growth on the rich medium, Luria broth (LB20) (Appendix I) and Tryptone soy broth (TSB)

(Oxoid), was also assessed. The marine minimal medium (MMM) (Appendix I) was used

during tests for growth of isolates on different substrates as sole carbon and energy sources at

the concentration of 4 g l-1.

The susceptibility of UL12T and UL13 to the antibiotics gentamicin, tetracycline, ampicillin,

kanamycin, streptomycin, carbenicillum, chloramphenicol, spectinomycin and penicillin G was

tested at concentrations of 50 µg ml-1 and 100 µg ml-1 in VNSS medium. Sensitivity to the

vibriostatic agent 0/129 was tested using disks at a concentration of 150 µg ml-1.

Arginine dihydrolase, tryptophane desaminase, lysine decarboxylase and ornithine

decarboxylase activities were determined using the API 20E system as described by the

manufacturer (bioMérieux). Exponential-phase bacterial cells were washed three times with

MMM before being inoculated into the test cupules.

Motility was determined by visualisation of cells under phase microscopy with a 100 x oil-

immersion objective.

7.2.3. Negative staining and electron microscopy

Cell morphology, size and flagella characteristics were determined by transmission electron

microscopy. One drop of cell suspension from an overnight bacterial culture was mixed with

a drop of sodium phosphotungstate (2 % aqueous) for 30 sec on a Formvar 300-square

copper grid. The grid was blotted using filter paper and dried for 10 min before examination

on a Hitachi H7000 electron microscope at 10 000 x magnification.

7.2.4. 16S rDNA amplification, sequencing and phylogenetic analysis

Extraction of genomic DNA, 16S rDNA amplification and sequencing of this gene was

performed as indicated in section 6.2.3. DNA sequences were aligned using the multiple

sequence alignment tools CLUSTAL W and PILEUP (GCG software packages) (Wisconsin,

1992). Ambiguous gap positions were manually deleted and the alignment was confirmed

and checked against both primary and secondary structure considerations of the 16S rRNA

214

molecule. The aligned sequences were applied to genetic distance and maximum parsimony

methods for phylogenetic inference. Genetic distances were calculated using the formula of

Jukes and Cantor (1969), Kimura (1980) and maximum likelihood (Felsenstein, 1981).

Phylogenetic inference protocols, EDNAML, EDNADIST, NEIGHBOR, EDNAPARS,

CONSENSE and SEQBOOT were supplied by the PHYLIP packages (version 3.57c)

(Felsenstein, 1989). All sequence manipulation and phylogeny programs were made available

through the Australian National Genome Information Service (ANGIS, Sydney, Australia).

7.2.5. Nucleotide sequence accession numbers

EMBL/GenBank/RDP accession numbers (in parentheses) for all small subunit rDNA

sequences of strains other than UL12T and UL13 used in this study are as follows:

Alteromonas macleodii IAM 12920T (X82145), “Marinobacter articus” (AF148811),

Moritella japonica strain DSKI (D21224), Pseudoalteromonas antarctica CECT 4664T

(X98336), Pseudoalteromonas atlantica IAM12927 (X82134), Pseudoalteromonas aurantia

ATCC 33046T (X82135), Pseudoalteromonas bacteriolytica IAM 14594T (D89929),

Pseudoalteromonas carrageenovora IAM 12662T (X82136), Pseudoalteromonas citrea

NCIMB 1889T (X82137), Pseudoalteromonas denitrificans ATCC 43337T (X82138),

Pseudoalteromonas distincta KMM 638T (X82142), Pseudoalteromonas elyakovii KMM

162T (AF082562), Pseudoalteromonas espejiana NCIMB 2127T (X82143),

“Pseudoalteromonas gracilis” strain B9 (AF038846), Pseudoalteromonas haloplanktis

subsp. haloplanktis ATCC 14393T (X67024), Pseudoalteromonas luteoviolacea NCIMB

1893T (X82144), Pseudoalteromonas nigrifaciens NCIMB 8614T (X82135),

Pseudoalteromonas peptidolytica F12-50-A1T (AF007286), Pseudoalteromonas piscicida

ATCC 15057T (X82215), Pseudoalteromonas prydzensis ACAM 620T (U85855),

Pseudoalteromonas rubra ATCC 29570T (X82147), Pseudoalteromonas haloplanktis subsp.

tetraodonis IAM 14160T (X82139), Pseudoalteromonas tunicata CCUG 26757T (Z25522),

Pseudoalteromonas undina NCIMB 2128T (X82140), Pseudoalteromonas sp. IC006

(U85856), Pseudoalteromonas sp. IC013 (U85859), Pseudoalteromonas sp. MB6-05

(U85860), Pseudoalteromonas sp. MB6-03 (U85857), Pseudoalteromonas sp. MB8-02

(U85858), Pseudoalteromonas sp. SW08 (U85861), Pseudoalteromonas sp. S9 (U80834),

Pseudoalteromonas sp. Y (AF030381), Pseudoalteromonas sp. ANG.RO2 (AF022407),

Shewanella putrefaciens ATCC 8071T (X82133), Aeromonas hydrophila ATCC 7966T

(X60408), Photobacterium phosphoreum ATCC 11040T (X74687), Vibrio fischeri ATCC

7744T (X74702), Vibrio alginolyticus ATCC 17749T (X74690) and Salinivibrio costicola

215

NCIMB 701T (X95527). Culture collection designations are: ATCC, American Type Culture

Collection, Rockville, MD; ACAM, Australian Collection of Antartic Microorganisms,

Antartic CRC, Hobart, Australia; NCIMB, National Collection of Industrial and Marine

Bacteria, Aberdeen, Scotland; IAM, Institute of Applied Microbiology, Tokyo, Japan; KMM,

Collection of Marine Microorganisms, Pacific Institute of Bioorganic Chemistry, Vladivostok,

Russia; CCUG, Culture Collection of the University of Göteborg, Sweden.

7.2.6. DNA-DNA hybridisation.

Levels of genomic relatedness were determined by performing DNA-DNA dot blot

hybridisations with radioactively labelled genomic DNA. Target genomic DNA was

denatured by boiling for 10 min and then quickly chilling on ice. Duplicate aliquots

containing 50 ng of denatured genomic DNA from P. aurantia, P. citrea, P. luteoviolacea, P.

piscicida, P. rubra, P. tunicata and the isolates UL12 T and UL13 were dotted onto Hybond-

N+ nylon membranes (Amersham Pharmacia Biotech). Membranes were allowed to air dry

and the DNA was subsequently fixed by UV cross-linking. Prehybridisation was performed

at 42 oC for 1 h in Rapid-hyb buffer (Amersham Pharmacia Biotech). Genomic DNA of

strain UL12 T was labelled by nick translation (Rigby et al., 1977) using a Nick translation kit

(Roche) and Redivue [α32 P] dCTP (Amersham Pharmacia Biotech). Hybridisations were

performed in the prehybridisation buffer with 10 ng ml-1 labelled probe at 42 oC for 16 h.

