Autolysis in the development and dispersal of biofilms formed by the marine bacterium Pseudoalteromonas tunicata
Anne Mai-Prochnow
A thesis in fulfilment of the requirements for the degree of
Doctor of Philosophy
School of Biotechnology and Biomolecular Sciences
Faculty of Science
The University of New South Wales,
Sydney, Australia
August, 2006 Table of Contents Table of Contents ...... 2
Acknowledgements...... 8
Abstract...... 10
Originality Statement...... 12
List of Publications...... 13
List of Figures...... 14
List of Tables ...... 16
List of Abbreviations ...... 17
1. General introduction and literature review ...... 19
1.1 Biofilm development ...... 21
1.1.1 Biofilm attachment...... 21
1.1.2 Biofilm maturation and differentiation ...... 24
1.1.2.1 Cell death during biofilm differentiation ...... 25
1.1.3 Biofilm dispersal...... 26
1.1.4 The affect of cyclic di-GMP on biofilm development...... 28
1.2 Genetic diversification of biofilm cells ...... 29
1.3 Cell to cell communication within biofilms – quorum sensing ...... 32
1.3.1 The involvement of quorum sensing systems in the formation of a
differentiated biofilm architecture ...... 33
1.4 Are biofilms multicellular communities?...... 34
1.4.1 Cell death during differentiation processes in prokaryotes...... 36
1.4.1.1 Cell death during fruiting body formation of Myxococcus xanthus....36
1.4.1.2 Cell death during sporulation of Bacillus subtilis...... 39
2 1.4.1.3 Cell death during mycelium differentiation of Streptomyces
antibioticus...... 41
1.5 The genus Pseudoalteromonas...... 43
1.5.1 Pseudoalteromonas species are associated with eukaryotic hosts...... 44
1.5.2 Production of secondary metabolites in Pseudoalteromonas species.....45
1.5.3 Pseudoalteromonas tunicata...... 47
1.5.3.1 The antibacterial and autolytic protein produced by P. tunicata ...... 49
1.6 Aims and objectives of this study...... 50
2. Biofilm development and cell death in Pseudoalteromonas tunicata.
...... 52
2.1 Introduction...... 52
2.2 Material and Methods...... 55
2.2.1 Bacterial strains and culture media ...... 55
2.2.2 Biofilm experiments...... 55
2.2.3 Biofilm staining...... 56
2.2.3.1 Confocal laser scanning microscopy...... 57
2.2.3.2 Biofilm structure analysis with COMSTAT ...... 57
2.2.4 Site-directed mutagenesis of alpP...... 59
2.2.5 Characterization of ǻAlpP mutant ...... 60
2.2.5.1 Testing ǻAlpP mutant supernatant for antibacterial activity...... 60
2.2.5.2 Assessment of ǻAlpP mutant growth ...... 60
2.2.5.3 Acrylamide gel analysis of P. tunicata wild-type and ǻAlpP mutant 60
2.2.5.3.1 Preparation of concentrated supernatant and partial AlpP
purification ...... 60
2.2.5.3.2 Native PAGE analysis and overlay assay ...... 61
3 2.2.5.4 Liquid chromatography tandem mass spectrometry (LC-MSMS)
analysis ...... 62
2.2.5.5 Characterization of ǻAlpP mutant biofilm formation...... 63
2.2.5.6 Add-back of purified AlpP to the ǻAlpP mutant and wild-type
biofilms ...... 63
2.3 RESULTS...... 64
2.3.1 P. tunicata biofilm development and cell death...... 64
2.3.2 Characterization of the ǻAlpP mutant ...... 66
2.3.2.1 Assessment of ǻAlpP mutant growth ...... 66
2.3.2.2 Drop test assay for ǻAlpP mutant...... 67
2.3.2.3 PAGE analysis of P. tunicata wild-type and ǻAlpP mutant supernatant
...... 68
2.3.2.3.1 LC-MSMS analysis...... 69
2.3.3 Biofilm development of the P. tunicata ǻAlpP mutant...... 69
2.3.4 Characterization of the biofilm structure with COMSTAT ...... 70
2.3.5 Add-back of purified AlpP to P. tunicata biofilms...... 72
2.4 Discussion...... 74
3. The mode of action of AlpP in P. tunicata ...... 77
3.1 Introduction...... 77
3.2 Materials and Methods...... 79
3.2.1 Bacterial strains and culture conditions ...... 79
3.2.2 AlpP purification...... 79
3.2.2.1 Sodium dodecyl sulphate polyacrylamide gel (SDS PAGE) analysis of
purified AlpP...... 80
3.2.2.2 Determination of protein concentration ...... 80
4 3.2.3 Drop test assay for AlpP activity testing...... 81
3.2.4 Amplex Red assay...... 81
