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
2
To my family and friends
- Cheers !
3
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
4
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
5
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
6
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
7
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
8
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
9
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
10
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.
11
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
14
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
80
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).
81
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
82
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.
83
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).
84
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
85
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
86
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)
87
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
88
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.
90
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.
91
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).
92
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.
94
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).
114
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.
115
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.
117
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).
122
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
170
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
171
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).
172
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
173
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.
174
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.
176
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.
180
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.
190
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).
192
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).
194
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).
195
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.
200
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).
201
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.
202
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).
205
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.
206
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.
211
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.
216
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).
217
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.
218
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.
219
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
223
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.
224
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
228
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
References
Abarzua, S., Jakubowski, S., Eckert, S., and Fuchs, P. (1999) Biotechnological
investigation for the prevention of marine biofouling II. Blue-green algae as potential
producers of biogenic agents for the growth inhibition of microfouling organisms. Bot. Mar.
42: 459-465.
Akagawa-Matsushita, M., Matsuo, M., Koga, M., and Yamasato, K. (1992)
Alteromonas atlantica sp.nov. and Alteromonas carrageenovora sp.nov., bacteria that
decompose algal polysaccharides. Int. J. Syst. Bacteriol. 42: 621-627.
Altschul, S.F., Gish, W., Miller, W., Myers, E.W., and Lipman, D.J. (1990) Basic local
alignment search tool. J. Mol. Biol. 215: 403-410.
Andersen, J.O., Sternberg, C., Poulsen, L.K., Bjorn, S.P., Givskov, M., and Molin, S.
(1998) New unstable varients of green fluorescent protein for studies of transient gene
expression in bacteria. Appl. Environ. Microbiol. 64: 2240-2246.
Andersen, R.J., Wolfe, M.S., and Faulkner, D.J. (1974) Auto-toxic antibiotic production
by marine Chromobacterium. Mar. Biol. 27: 281-285.
Anthoni, U., Nielsen, P.H., Pereira, M., and Christophersen, C. (1990) Bryozoan
secondary metabolites: a chemotaxonomical challenge. Comp. Biochem. Physiol. 96B: 431-
437.
Arima, K.F., Imanaka, H., Kousaka, M., Fukuda, A., and Tamura, C. (1964)
Pyrrolnitrin, a new antibiotic subatance, produced by Pseudomonas. Agric. Biol. Chem. 28:
575-576.
Bally, M., Filloux, M., Akrim, G., Ball, A., Lazdunski, A., and Tommassen, J. (1992)
Protein secretion in Pseudomonas aeruginosa: characterisation of seven xcp genes and
processing of secretory apparatus componenents by prepilin peptidase. Mol. Microbiol. 6:
1121-1131.
242
Baloun, A.J., and Morse, D.E. (1984) Ionic control of settlement and metamorphosis in
larval Haliotis rufescens (Gastropoda). Bio. Bull. 167: 124-138.
Banin, E., Israely, T., Kushmaro, A., Loya, Y., Orr, E., and Rosenberg, E. (2000)
Penetration of the coral-bleaching bacterium Vibrio shiloi into Oculina patagonica. Appl.
Environ. Microbiol. 66: 3031-3036.
Barber, C., Tang, J., Feng, J., Pan, M., Wilson, T., Slater, H., Dow, J., Williams, P.,
and Daniels, M. (1997) A novel regulatory system required for pathogenicity of
Xanthomonas campestris is mediated by small diffusible signal molecules. Mol. Microbiol.
24: 555-566.
Bartlett, D.H., Wright, M.E., and Silverman, M. (1988) Variable expression of
extracellular polysaccharide in the marine bacterium Pseudomonas atlantica is controlled by
genome rearrangement. Proc. Natl. Acad. Sci. 85: 3923-3927.
Bassler, B.L., Wright, M., and Silverman, M.R. (1994) Multiple signalling systems
controlling expression of luminescence in Vibrio harveyi: sequence and function of genes
encoding a second sensory pathway. Mol. Microbiol. 13: 273-286.
Bassler, B.L., Greenberg, E.P., and Stevens, A.M. (1997) Cross-species induction of
luminescence in the quorum-sensing bacterium Vibrio harveyi. J. Bacteriol. 179: 4043-4045.
Bassler, B.L. (1999) A multichannel two-component signaling relay controls expression by
quorum sensing in Vibrio harveyi. In cell-cell signaling in bacteria. Dunny, G. M. and
Winans, S. C. (eds). Washington, D.C. : American Society for Microbiology, pp. 259-273.
Battilani, P., Chiusa, G., and Ceri, C. (1996) Fungal growth and erogsterol content in
tomato fruits infected by fungi. Italian Journal of Food Science 8: 283-289.
Baumann, P., Gauthier, M.J., and Baumann, L. (1984) Genus Alteromonas Baumann,
Mandel and Allen 1972. In Bergey's Manual of Systematic Bacteriology. Krieg, N. R. and
Holt, J. G. (eds). Baltimore : Williams and Wilkins, pp. 343-352.
Bein, S. (1954) A study of certain chromogenic bacteria isolated from "Red Tide" water with
a description of a new species. Bull. Mar. Sci. Gulf. Carribb. 4: 110-119.
243
Beiras, R., and Widdows, J. (1995) Induction of metamorphosis in larvae of the oyster
Crassostrea gigas using neuroactive compounds. Mar. Biol. 123: 327-334.
Ben-Haim, Y., Banim, E., Kushmaro, A., Loya, Y., and Rosenberg, E. (1999) Inhibition
of photosynthesis and bleaching of zooxanthellae by the coral pathogen Vibrio shiloi.
Environ. Microbiol. 1: 223-229.
Berland, B.R., Bonin, D.J., and Maestrini, S.Y. (1972) Are some bacteria toxic for
marine algae? Mar. Biol. 12: 189-193.
Bidle, K.A, and Bartlett, D.H. (2001) An RNA arbitrarily primed PCR survey of genes
regulated by ToxR/S in the deep-sea bacterium Photobacterium profundum strain SS9. J.
Bacteriol. 183: 1688-1693
Black, P.N., DiRusso, C.C., Metzger, A.K., and Heimert, T.L. (1992) Cloning,
sequencing, and expression of the fadD gene of Escherichia coli encoding acyl Coenzyme A
synthetase. J. Biol. Chem. 267: 25513-25520.
Black, P.N., and DiRusso, C.C. (1994) Molecular and biochemical analyses of fatty acid
transport, metabolism, and gene regulation in Escherichia coli. Biochem. Biophys. Acta 1210:
123-145.
Black, P.N., Zhang, Q., Weimar, J., and DiRusso, C.C. (1997) Mutational analysis of a
fatty acyl-Coenzyme A synthetase signature motif identifies seven amino acid residues tha
modulate fatty acid substrate specificity. J. Biol. Chem. 272: 4896-4903.
Bonar, D.B., Coon, S.L., Walch, M., Weiner, R.M., and Fitt, W. (1990) Control of
oyster settlement and metamorphosis by endogenous and exogenous chemical cues. Bull.
Mar. Scien. 46: 484-498.
Bower, S., Perkins, J.B., Yocum, R.R., Howitt, C.L., Rahaim, P., and Pero, J. (1996)
Cloning, sequencing, and characterisation of the Bacillus subtilis biotin biosynthetic operon. J.
Bacteriol. 178: 4122-4130.
244
Braten, T. (1971) The ultratructure of fertilization and zygote formation in the green alga
Ulva Mutabilis Foyn. J. Cell Sci. 9: 621-635.
Bryan, P.J., Qian, P.Y., Krider, J.L., and Chia, F.S. (1997) Induction of larval settlement
and metamorphosis by pharmacological and conspecific associated compounds in the serpulid
polychaete Hydroides elegans. Mar. Ecol. Prog. Ser. 146: 81-90.
Burke, R.D. (1984) Pheromonal control of metamorphosis in the Pacific sand dollar,
Dendraster excentricus. Science 225: 442-443
Callow, M., Callow, J., Pickett-Heaps, J., and Wetherbee, R. (1997) Primary adhesion
of enteromorpha (Chlorophyta, Ulvales) propagules: Quantitative settlement studies and video
microscopy. J. Phycol. 33: 938-947.
Callow, M. (1999) The status and future of biocides in marine biofouling prevention. In
Recent advances in marine biotechnology, Volume 3, Fingerman, M., Nagabhushanam, R.,
and Thompson, M. (eds). New Hampshire: Science publishers, Inc, pp.109-126.
Chakrabortty, A., Das, S., Majumdar, S., Mukhopadhyay, K., Roychoudhury, S.,
Chaudhuri, K. (2000) Use of RNA arbitrarily primed-PCR fingerprinting to identify Vibrio
cholerae genes differentially expressed in the host following infection. Infect. Immun. 68:
3878-3887.
Champion, G., Needly, M., Brennan, M., and DiRita, V. (1997) A branch in the ToxR
regulaory cascade of Vibrio cholerae revealed by characterisation of toxT mutant strains. Mol.
Microbiol. 23: 323-331.
Chan, H.Y.E., Harris, S.J., and O'Kane, C.J. (1998) Identification and characterisation of
kraken, a gene encoding a putative hydrolytic enzyme in Drosophila melanogaster. Gene
222: 195-201.
Characklis, W.G., and Cooksey, K.E. (1983) Biofilms and microbial fouling. Adv. Appl.
Microbiol. 29: 93-138.
245
Chen, C.C., L., Chang, Y., Liu, S., and Tschen, J. (1995) Transposon mutagenesis and
cloning of the genes encoding the enzymes of fengycin biosynthesis in Bacillus subtilis.
Molecular and General Genetics 248: 121-125.
Clare, A.S., Rittschof, D., Gerhart, D.J., and Maki, J.S. (1992) Molecular approaches to
nontoxic antifouling. Invertebrate Reproduction and Development 22: 67-76.
Clayton, M.N. (1992) Propagules of marine macroalgae: structure and development. British
Phycological Journal 27: 219-232.
Conover, J.T., and Seiburth, J.M. (1964) Effects of Sargassum distribution on its epibiota
and antibacterial activity. Bot. Mar. 6: 147-157.
Cooksey, B., Cooksey, K.E., Millar, C.A., Paul, J.H., Rubin, R., and Webster, D.
(1984) The attachment of microfouling diatoms. In Marine biodeterioration: an
interdisciplinary study. Costlow, J. D. and Tipper, R. C. (eds). London : E & FN Spon Ltd,
pp. 167-172.