After hybridisation the membranes were washed once in 2 x SSC (1 x SSC is 0.15 M, NaCl,

0.015 M sodium citrate pH 7), 0.1 % (w/v) SDS at room temperature for 20 min, followed by

twice in 0.5 x SSC, 0.1 % (w/v) SDS at 55 oC for 15 min and a final high stringency wash in

0.5 x SSC, 0.1 % (w/v) SDS at 65 oC for 15 min. The degree of probe binding was

determined by exposing the membrane to a phosophoimager-screen (BioRad) overnight,

thereafter the images were captured with a BioRad GS425 imager (greater than three log

signal response linearity). Image analysis was performed using the BioRad software package

Multi-Analyst. The signal produced by hybridisation of the probe to the homologous target

DNA was taken to be 100 % and the percentage hybridisation for each of the test species were

calculated for the duplicate dots. Hybridisation experiments were repeated twice.

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7.3. Results and Discussion

7.3.1. Biochemical and phenotypical characterisation of UL12 T and UL13

The two strains (UL12T and UL13) isolated from the surface of the marine alga Ulva lactuca,

appeared as small, regular dark purple colonies on VNSS agar and were Gram negative, motile

rods as viewed under phase microscopy. The cells examined by electron microscopy were

1.7-2.5 µm in long, 1-1.5 µm wide and possessed a single polar flagellum (Figure 7.1).

Isolates UL12 T and UL13 were found to be identical with respect to specific physiological

and biochemical features (Table 7.1). Both isolates are strict aerobes that are capable of

growth at 4 oC but not at 35 oC. The optimum temperature for growth was found to be 23 oC

and both isolates could grow within a pH range of 5.5 to 12 (optimum at pH 8). The isolates

required sodium ions at a concentration of 0.1 % (w/v) NaCl with the optimum concentration

for growth being 1-2.5 % (w/v) NaCl.

The isolates exhibited gelatinase and tryptophane desaminase activity while β-galactosidase,

arginine dihydrolase, lysine-ornithine decarboxylase and urease activity were not detected.

Growth on different carbon and energy sources showed that the bacterium utilises citrate,

maltose and Tween 20 within 2 days incubation, L- proline after 4 days incubation, and

glucose and mannose after 7 days incubation. UL12 T and UL13 were unable to ferment

sugar and displayed little or no oxidative acid production as demonstrated by the Hugh and

Leifson test. In addition the isolates were positive for both catalase and oxidase activity.

Cells were sensitive to gentamicin, tetracycline, ampicillin, kanamycin, streptomycin,

carbenicillium, chloramphenicol and spectinomycin at concentrations of 50 µg ml-1 but was

resistant to penicillin G at concentrations up to 100 µg ml-1. Both isolates were sensitive to

the vibriostatic agent 0/129 at a concentration of 150 µg ml-1.

After 3 days incubation on LB20 and TSB media the isolates UL12T and UL13 grew as small

white colonies. Streaking the white colonies from an LB20 or TSB agar plate onto an agar

plate containing VNSS media resulted in the formation of dark purple colonies after 24 hours

incubation. Variations in the level of pigment expression depending on the growth medium is

a characteristic shared by other related species such as Pseudoalteromonas tunicata

(Holmström et al., 1998), P. nigrifaciens (Ivanova et al., 1996), P. denitrificans (Enger et al.,

1987) and Shewanella hanedai (Baumann et al., 1984).

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Table 7.1: Phenotypic characterisation of Pseudoalteromonas ulvae UL12T

Characteristic Phenotype

Pigmentation PurpleGrowth at 4 °C +Growth at 37 °C -Optimal NaCl concentration (%) 1-2.5Optimal pH 8Hugh-Leifson test no reaction or

oxidativeProduction of: Beta-galactosidase activity - Tryptophane desaminase activity * + Arginine dihydrolase activity * - Lysine decarboxylase activity * - Ornithine decarboxylase activity * -Hydrolysis of: Urease - Gelatin +Utilisation of: Citrate, maltose + Mannose, glucose +§ Trehalose, fructose, xylose, arabinose, - lactose, raffinose, melibiose, glycerol, sucrose, sorbitol, erythritol, rhamnose, cellobiose DL-serine - L-glutamine - L-proline + Tween 20 +Oxidase +Catalase +Motility +

-, negative; +, positive.* Tests using the API 20E system.§ Growth after 7 days.

Growth of isolates UL12T and UL13 in liquid culture often results in the aggregation of cells.

While this phenotype was not studied in greater detail it is likely to be related to the

production of an extracellular polysaccharide. It is interesting to speculate the role of

extracellular polysaccharide production for this organism in its natural environment.

Extracellular polysaccharides are known to be involved in bacterial adhesion and biofilm

formation (see chapter 1) and perhaps UL12T and UL13 strains utilise this phenotype for the

establishment and growth as biofilms on surfaces in the marine environment.

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7.3.2. Genotypic characterisation

The DNA sequencing strategy used in this investigation generated 1355 and 1403 bases of

the 16S rRNA gene for UL12T and UL13, respectively (see section 6.3.6). The resulting

sequences were aligned with other closely related 16S rRNA gene sequences within the

EMBL, GenBank, RDP databases. The derived multiple sequence alignment (1191

characters), was used to generate pair-wise sequence identity and genetic distances between

UL12T, UL13 and related bacteria. Several phylogenetic trees were constructed using

different methods, including genetic distance matrices and maximum parsimony. Statistical

evaluation of the derived genetic divergences was performed by bootstrap resampling (100

replicates) of the sequence data.

The tree topology shown in Figure 7.2 was identical to other statistical representations of the

sequence data. The strains UL12T and UL13 were found to belong to the gamma-3 subclass

of Proteobacteria and in a lineage with members of the genus Pseudoalteromonas as

supported by high bootstrap values for the cluster (Figure 7.2). The isolates shared 16S

rRNA similarity of 99.8%, while within the Pseudoalteromonas genus both had a range of

sequence identities between 91% and 97% with other members included in this analysis. The

highest sequence identity outside of this genus were 92% for members of the genera Vibrio

and Photobacterium and 91% identical to the genus Shewanella. The highest 16S rDNA

sequence identity that the isolates shared with other species was 97% with P. piscicida.

However, phenotypically UL12T shares a greater number of characteristics with P. tunicata.

When comparing a number of different taxonomic parameters (Table 7.2) both UL12T and

UL13 differed form the P. tunicata type strain by only 4 traits. They include the inability to

grow at 35 oC, differences in pigmentation, the ability to utilise citrate and the inability to use

trehalose as a sole carbon source. The comparison with UL12T and other strains of

Pseudoalteromonas in Table 7.2 shows that most strains differed by 4 to 8 characteristics.