3.2.5 Determination of the minimum inhibitory catalase concentration...... 82
3.2.6 Assessment of storage conditions on the stability of AlpP...... 82
3.2.7 Biofilm experiments...... 82
3.2.7.1 Detecting hydrogen peroxide in biofilms...... 82
3.2.7.2 Removing hydrogen peroxide from biofilms...... 83
3.2.8 Assessment of catalase on growth of P. tunicata...... 83
3.3 Results ...... 84
3.3.1 AlpP purification...... 84
3.3.1.1 Determination of AlpP concentration and activity ...... 84
3.3.1.1.1 AlpP storage conditions ...... 85
3.3.2 AlpP mode of action investigation...... 86
3.3.2.1 Minimum catalase concentration to abolish AlpP activity ...... 87
3.3.3 Hydrogen peroxide detection in P. tunicata biofilms ...... 88
3.3.3.1 Hydrogen peroxide detection in ǻAlpP mutant biofilms...... 89
3.3.4 The effect of catalase on microcolony formation in P. tunicata wild-type
biofilms ...... 91
3.3.5 Assessment of catalase on growth of P. tunicata...... 93
3.4 Discussion...... 94
4. Ecological advantages of autolysis during biofilm development and dispersal of P. tunicata...... 97
4.1 Introduction...... 97
4.2 Materials and Methods...... 100
4.2.1 Bacterial strains and culture media ...... 100
5 4.2.2 Biofilm experiments...... 100
4.2.3 Phenotypic characterization of biofilm dispersal cells...... 100
4.2.4 Phenotypic variation of batch culture cells ...... 101
4.2.5 Calculating the variation coefficient ...... 101
4.2.6 Add back experiments...... 102
4.2.7 Analysing metabolic activity of biofilm effluent...... 103
4.2.7.1 Fluorescent staining of biofilm effluent...... 103
4.2.7.2 Flow cytometry ...... 103
4.2.8 Starvation experiments...... 104
4.3 Results ...... 105
4.3.1 Quantification of biofilm dispersal cells...... 105
4.3.1.1 AlpP add back to induce dispersal of ǻAlpP mutant biofilms...... 106
4.3.2 Metabolic activity of biofilm dispersal cells...... 107
4.3.3 Phenotypic variation of biofilm dispersal cells...... 109
4.3.4 AlpP add back to induce phenotypic variation in ǻAlpP mutant dispersal
cells ...... 115
4.3.5 Phenotypic variation in batch culture...... 115
4.3.6 Influence of nutrient starvation on metabolic activity within biofilms.119
4.4 Discussion...... 122
5. AlpP homologues appear to have a conserved function during biofilm development and dispersal of several Gram negative organisms
...... 127
5.1 Introduction...... 127
5.2 Materials and Methods...... 131
5.2.1 Strains and culture conditions ...... 131
6 5.2.2 Purification of LodA from Marinomonas mediterranea ...... 132
5.2.3 Biofilm experiments...... 133
5.2.3.1 Biofilm staining...... 133
5.2.3.2 Removal of hydrogen peroxide from biofilms...... 134
5.2.3.3 Add back of LodA protein to SB1 mutant biofilms...... 134
5.2.4 Phenotypic variation of M. mediterranea dispersal cells...... 134
5.3 Results ...... 135
5.3.1 AlpP homologues in Gram negative organisms...... 135
5.3.2 Biofilm development and cell death in M. mediterranea ...... 137
5.3.3 Biofilm development of the SB1 mutant strain ...... 137
5.3.4 Phenotypic variation of M. mediterranea biofilm dispersal cells...... 140
5.3.5 Biofilm formation and cell death in C. violaceum, C. crescentus and M.
degradans...... 142
5.3.6 Detection of hydrogen peroxide in M. mediterranea, C. violaceum, C.
crescentus and M. degradans biofilms ...... 143
5.4 Discussion...... 145
6. General discussion and future outlooks...... 149
6.1 Cell death promotes biofilm dispersal and phenotypic variation...... 149
6.2 Biofilm dispersal displays similarities to dispersal of sessile invertebrates.
...... 151
6.3 A novel mechanism mediates biofilm cell death in P. tunicata and other
Gram negative organisms...... 152
6.4 Future directions ...... 154
Appendix ...... 156
References ...... 160
7 Acknowledgements
Foremost, I would like to give a big thanks to my supervisor Staffan Kjelleberg for giving me the opportunity of undertaking a PhD in his lab. Your ideas and enthusiasm are always inspiring and encouraging and made me enjoy science and doing a PhD very much.