Coon, S.L., and Bonar, D.B. (1985) Induction of settlement and metamorphosis of the
pacific oyster, Crassostrea gigas, by L-DOPA and catecholamines. J. Exp. Mar. Biol. Ecol.
94: 211-221.
Coon, S.L., Fitt, W.K., and Bonar, D.B. (1990) Competence and delay of metamorphosis
in the pacific oyster Cassastrea gigas. Mar. Biol. 106: 379-387.
Coon, S.L., Walsh, M., Fitt, W.K., Weiner, R.M., and Bonar, D.B. (1990) Ammonia
induces settlement behaviour in oyster larvae. Biol. Bull. 179: 297-303.
Costa, J.M., and Loper, J.E. (1997) EcbI and EcbR: homologs of LuxI and LuxR affecting
antibiotic and exoenzyme production by Erwinia carotovora subsp. betavasculorum. Can. J.
Microbiol. 43: 1164-1171.
Costerton, J.W., Lewandowski, Z., Caldwell, D.E., Korber, D.R., and Lappin-Scott,
H.M. (1995) Microbial biofilms. Annu. Rev. Microbiol. 49: 711-745.
246
Costerton, J.W., Stewart, P.S., and Greenberg, E.P. (1999) Bacterial biofilms: A common
cause of persistent infection. Science 284: 1318-1322.
Cray, K.M., Pearson, J.P., Downie, J.A., Boboye, B.A., and Greenberg, E.P. (1996)
Cell-to-cell signaling in the symbiotic nitrogen-fixing bacterium Rhizobium leguminosarum:
autoinduction of stationary phase and rhizophere-expressed genes. J. Bacteriol. 178: 372-
376.
Crisp, D. (1984) Overview of research on marine invertebrate larvae 1940-1980. In Marine
biodeterioration: an interdisciplinary study. Costlow, J. D. and Tipper, R. C. (eds).
Annapolis, Maryland : Naval institute press, pp. 103-126.
Dalley, R. (1989) Legislation affecting tributyltin antifoulings. Biofouling 1: 363-366.
Dalton, H.M., Poulsen, L.K., Halasz, P., Angles, M.L., Goodman, A.E., and Marshall,
K.C. (1994) Substratum-induced morphological changes in a marine bacterium and their
relevance to biofilm structure. J. Bacteriol. 176: 6900-6906.
Davidson, S.K. and Haygood, M.G. (1999) Identification of sibling species of the
Bryozoan Bugula neritina that produce different anticancer bryostatins and harbor distinct
strains of the bacterial symbiont "Candidatus Endobugula sertula". Biol. Bull. 196: 273-280.
Davies, D., Parsek, M., Pearson, J., Iglewski, B., Costerton, J., and Greenberg, E.
(1998) The involvement of cell-cell signals in the development of a bacterial biofilm. Science
280: 295-298.
Davis, A.R., and Wright, A.E. (1990) Inhibition of larval settlement by natural products
from the ascidian, Eudistoma olivaceum (Van Name). J. Chem. Ecol. 16: 1349-1357.
Davis, A.R., Butler, A.J., and van Altena, I. (1991) Settlement behaviour of ascidian
larvae: preliminary evidence for inhibition by sponge allelochemicals. Mar. Ecol. Prog. Ser.
72: 117-123.
de Beer, D., Stoodley, P., and Lewandowski, Z. (1994) Liquid flow in heterogeneous
biofilms. Biotechnol. Bioeng. 44: 636-641.
247
de Lorenzo, V., Herrero, M., Jakubzik, U., and Timmis, K.N. (1990) Mini-Tn5
transposon derivatives for insertion mutagenesis, promotor probing, and chromosomal
insertion of cloned DNA in Gram-negative eubacteria. J. Bacteriol. 172: 6568-6572.
de Nys, R., Steinberg, P.D., Willemsen, P., Dworjanyn, S.A., Gabelish, C.L., and
King, R.J. (1994) Broad spectrum effects of secondary metabolites from the red algal
Delisea pulchra in antifouling assays. Biofouling 8: 259-271.
de Nys, R., Dworjanyn, S.A., and Steinberg, P.D. (1998) A new method for determining
surface concentrations of marine natural products on seaweeds. Mar. Ecol. Prog. Ser. 162:
79-87.
Decho, A.W., Browne, K.A., and Zimmer-Faust, R.K. (1998) Chemical cues: why basic
peptides are signal molecules in marine environments. Limnol. Oceanogr. 1410-1417.
Dekkers, L.C., Bloemendaal, C.J.P., de Weger, L.A., Wijffelman, C.A., Spaink, H.P.,
and Lugtenberg, B.J.J. (1998) A two-component system plays an important role in the root
colonising ability of Pseudomonas fluorescens strain WCS365. Mol. Plant-Microbe Inter.
11: 45-56.
Dell, C.L., Needly, M.N., and Olson, E.R. (1994) Altered pH and lysine signalling
mutants of cadC, a gene encoding a membrane-bound transcriptional activator of the
Escherichia coli cadBA operon. Mol. Microbiol. 14: 7-16.
Dexter, S.C. (1978) Influence of substratum critical surface tension on bacterial adhesion-in
situ studies. J. Coll. Interf. Sci. 70: 346-354.
Dillon, P.S., Maki, J.S., and Mitchell, R. (1989) Adhesion of Enteromorpha swarmers to
microbial films. Microb. Ecol. 17: 39-47.
DiRita, V., and Melkalanos, J. (1991) Periplasmic interaction between two membrane
regulatory proteins, ToxR and ToxS, results in signal transduction and transcriptional
activation. Cell 64: 29-37.
DiRita, V., Parsot, C., Jander, G., and Mekalanos, J. (1991) Regulatory cascade controls
virulence in Vibrio cholerae. Proc. Natl. Acad. Sci. USA 88: 5403-5407.
248
DiRita, V. (1992) Co-ordinate expression of virulence genes by ToxR in Vibrio cholerae.
Mol. Microbiol. 6: 451-458.
Dorel, C., Vidal, O., Prigent-Combaret, C., Vallet, I., and Lejeune, P. (1999)
Involvemant of the Cpx signal transduction pathway in Escherichia coli in biofilm formation.
FEMS Microbiol. letters 178: 169-175.
Doster, M., and Michailides, T. (1995) The relationship between date of hullsplitting and
decay of pistachio nuts by Aspergillus species. Plant Disease 79: 766-769.
Downard, J., and Toal, D. (1995) Branched-chain fatty acids: the case for a novel form of
cell-cell signaling during Myxococcus xanthus development. Mol. Microbiol. 16: 171-175.
Dums, F., Dow, J.M., and Daniels, M.J. (1991) Structural characterisation of protein
secretion genes of the bacterial phytopathogen Xanthomonas campestris pathovar campestris:
relatedness to secretion systems of other Gram-negative bacteria. Mol. Gen. Genet. 229: 357-
364.
Dworjanyn, S.A., de Nys, R., and Steinberg, P.D. (1999) Localisation and surface
quantification of secondary metabolites in the red alga Delisea pulchra. Mar. Biol. 727-736.
Eberl, L., Winson, M.K., Sternberg, C., Stewart, G.S.A.B., Christiansen, G.,
Chhabra, S.R., Bycroft, B., Williams, P., Molin, S., and Givskov, M. (1996)
Involvement of N-acyl-L-homoserine lactone autoinducers in controlling the multicellular
behaviour of Serratia liquefaciens. Mol. Microbiol. 20: 127-136.
El-Gedaily, A., Paesold, G., Chen, C.-Y., Guiney, D.G., and Krause, M. (1997) Plasmid
virulence gene expression induced by short-chain fatty acids in Salmonella dublin:
Identification of rpoS-dependent and rpoS-independent mechanisms. J. Bacteriol. 179: 1409-
1412.
Elvers, K.T., Leeming, K., Moore, C.P., and Lappin-Scott, H.M. (1998) Bacterial-fungal
biofilms in flowing water photo-processing tanks. J. Appl. Microbiol. 84: 607-618.
249
Engebrecht, J., and Silverman, M. (1984) Identification of gene and gene products
necessary for bioluminescence. Proc. Natl. Acad. Sci. USA. 81: 4154-4158.
Enger, O., Nygaard, H., Solberg, M., Schei, G., Nielsen, J., and Dundas, I. (1987)
Characterisation of Alteromonas denitrificans sp.nov. Int. J. Syst. Bacteriol. 37: 416-421.
Evans, L.V., and Christie, A.O. (1970) Studies on the ship-fouling alga Enteromorpha. I.
Aspects of fine structure and biochemistry of swimming and newly settled zoospores. Ann.
Bot. 34: 451-466.
Evans, L.V. (1981) Marine algae and fouling: A review, with particular reference to ship
fouling. Bot. Mar. 24: 167-171.
Fath, M., and Kolter, R. (1993) ABC transporters: Bacterial exporters. Microbiol. Rev. 57:
995-1017.
Felsenstein, J. (1981) Evolutionary trees from DNA sequences: a maximum likelihood
approach. J. Mol. Evol. 17: 368-376.
Felsenstein, J. (1989) PHYLIP: phylogeny inference package. Cladistics 5: 164-166.
Fisher, M.M., Wilcox, L.W., and Graham, L.E. (1998) Molecular characterisation of
epiphytic bacterial communities on Charophycean green algae. Appl. Environ. Microbiol. 64:
4384-4389.
Fitt, W.K., and Coon, S.L. (1989) Factors influencing bacterial production of inducers of
settlement behaviour of larvae of the oyster Crassostrea virginica. Micro. Ecol. 17: 287-298.
Flavier, A.B., Clough, S.J., Schell, M.A., and Denny, T.P. (1997) Identification of 3-
hydroxypalmitic acid methyl ester as a novel autoregulator controlling virulence in Ralstonia
solanacearum. Mol. Microbiol. 26: 251-259.
Flavier, A.B., Ganova-Raeva, L.M., Schell, M.A., and Denny, T.P. (1997) Hierarchical
autoinduction in Ralstonia solanacearum: control of acyl-homoserine lactone production by a
novel autoregulatory system responsive to 3-hydroxypalmitic acid methyl ester. J. Bacteriol.
179: 7089-7097.
250
Flavier, A.B., Schell, M., and Denny, T. (1998) An RpoS homologue regulates
acylhomoserine lactone-dependent autoinduction in Ralstonia solanacearum. Mol. Microbiol.
28: 475-486.