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Figure 7.1: Electron micrograph of strain UL12T (= UNSW 095600). Bar equals 1000 nm

220

Table 7.2: Differential characteristics of Pseudoalteromonas species

Characteristic UL

12T

P.

tunicata

P.

piscicida

P.

undina

P.

rubra

P.

citrea

P.

aurantia

P.

luteoviolacea

Growth 4 ˚C + + - d - - + -

Growth 35 ˚C - + + - + d - +

Pigmentation P G Y - R Y Y P

Utilisation of:

Mannose + + + - + + + -

Sucrose - - + + - - - -

Maltose + + + + - - d +

Sorbitol - - ND - - - - -

Fructose - - + - - + + -

Citrate + - + - - - - -

Glycerol - - - - - - - -

Lactose - - ND - - - - -

Melibiose - - ND - ND - - -

Trehalose - + ND + + + + +

L-Proline + + ND d ND - - +

Data from (Baumann et al., 1984; Gauthier, 1982; Hansen et al., 1965; Holmström et al., 1998).

+,positive; -, negative; d, 11-89% of the strains are positive; ND, not determined. P = purple pigmentation;

G = green pigmentation; Y = yellow pigmentation; R = red pigmentation. Strain UL13 has the same

characteristics as UL12T. See text for type strain numbers.

In view of the high 16S rDNA sequence similarity between UL12 T and UL13 with other

Pseudoalteromonas species, DNA-DNA hybridisation studies were performed between these

isolates and close phylogenetic neighbours. Both UL12 T and UL13 were found to have low

hybridisation levels (13.6 to 28.5 %) with other Pseudoalteromonas species including P.

aurantia ATCC 33046T, P. citrea NCIMB 1889T, P. luteoviolacea NCIMB 1893T, P.

piscicida ATCC 15057T, P. rubra ATCC 29570T and P. tunicata CCUG 26757T. These

values are below the currently accepted limit of DNA-relatedness (70%) for the phylogenetic

definition of a species (Stackebrandt and Goebel, 1994) and therefore give further evidence

that the isolates represent a novel species with the genus Pseudoalteromonas. In contrast,

hybridisation levels of 68.5 % were found between the isolates UL12 T and UL13. This value,

while being slightly below that of 70% limit for species delineation when taken together with

221

the high 16S rDNA sequence similarity and the phenotypic similarities between the isolates,

suggests that UL12 T and UL13 may belong to the same species.

7.3.3. Assignment of UL12T and UL13 for a new species

The marine isolates UL12T and UL13 are different from previously characterised

Pseudoalteromonas species. The 16S rDNA sequence together with the DNA relatedness

values clearly shows that isolates UL12T and UL13 make up a novel species within the

Pseudoalteromonas genus. Both isolates display phenotypic and biochemical characteristics

typical for Pseudoalteromonas species, including requirement for sodium ions, motility by a

single flagellum, oxidase positive, catalase positive, gelatinase activity and oxidative

metabolism. The main phenotypic features of isolates UL12T and UL13 closely resemble

those of P. tunicata. However, in addition to the features listed in Table 7.2, both isolates can

be distinguished from P. tunicata by the lack of a sheathed flagellum and a strict aerobic

metabolism in isolates UL12 T and UL13 (Holmström et al., 1998). Therefore, on the basis of

phenotypic and genetic characterisation isolates UL12T and UL13 can be considered as a

distinct new species for which the name Pseudoalteromonas ulvae sp.nov. is proposed.

Figure 7.2: Distance matrix tree based on a 1191 base pair sequence alignment of the 16S

rDNA gene of the isolates UL12T and UL13 (P. ulvae sp. nov.), with members of the genus

Pseudoalteromonas and other closely related bacteria. Distances were calculated according to

the algorithm of Jukes and Cantor (1969) and trees constructed by the Neighbor-Joining

method of Saitou and Nei (1987). Marinobacter arcticus was chosen as the outgroup. Bar

indicates 1 substitution per 100 nucleotide positions. Bootstrap values (100 replicates) are

indicated at branching points.

222

0.01

Shewanella putrefaciens

Aeromonas hydrophila

Moritella japonica

Alteromonas macleodii

Salinivibrio costicola

Vibrio alginolyticus

Vibrio fischeri

Photobacterium phosphoreum

Pseudoalteromonas denitrificans

Pseudoalteromonas tunicata

Pseudoalteromonas ulvae UL12T

Pseudoalteromonas ulvae UL13

Pseudoalteromonas aurantia

Pseudoalteromonas citrea

Pseudoalteromonas sp. S9

Pseudoalteromonas luteoviolacea

Pseudoalteromonas rubra

Pseudoalteromonas peptidolytica

Pseudoalteromonas sp. Y

Pseudoalteromonas piscicida

Pseudoalteromonas sp. ANG. RO2

Pseudoalteromonas prydzensis

Pseudoalteromonas sp. MB8-02

Pseudoalteromonas undina

Pseudoalteromonas sp. SW08

Pseudoalteromonas nigrifaciens

Pseudoalteromonas haloplanktis subsp. haloplanktis

“Pseudoalteromonas gracilis”

Pseudoalteromonas sp. MB6-05

Pseudoalteromonas sp. IC006

Pseudoalteromonas antarctica

Pseudoalteromonas haloplanktis subsp. tetraodonis

Pseudoalteromonas atlantica

Pseudoalteromonas espejiana

Pseudoalteromonas carrageenovora

Pseudoalteromonas elyakovii

Pseudoalteromonas distincta

Pseudoalteromonas sp. IC013

Pseudoalteromonas sp. MB6-03

Marinobacter articus

Pseudoalteromonas bacteriolytica

100100

100

6482

69 100

58

47

57

99

100

69

48

51

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7.3.4. Description of Pseudoalteromonas ulvae sp. nov.