Another big thanks must go to Jeremy, who has been an excellent co-supervisor and friend. You have a great passion for science and particularly biofilms. You have been with this project from the beginning and taught me everything about biofilms as well as scientific writing in English. Thank you also for the BBQ and movie nights with Sarah.
To all the people who are working or have worked with D2 (P. tunicata) and your helpful discussions: Su, Sacha, Dhana, Doralyn, Flavia, Carola, Brendan and Cathy. D2 is a great bacterium with endless interesting, always surprising characteristics. I am sure, that lots of exciting things will be discovered with the new genome data.
All the other people in Lab 304 who made my PhD years so enjoyable: Kirsty, Sharon,
Ani, Bec, Niina, Megan, Maria, Vicky; and the people from CMBB with all the nice equipment: Ana-Maria, Johnny, Krager, Nidhi, Nicolas, Leena, Scott and Diane.
A special thanks to Patricia who helped me a lot with the protein mode of action work and Marinomonas biofilms. It was so much fun to have you here and I am very happy to have you as a friend.
Thanks to Belinda Ferrari for helping me with the flow cytometry at Macquarie
University. Thanks also to Mark Raftery for collaborating with the mass spectrometry at
BMSF and to Torsten Thomas for help with the AlpP purification work.
8 Big thanks to Adam and Evi for knowing all the administration things. You guys are a great help for all of us and cannot be appreciated enough.
Thank you to Bill O’Sullivan for fast and thorough proof reading of my thesis.
To Joyce and Anne-Dorothee for the nice coffee and lunch breaks when I needed to talk to people outside my lab.
A very special thanks goes to the girls from the “breakfast crew”: Evi, Ana-Maria,
Lyndal and Su. Your friendship and our chats about life and especially pregnancy always helped me so much. I can’t wait till all our little ones are born. This is for our children.
I also want to thank my family, especially my parents and my brother Martin, who are always so supportive and proud of me, which makes me feel very special. Thanks for making the long trips to Australia to visit us. I could not have done this without you.
To the love of my life, René, who is always there for me and helps me in every possible way: discussing science stuff and spending endless hours helping me with computer issues. But most of all, I am grateful for the wonderful relationship we have and that we will have a little daughter soon. It will be so exciting to have our own little family and doing things other than science for a while.
9 Abstract
The marine bacterium Pseudoalteromonas tunicata produces target-specific inhibitory compounds against bacteria, algae, fungi and invertebrate larvae and is frequently found in association with living surfaces in the marine environment. This study examined the ability of P. tunicata to form biofilms under continuous culture conditions within the laboratory. P. tunicata biofilms exhibited a characteristic architecture consisting of differentiated microcolonies surrounded by water-channels. Interestingly, a repeatable pattern of cell death in the centre of microcolonies was observed. The antibacterial and autolytic protein, AlpP, produced by P. tunicata was found to be involved in this biofilm killing and a ǻAlpP mutant derivative of P. tunicata did not show cell death during biofilm development. AlpP mediated biofilm killing occurred via the production of hydrogen peroxide due to lysine oxidase activity. Significantly, the process of biofilm cell death was found to be linked to two parameters of importance to the ecology of P. tunicata: dispersal and phenotypic variation. Cell death in P. tunicata wild-type biofilms led to a major reproducible dispersal event after 192 h of biofilm development.