Fletcher, M. (1996) Bacterial attachment in aquatic environments: a diversity of surfaces and
adhesion strategies. In Bacterial adhesion: molecular and ecological diversity. Fletcher, M.
and Belle, W. (eds). New York : John Wiley and Sons, pp. 1-24.
Fletcher, R.L. (1975) Heteroantagonism observed in mixed algal cultures. Nature 253: 534-
535.
Fletcher, R.L., and Callow, M. (1992) The settlement, attachment and establishment of
marine algal spores. British Phycological Journal 27: 303-329.
Freeman, J.A., and Bassler, B.L. (1999) A genetic analysis of the function of LuxO, a two-
component response regulator involved in quorum sensing in Vibrio harveyi. Mol. Microbiol.
31: 665-677.
Freeman, J.A., and Bassler, B.L. (1999) Sequence and functions of LuxU: a two-
component phosphorelay protein that regulates quorum sensing in Vibrio harveyi. J.
Bacteriol. 181: 899-906.
Fries, L. (1975) Some observations on the morphology of Enteromorpha linza (L.) J. Ag.
and Enteromorpha compressa (L.) Grev, in axenic culture. Bot. Mar. 18: 251-253.
Fuqua, W.C., and Winans, S.C. (1994) A LuxR-LuxI type regulatory system activates
Agrobacterium Ti plasmid conjugal transfer in the presence of a plant tumor metabolite. J.
Bacteriol. 176: 2796-2806.
Gaffney, T.D., Lam, S.T., Ligon, J., Gates, K., Frazelle, A., Di Maio, J., Hill, S.,
Goodwin, S., Torkewitz, N., Allshouse, A.M., Kempf, H.J., and Becker, J.O. (1994)
Global regulation of expression of anti-fungal factors by a Pseudomonas fluorescens
biological control strain. Mol. Plant-Microbe Interact. 7: 455-463.
251
Gates, R.D., Baghdasarian, G., and Muscatine, L. (1992) Temperature stress causes host
cell detachment in sybiotic cnidarians: implications for coral bleaching. Biol. Bull. 182: 324-
332.
Gauthier, M.J., and Flatau, G.N. (1976) Antibacterial activity of marine violet-pigmented
Alteromonas with special reference to the production of brominated compounds. Can. J.
Microbiol. 22: 1612-1619.
Gauthier, M.J. (1977) Alteromonas citrea, a new Gram-negative, yellow pigmented species
from seawater. Int. J. Syst. Bacteriol. 27: 349-354.
Gauthier, M.J. (1979) Alteromonas rubra sp.nov., a new marine antibiotic-producing
bacterium. Int. J. Syst. Bacteriol. 26: 459-466.
Gauthier, M.J., and Breittmayer, V.A. (1979) A new antibiotic-producing bacterium from
seawater: Alteromonas aurantia sp.nov. Int. J. Syst. Bacteriol. 29: 366-372.
Gauthier, M.J. (1982) Validation of the name Alteromonas luteoviolacea. Int. J. Syst.
Bacteriol. 32: 82-86.
Gauthier, G., Gauthier, M.J., and Christen, R. (1995) Phylogenetic analysis of the
Genera Alteromonas, Shewanella, and Moritella using genes coding for small-subunit rRNA
sequences and division of the genus Alteromonas into two genera, Alteromonas (emended)
and Pseudoalteromonas gen. nov., and proposal of twelve new species combinations. Int. J.
Syst. Bacteriol. 45: 755-761.
Gerhart, D.J., Rittschof, D., and Mayo, S.W. (1988) Chemical ecology and the search for
marine antifoulants. J. Chem. Ecol. 14: 1905-1917.
Gibbs, P.E., Bryan, G.W., Pascoe, P.L., and Burt, G.R. (1990) Reproductive
abnormalities in female Ocenebra erinacea (Gastropoda) resulting from tributyltin-induced
imposex. J. Mar. Biol. Assoc. U.K. 70: 639-656.
Gil-Turnes, M.S., Hay, M.E., and Fenical, W. (1989) Symbiotic marine bacteria defend
crustacean embryos from a pathogenic fungus. Science 240: 116-118.
252
Gil-Turnes, M.S., and Fenical, W. (1992) Embryos of Homarus americanus are protected
by epibiotic bacteria. Biol. Bull. 182: 105-108.
Gillan, D., Speksnijder, A., Zwart, G., and Ridder, C. (1998) Genetic diversity of the
biofilm covering Montacuta ferruginosa (mollusca, bivalvia) as evaluated by denaturing
gradient gel electrophoresis analysis and cloning of PCR-amplified gene fragments coding for
16S rDNA. Appl. Environ. Microbiol. 64: 3464-3472.
Gitlitz, M.H. (1981) Recent developments in marine antifouling coatings. J. Coatings
Technol. 53: 46-52.
Givskov, M., de Nys, R., Manefield, M., Gram, L., Maximilien, R., Eberl, L., Molin,
S., Steinberg, P., and Kjelleberg, S. (1996) Eucaryotic interference with homoserine
lactone mediated procaryotic signalling. J. Bacteriol. 178: 6618-6622.
Glick, B.R., and Bashan, Y. (1997) Genetic manipulation of plant growth-promoting
bacteria to enhance biocontrol of phytopathogens. Biotechnology Advances 15: 353-378.
Glick, B.S., and Rothman, J.E. (1987) Possible role for fatty acyl-coenzyme A in
intracellular protein transport. Nature 326: 309-312.
Glynn, P.W. (1991) Coral reef bleaching in the 1980s and possible connections with global
warming. Trends Ecol. Evol. 6: 175-179.
Godchaux, W., Lynes, M.A., and Leadbetter, E.R. (1991) Defects in gliding motility in
mutants of Cytophaga johnsonae lacking a hight-molecular-weight cell surface
polysaccharide. J. Bacteriol. 173: 7607-7614.
Gonzalez, J.M., Sherr, E.B., and Sherr, B.F. (1993) Differential feeding by marine
flagellates on growing versus starving, and on motile versus nonmotile, bacterial prey. Mar.
Ecol. Prog. Ser 102: 257-267.
Goto, R., Kado, R., Muramoto, K., and Kamiya, H. (1992) Fatty acids as antifoulants in a
marine sponge. Biofouling 6: 61-68.
253
Hammer, P., Hill, D., Lam, S., Van Pee, K., and Ligon, J. (1997) Four genes from
Pseudomonas fluorescens that encode the biosynthesis of pyrrolnitrin. Appl. Environ.
Microbiol. 63: 2147-2154.
Hansen, A.J., Weeks, O.B., and Colwell, R.R. (1965) Taxonomy of Pseudomonas
piscicida (Bein) Buck, Meyers and Leifson. J. Bacteriol. 89: 752-761.
Harlin, M., and Lindberg, J.M. (1977) Selection of substratum by seaweeds: optimal
surface relief. Mar. Biol. 40: 33-40.
Harrison, P.G. (1982) Control of microbial growth and of amphipod grazing by water-
soluble compounds from leaves of Zostera marina. Mar. Biol. 67: 25-230.
Harshey, R.M. (1994) Bees aren't the only ones: swarming in Gram-negative bacteria. Mol.
Microbiol. 13: 389-394.
Häse, C., and Mekalanos, J. (1998) TcpP protein is a positive regulator of virulence gene
expression in Vibrio cholerae. Proc. Natl. Acad. Sci. USA 95: 730-734.
Hauser, D.C.R., Levandowsky, M., Hunter, S.H., Chunosoff, L., and Hollwitz, J.S.
(1975) Chemosensory responses by the heterotrophic marine dinoflagellate Crypthecodinium
cohnii. Microb. Ecol. 1: 246-254.
Hay, M.E., and Fenical, W. (1988) Marine plant herbivore interactions: the ecology of
chemical defense. Annu. Rev. Ecol. Sys. 19: 111-145.
Henry, E.C., and Cole, K.M. (1982) Ultrastructure of swarmers in the Laminariales. I.
zoospore. J. Phycol. 18: 550-569.
Herrero, M., de Lorenzo, V., and Timmis, K. (1990) Transposon vectors containing non-
antibiotic resistance selection markers for cloning and stable chromosomal insertion of
foreign genes in Gram-negative bacteria. J. Bacteriol. 172: 6557-6567.
Higgins, C. (1992) ABC transporters: from microorganisms to man. Annu. Rev. Cell Biol. 8:
67-113.
254
Higgins, D.E., Nazareno, E., and DiRita, V.J. (1992) The virulence gene activator ToxT
from Vibrio cholerae is a member of the ArcC family of transcriptional activators. J.
Bacteriol. 174: 6974-6980.
Hills, J.M., and Thomason, J.C. (1998) The effect of scales of surface roughness on the
settlement of barnacle (Semibalanus balanoides) cyprids. Biofouling 12: 57-69.
Holmström, C., Rittschof, D., and Kjelleberg, S. (1992) Inhibition of attachment of larval
barnacles, Balanus amphitrite by Ciona intestinalis a surface colonizing marine bacterium.
Appl. Environ. Microbiol. 58: 2111-2115.
Holmström, C., James, S., Egan, S., and Kjelleberg, S. (1996) Inhibition of common
fouling organisms by marine bacterial isolates with special reference to the role of pigmented
bacteria. Biofouling 10: 251-259.
Holmström, C., James, S., Neilan, B., White, D., and Kjelleberg, S. (1998)
Pseudoalteromonas tunicata sp. nov., a bacterium that produces antifouling agents. Int. J.
Syst. Bacteriol. 48: 1205-1212.
Holmström, C., and Kjelleberg, S. (1999) Marine Pseudoalteromonas species are
associated with higher organisms and produce active extracellular agents. FEMS Microbiol
Ecol. 30: 285-293.
Holmström, C., Steinberg, P., Christov, V., and Kjelleberg, S. (2000) Bacteria
immobilised in hydrogels: a novel concept to prevent development of biofouling communities.
Biofouling 15:109-117
Horinouchi, S., and Beppu, T. (1992) Autoregulatory factors and communication in
actinomycetes. Annu. Rev. Microbiol. 46: 377-398.
Howell, W.J., and Stipanovic, R.D. (1980) Supression of Pythium ultimum-induced
damping off of cotton seedlings by Pseudomonas fluorescens and its, antibiotic, pyoluteorin.
Phytopathology 70: 712-715.
255
Hrabak, E.M., and Willis, D.K. (1992) The lemA gene required for pathogenicity of
Pseudomonas syringae on bean is a member of a family of two-component regulators. J.
Bacteriol. 1992: 3011-3020.