Pseudoalteromonas ulvae (ul.’vae. L. gen. n. ulvae of Ulva, the generic name of the host alga

U. lactuca.)

Strict aerobic, Gram-negative rod shaped cells that are 1.75-2.5 µm in length and 1-1.5 µm in

width. Cells are motile by means of a single polar flagellum. Growth on VNSS medium

results in small dark purple colonies, while growth on LB20 or TSB with 2 % NaCl results in

the formation of white colonies. Does not ferment sugar in the Hugh-Leifson test. Sodium

ions are required for growth with the optimum NaCl concentration being 1-2.5 %. Can grow

within a pH range of pH 5.5 to 12; the optimum pH for growth is pH 8. Slow growth occurs

at 4 oC and no growth is detectable at 35 oC. Oxidase and catalase positive. Utilises citrate,

maltose, L- proline, glucose, mannose and Tween 20 but can not use trehalose, lactose,

sucrose, fructose, glycerol, raffinose, sorbitol, melibiose, xylose, cellobiose, erythritol, L-

glutamine, arabinose, rhaminose or DL-serine as sole carbon and energy sources. Positive for

hydrolysis of gelatin and displays tryptophane desaminase activity. Strains UL12T and UL13

are negative for H2S production and no β-galactosidase, arginine dihydrolase, lysine

decarboxylase, ornithine decarboxylase or urease activities have been detected. Sensitive to

tetracycline, ampicillin, kanamycin, streptomycin, carbenicillium, chloramphenicol and

spectinomycin at concentrations of 50 µg ml-1 and sensitive to the vibriostatic agent 0/129 at a

concentration of 150 µg ml-1. Cells were resistant to Penicillin G up to 100 µg ml-1

concentration. Isolated from the surface of a marine alga, Ulva lactuca, collected from the

rocky intertidal zone off the eastern coast of Australia. The type strain UL12T has been

deposited in the Culture Collection of the University of New South Wales, Sydney, Australia

as strain UNSW 095600T and the National Collection of Industrial and Marine Bacteria,

Aberdeen, Scotland as strain NCIMB 13762 T. Nucleotide sequences for the 16S rDNA

genes of isolates UL12T and UL13 have been deposited in the DDBJ/EMBL/GenBank

database under the accession numbers AF172987 and AF172988 respectively.

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8. General discussion

The work presented in this thesis has investigated the anti-fouling properties of marine

bacteria. As the primary colonisers of a surface, bacteria play an important role in the

development and maintenance of a biofouling community. The marine bacterium

Pseudoalteromonas tunicata was chosen as the model organism for these studies. P. tunicata

cells inhibit the settlement and growth of a number of common fouling organisms including,

invertebrate larvae, bacteria, algal spores and fungi (Holmström et al., 1998). The means by

which P. tunicata cells inhibit algal spore germination and fungal growth were addressed in

chapters 2 and 3 of this thesis. These studies were successful in characterising these activities

and provided evidence that each of the antifouling activities are due to the production of

separate and target specific molecules. The correlation between the production of pigments

and inhibitory compounds in P. tunicata was investigated in chapters 4 and 5 by the

generation and analysis of transposon mutants altered in wild-type pigmentation. The study

of these mutants lead to the identification of genes involved in the synthesis and regulation of

pigment and specific inhibitors. Finally, while is it widely accepted that marine surface

bacteria can influence the colonisation of other fouling organisms, little information is

available regarding the prevalence and diversity of these bacteria. To examine this, a collection

of bacterial isolates from different marine surfaces was studied for their antifouling activity

(chapter 6). Interestingly, a number of the inhibitory isolates were found to belong to the

genus Pseudoalteromonas, being most closely related to P. tunicata. Two of these isolates

were characterised as strains of a new species, Pseudoalteromonas ulvae (chapter 7). This

final chapter will summarise and discuss the major findings presented in this thesis and

suggests of the directions for future work.

8.1. Antifouling and biocontrol properties of Pseudoalteromonas

tunicata

P. tunicata has been previously studied for its ability to inhibit a range of common fouling

organisms by way of specific extracellular inhibitory molecules. These inhibitors include a

polar, heat stable, anti-larval molecule of less than 500 Da (Holmström et al., 1992) and a 190

kDa anti-bacterial protein (James et al., 1996). In addition to these compounds P. tunicata is

225

known to be inhibitory towards diatoms, algal spores and fungi (Holmström et al., 1996;

James, 1998). The first and second aims of this thesis were to investigate the anti-algal and

anti-fungal activities, respectively and provide information on the nature of the active

compounds.

With respect to the anti-algal activity it was demonstrated that a high proportion (23 %) of

marine surface-associated bacteria are able to inhibit the germination of algal spores. The

level of inhibition varied between different bacterial isolates, with P. tunicata being the most

effective. Other authors have also demonstrated that marine bacteria are capable of inhibiting

the settlement of marine plants and animals (Berland et al., 1972; Maki et al., 1988; Mary et

al., 1993; Thomas and Allsopp, 1983), however few studies have attempted to determine the

cause of inhibition. A possible reason for this is the difficulty associated with the bioassay

used to determine algal spore settlement/ germination. The major limitation of the bioassay is

its dependence on seasonal and field conditions. Spores are collected from fertile algae taken

directly from the environment. Therefore, the success of the bioassay is dependent on weather

conditions and the availability of fertile algae. Even when fertile algae have been collected,

sporulation may not occur when the algae are taken into the laboratory. Despite these

limitations this study has successfully characterised the anti-algal compound from P. tunicata

as an extracellular, heat sensitive and polar compound between 3 and 10 kDa in size. These

results also provide the first evidence that the anti-algal compound is distinct from the

compounds that are active against larval settlement and bacterial growth. Future studies will

involve further characterisation of the active component to elucidate its exact chemical

structure.

The anti-fungal activity of P. tunicata was examined in chapter 3. The effectiveness of the

compound as a broad-spectrum fungicide was demonstrated by its ability to inhibit the growth

of a variety of both ecologically and medically important fungi. To characterise the active

compound a multi-disciplinary approach was used, including genetic and chemical analysis

together with established bioassays.

Transposon mutants of P. tunicata defective in the ability to inhibit fungal growth were

generated and these mutants were used to both guide the chemical analysis and to identify

genes involved in the synthesis of the active compound. DNA-sequence analysis indicated

that the transposon had disrupted a gene (afaA) with similarity to the gene encoding a long-

chain fatty-acid CoA ligase (FadD) from E. coli. This protein is responsible in E. coli for the

activation and transport of exogenous fatty-acids into metabolically active CoA thioesters.

226

The activated fatty-acids can then be used in beta-oxidation or are incorporated into cellular

phospholipids.

The anti-fungal compound has successfully been purified and chemical analysis thus far

suggests that the compound consists of a carbon ring bound to a fatty-acid side chain. These

results are exciting as they provide a direct link to the genetic analysis and have allowed for

the establishment of a model for the role of AfaA in the synthesis of the active compound.

The model (detailed in section 3.3) proposes that AfaA is required to activate and transport a

particular fatty-acid from the environment which then forms the fatty-acid side chain of the

active compound. It should be emphasised that the non anti-fungal mutant remains green in

pigmentation and preliminary chemical analysis of its yellow pigment suggests that it is a

similar compound, the only difference being that it has a slightly greater molecular weight than

the wild-type compound. It is possible that without the uptake of exogenous fatty acids, a

different side chain is added to the carbon ring giving the compound its yellow colour but not

the anti-fungal activity.