The dispersal was not observed in the ǻAlpP mutant strain. Furthermore, it was shown that P. tunicata wild-type biofilm dispersal cells have enhanced metabolic activity and considerably greater phenotypic variation compared to those cells that disperse from
ǻAlpP mutant biofilms. Wild-type dispersal cells showed significant increases in variation in growth, motility and biofilm formation which may be important for successful colonization of new surfaces. Moreover, AlpP homologues were identified in a range of other Gram negative organisms. These homologues are proposed to have a similar function as they also cause biofilm cell death in at least four of these organisms:
Marinomonas mediterranea, Chromobacterium violaceum, Caulobacter crescentus and
Microbulbifer degradans. In conclusion, the findings of this study suggest that autocidal
10 events mediated by an antibacterial protein can confer ecological advantages to the species by generating a metabolically active and phenotypically diverse subpopulation of dispersal cells. It is proposed that AlpP-mediated biofilm killing and its subsequent role in dispersal and colonization is a conserved mechanism of importance to the biofilm lifestyle across a range of Gram negative microorganisms.
11 Originality Statement
‘I hereby declare that this submission is my own work and to the best of my knowledge it contains no materials previously published or written by another person, or substantial proportions of material which have been accepted for the award of any other degree or diploma at UNSW or any other educational institution, except where due acknowledgement 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.’
Anne Mai-Prochnow
12 List of Publications
Mai-Prochnow, A., F. Evans, D. Dalisay-Saludes, S. Stelzer, S. Egan, S. James, J. S.
Webb and S. Kjelleberg (2004). "Biofilm development and cell death in the marine bacterium Pseudoalteromonas tunicata." Appl Environ Microbiol 70 (6): 3232-8.
Mai-Prochnow, A., B. C. Ferrari, J. S. Webb and S. Kjelleberg (2006). “Ecological advantages of autolysis during the development and dispersal of Pseudoalteromonas tunicata biofilms.” Appl Environ Microbiol 72 (8): in press
Submitted:
Mai-Prochnow A., P. Lucas-Elio, S. Egan, T. Thomas, J. S. Webb, A. Sanchez-Amat, and S. Kjelleberg “The autolytic protein, AlpP and its homologues play a similar role during biofilm differentiation and dispersal of several Gram negative organisms”
13 List of Figures
Figure 1.1: Steps of biofilm formation...... 21
Figure 1.2: Variant colonies on agar produced by a 5-day-old Pseudomonas aeruginosa
biofilm ...... 30
Figure 1.3: Structural similarities between microcolonies from bacterial biofilms and
fruiting bodies formed by social Myxococcus spp...... 38
Figure 1.4: Cell death during differentiation of S. antibioticus ...... 43
Figure 2.1: Flow cell setup...... 56
Figure 2.2: Biofilm development and cell death of the P. tunicata wild-type strain...... 65
Figure 2.3: Growth curve of P. tunicata wild-type (Ƈ) and ǻAlpP mutant (Ŷ) ...... 66
Figure 2.4: P. tunicata drop-plate assay...... 67
Figure 2.5: Native PAGE analysis (A) and overlay of P. tunicata wild-type and ǻAlpP
mutant (B)...... 68
Figure 2.6: Biofilm development of the P. tunicata ǻAlpP mutant ...... 70
Figure 2.7: Addition of purified of AlpP to P. tunicata biofilms ...... 73
Figure 3.1: SDS PAGE analysis of AlpP purification steps ...... 84
Figure 3.2: Drop plate assay for purified AlpP...... 85
Figure 3.3: Amplex Red assay for purified AlpP...... 87
Figure 3.4: Catalase effect on AlpP activity ...... 88
Figure 3.5: Hydrogen peroxide detection in P. tunicata wild-type biofilms ...... 89
Figure 3.6: Hydrogen peroxide detection in ǻAlpP mutant biofilms ...... 90
Figure 3.7: Catalase effect on microcolony formation in P. tunicata wild-type...... 92
Figure 3.8: Growth curve P. tunicata wild-type in the presence of catalase ...... 93
Figure 4.1: Dispersal of viable cells from P. tunicata wild-type and ǻAlpP mutant
biofilms...... 106
14 Figure 4.2: Add back of purified AlpP (Ŷ) and Tris (20 mmol) buffer control (Ÿ) ....107
Figure 4.3: Flow cytometry analysis of biofilm effluent ...... 109
Figure 4.4: Biofilm effluent of a 7 day old P. tunicata wild-type biofilm spread onto
VNSS agar...... 111