Hugh, R., and Leifson, E. (1953) The taxonomic significance of fermentative versus
oxidative metabolism ofcarbohydrates by various Gram-negative bacteria. J. Bacteriol. 66:
24-26
Hyde, K.D., Jones, E.B.G., Leano, E., Pointing, S.B., Poonyth, A.D., and Vrijmoed,
L.L.P. (1998) Role of fungi in marine ecosystems. Biodiversity and conservation 7: 1147-
1161.
Ilan, M., Jensen, R.A., and Morse D.E. (1993) Calcium control of metamorphosis in
polychaete larvae. J. Exp. Zool. 267: 193-207.
Imai, I., Ishida, Y., Sakaguchi, K., and Hata, Y. (1995) Algicidal marine bacteria isolated
from northern Hiroshima bay, Japan. Fish. Sci. 61: 628-636.
Ivanova, E.P., Kiprianova, E.A., Mikhailov, V.V., Levanova, G.F., Garagulya, A.D.,
Gorshkova, N.M., Yumoto, N., and Yoshikawa, S. (1996) Characterisation and
identification of marine Alteromonas nigrifaciens strains and emendation of the description.
Int. J. Syst. Bacteriol. 46: 223-228.
Ivanova, E.P., Kiprianova, E.A., Mikhailov, V.V., Levanova, G.F., Garagulya, A.D.,
Gorshkova, N.M., Vysotskii, M.V., Nicolau, D.V., Yumoto, N., Taguchi, T., and
Yoshikawa, S. (1998) Phenotypic diversity of Pseudoalteromonas citrea from different
marine habitats and emendation of the description. Int. J. Syst. Bacteriol. 48: 247-256.
Ivanova, E.P., Nicolau, D.V., Yumoto, N., Taguchi, T., Okamoto, K., Tatsu, Y., and
Yoshikawa, S. (1998) Impact of conditions of cultivation and adsorption on antimicrobial
activity of marine bacteria. Mar. Biol. 130: 545-551.
Jaeckle, W.B., and Manahan, D.T. (1989) Feeding by a nonfeeding larva: uptake of
dissolved amino acids from seawater by lecithotrophic larvae of the gastropod Haliotis
rufescens. Mar. Biol. 103: 87-94.
256
James, S., Holmström, C., and Kjelleberg, S. (1996) Purification and characterisation of a
novel anti-bacterial protein from the marine bacterium D2. Appl. Environ. Microbiol. 62:
2783-2788.
James, S. (1998), Antifouling and antibacterial effects of Pseudoalteromonas tunicata. PhD
Thesis, University of New South Wales, Sydney, Australia.
Janisiewicz, W.J. (1988) Biocontrol of post harvest diseases of apples with antagonist
mixtures. Phytopathology 78: 194-198.
Jennings, J.G., and Steinberg, P.D. (1997) Phlorotannins versus other factors affecting
epiphyte abundance on the kelp Ecklonia radiata. Oecologia 109: 461-473.
Jensen, R.A., Morse, D.E., Petty, R.L., and Hooker, N. (1990) Artificial induction of
larval metamorphosis by free fatty acids. Mar. Ecol. Prog. Ser. 67: 55-71.
Jensen, P. R., Jenkins, K. M., Porter, D., Fenical, W. (1998) Evidence that a new
antibiotic flavone glycoside chemically defends the sea grass Thalassia testudinum against
zoosporic fungi. Appl. Environ. Microbiol. 64: 1490-1496.
Jobling, M.G., and Holmes, R.K. (1997) Characterisation of hapR, a positive regulator of
the Vibrio cholerae HA/protease gene hap, and its identification as a functional homologue of
the Vibrio harveyi luxR gene. Mol. Microbiol. 26: 1023-1034.
Johnson, C.R., and Sutton, D.C. (1994) Bacteria on the surface of crustose coralline algae
induce metamorphosis of the crown of thoens starfish Acanthaster plani. Mar. Biofouling
120: 305-310.
Johnston, C., Pegues, D.A., Hueck, C.J., Lee, C.A., and Miller, S.I. (1996)
Transcriptional activation of Salmonella typhimurium invasion genes by a member of the
phosphorylated response-regulator superfamily. Mol. Microbiol. 22: 715-727.
Jones, E.B.G. (1994) Fungal adhesion. Mycol. Res. 98: 961-981.
Jukes, T.H., and Cantor, C.R. (1969) Evolution of protein molecules. In Mammalian
Protein Metabolism. Munro, H. N. (eds). New York : Academic Press, pp. 21-132.
257
Junker, F., Kiewitz, R., and Cook, A.M. (1997) Characterisation of the p-toluenesulfonate
operon tsaMBCD and tsaR in comamonas testosteroni T-2. J. Bacteriol. 179: 919-927.
Kaneko, T., Sato, S., Kotani, H., Tanaka, A., Asamizu, E., Nakamura, Y., Miyajima,
N., Hirosawa, M., Sugiura, M., Sasamoto, S., Kimura, T., Hosouchi, T., Matsuno, A.,
Muraki, A., Nakazaki, N., Naro, K., Okumura, S., Shimpo, S., Takeuchi, C., Wada,
T., Watanbe, A., Yamada, M., and Tabata, S. (1996) Sequence analysis of the genome of
the unicellular cyanobacterium Synechocystis sp. strain PCC6803. II. Sequence determination
of the entire genome and asignment of potential protein-coding regions. DNA Res. 30: 190-
136.
Kaper, J.B., Morris, J.G., and Levine, M.M. (1995) Cholera. Clin. Microbiol. Rev. 8: 48-
86.
Karaolis, D.K., Johnson, J.A., Bailey, C.C., Boedeker, E.C., Kaper, J.B., and Reeves,
P.R. (1998) A Vibrio cholerae pathogenicity island associated with epidemic and pandemic
strains. Proc. Natl. Acad. Sci. USA 95: 3134-3139.
Karaolis, D.K.R., Somara, S., Maneval, D.R., Johnson, J.A., and Kaper, J.B. (1999) A
bacteriophage encoding a pathogenicity island, a type-IV pilus and phage receptor in cholera
bacteria. Nature 399: 375-379.
Kaspar, H., and Mountfort, D. (1995) Microbial production and degradation of Gamma-
aminobutyric acid (GABA) in the abalone larval settlement habitat. FEMS Microbiol. Ecol. 17:
205-212.
Kato, J., Ikeda, T., Kuroda, A., Takiguchi, A., and Ohtake, H. (1999) Influence of
Pseudoalteromonas sp. A28 extracellular products on the synthesis of proteases exhibiting
lytic activity towards marine algae. In Abstracts of the 99th general meeting of American
Society for Microbiology. (ed). Washington, DC.
Kawagishi, I., Imagawa, M., Imae, Y., L., M., and Homma, M. (1996) The sodium -
driven polar flagellar motor of marine Vibrio as the mechanosensor that regulates lateral
flagellar expression. Mol. Micobiol. 20: 693-699.
258
Keel, C., Wirthner, P., Oberhansli, T., Voisard, C., Burger, U., Haas, D., and Defago,
G. (1990) Pseudomonads as antagonists of plant pathogens in the rhizosphere: role of the
antibiotic 2, 4-diacetylphloroglucinol in the suppression of black rot of tobacco. Symbiosis 9:
327-341.
Kimura, M. (1980) A simple method for estimating evolutionary rates of base substitutions
through comparative studies of nucleotide sequences. J. Mol. Evol. 16: 111-120.
Kinscherf, T.G., and Willis, D.K. (1999) Swarming by Pseudomonas syringae B728a
requires gacS (lemA) and gacA but not the acyl-homoserine lactone biosynthetic gene ahlI. J.
Bacteriol. 181: 4133-4136.
Kirchman, D.L., Graham, D.S., Reish, D., and Mitchell, R. (1982a) Bacteria induce the
settlement and metamorphosis of Janua (dexiospira) basiliensis grube (polychaete:
Spirorbidae). J. Exp. Mar. Biol. Ecol. 56: 153-163.
Kirchman, D.L., Graham, D.S., Reish, D., and Mitchell, R. (1982b) Lectins may
mediate in the settlement and metamorphosis of Janua (Dexiospira) brasiliensis (Grube).
Mar. Biol. Lett. 3: 1-12.
Kita-Tsukamota, K., Oyaizu, H., Nanba, K., and Simidu, U. (1993) Phylogenetic
relationships of marine bacteria, mainly members of the family Vibrionaceae determined on
the basis of 16S rRNA sequences. Int. J. Syst. Bacteriol. 43: 8-19.
Kitamura, H., Kitahara, H., and Koh, B. (1993) The induction of larval settlement and
metamorphosis of two sea urchins, Pseudocentrotus depressus and Anthocidaris crassispina,
by free fatty acids extracted from the coralline red alga Corallina pilulifera. Mar. Biol. 115:
387-392.
Kitten, T., Kinscherf, T.G., McEvoy, J.L., and Willis, D.K. (1998) A newly-identified
regulator is required for virulence and toxin production in Pseudomonas syringae. Mol.
Microbiol. 28: 917-930.
Kjelleberg, S., Steinberg, P., Givskov, M., Gram, L., Manefield, M., and de Nys, R.
(1997) Do marine natural products interfere with prokaryotic AHL regulatory systems?
Aquat. Microbiol. Ecol. 13: 85-93.
259
Kleckner, N., Bender, J., and Gottesman, S. (1991) Uses of transposons with emphasis
on Tn10. In Methods in Enzymology: Bacterial Genetic Systems. Miller, J. H. (eds). San
Diego, California : Academic Press Inc, pp. 139-180.
Kleerbezem, M., Quadri, L., Kuipers, O., and de Vos, W. (1997) Quorum sensing by
peptide pheromones and two-component signal-transduction systems in Gram-positive
bacteria. Mol. Microbiol. 24: 895-904.
Klut, E.M., Bisalputra, T., and Antia, N.J. (1983) Agglutination of the chlorophycean
flagellate Dunaliella tertiolecta by treatment with lectins or divalent cations at alkaline pH. J.
Phycol. 19: 112-115.
Köhler, J., Hansen, P.D., and Wahl, M. (1999) Colonisation patterns at the substratum-
water interface: how doues surface microtopography influence recruitment patterns of sessile
organisms? Biofouling 14: 237-248.
Kon-ya, K., Shimidzu, N., Adachi, K., and Miki, W. (1994) 2,5,6-Tribromo-1-
methylgramine, an antifouling substance from the marine bryozoan Zoobotryon pellucidum.