The data obtained during this study has opened the field for a number of new projects. It will

be most important to define the exact chemical structure of the anti-fungal compound and to

compare this to the non-active compound of the AfaA− mutant. This will not only provide

information regarding the role of AfaA in the formation of the active anti-fungal compound

but will also be useful for commercial applications of this novel compound in the medical or

agricultural industries. In addition, knowledge of the differences between the mutant

compound and the wild-type compound would benefit any future developments that may

involve making chemical variations of the natural compound to improve activity and/ or

stability.

Studies directed towards the elucidation of the mode of action of the anti-fungal compound

are also of interest. One method would be to generate random mutants in a fungal strain and

isolate those mutants that have become resistant to the anti-fungal component.

Characterisation of the disrupted gene will provide information concerning the target of the

anti-fungal compound. For example, a disruption in a cell wall protein may suggest that the

anti-fungal compound is targeting the fungal cell wall. These experiments could be

performed easily with the use of the Saccharomyces cerevisiae mutant library, which has

recently become available (Winzeler et al, 1999). This library, in which all non-essential open

reading frames have been deleted, would prove useful as a tool to screen for the action of the

227

anti-fungal component. If a mutant clone is discovered with increased resistance to the

compound the corresponding gene representing a potential target for the anti-fungal

compound is already known. However, if the anti-fungal compound targets an essential

protein, other methods will need to be applied. For example, labelling the compound and

localisation/ co-purification of the target may be a possible way to initially define the mode of

action of the anti-fungal compound.

The negative environmental impact caused by the use of chemicals to control pests and

disease has stimulated a great interest in the development of biocontrol agents, such as natural

products produced by bacteria. For example, in the agriculture industry strains of

Pseudomonas and Bacillus have been studied and field trials undertaken demonstrating their

effectiveness as biological control agents for fungal plant diseases (Glick and Bashan, 1997;

Ryder and Rovira, 1993). However, in addition to the obvious aim to develop the anti-fungal

compound into a commercial product, an investigation into the ecological relevance for the

synthesis of an anti-fungal metabolite by P. tunicata cells is also of major interest. Fungi are

present in most marine habitats and marine fungi comprise of an estimated 1500 different

species (Hyde et al., 1998). Fungi are important decomposers of woody substrates in marine

ecosystems and may also be important as decomposers of dead organisms. In addition,

many marine fungi have been described as important pathogens of both plants and animals

and to form symbiotic relationships with other marine organisms (Hyde et al., 1998). Given

the prevalence of fungi in the marine habitat it is not surprising that bacteria have developed

methods to control fungal growth. As such the expression of an anti-fungal compound by P.

tunicata may be a mechanism by which the host organism (i.e. tunicates) defends itself from

fungal disease. In addition to the other antifouling molecules expressed by P. tunicata, the

anti-fungal compound may give the bacterium a competitive advantage in the acquisition of

living space and nutrient in the marine surface environment.

Understanding the biological interactions between different marine surface-associated

organisms and the identification of natural biologically-active metabolites will be of great

benefit to applications such as the development of environmentally benign antifouling

methodologies as well as novel biocontrol agents. Specific inhibitory molecules may be

incorporated separately or in combination into paints or coatings applied to water-submerged

structures (eg. ship hulls, fishing nets and cages). Alternatively it may be possible to

immobilise living bacteria into a suitable matrix which can be used as a antifouling coating in

both marine and fresh water system, food preservation or medicine. In a recent study

performed by Holmström et al (2000), E. coli cells maintained their viability for over two

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months when incorporated into a polyvinylalcohol matrix. Given these results it is possible

that “living coatings” will be commonly used in the future to prevent surface fouling. In the

situation where bacterial cells are incorporated into coatings, the approach of identifying the

individual compounds would be complementary and could advance future pesticide and drug

design.

While many potent inhibitory components directed towards fouling organisms have been

isolated from eukaryotic organisms (de Nys et al., 1994; Mizobuchi et al., 1996; Tsukamoto

et al., 1997), the use of bacteria as a source of natural antifouling and biocontrol agents has

many advantages. Most importantly, bacteria are easy to culture at low cost and through the

use of various fermentation technologies the inhibitory agent may be obtained in large

amounts. In terms of molecular biology, most recombinant-DNA techniques can be easily

applied to marine bacteria. Thus, genetic engineering may be employed to further increase the

production of the inhibitory compound either through stimulating the production in the

original organism or by cloning the genes into a new host organism.

8.2. A model for the synthesis and regulation of pigmentation and

fouling inhibitors in P. tunicata

The dark green colour of P. tunicata cells is due to the production of both a yellow and a

purple pigment. A correlation between the expression of pigmentation and the production of

fouling inhibitors lead to the hypothesis that both are tightly linked (see chapter 4). The third

aim of this study was to investigate the regulation of expression of fouling inhibitors and

pigmentation in this organism through the use of transposon mutagenesis. Transposon

mutants altered in wild-type pigmentation were generated and phenotypic characterisation of

these mutants with respect to their antifouling activity demonstrated a link between the

production of the yellow pigment and the inhibitory activities against different target

organisms. Four different categories of pigmented mutants were isolated including yellow,

dark purple, light purple and white phenotypes.

Sequencing and analysis of the genes disrupted by the transposon to give a purple phenotype

revealed open reading frames encoding for enzymes potentially involved in the synthesis (eg.

oxidase, transferase) of the yellow pigment and fouling inhibitors. These enzymes appear to

be clustered in a single operon that also contains a putative ABC transporter. ABC

transporters may function as either exporters or importers (Higgins, 1992), thus it is tempting

229

to speculate that they are involved in the export of the yellow pigment from the cell or may

import a precursor for the synthesis of the yellow pigment.

Two white mutants that do not express pigments and fouling inhibitors were also analysed.

One of these mutants (W3) had been disrupted in an operon encoding for proteins in the

general secretion pathway (GSP). In P. tunicata the GSP may be important for the secretion

of surface-structures or extracellular enzymes required to sense environmental cues or obtain

specific metabolites needed as precursors for pigment/ fouling inhibitor production. A

comparison of the proteins secreted by this mutant and by the wild-type strain demonstrated

differences in the protein profiles, which supports the proposed function of these genes.