Figure 4.11 (previous page): Starvation influence on 72 h P. tunicata biofilms...... 121
Figure 5.1: Biofilm development of M. mediterranea wild-type and SB1 mutant...... 139
Figure 5.2: Add back of the LodA to M. mediterranea SB1 mutant ...... 140
Figure 5.3: Phenotypic variation in M. mediterranea wild-type and SB1 mutant biofilm
dispersal cells...... 141
Figure 5.4: Mature biofilms of other organisms containing the AlpP homologue ...... 142
15 List of Tables
Table 2.1: Characterization of P. tunicata wild-type and ǻAlpP mutant biofilm structure
using the biofilm image analysis software COMSTAT...... 72
Table 5.1: Bacterial strains and culture media...... 132
Table 5.2 AlpP homologues in Gram negative organisms...... 136
Table 5.3: Detection of hydrogen peroxide and cell death in biofilms with and without
the addition of catalase...... 144
16 List of Abbreviations
A: Ampere AHL : Acylated homoserine lactone AR: Amplex red BLAST: Basic Local Alignment Search Tool C: Celsius c-di-GMP: Bis-(3’-5’)-cyclic dimeric guanosine monophosphate CFU: Colony forming unit CLSM: Confocal laser scanning microscopy CTC: 5-cyano-2,3,-ditolyl tetrazolium chloride CV: Coefficient of variation DGC: Diguanylate cyclase
DiBAC4(3): Bis-(1,3 -dibutylbarbituric acid) trimethine oxonol DNA: Deoxyribonucleic acid EDTA: Ethylene diamine tetraacetic acid, trisodium salt EPS: Exopolysaccharide FCM: Flow cytometry FSC: Forward scatter channel g: gram h: hour(s) HPR: Horseradish peroxidase kb: kilobase(s), 1000 bp kDa: kilodalton(s), 1000 Da Km: Kanamycin l: litre LPS: Lipopolysaccharide m: milli (10-3) µ: micro (10-6) M: Molar (= mole per litre) min: minute MMM: Marine minimal media MOPS: Morpholinepropanesulfonic acid MW: Molecular weight
17 NCBI: National Center for Biotechnology Information NSS: Nine salts solution OD: Optical density ORF: Open reading frame PAGE: Polyacrylamide gel electrophoresis PCD: Programmed cell death PCR: Polymerase chain reaction PDE: Phosphodiesterase PI: Propidium iodide QS: Quorum sensing SCV: Small colony variant SDS: Sodium dodecyl sulphate SSC: Side scatter channel Sm: Streptomycin sec: second v/v: volume per volume VNSS: V-medium modified from väätänen w/v: weight per volume
18 Chapter 1 General introduction and literature review
1. General introduction and literature review
Biofilms were first described in 1936 by Zobell and Anderson who reported that bacterial numbers could increase more when a large surface area was available.
However, it is only in recent years that biofilms have been the focus of intense research.
This reflects their importance in both clinical and environmental situations but also that they follow a developmental program that may represent a prokaryotic based precursor to multicellularity. The importance of biofilms has been emphasised by the proposal that they may be the predominant mode of growth for bacteria (Costerton et al., 1995).
Controlling the survival and spread of biofilms is of significance since biofilms can cause many problems. They are a primary source for medical infections (Costerton et al., 1999; Chen, 2001; Tenke et al., 2006), cause damage to industrial equipment
(Lappin-Scott and Costerton, 1989) and contaminate food products (Zottola and
Sasahara, 1994) leading to large expenses. Conventional methods of killing bacteria, such as the use of antibiotics or disinfectants are often not practical, since biofilm bacteria demonstrate an increased resistance to toxic compounds. Thus the development of alternative methods to eradicate biofilms is important.
Biofilm formation has been well studied in a variety of pathogens, including
Pseudomonas aeruginosa (Davies et al., 1998; Sauer et al., 2002), Vibrio cholerae
(Watnick and Kolter, 1999; Hammer and Bassler, 2003), Escherichia coli (Danese et al., 2000; Jackson et al., 2002; Reisner et al., 2003), Serratia marcescens (Labbate et al., 2004; Rice et al., 2005) and Staphylococcus epidermidis (Vuong et al., 2003;
Kaplan et al., 2004). Apart from the intrinsic interest in the clinical relevance of such pathogens, the limited access of antibiotics to cells embedded in a biopolymer matrix
19 Chapter 1 General introduction and literature review
has been studied by many research teams and hence generated considerable insight into the structural nature of such biofilms.