Fish. Sci. 60: 773-775.
Kon-ya, K., Shimidzu, N., Otaki, N., Yokoyama, A., Adachi, K., and Miki, W. (1995)
Inhibitory effect of bacterial ubiquinones on the settling of barnacle, Balanus amphitrite.
Experientia 51: 153-155.
Korchak, H.M., Kane, L.H., Rossi, M.W., and Corkey, B.E. (1994) Long chain acyl-
coenzyme A and signaling in neutrophils. An inhibitor of acyl coenzyme A synthetase, triacsin
C, inhibits superoxide anion generation and degranulation by human neutrophils. J. Biol.
Chem. 269: 30281-30287.
Kovacikova, G., and Skorupski, K. (1999) A Vibrio cholerae LysR homolog, AphB,
cooperates with AphA at the tcpPH promoter to activate expression of the ToxR virulence
cascade. J. Bacteriol. 181: 4250-4256.
Kovacs, N. (1956) Identification of Pseudomonas pyocyanea by the oxidase reaction.
Nature 178: 703.
260
Kuo, A., Callahan, S.M., and Dunlap, P.V. (1996) Modulation of luminescence operon
by N-octanoyl-L-homoserine lactone in ainS mutants of Vibrio fischeri. J. Bacteriol. 178:
971-976.
Kushmaro, A., Rosenberg, E., Fine, M., and Loya, Y. (1997) Bleaching of the coral
Oclina patagonica by Vibrio AK-1. Mar. Ecol. Prog. Ser. 147: 159-167.
Kyte, J., and Doolittle, R.F. (1982) A simple method for displaying the hydrophobic
character of a protein. J. Mol. Biol. 157: 105-202.
Laemmli, U.K. (1970) Cleavage of structural proteins during the assembly of the head of the
bacteriophage T4. Nature 227: 680-685.
Lau, S.C.K., and Qian, P. (1997) Phlorotannins and related compounds as larval settlement
inhibitors of the tube-building polychaete Hydroides elegans. Mar. Ecol. Prog. Ser. 159:
219-227.
Laville, J.C., Voisard, C., Keel, C., Maurhofer, M., Défago, G., and Haas, D. (1992)
Global control of Pseudomonas fluorescens mediating antibiotic synthesis and suppression
of black root rot of tobacco. Proc. Natl. Acad. Sci. USA 89: 1562-1566.
Lee, S., Kato, J., Takiguchi, N., Kuroda, A., Ikeda, T., Mitsutani, A., and Ohtake, H.
(2000) Involvement of an extracellular protease in algicidal activity of the marine bacterium
Pseudoalteromonas sp. strain A28. Appl. Environ. Microbiol. 66: 4334-4339.
Lee, S.E., Shin, S.H., Kim, S.Y., Kim, Y.R., Shin, D.H., Chung, S.S., Lee, Z.H., Lee,
J.Y., Jeong, K.C., Choi, S.H., and Rhee, J.H. (2000) Vibrio vulnificus has the
transmembrane transcriptional activator ToxRS stimulating the expression of Hemolysin gene
vvhA. J. Bacteriol. 182: 3405-3415.
Leitz, T., and Wagne, R.T. (1993) The marine bacterium Alteromonas espejiana induces
metamorphosis of the hydroid Hydractinia echinata. Mar. Biol. 115: 173-178.
Levantine, P.L., and Bonar, D.B. (1986) Metamorphosis of Ilyanassa obsoleta: natural
and artificial inducers. Am. Zool. 26: (Abstract).
261
Lewenza, S., Conway, B., Greenberg, E.P., and Sokol, P.A. (1999) Quorum sensing in
Burkholderia cepacia: identification og the LuxRI homologs CepRI. J. Bacteriol. 181: 748-
756.
Lewis, T. (1994) Impact of biofouling on the aquaculture industry. In Biofouling : Problems
and Solutions. Kjelleberg, S., Steinberg, P. (ed). UNSW, Sydney, Austraila.
Li, C.C., Crawford, J.A., DiRita, V.J., and Kaper, J.B. (2000) Molecular cloning and
transcriptional regulation of ompT, a ToxR-repressed gene in Vibrio cholerae. Mol.
Microbiol. 35: 189-203.
Lichstein, H.C. and van de Sand, V.F. (1945) Violacein, an antibiotic pigment produced
by Chromobacterium violaceum. J. Infect. Dis. 76: 47-51.
Lin, Z., Kumagal, K., Baba, K., Mekalanos, J., and Nishibuchi, M. (1993) Vibrio
parahaemolyticus has a homolog of the Vibrio cholerae toxRS operon that mediates
environmentally induced regulation of the thermostable directed hemolysin gene. J. Bacteriol.
175: 3844-3855.
Lindum, P.W., Anthoni, U., Christophersen, C., Eberl, L., Molin, S., and Givskov,
M. (1998) N-acyl-L-homoserine lactone autoinducers control production of an extracellular
lipopeptide biosurfactant required for swarming motility of Serratia liquefaciens MG1. J.
Bacteriol. 180: 6384-6388.
Little, B. (1985) Factors influencing the adsorption of dissolved organic material from
natural waters. J. Coll. Interf. Sci. 108: 331-340.
Lobban, C.S., and Harrison, P.J. (1994) Seaweed ecology and physiology. Cambridge :
Cambridge University Press.
Loeb, G.J., and Neihof, R.A. (1975) Marine conditioning films. Adv. Chem. Ser. 145:
319-335.
262
Lovejoy, C., Bowman, J., and Hallegraeff, G. (1998) Algicidal effects of a novel marine
Pseudoalteromonas isolate (class Proteobacteria, gamma subdivision). Appl. Environ.
Microbiol. 64: 2806-2813.
Maeda, M., Nogami, K., Kanematsu, M., and Hirayama, K. (1997) The concept of
biological control methods in aquaculture. Hydrobiologia 358: 285-290.
Maki, J.S., Rittschof, D., Costlow, J.D., and Mitchell, R. (1988) Inhibition of attachment
of larval barnacles, Balanus amphitrite, by bacterial surface films. Mar. Biol. 97: 199-206.
Maki, J.S., Rittschof, D., Samuelsson, M.O., Szewzyk, U., Yule, A.B., Kjelleberg, S.,
Costlow, J.D., and Mitchell, R. (1990) Effects of marine bacteria and their exopolymers on
the attachment of barnacle cypris larvae. Bull. Mar. Sci. 46: 499-511.
Manefield, M., de Nys, R., Kumar, N., Read, R., Givskov, M., Steinberg, P., and
Kjelleberg, S. (1999) Evidence that halogenated furanones from Delisea pulchra inhibit
acylated-homoserine lactone (AHL)- mediated gene expression by displacing the AHL signal
from its receptor protein. Microbiology 145: 283-291.
Marden, P., Tunlid, A., Malmcrona-Friberg, K., Odham, G., and Kjelleberg, S. (1985)
Physiological and morphological changes during short term starvation of marine bacterial
isolates. Arch. Microbiology 142: 326-332.
Margalith, P. (1992) Pigment microbiology. London : Chapman and Hall.
Marshall, K.C., Stout, R., and Mitchell, R. (1971) Mechanisms of the initial events in the
sorption of marine bacteria to surfaces. J. Gen. Microbiol. 68: 337-348.
Marshall, K.C. (1994) Biofouling-what is the problem? In Biofouling: problems and
solutions. Kjelleberg, S. and Steinberg, P. (ed). Sydney, Austraila.
Mary, A., Mary, V., Rittschof, D., and Nagabhushanam, R. (1993) Bacterial-barnacle
interaction: potential for using juncellins and antibiotica to alter structure of bacterial
communities. J. Chem. Ecol. 19: 2155-2167.
263
Mathews, C.K., and van Holde, K.E. (1990) Biochemistry. Redwood city, California : The
Benjamin/ Cummings Publishing Company.
Matsuyama, T., Kaneda, K., Nakagawa, Y., Isa, K., Hara-Hotta, H., and Yano, I.
(1992) A novel extracellular cyclic lipopeptide which promotes flagellum-dependent and -
independent spreading growth of Serratia marcescens. J. Bacteriol. 174: 1769-1776.
Maximilien, R., de Nys, R., Holmström, C., Gram, L., Givskov, M., Crass, K.,
Kjelleberg, S., and Steinberg, P. (1998) Chemical mediation of bacterial surface
colonisation by secondary metabolites from the red alga Delisea pulchra. Aquat. Microb.
Ecol. 15: 233-246.
McCaffrey, E.J., and Endean, R. (1985) Antimicrobial activity of tropical and subtropical
sponges. Mar. Biol. 89: 1-8.
McCarter, L.L., Showalter, R., and Silverman, M. (1992) Genetic analysis of surface
sensing in Vibrio parahaemolyticus. Biofouling 5: 163-175.
McCarter, L.L. (1998) OpaR, a homolog of Vibrio harveyi LuxR, controls opacity of Vibrio
parahaemolyticus. J. Bacteriol. 180: 3166-3173.
McCarthy, S.A., Johnson, R.M., and Kakimoto, D. (1994) Characterisation of an
antibiotic produced by Alteromonas luteoviolacea Gauthier 1982, 85 isolated from Kinko bay,
Japan. J. Appl. Bacteriol. 77: 426-432.
McClean, K.H., Winson, M.K., Fish, L., Taylor, A., Chhabra, S., Camara, M.,
Daykin, M., Lamb, J.H., Swift, S., Byrcoft, B.W., Stewart, G.S.A.B., Williams, P.
(1997) Quorum sensing and Chromobacterium violaceum: exploitation of violacein
production and inhibition for the detection of N-acylhomoserine lactones. Microbiology, 143:
3703-3711.
McClintock, J.B. (1994) Trophic biology of Antartic shallow-water echinoderms. Mar.
Ecol. Prog. Ser. 121: 191-202.
264
McDonald, M., Wilkinson, B., Van't Land, C.W., Mocek, U., Lee, S., Floss, H.G.
(1999) Biosynthesis of phenazine antibiotics in Streptomyces antibioticus: Stereochemistry of
Methyl Transferase from Carbon-2 of Acetate. J. Am. Chem. Soc. 121: 5619-5624.
McDougald, D., Rice, S.A., and Kjelleberg, S. (2000) The marine pathogen Vibrio
vulnificus encodes a putative homologue of the Vibrio harveyi regulatory gene, luxR: a genetic
and phylogenetic comparison. Gene 248: 213-221.