The second white mutant (W2) had been disrupted in a gene, (designated wmpR) with

sequence similarity to common transcriptional regulators such as CadC from E. coli and

ToxR from Vibrio cholerae. Both CadC and ToxR are transmembrane DNA-binding

proteins, which function as transcriptional activators allowing for the coordinated control of

protein expression in response to environmental signals. Analysis of global protein

expression using two-dimensional gel electrophoresis (2DGE) in the W2 mutant and in the

wild-type provides the first evidence that WmpR functions as a regulator of protein

expression during stationary phase growth in P. tunicata. Future studies will be aimed at

identifying the proteins regulated by WmpR. While 2DGE is a powerful tool used to study

global changes in protein expression there are certain limitations. In the case of P. tunicata,

when the protein concentration was increased (above 100 µg of total protein) for preparative

2DGE, distortion of the gel image was observed (data not shown). This may be due to the

production of interfering substances in the wild-type, as a scale up of the white mutant protein

sample remains unaffected. Furthermore, while the methods for identifying proteins from a

gel sample have improved greatly, large quantities are often still required. Taken together

these limitations may require that multiple gels will need to be run and samples pooled for the

successful identification of the WmpR controlled proteins. An alternative approach to identify

regulated genes is RNA arbitrarily primed polymerase chain reaction (RAP-PCR) which has

been used to successfully identify genes controlled by the ToxR homologue in

Photobacterium profundum (Bidle and Bartlett, 2001). This method employs random

oligonucleotide priming to create a cDNA fingerprint for a particular bacterial strain in a

certain physiological state. Bands that differ between samples can be isolated and sequenced,

thus differences in the gene expression can easily be identified. However, RAP-PCR also has

limitations, in particular this method is limited by the number of random primers and primer

combinations used (Chakrabortty et al., 2000; Bidle and Bartlett, 2001). Therefore using both

230

2DGE and RAP-PCR may allow for the identification of all genes and proteins regulated by

WmpR.

In addition to identifying the genes and proteins regulated by WmpR it will also be important

to understand the nature of the stimuli required for WmpR activity. The 2DGE analysis

indicates that growth phase plays a role, as the phenotypic expression of pigment and

inhibitors corresponds well with the up-regulation of specific proteins during early-stationary

phase growth. Based on sequence similarity and secondary structure predictions WmpR is

located in the cytoplasmic membrane with a large periplasmic domain, that could potentially

function as a sensor of changes in the external environment. The signals sensed by WmpR

could include pH, nutrient levels or specific environmental cues generated from the host

organism or from neighbouring bacterial cells. In order to elucidate the environmental signals

needed for the activation of WmpR, expression studies can be performed using a green

fluorescent protein (GFP) reporter gene fused to the promoter of genes regulated by WmpR

(identified through the 2DGE studies). By using an unstable derivative of GFP, temporal

changes in the levels of protein expression can be easily monitored under a variety of

environmental conditions (Andersen et al., 1998). Studies with CadC indicate that the activity

and not the expression of the regulator is altered between inducing and non-inducing

conditions (Watson et al., 1992). While experimental evidence is yet to be obtained, it is

possible that this may also apply to WmpR. In similar experiments as described above,

expression studies using a WmpR-GFP fusion system will be useful in determining if

WmpR expression also varies with different environmental conditions.

The study of transposon mutants has lead to the hypothetical model shown in Figure 8.1.

This model proposes that the transcriptional regulator WmpR is located in the cytoplasmic

membrane and is able to sense and respond to environmental cues/ stimuli by up-regulating

the genes for the production of the yellow pigment, fouling inhibitors and purple pigment.

The general secretion pathway (GSP), encoded in part by wmpC and wmpD is required to

secrete extracellular enzymes that are able to degrade substrates in the environment resulting

in the formation of precursors needed for yellow pigment synthesis. These hypothetical

precursors are then taken back into the cell via the ABC transporter encoded by the genes

dppA and dppB. Once in the cytoplasm the precursors are then metabolised by the proposed

biosynthetic proteins (DppC, DppD, LppA and LppB) to yield the yellow pigment and fouling

inhibitors. A number of these biosynthetic proteins have putative oxidase and transferase

functions and may be encoded in one or several gene clusters within the P. tunicata genome.

231

Although not explicitly incorporated into the model, further extension and testing of it will

explain why the light purple mutants have less anti-bacterial protein and how the GSP affects

purple pigment production. One proposal is that an intermediate of the yellow pigment

pathway acts as a regulator by providing a positive feedback loop to WmpR. Thus a

reduction or loss of this intermediate due to a disruption in the biosynthetic pathway or in the

GSP would result in a reduced level of WmpR activation and less anti-bacterial protein and

purple pigment. An alternative explanation is that the yellow and purple pigments have part of

their biosynthetic pathway in common.

Cytoplasm

Extracellular environment

Anti-bacterial protein

Purple pigment

Extracellular enzymes

Precursors for yellow pigment and fouling inhibitors

To yellow pigment and fouling inhibitors

WmpR

ABC transporter

GSP

wmpC wmpD

dppA dppB dppC dppD lppA lppB? ?

? TT O

Figure 8.1: Hypothetical model for the regulation of yellow pigment and fouling inhibitors

in P. tunicata. ? = Unknown gene or protein; T = putative transferase; O = putative oxidase.

See text for further description.

232

8.3. The ecological significance of Pseudoalteromonas species

In the marine environment the competition for living space is intense and as a consequence all

surfaces are potential sites for colonisation by a variety of organisms. To protect themselves

from biofouling, sessile invertebrates and algae have been reported to use a variety of physical

and chemical defences. Physical defences such as the production of mucus and sloughing of

epidermal tissue are common. With respect to the chemical defences many marine animals

and plants are known to use a wide range of secondary metabolites that inhibit surface

colonisation of other organisms (Davis et al., 1991; de Nys et al., 1994; Kon-ya et al., 1994;

Slattery et al., 1997). In addition, they may also rely on the secondary metabolites produced

by surface associated bacteria (Anthoni et al., 1990; Holmström et al., 1992; Kon-ya et al.,

1995; Maki et al., 1990). Thus, biological, physical and chemical interactions between

different surface associated organisms play a major role in the development and maintenance

of a marine biofouling community.

It is well established that bacteria influence the colonisation of fouling organisms however

little is known about the diversity or distribution of these bacteria. The final aim of this thesis

was to assess the frequency of antifouling bacteria isolated from different marine surfaces and

to determine the phylogenetic relationship of the inhibitory isolates. Results of these

experiments demonstrated that bacterial isolates from different surfaces could vary with

respect to their antifouling properties. In general, isolates from living surfaces were found to

be more active in preventing the colonisation of other organisms, indicating that the marine

surface environment is very niche-specific. In addition, a large proportion of the dark-

pigmented isolates had antifouling activity. This is in agreement with the correlation between

pigmentation and the production of fouling inhibitors in P. tunicata and suggests that the link

between pigmentation and antifouling capabilities may not be restricted to P. tunicata. This

finding could be useful for future studies that involve screening for antifouling bacteria.

Five of the dark pigmented isolates originating from the green alga U. lactuca and with strong

antifouling activity were further studied. Phylogenetic analysis based on the 16S RNA gene

revealed that the isolates were members of the genus Pseudoalteromonas and were closely

related to P. tunicata. Detailed phenotypic and genotypic characterisation of two of these

isolates lead to the taxanomic definition of a new species, P. ulvae. P. tunicata was originally

isolated from the surface of a tunicate (Ciona intestinalis) in waters off the coast of Sweden.