In contrast, biofilms produced by environmental strains have received relatively little attention. Pseudoalteromonas tunicata is a ubiquitous, marine bacterium that is of particular interest. It has been isolated from various marine eukaryotes, suggesting that it forms biofilms in the environment (Holmström et al., 1998). Furthermore, P. tunicata produces a suite of inhibitory compounds with specific activity against other prokaryotic and eukaryotic organisms (Holmström et al., 1998). One of these compounds is the antibacterial and autolytic protein, AlpP (James et al., 1996), which has been shown to cause lysis of a subpopulation of cells during biofilm development of
P. tunicata (Chapter 2) (Mai-Prochnow et al., 2004). Reproducible events of cell death have recently been described for several other biofilm forming organisms, including P. aeruginosa (Webb et al., 2003b), V. cholerae (D. McDougald, J. S. Webb and S.
Kjelleberg, unpublished data), S. marcescens (K.W. Lam, S.A. Rice and S. Kjelleberg, unpublished) and Caulobacter crescentus (Entcheva-Dimitrov and Spormann, 2004).
This process displays striking similarities with higher organisms, where cell death is a normal part of the development. Thus it was proposed that bacterial biofilms comprise a certain degree of multicellularity and cell death may play an important role during biofilm development (Webb et al., 2003a).
This thesis addresses the hypothesis that cell death during biofilm formation is of significance to the biofilm lifestyle in P. tunicata and other organisms containing an
AlpP-homologue.
20 Chapter 1 General introduction and literature review
1.1 Biofilm development
Biofilm formation follows a cycle of events: A) attachment of single cells to the surface,
B) multiplication C) maturation and differentiation into three-dimensional structures
(microcolonies) D) formation of hollow microcolonies and E) dispersal of single cells and/or cell clusters to colonize fresh surfaces (Figure 1.1). During these stages of the biofilm life cycle, altered gene expression and multifactorial genetic control systems have been identified, leading to the hypothesis that biofilm formation follows a developmental program similar to eukaryotic differentiation (Watnick and Kolter, 1999;
O'Toole et al., 2000; Danese et al., 2001; Webb et al., 2003a; Kolter, 2005; Southey-
Pillig et al., 2005). However, others have proposed that these physiological cell changes may simply be the result of the cumulative effect of single cells responding to changes in their environment (Kjelleberg and Molin, 2002). Results and experiments addressing both proposals are discussed below (see Section 1.1.1 to 1.4).
AB C DE
Figure 1.1: Steps of biofilm formation. (A) Attachment, (B) Multiplication, (C) Maturation and differentiation of microcolonies, (D) Formation of hollow microcolonies and (E) Dispersal
1.1.1 Biofilm attachment
Bacteria sense certain environmental parameters that trigger the transition from planktonic growth to life on a surface. The environmental cues that control this
21 Chapter 1 General introduction and literature review
transition vary greatly among organisms. The conditions that influence the initial attachment are surface properties, osmolarity, pH, iron availability, oxygen tension and temperature (reviewed by Davey and O’Toole (2000)).
The first step during microbial adhesion to a surface within a liquid is the transport to the surface (van Loodsrecht et al., 1990). A microorganism in a liquid environment can reach a surface by several different mechanisms. The first is diffusive transport, where cells exhibit Brownian motion (van Loodsrecht et al., 1990). The second mechanism is gravitational settling, which is more relevant for larger cells. Thirdly, cells may be transported by convective liquid flow, which can occur much faster than diffusive transport (Characklis, 1981).
After a planktonic bacterium has contacted the substratum it becomes transiently fixed.
This initial attachment is reversible, since some cells are observed to continue to exhibit
Brownian motion or be removed from the surface by mild shear forces (van Loodsrecht et al., 1990). Depending on the organism, surface structures such as flagella, pili and outer membrane proteins are often required for this initiation of biofilm formation.
Mutants defective in initial attachment have been used to identify several factors required for surface attachment. P. aeruginosa strains defective in flagellum-mediated motility appeared to be blocked in their initial stages of interaction with the surface
(O'Toole and Kolter, 1998). E. coli has also been found to require flagella and pili to initiate the early attachment processes (Genevaux et al., 1996). Flagellum mediated motility is required for movement parallel to the surface in addition to bringing the bacterium into proximity to the surface. Similar to E. coli and P. aeruginosa it has also been shown that flagella are important in the initial attachment of V. cholerae biofilm attachment (Davey and O'Toole, 2000). However, other factors besides motility seem to play a role in initial attachment as motility-independent, attachment-deficient mutants 22 Chapter 1 General introduction and literature review
have been identified in P. aeruginosa (Ramsey and Whiteley, 2004). The genes involved in sensing and responding to external stimuli were disrupted in these mutants, implying a significant impact of external factors on the biofilm developmental pathway
(Ramsey and Whiteley, 2004). Lipopolysaccharide (LPS), an important component of the bacterial outer membrane, also plays a role in initial surface attachment for P. aeruginosa (Davey and O'Toole, 2000) as well as for E. coli (Genevaux et al., 1999). A study by Ryu et al. showed that exopolysaccharide (EPS) overproduction can inhibit attachment of E. coli O157:H7 to stainless steel coupons (Ryu et al., 2004).