McLean, R.J.C., Whiteley, M., Stickler, D.J., and Fuqua, W.C. (1997) Evidence of
autoinducer activity in naturally occuring biofilms. FEMS Microbiol. lett. 154: 259-263.
Miller, V., Taylor, R., and Mekalanos, J.J. (1987) Cholera toxin transcriptional activator
ToxR is a transmembrane DNA binding protein. Cell 48: 271-279.
Miller, V.L., and Mekalanos, J.J. (1988) A novel suicide vector and its use in construction
of insertional mutations: osmoregulation of outer membrane proteins and virulence
determinants in Vibrio cholerae requires toxR. J. Bacteriol. 170: 2575-2583.
Miller, V.L., DiRita, V.J., and Mekalanos, J.J. (1989) Identification of toxS, a regulatory
gene whose product enhances ToxR mediated activation of the cholera toxin promoter. J.
Bacteriol. 171:
Mitchell, R. (1984) Colonization by higher organisms. In Microbial adhesion and
aggreation. Marshall, C. (eds). : Springer-verlag, pp. 189-200.
Mizobuchi, S., Abachi, K., and Miki, W. (1996) Antifouling polyhydroxysterols isolated
from a Palauan octocoral of Sinularia sp. Fish. Sci. 62: 98-100.
Moat, A., and Foster, J. (1995) Regulation of prokaryotic gene expression. In Microbial
physiology. Moat, A. and Foster, J. (eds). New York : Wiley-liss, pp. 152-202
Montgomery, M., and Kirchman, D. (1993) Role of chitin-binding proteins in the specific
attachment of the marine bacterium Vibrio harveyi to chitin. Appl. Environ. Microbiol. 59:
373-379.
265
Montgomery, M.T., and Kirchman, D.L. (1994) Induction of chitin-binding proteins
during specific attachment of the marine bacterium Vibrio harveyi to chitin. Appl. Environ.
Microbiol. 60: 4284-4288.
Moriarty, D.J.W., and Bell, R.T. (1993) Bacterial growth and starvation in aquatic
environments. In Starvation in bacteria. Kjelleberg, S. (eds). New York : Plenum press, pp.
25-48.
Morisaki, H., Nagai, S., Ohshima, H., Ikemoto, E., and Kogure, K. (1999) The effect of
motility and cell-surface polymers on bacterial attachment. Microbiology 145: 2797-2802.
Morse, A.N.C., and Morse, D.E. (1984) Reruitment and metamorphosis of Haliotis larvae
induced by molecules uniquely available at the surfaces of crustoses red algae. J. Exp. Mar.
Biol. Ecol. 75: 191-215.
Morse, D.E. (1985) Neurotransmitter-mimetic inducers of larval settlement and
metamorphosis. Bull. Mar. Sci. 37: 697-706.
Morse, D.E. (1990) Recent progress in larval settlement and metamorphosis: closing the gaps
between molecular biology and ecology. Bull. Mar. Sci. 46: 465-483.
Nakanishi, K., Nishijima, M., Nishimura, M., Kuwano, K., and Saga, N. (1996)
Bacteria that induce morphogenesis in Ulva pertusa (Chlorophyta) grown under axenic
conditions. J. Phycol. 32: 479-482.
Nealson, K. (1977) Autoinduction of bacterial luciferase. Occurance, mechanism and
significance. Arch. Microbiol. 112: 73-79.
Neely, M.N., and Olson, E.R. (1996) Kinetics of expression of the Escherichia coli cad
operon as a function of pH and Lysine. J. Bacteriol. 178: 5522-5528.
Nelson, E.J., and Ghiorse, W.C. (1999) Isolation and identification of Pseudoalteromonas
piscicida strain Cura-d associated with diseased damselfish (Pomacentridae) eggs. Journal of
Fish Diseases 22: 253-260.
266
Neu, T.R. (1996) Significance of bacterial surface-active compounds in interaction of bacteria
with interfaces. Microbiol. Rev. 60: 151-166.
Ninfa, A.J. (1996) Regulation of gene transcription by extracellular stimuli. In Escherichia
coli and Salmonella: Cellular and Molecular Biology. Neidhardt, F. C. (eds). Washington, D.
C.: American Society for Microbiology Press, pp. 1246-1286.
Nyström, T. (1993) Global systems approach to the physiology of the starved cell. In
Starvation in bacteria. Kjelleberg, S. (eds). New York : Plenum Press.
O'Toole, G., and Kolter, R. (1998a) Flagellar and twitching motility are necessary for
Pseudomonas aeruginosa biofilm development. Mol. Microbiol. 30: 295-304.
O'Toole, G., and Kolter, R. (1998b) Initiation of biofilm formation in Pseudomonas
fluorescens WCS365 proceeds via multiple, convergent signalling pathways: a genetic
analysis. Mol. Microbiol. 28: 449-461.
Odagami, T., Suzuki, S., Takama, K., Azumi, K., and Yokosawa, H. (1993)
Characterization of extracellular protease produced by the marine putrefactiva bacteria,
Alteromonas haloplanktis S5B. J. Mar. Biotech. 1: 55-58.
Osorio, C.R., and Klose, K.E. (2000) A region of the transmembrane regulatory protein
ToxR that tethers the transcriptional activation domain to the cytoplasmic membrane displays
wide divergence among Vibrio species. J. Bacteriol. 182: 526-528.
Overbye, L., Sandkvist, M., and Bagdasarian, M. (1993) Genes required for extracellular
secretion of enterotoxin are clustered in Vibrio cholerae. Gene 132: 101-106.
Parkinson, J.S., and Kofoid, E.C. (1992) Communication modules in bacterial signaling
proteins. Annu. Rev. Genet. 26: 71-112.
Pawlik, J.R. (1986) Chemical induction of larval settlement and metamorphosis in the reef-
building tube worm Pragmatopoma californica (Sabellariidae: Polychaeta). Mar. Biol. 91:
59-68.
267
Pawlik, J.R. (1990) Natural and artifical induction of metamorphosis of Phragmatopoma
lapidosa californica (Polychaeta: Sabellariidae), with a critical look at the effect of bioactive
compounds on marine invertebrate larvae. Bull. Mar. Scien. 46: 176-188.
Pawlik, J.R. (1992) Chemical ecology of the settlement of bethic marine invertebrates.
Oceanography, Mar. Biol. Annu. Rev. 30: 273-335.
Person, J.P., Pesci, E.C., and Iglewski, B.H. (1997) Roles of Pseudomonas aeruginosa
las and rhl quorum-sensing systems in control of elastase and rhamnolipid biosynthesis
genes. J. Bacteriol. 179: 5756-5767.
Pesci, E.C., and Igleweski, B.H. (1997) The chain of commands in Pseudomonas quorum
sensing. Trends Microbiol. 5: 132-134.
Polz, M.F., Harbison, C., and Cavanaugh, C.M. (1999) Diversity and heterogeneity of
epibiotic bacterial ccommunities on the marine nematode Eubostrichus dianae. Appl. Environ.
Microbiol. 65: 4271-4275.
Pratt, L., and Kolter, R. (1998) Genetic analysis of Escherichia coli biofilm formation:
roles of flagella, motility, chemotaxis and type 1 pili. Mol. Microbiol. 30: 285-293.
Prigent-Combaret, C., Prensier, G., Le Thi, T.T., Vidal, O., Lejeune, P., and Dorel, C.
(2000) Developmental pathway for biofilm formation in curli-producing Escherichia coli
strains: role of flagella, curli and colanic acid. Environ. Microbiol. 2: 450-465.
Provasoli, L., and Pinter, I.J. (1964) Symbiotic relationship between microorganisms and
seaweeds. Am. J.Bot. 51: 681.
Provasoli, L., and Pintner, I.J. (1980) Bacteria induced polymorphism in an axenic
laboratory strain of Ulva lactuca (Chlorophyceae). J. Phycol. 16: 196-201.
Pruzzo, C., Crippa, A., Bertone, S., Pane, L., and Carli, A. (1996) Attachment of Vibrio
alginolyticus to chitin mediated by chitin-binding proteins. Microbiology 142: 2181-2186.
Pugsley, A. (1993) The complete general secretory pathway in Gram-negative bacteria.
Microbiol. Rev. 57: 50-108.
268
Reeves, P.J., Whitecombe, D., Wharam, S., Gibson, M., Allison, G., Bunce, N.,
Barallon, R., Douglas, P., Mulholland, V., Stevens, S., Walker, D., and Slamond,
G.P.C. (1993) Molecular cloning and characterization of 13 out genes from Erwinia
carotovora subspecies carotovora: genes encoding members of a general secretion pathway
(GSP) widespread in Gram-negative bacteria. Mol. Microbiol. 8: 443-456.
Reich, K., and Schoolnik, G. (1994) The light organ symbiont Vibrio fischeri possesses a
homolog of the Vibrio cholerae transmembrane transcriptional activator ToxR. J. Bacteriol.
176: 3085-3088.
Reimmann, C., Beyeler, M., Latifi, A., Winteler, H., Foglino, M., Lazdunski, A., and
Haas, D. (1997) The global activator GacA of Pseudomonas aeruginosa PAO positively
controls the production of the autoinducer N-butyryl-homoserine lactone and the formation of
virulence factors pyocyanin, cyanide and lipase. Mol. Microbiol. 24: 309-319.
Reizer, J., Reizer, A., and Saier, M.H.J. (1992) A new subfamily of ABC-transport
systems catalyzing export of drugs and carbohydrates. Protein. Sci. 1: 1326-1332.
Rigby, P.W.J., Diekmann, M., Rhodes, C., and Berg, P. (1977) Labelling
deoxyribonucleic acid to high specific activity in vitro by nick translation with DNA
polymerase I. J. Mol. Biol. 113: 237-245.
Ritchie, K.B., Nagelkerken, I., James, S., and Smith, G.W. (2000) A tetrodotoxin-
producing marine pathogen. Nature 404: 354.
Rittschof, D., Hooper, I.R., and Costlow, J.D. (1986) Barnacle settlement inhibitors from
sea pansies, Renilla reniforms. Bull. Mar. Sci. 39: 376-382.
Rittschof, D., Maki, J.S., Mitchell, R., and Costlow, J.D. (1986) Ion and
neuropharmacological studies of barnacle settlement. Neth. J. Sea. Res. 20: 269-275.
Rittschof, D., Branscomb, E.S., and Costlow, J.D. (1989) Bryozoan and barnacle
settlement in relation to initial surface wettability: a comparison of laboratory and field studies.