The data presented in this thesis indicate that close relatives of P. tunicata can also be isolated

233

from green algae in Australia, suggesting that this group of bacteria may be widely distributed

in a range of marine environments and in association with higher organisms.

Interestingly, both C. intestinalis and U. lactuca have not been reported to produce any

chemical defences for their protection against fouling. It is possible that these organisms are

able to remain free of fouling due to the colonisation of antifouling bacteria such as P.

tunicata, P. ulvae and other isolates identified in this study. The production of fouling

inhibitors by epibiotic bacteria potentially gives them a competitive advantage in the marine

surface environment. Expression of anti-bacterial and anti-fungal compounds will help the

bacterium to out-compete other micro-foulers during the colonisation process. The host

organism may also benefit from the bacterial production of fouling inhibitors, therefore once

established, the production of additional inhibitors that target macro-foulers (eg. invertebrates

and algae) may prevent the bacteria from being removed by these higher organisms. Thus, a

symbiotic relationship may exist between the host and its epibiotic bacteria. In return for

protection from fouling organisms the host provides the bacterium with access to nutrients

and to living space. Other authors have made similar observations and active compounds first

thought to be of eukaryotic origin are now known to be the product of surface-associated

bacteria. For example, many of the active metabolites from bryozoans are produced by their

associated bacteria (Anthoni et al., 1990).

In addition to the isolation and description of a novel Pseudoalteromonas species (P. ulvae),

this study has added to the understanding of the diversity and abundance of antifouling

bacteria and in particular with respect to Pseudoalteromonas species in the marine habitat.

The ecological role of bacteria that produce antifouling compounds is diverse and this is

reflected in the distribution of these bacteria in the marine environment. The discovery that

bacterial strains similar to P. tunicata exist in association with different eukaryotic hosts and

in different geographical waters suggests that they may be present in marine habitats as

successful and beneficial colonisers of living surfaces. Other researchers have provided

evidence for the presence of P. tunicata-like strains in diverse environments. For example, a

bacterial strain closely related to P. tunicata has been isolated from the Huon estuary in

Tasmania, Australia and was identified as having strong algicidal activity against harmful algal

bloom species (Skerratt et al., 1998). In addition, one of the two major phylotypes (based on

16S clone libraries) identified from the analysis of the bacterial community in microbial

biofilm formations in the submerged cave systems of the Nullarbor region of Australia,

showed high sequence identity to P. tunicata (Holmes et a., unpubl.). While the bacterium

corresponding to this phylotype has not been cultured, these results indicate that P. tunicata-

234

like strains can potentially be isolated from much more diverse habitats then previously

thought.

The prevalence of P. tunicata and related strains in a variety of environmental niches has lead

to the development of future projects aimed to further investigate the ecological role of

Pseudoalteromonas species in the marine environment. The first of these projects will involve

a comprehensive study of the distribution of Pseudoalteromonas species on living and

inanimate surfaces on a global scale using both culturing and molecular techniques such as

denaturing gradient gel electrophoresis (DGGE) and fluorescent in situ hybridisation (FISH).

As suggested previously it is possible that the production of extracellular inhibitors by P.

tunicata cells gives this bacterium a selective advantage over other strains during the

colonisation process. It will be of interest in future studies to determine if P. tunicata is able

to influence the structure of the bacterial community both in simple co-culture biofilm

experiments and when introduced to an established complex biofilm community. Preliminary

data from co-culture biofilm experiments with P. tunicata and other marine bacteria suggest

that P. tunicata cells are able to out-compete other strains. Five out of seven unidentified

marine isolates were killed and removed from the biofilm after exposure to P. tunicata

(Holmström et al., unpubl.). Given these results, such experiments could be expanded to

examine the ecological importance of each of the fouling inhibitors by including the various

transposon mutants of P. tunicata.

The diversity of specific biologically active metabolites expressed by P. tunicata and closely

related strains have not been reported for other bacteria and thus may suggest that they are

unique to P. tunicata and related strains. However, the lack of reports may equally reflect that

laboratories generally do not have access to a broad range of bioassays. Recently, a collection

of ten Pseudoalteromonas species have been tested for activity against a variety of bacteria,

fungi, invertebrate larvae and algal spores. Results of these screens show that while several

strains did display antifouling activity, only P. tunicata had an exceptional broad range activity

(Holmström et al., unpubl.).

235

Appendix I

Solutions and Buffers

I.I Nine Salts Solution (NSS) (per litre)

17.6 g NaCl,

1.47 g Na2SO4,

0.08 g NaHCO3,

0.25 g KCl,

0.04 g KBr,

1.87 g MgCl2.6 H2O,

0.41 g CaCl2.2H2O,

0.008 g SrCl.6 H2O,

0.008 g H3BO3,

- Adjust to pH 7

I.II VNSS (per litre NSS) (Marden et al., 1985)

1.0 g peptone,

0.5 g yeast extract,

0.5 g glucose,

0.01 g Fe SO4.7H2O,

0.01 g Na2HPO4

- For agar plates add 15 g agar before autoclaving

I.III Marine Minimal Medium (MMM)

920 ml 1.1 x NSS (i.e. salts for one litre in 920 ml H2O) autoclaved

40 ml 1 M MOPS (pH 8.2) sterile filtered

10 ml 0.4 M Tricine +1mM FeSO4.7H2O (pH 7.8) sterile filtered

10 ml 132 mM K2HPO4 autoclaved (add slowly while stiring)

10 ml 952 mM NH4Cl (pH 7.8) autoclaved

10 ml 400 g / L carbon source stock solution, sterile filtered

236

I.IV Luria Broth (LB) medium (per litre)

LB 10

10 g NaCl,

10 g tryptone,

5 g yeast extract

- Adjust to pH 7.5

- For agar plates add 15 g agar before autoclaving

LB 20

20 g NaCl,

10 g tryptone,

5 g yeast extract

- Adjust to pH 7.5

- For agar plates add 15 g agar before autoclaving

I.V 5 x TBE buffer (per litre)

54 g Tris base

27.5 g boric acid

20 ml 0.5M EDTA solution (pH 8.0)

I.V. 6 x Agarose gel loading buffer

0.25 % bromophenol blue

0.25 % xylene cyanol

30 % glycerol in H2O

- Store at 4 oC

237

Appendix II

Primers (5'- 3')