The next stage of the biofilm development is irreversible attachment to the surface.
After this stage cells cannot be removed from the surface by simple washing procedures, indicating irreversible adhesion (Oliveira, 1992). Cells utilize extra-cellular substances, fimbriae or other surface structures, allowing the cell and the substratum to interact directly by short range hydrophobic and specific interactions (Busscher and
Weerkamp, 1987). Mutants are available which are capable of initial (reversible) attachment but are deficient in the transition to irreversible attachment. Different factors have been identified to play a role in this transition, including a large adhesion protein
(Lap) secreted by an ABC transporter in Pseudomonas fluorescens (Hinsa et al., 2003) and a protein of unknown function SadB (surface attachment defective) in P. aeruginosa. Interestingly, it was demonstrated for E. coli K12 that while temporary attachment normally precedes permanent attachment, both reversible and irreversible attached cells can coexist in a population. For E. coli, temporarily attached cells were mainly surface associated with the cell pole and permanent attachment of cells occurred via the lateral cell surface (Agladze et al., 2003; Agladze et al., 2005). It is clear that, factors important for attachment can differ greatly from species to species and also depend on the substratum and the environment, where the attachment takes place.
23 Chapter 1 General introduction and literature review
1.1.2 Biofilm maturation and differentiation
After irreversible attachment has taken place cells adjust to an immobile lifestyle on the surface. Cells are embedded in an extracellular matrix, which is produced with a species specific chemical composition (Sutherland, 1985). In P. aeruginosa, it has been suggested that the EPS alginate is important for biofilm maturation, because strains from cystic fibrosis patients over-express alginate (Garrett et al., 1999). Further, a down-regulation of flagellum synthesis linked with the up-regulation of alginate occurs in P. aeruginosa biofilms after attachment (Garrett et al., 1999). However, in a recent study no significant difference in biofilm architecture could be identified in alginate overproducing and alginate deficient mutants compared to the wild-type, indicating that alginate is not critical for biofilm formation in P. aeruginosa (Stapper et al., 2004). In contrast, EPS production regulated by a gene called mbaA was shown to be critical for the formation of the characteristic biofilm architecture in V. cholerae E1 Tor (Bomchil et al., 2003). Another study also demonstrated that EPS mutants of V. cholerae E1 Tor and V. cholerae 0139 form thin monolayer biofilms, further suggesting that EPS is critical for the formation of a voluminous, three-dimensional biofilm (Watnick and
Kolter, 1999; Watnick et al., 2001).
An important step in biofilm development is the formation of microcolonies, which occurs in most bacteria (Figure 1.1). First, cell multiplication results in the development of immature microcolonies smaller than 10 µm and these cell clusters become progressively layered (Sauer et al., 2002). As defined by Davies et al. (1998) the fourth stage of biofilm development is reached when cell clusters attain their maximum average thickness. During this stage of maturation, cells within microcolonies are non motile, the microcolonies reach their maximum dimensions and the majority of the cells are segregated within cell clusters (Sauer et al., 2002). Some microcolonies are simple 24 Chapter 1 General introduction and literature review
conical structures, while others are mushroom shaped and they are usually interspersed with less dense regions of the matrix that include highly permeable water-channels
(Lawrence et al., 1991). These water-channels can be seen as a primitive “circulatory system” which delivers nutrients from the bulk fluid to the microcolonial niche and removes metabolic products by the same process (Costerton et al., 1994). Bacteria are observed to swim freely within the void spaces, indicating the absence of dense polymer or other gel-like material in the void space (Sauer et al., 2002).