Scien. Mar. 53: 411-416.
269
Roberts, S.K., Lappin-Scott, H., and Leeming, K. (1999) The control of bacterial-fungal
biofilms. In Proceedings from the fourth meeting of the biofilm club. Wimpenny, J., Gilbert,
P., Walker, J., Brading, M. and Bayston, R. (ed). Powys, UK.
Robins-Browne, R. (1994) Escherichia coli strains that cause diarrhoea: models of bacterial
pathogenisis. In Recent advances in microbiology. Gilbert, G. L. (eds). Melbourne : The
Austrailan Society for Microbiology Inc., pp. 292-375.
Rodriguez, S., Ojeda, P., and Inestrosa, N. (1993) Settlement of bethnic marine
invertebrates. Mar. Ecol. Prog. Ser. 97: 193-207.
Rost, B. (1996) P.H.D. Predicting one-dimensional protein structure by profile based neural
networks. Methods in Enzymology 266: 525-539.
Ruby, E.G., and Lee, K.H. (1998) The Vibrio fischeri-Euprymna scolopes light organ
association: current ecological paradigms. J. Bacteriol. 64: 805-812.
Rumrill, S.S., and Cameron, R.A. (1983) Effects of gamma-aminobutyric acid on the
settlement of larvae of the black chiton Katharina tunicata. Mar. Biol. 72: 243-247.
Ryder, M., and Rovira, A. (1993) Biological control of Take-all of glasshouse-grown
wheat using strains of Pseudomonas corrugata isolated from wheat field soil. Soil Biol.
Biochem. 25: 311-320.
Sacherer, P., Defago, G., and Haas, D. (1994) Extracellular protease and phospholipase C
are controlled by the global regulatory gene gacA in the biocontrol strin Pseudomonas
fluorescens CHAO. FEMS Microbiol. Lett. 116: 155-160.
Saitou, N., and Nei, M. (1987) The neighbour-joining method: a new method for
reconstructing phylogenetic trees. Mol. Biol. Evol. 4: 406-425.
Saraste, M., Sibbald, P.R., and Wittingholfer, A. (1990) The P-loop: a common motif in
ATP- and GTP- binding proteins. Trends Biochem. Sci. 15: 430-434.
Sawabe, T., Makino, H., Tatsumi, M., Nakano, K., Tajima, K., Iqbal, M., Yumoto, I.,
Ezura, Y., and Christen, R. (1998) Pseudoalteromonas bacteriolytica sp.nov., a marine
270
bacterium that is the causative agent of red spot disease of Laminaria japonica. Int. J. Syst.
Bacteriol. 48: 769-774.
Sawabe, T., Tanaka, R., Iqbal, M.M., Tajima, K., Ezura, Y., Ivanova, E., and
Christen, R. (2000) Assignment of Alteromonas elyakovii KMM 162T and five strains
isolated from spot-wounded fronds of Laminaria japonica to Pseudoalteromonas elyakovii
comb. nov. and the extended description of the species. Int. J. Syst. Evol. Micro. 50: 265-
271.
Schneider, R.P., Chadwick, B.R., Pembrey, R., Jankowski, J., and Acworth, I. (1994)
Retention of the Gram-negative bacterium SW8 on surfaces under conditions relevant to the
subsurface environment: Effects of conditioning films and substratum nature. FEMS
Microbiol. Ecol. 14: 243-254.
Schumacher, D., and Klose, K. (1999) Environmental signals modulate ToxT-dependent
virulence factor expression in Vibrio cholerae. J. Bacteriol. 181: 1508-1514.
Seiburth, J., and Tootle, J.L. (1981) Seasonality of microbial fouling on Ascophyllum
nodosum (L.) Lejol, Fucus vesiculosus L., Polysiphonia lanosa (L.) Tandy and Chondrus
crispus Stackh. J. Phycol. 17: 57-64.
Siebert, P., Chenchik, A., Kellogg, D., Lukyanov, K., and Lukyanov, S. (1995) An
improved PCR method for walking in uncloned genomic DNA. Nucleic Acids Res. 23: 1087-
1088.
Simidu, U., Kita-Tsukamoto, K., Yasumoto, T., and Yotsu, M. (1990) Taxonomy of
four marine bacterial strains that produce tetrodotoxin. Int. J. Syst. Bacteriol. 40: 331-336.
Skerman, V.B.D. (1967) A guide to the identification of the genera of bacteria. In Abstracts
of microbiological methods. Skerman, V. B. D. (eds). New York : Wiley, pp. 147.
Skerratt, J., Bowman, J.P., Nichols, P.D., Lovejoy, C., and Hallegraef, G. (1998)
Algicidal bacteria: Algal bacterial interactions. In The Australian Society for Microbiology,
annual scientific meeting and exhibition, conference proceedings. The Australian Society for
Microbiology (eds) Hobart, Tasmania.
271
Skorupski, K., and Taylor, R. (1999) A new level in the Vibrio cholerae ToxR virulence
cascade: AphA is required for transcriptional activation of the tcpPH operon. Mol. Microbiol.
31: 763-771.
Slattery, M., Hamann, M., McClintock, J., Perry, T., Puglisi, M., and Yoshida, W.
(1997) Ecological roles for water-borne metabolites from Antartic soft corals. Mar. Ecol.
Prog. Ser. 161: 133-144.
Sperandio, V., Giron, J.A., Silveira, W.D., and Kaper, J.B. (1995) The OmpU outer
membrane protein, a potential adherence factor of Vibrio cholerae. Infect. Immun. 63: 4433-
4438.
Stackebrandt, E., and Goebel, B.M. (1994) Taxonomic note: a place for DNA-DNA
reassociation and 16S rRNA sequence analysis in the present species definition in
bacteriology. Int. J. Syst. Bacteriol. 44: 846-849.
Standing, J.D., Hooper, I.R., and D., C.J. (1984) Inhibition and induction of barnacle
settlement by natural products present in octocorals. J. Chem. Ecol. 10: 823-833.
Steinberg, P.D., de Nys, R., and Kjelleberg, S. (2001) Chemical mediation of surface
colonisation. In. Marine Ecology. McClinton, J., Backer, B. (eds) CRC Press LLC, Boca
Ratun, Florida (In Press)
Stelzer, S. (1999) Investigation of the antibacterial protein produced by Pseudoalteromonas
tunicata, Honours Thesis. University of New South Wales, Sydney, Australia.
Stewart, C., and de Mora, S.J. (1990) A review of the degredation of tri(n-butyl)tin in the
marine environment. Env. Tech. 11: 565-570.
Stock, J.B., Ninfa, A.J., and Stock, A.M. (1989) Protein phosphorylation and regulation
of adaptive responses in bacteria. Microbiol. Rev. 53: 450-490.
Surette, M.G., Miller, M.B., and Bassler, B.L. (1999) Quorum sensing in Escsherichia
coli, Salmonella typhimurium and Vibrio harveyi: a new family of genes responsible for
autoinducer production. Proc. Natl. Acad. Sci. USA 96: 1639-1644.
272
Swift, S., Bainton, J., and Wison, M. (1994) Gram-negative bacterial communication by
N-acyl homoserine lactones: a universal language? Trends in microbiology 2: 193-197.
Swift, S., Karlyshev, A.V., Fish, L., Durant, E.L., Winson, M.K., Chhabra, S.R.,
Williams, P., MacIntyre, S., and Stewart, G.S.A.B. (1997) Quorum sensing in
Aeromonas hydrophila and Aeromonas salmonicida: identification of the LuxRI homologs
AhyRI and AsaRI and their cognate N-acylhomoserine lactone signal molecules. J. Bacteriol.
179: 5271-5281.
Szewzyk, U., Holmström, C., Wangstadh, M., Samuelson, M.O., Maki, J.S., and
Kjelleberg, S. (1991) Relevance of the exoploysaccharide of marine Pseudomonas sp. strain
S9 for the attachment of Ciona intestinalis larvae. Mar. Ecol. Prog. Ser. 75: 259-265.
Targett, N.M., Bishop, S.S., McConnell, O.J., and Yoder, J.A. (1983) Antifouling agents
against the bethnic marine diatom, Navicula salinicola: homerine from the gorgonians
Leptogorgia virgulata and L. setacea and analogs. J. Chem. Ecol. 9: 817-828.
Tatewaki, M., Provasoli, L., and Pinter, I.J. (1983) Morphogenesis of Monostroma
oxyspermum (Kutz) Doty (Chlorophyceae) in axenic culture, especially in bialgal culture. J.
Phycol. 19: 409-416.
Tegtmeyer, K., and Rittschof, D. (1989) Synthetic peptide analogs to barnacle settlement
pheremone. Peptides 9: 1403-1406.
Thomas, R.W.S.P., and Allsopp, D. (1983) The effects of certain periphytic marine
bacteria upon the settlement and growth of Enteromorpha, a fouling alga. Biodeterioration 5:
348-357.
Thomashow, L.S., and Weller, D.M. (1988) Role of phenazine antibiotic from
Pseudomonas fluorescens in biological control of Gaeumannomyces graminis var. tritici. J.
Bacteriol. 170: 3499-3508.
Tillett, D., and Neilan, B.A. (2000) Rapid nucleic acid isolation from cultured and
environmental cyanobacteria: Novel techniques based on xanthogenate. J. Phycol. 36: 251-
258.
273
Todd, J.S., Zimmerman, R.C., Crews, P., and Alberte, R.S. (1993) The antifouling
activities of natural and synthetic phenolic sulphate esters. Phytochemistry 34: 401-404.
Toren, A., Landau, L., Kushmaro, A., Loya, Y., and Rosenberg, E. (1998) Effect of
temperature on adhesion of Vibrio strain AK-1 to Oculina patagonia and coral bleaching.
Appl. Environ. Microbiol. 64: 1379-1384.
Tosato, V., Albertini, A., Zotti, M., Sonda, S., and Bruschi, C. (1997) Sequence
completion, identification and definition of the fengycin operon in Bacillus subtilis 168.
Microbiology 143: 3443-3450.
Tosteson, T.R., and Corpe, W.A. (1975) Enhancement of adhesion of the marine Chorella
vulgaris to glass. Can. J. Microbiol. 21: 1025-1031.
Trapido-Rosenthal, H., and Morse, D.E. (1986) Regulation of receptor mediated
settlement and metamorphosis in larvae of a gastropod mollusc (Haliotis Rufescens). Bull.
Mar. Sci. 39: 383-392.