Ad1 CTA ATA CGA CTC ACT ATA GGG CTC GAG CGG CCG

CCC GGG CAG GT

Ad2 P- ACC TGC CC -NH2

Ap1 GGA TCC TAA TAC GAC TCA CTA TAG GGC

Ap2 AAT AGG GCT CGA GCG GC

Tn10D CCT CGA GCA AGA CGT TTC CCG

Tn10C GCT GAC TTG ACG GGA CGG CG

S1 GGG TAT TCA GGC TGA CCC

FMTnC-S2 ATA CTG TAC TTG ATC GCG G

FMTnC-S4 GGT TTA CCA GCA CCT AGC

FMTnC-S8 TCT TGG CCA TCT TCA CCC

FMTnD-S3 TTT CAC ACC CGT TTT GCC

FMTnD-S5 CAA CCA CAA CGG CTT GCC

FMTnD-S6 TCT GGA AAC CTG TTT AGC

FMTnD-S7 AGT GGC TGT TAT GAT GCC

FMpan1 AGA AGT TGC AAA AGG TGA AGC GG

FMpan1-S2 AAA GGG GCT CAC ACT TGC

FMpan1-S3 AAC TTG TTC ACC ACT GAC C

FMpan1-S4 CAA ACA CTT GGA TAA GGG C

FMpan2 AAC CAG CAC TAT TGG AGC TGG C

FMpan2-S2 ATG TGC TGA CGA ATG GCG

FMpan2-S3 TTC GAT TCT ATT TTG ACC GG

FMpan2-S4 ATG GAC TCT CTG ACT GGC

FMpan2-S5 TGT ACT TAG GGC TGT CGC

Lp2TnC-S7 ATA TGT GCC GAA TTG AGC G

238

Lp2TnC-S8 GCG GGT CGT TAG CTA ACC

Lp2TnC-S9 ACT CCT GCG TCT GAT AGC

Lp2TnC-S10 ATT GAG CAA ATA CAC GCC C

Lp2TnC-S12 ACA GCG TTC AAC TCA GGC

Lp2TnC-S13 TGG ATT AGA CTT GGC AAG C

Lp2TnD-S2 TGC GGT ATC ATC TGG AC

Lp2TnD-S4 GCA TCC CAG CCA TAA TAG G

Lp2TnD-S5 CGC ATA CCA TTG ATT AGG G

Lp2TnD-S6 GCC ACG GTT GAT GAG AGC

Lp2TnD-S11 TTG TGC CAG TTT ATC GCA C

Lp2TnD-S14 TGT CTT GAT GAT CGT TGC C

Lp3TnC-S2 ATC CAA GTT TGC GGT CGG

Lp3TnD-S2 CGT GAT GTT ACC GAT CGG

Lp3TnD-S3 ATC GAC CAG CCG ATC GG

Dp3TnC-S3 TGC GAA TTG GAG ACA CGG

Dp3TnC-S5 AAA GCA CTG AGG TAA ACC G

Dp3TnC-S6 CAA GTA GCC TTT GCA GCG

Dp3TnD-S2 AGA GGT CAG TAT TGA ACG G

Dp3TnD-S4 AGC CGT TGG TGC AAG GG

Dp3TnD-S7 GTT GTC TGA TGG AGG CC

Lp3pan2 CAG CAA TTC GTG AAG AGC AGC G

Lp3pan2-S2 GCA CTA CCT CAG ACT ACG

Lp3pan2-S3 GTT CTG GTG TCC ACG CC

Lp3pan2-S4 TGA CCG CCG CAA ACT CC

W2TnC-S3 CAG CTG TGA CGA CCA GC

W2TnC-S4 TAA CGG TTA TGA TGC TGG C

W2pan1 CGA GGC TAT AAG CTG ATC TGC C

W2pan1-S2 TTA GGC CGA TAA CGC ACC

W2pan1-S3 CTC CAA TTC GCC AAT GGG

W2pan3 TTG GCT GTC TTA GAT GTT GCT CC

239

W2pan3-S2 CTG AGT TGG AAG TGC AGC

W2pan3-S3 TCG TTT TAG CCA AAG CCG

W2pan3-S4 AAG TTA CAA CGG CTT TGG C

W2pan3-S5 TGC ACT TCC AAC TCA GGC

W2pan4 TGG TCG TCA CAG CTG AGC ACC

W2pan4-S2 CTG TCA CCA ATA TGC CGC

W3TnC-S2 TTC GCT TAG TTG ACC AAG C

W3TnC-S3 CTA ATT CGA ACT CGC TCG

W3TnC-S4 ACC GGT TCG CAA ACG GG

W3TnC-S5 AAG CAA ACT AGC AAG GGC

W3TnC-S7 AAT TGC ACG CTC ACG TGC

W3TnD-S2 ATC ATC GCC CAA TAC CGG

W3TnD-S3 ATT TGC TCA CCA TCA CGC

W3TnD-S4 AAC CCA TCG AGC ATT AAG G

W3TnD-S5 CAG GTT TCT CCG GAG CG

W3TnD-S6 CAC CAT CAC GCT CAA TGG

W3TnD-S7 GTT TAG TGC GGC AAG CCC

W3pan2 CAG TGC TGG CAG ATG ATG GCG

F27 GAG TTT GAT CCT GGC TCA G

FD2L1 TGT GAA GAA GGC CTT CGG

R1492 ACG GTT ACC TTG TTA CGA CTT

240

Appendix III

Routine molecular biology methods

III.I RNase treatment of DNA

- To prepare a of stock solution of RNase dissolve RNase A at a concentration of 10 mg/ml in

10 mM Tris-HCl (pH 7.5) and 15 mM NaCl. Boil for 15 min and cool slowly to room

temperature. Store at -20 oC.

- Prior to use boil stock solution for 5 min and cool to room temperature.

- Add 1 µl of the stock solution for every 100 µl of DNA solution to be treated.

- Incubate at room temperature for 30 min.

- Follow with phenol : chloroform : isolamylalcohol extraction and ethanol precipitation.

III.II Phenol : chloroform : isolamylalcohol extraction

- In an Eppendorf tube increase the volume of the DNA solution to a minimum of 300 µl with

milli-Q water.

- Add a equal volume of phenol : chloroform : isolamylalcohol (25:24:1 (v/v/v)), mix

thoroughly.

- Separate the phenol and aqueous phases by centrifugation at 14 000 x g for 5 min at room

temperature.

- Transfer the upper aqueous phase into a fresh Eppendorf tube.

- Repeat extraction until the interface is no longer visible.

III.III Ethanol precipitation

- Add 1/10 th volume of a 3 M sodium acetate (pH 5.2) solution. Mix well.

- Add exactly 2.5 volumes of ice-cold absolute ethanol. Mix well.

- Chill at -20 oC for 60 min (longer times for small concentrations or small fragments of

DNA).

- Pellet the DNA by centrifugation at 14 000 x g at 4 oC for 15 min (longer times for smaller

fragments).

- Discard supernatant and wash pellet in 70 % ethanol to remove salts.

- Invert tube and dry the pellet or use a vacuum desiccator.

- Resuspend DNA in appropriate volume of milli-Q water or TE Buffer (10 mM Tris-HCl, 1

mM EDTA, pH 8). If necessary heat at 37 oC to assist.

241

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