Tsukamoto, S., Kato, H., Hirota, H., and Fusetani, N. (1997) Antifouling terpenes and
steroids against barnacle larvae from marine sponges. Biofouling 11: 283-291.
Tsukamoto, S., Kato, H., Hirota, H., and Fusetani, N. (1999) Lumichrome: a larval
metamorphosis-inducing substance in the acidian Halocynthia roretzi. Eur. J. Biochem. 264:
785-789
Turgay, K., Krouse, M., and Marahiel, M.A. (1992) Four homologous domains in the
primary structure of GrsB are related to domains in a superfamilty of adenylate-forming
enzymes. Mol. Microbiol. 529-546.
Uchida, M., Nakata, K., and Maeda, M. (1997) Conversion of Ulva fronds to a hatchery
diet for Artemia naupli utilizing the degrading and attaching abilities of Pseudoalteromonas
espejiana. J. Appl. Phycol. 9: 541-549.
Upton, C., and Buckley, J.T. (1995) A new family of lipolytic enzymes? Trends Biochem.
Sci. 20: 178-179.
274
Van Landschoot, A., and De Ley, J. (1983) Intra- and intergeneric similarities of the rRNA
cistrons of Alteromonas, Marinomonas (gen. nov.) and some other Gram-negative bacteria. J.
Gen. Microbiol. 129: 3057-3074.
Vanittanakom, N., Loeffler, W., Koch, U., and Jung, G. (1986) Fengycin- A novel
antifungal lipopeptide antibiotic produced by Bacillus subtilis F-29-3. The Journal of
Antibiotics 34: 888-901.
Venkateswaran, K., and Dohmoto, N. (2000) Pseudoalteromonas peptidolytica sp. nov., a
novel marine mussel-thread-degrading bacterium isolates from the sea of Japan. Int. J. Syst.
Evol. Micro. 50: 565-574.
Vera, J., Alvarez, R., Murano, E., Slebe, J.C., and Leon, O. (1998) Identification of a
marine agarolytic Pseudoalteromonas isolate and characterisation of its extracellular agarase.
Appl. Environ. Microbiol. 64: 4378-4383.
Vidal, O., Longin, R., Prigent-Combaret, C., Dorel, C., Hooreman, M., and Lejeune,
P. (1998) Isolation of an Escherichia coli K-12 mutant strain able to from biofilms on inert
surfaces: Involvement od a new ompR allele that increase curli expression. J. Bacteriol. 180:
2442-2449.
Vincent, M., Harrison, L., Brackin, J., Kovacevich, P., Mukerji, P., Weller, D., and
Pierson, E. (1991) Genetic analysis of the antifungal activity of a soilborne Pseudomonas
aureofaciens strain. Appl. Environ. Microbiol. 57: 2928-2934.
Vincent, P., Pignet, P., Talmont, F., Bozzi, L., Fournet, B., Guezennec, J., Jeanthon,
J., and Prieur, D. (1994) Production and characterization of an exopolysaccharide excreted
by a deep-sea hydrothermal vent bacterium isolated from the polychaeta annelid Alvinella
pompejana. Appl. Environ. Microbiol. 60: 4134-4141.
Voisard, C., Keel, C., Haas, D., and Defago, G. (1989) Cyanide production by
Pseudomonas fluorescens helps suppress black rot of tobacco under gnotobiotic conditions.
EMBO J. 8: 351-358.
Wahl, M. (1989) Marine epibiosis.1. Fouling and antifouling : some basic aspects. Mar.
Ecol. Progr. Ser. 58: 175-186.
275
Wahl, M., Jensen, R. A., and Fenical, W. (1994) Chemical control of bacterial epibiosis
on ascidians. Mar. Ecol. Progr. Ser. 110: 45-57.
Waldor, M.K., and Melkalanos, J.J. (1996) Lysogeneic conversion by a filamentous
phage encoding cholera toxin. Science 272: 1910-1914.
Walker, J.E., Saraste, M., Runswick, M.J., and Gay, N.J. (1982) Distantly related
sequences in the α and β- subunits of ATP synthase, myosin, kinases and other ATP-
requiring enzymes and a common nucleotide binding fold. EMBO J 1: 945-951.
Walls, J.T., Ritz, D.A., and Blackman, A.J. (1993) Fouling, surface bacteria and anti-
bacterial agents of four bryozoan species found in Tasmaina, Australia. J. Exp. Mar. Biol.
Ecol. 169: 1-13.
Wandersman, C. (1996) Secretion across the bacterial outer membrane. In Escherichia coli
and Salmonella: cellular and molecular biology. Neidhart, F. C., Curtiss III, R., Ingraham, J.
L., Lin, E. C. C., Low, K. B., Magasanik, B., Reznikoff, W. S., Riley, M., Schaechter, M. and
Umbarger, H. E. (eds). Washington, D.C : American Society for Microbiology, pp. 955-966.
Watnick, P., and Kolter, R. (1999) Steps in the development of a Vibrio cholerae EI Tor
biofilm. Mol. Micriobiol. 34: 586-595.
Watson, N., Dunyak, D., Rosey, E., Slonczewski, J., and Olson, E. (1992) Identification
of elements involved in transcripitional regulation of the Escherichia coli cad operon by
external pH. J. Bacteriol. 174: 530-540.
Way, J.C., Davis, M.A., Morisato, D., Roberts, D.E., and Kleckner, N. (1984) New
Tn10 derivatives for transposon mutagenesis and for construction of lacZ operon fusions by
transposition. Gene 32: 369-379.
Weidner, S., Arnold, W., and Puhler, A. (1996) Diversity of uncultured microorganisms
associated with the seagrass Halophila stipulacea estimated by restriction fragment lenght
polymorphism analysis of PCR-amplified 16S rRNA genes. Appl. Environ. Microbiol. 62:
766-771.
276
Weiner, R.M., Segall, A.M., and Colwell, R.R. (1985) Characterization of a marine
bacterium associated with Crassostrea virginica (the eastern oyster). App. Environ.
Microbiol. 49: 83-90.
Weiner, R.M., Coyne, V.E., Brayton, P., and Raiken, R. (1988) Alteromonas
colwelliana sp. nov., and isolate from oyster habitats. Int. J. Syst. Bacteriol. 38: 240-244.
Welch, T., and Bartlett, D. (1998) Identification of a regulatory protein required for
pressure-responsive gene expression in the deep-sea bacterium Photobacterium species strain
SS9. Mol. Microbiol. 27: 977-985.
Whitchurch, C.B., Alm, R.A., and Mattick, J.S. (1996) The alginate regulator AlgR and
an associated sensor FimS are required for twiching motility in Pseudomonas aeruginosa.
Proc. Natl. Acad. Sci. USA 93: 9839-9843.
Wieczorek, S., Clare, A., and Todd, C. (1995) Inhibitory and facilitatory effects of
microbial films on settlement of Balanus amphitrite amphitrite larvae. Mar. Ecol. Progr. Ser.
119: 221-228.
Wieczorek, S., and Todd, C. (1997) Inhibition and facilitation of bryozoan and ascidian
settlement by natural multi-species biofilms: effects of film age and the roles of active and
passive larval attatchment. Mar. Biol. 128: 463-473.
Williamson, J.E., de Nys, R., Kumar, N., Carson, D.G., and Steinberg, P.D. (2000)
Induction of metamorphosis in the sea urchin Holopneustes purpurascens by a metabolite
complex from the algal host Delisea pulchra. Biol. Bull. 198: 332-345.
Winzeler, E. A., Shoemaker, D.D, Astromoff, A., Anderson, K., Andre, B., Bangham,
R., Benito, R., Boeke, J. D., Bussy, H., Chu, A.M., Connelly, C., Davis, K., Dietrich,
F., Dow, S.W., EL Bakkoury, M., Foury, F., Friend, S. H., Gentalen, E., Giaever, G.,
Hegemann, J.H., Jones, T., Laub., Liao. H., Leibundguth, N., Davis, R.W. et al (1999)
Functional characterization of the S. cerevisiae genome by gene deletion and parallel analysis.
Science. 285: 901-906.
277
Wong, S.M., Carroll, P.A., Rahme, L.G., Ausubel, F.M., and Calderwood, S.B. (1998)
Modulation of expression of the ToxR regulon in Vibrio cholerae by a member of the two-
component family of response regulators. Infect. Immun. 66: 5854-5861.
Wood, D.W., and Pierson, L.S. (1996) The phzI gene of Pseudomonas aureofaciens 30-84
is responsible for the production of a diffusable signal required for phenazine antibiotic
production. Gene 168: 49-53.
Woodin, S.A. (1991) Recruitment of infauna: positive and negative cues? Amer. Zool. 31:
797-807.
Ye, L., Buck, L.M., and Schaeffer, M.J. (1997) Cloning and sequencing of a cDNA for
firefly luciferase from Photuris pennsylvanica. Biochem. Biophys. Acta 1339: 39-52.
Yoshikawa, K., Takadera, T., Adachi, K., Nishijima, M., and Sano, H. (1997)
Korormicin, a novel antibiotic speciffically active against marine gram-negative bacteria,
produced by a marine bacterium. The Journal of Antibiotics 50: 949-953.
Yoshinaga, I., Kawai, T., and Ishida, Y. (1997) Analysis of algicidal ranges of the bacteria
killing the marine dinoflagellate Gymnodinium mikimotoi isolates from Tanabe bay,
Wakayama Pref. Fish. Sci. 63: 94-98.
Yu, C., Lee, A.M., Bassler, B.L., and Roseman, S. (1991) Chitin utilization by marine
bacteria: A physiological function for bacterial adhesion to immobilized carbohydrates. J.
Biol. Chem 25: 24260-24267.
Yu, R., and DiRita, V. (1999) Analysis of an autoregulatory loop controlling ToxT, cholera
toxin, and toxin-coregulated pilus production in Vibrio cholerae. J. Bacteriol. 181: 2584-
2592.
Zhang, J., and Deutscher, M.P. (1988) Escherichia coli RNase D: sequence of the rnd
structural gene and purification of the overexpressed protein. Nucleic acids res. 16: 6265-
6278.
Zimmer-Faust, R.K., and Tamburri, M.N. (1994) Chemical identity and ecological
implications of a waterborne, larval settlement cue. Limnol. Oceanogr. 39: 1075-1087.
278
ZoBell, C.E., and Allen, E.C. (1935) The significance of marine bacteria on the fouling of
submerged surfaces. J. Bacteriol. 29: 239-251.