1.1.2.1 Cell death during biofilm differentiation
Recently, it has been demonstrated that lysis of a subpopulations of cells occurs during biofilm differentiation in several model biofilm organisms. This phenomenon has been observed to occur in mono-species biofilms where cell death occurs localized within the centre of microcolonies (Webb et al., 2003b; Entcheva-Dimitrov and Spormann, 2004;
Mai-Prochnow et al., 2004). Furthermore, biofilm cell death also occurs in mixed species communities, including dental biofilms (Auschill et al., 2001; Hope et al., 2002) and river biofilm communities (Lawrence et al., 2004). The mechanism leading to biofilm cell death has been investigated in P. aeruginosa, where biofilm killing has been linked to the activity of a prophage (Webb et al., 2003b) and is controlled by specific regulatory determinants, such as RpoN (Webb et al., 2003b) and quorum sensing (J. S. Webb and S. Kjelleberg, unpublished data). Furthermore, it was suggested that cell death in P. aeruginosa biofilms is involved in the dispersal process of the organism. However, for most organisms the mechanism/s or role of cell death during biofilm formation has not been investigated and thus is unclear. One objective of the current study was to investigate the mechanism and a possible role for biofilm cell death in P. tunicata biofilms.
25 Chapter 1 General introduction and literature review
1.1.3 Biofilm dispersal
Another important aspect of the development and differentiation of biofilms on surfaces is the reversion of biofilm cells to the planktonic phase. This last stage in the biofilm life cycle is of particular importance because of the associated potential transmission of pathogens (Zottola and Sasahara, 1994; Walker et al., 1995; Piriou et al., 1997). For example, in medical devices the growth and subsequent dispersal of bacteria from biofilms have the potential to increase the risk of pathogen exposure to patients (Inglis,
1993; Putnins et al., 2001). Thus, recent years have seen a dramatic increase in studies that focus specifically on the process of biofilm dispersal.
Traditionally, biofilm dispersal was viewed as a passive process, such as shear-induced sloughing (Stoodley et al., 2002) and erosion of cells from the biofilm (Stoodley et al.,
2001). It was found that dispersal can occur via single cells and small cell cluster or larger aggregates (Stoodley et al., 2001; Fux et al., 2004). Passive dispersal largely depends on the hydrodynamic conditions surrounding the cells (e.g. change of flow rate) as well as the mechanical properties of the biofilm (Stoodley et al., 2002). For example,
Stoodley and co-workers demonstrated that biofilms grown under low shear or static conditions dislodge clumps of bacteria more easily when the flow rate changes
(Stoodley et al., 1999; Stoodley et al., 2002).
More recently it has become clear that biofilm dispersal can also be an active process used by bacteria to colonize new surfaces when conditions have become unfavourable.
Specific cell-surface molecules produced by bacteria as well as regulatory elements and environmental nutrient conditions have all been identified to modulate dispersal in different bacterial species. In E. coli K-12 the RNA binding global regulatory protein
CsrA (carbon storage regulator) serves as both a repressor of biofilm formation and an
26 Chapter 1 General introduction and literature review
activator of biofilm dispersal under a variety of culture conditions. It was demonstrated that the effect of CsrA is largely mediated through the regulation of intracellular glycogen biosynthesis and catabolism (Jackson et al., 2002). In P. aeruginosa it was also shown that a change in nutrient availability is a trigger for biofilm dispersal (Hunt et al., 2004; Sauer et al., 2004). It was also found that both the removal of nutrients
(Hunt et al., 2004) as well as a sudden increase in specific carbon sources can induce dispersal (Sauer et al., 2004). A study by Mitrakul et al. (2004) suggested that the trace nutrient copper influences biofilm detachment of the oral pathogen Streptococcus gordonii. Other factors that influence biofilm dispersal include cell signalling systems
(Dow et al., 2003; Crossman and Dow, 2004), quorum sensing (Rice et al., 2005), the production of endogenous enzymes (Boyd and Chakrabarty, 1994; Lee et al., 1996;
Kaplan et al., 2003a; Kaplan et al., 2003b) and surfactants (Davey et al., 2003; Boles et al., 2005).
The formation of cavities within microcolonies (hollow microcolonies) was often observed to precede the dispersal process in several organisms (Sauer et al., 2002;
Davey et al., 2003; Kaplan et al., 2003b; Webb et al., 2003b; Boles et al., 2005).
Kaplan et al. (2003b) showed that in Actinobacillus actinomycetemcomitans the formation of non-aggregated cells within the centre of microcolonies through the activity of a soluble ȕ-N-acetylglucosaminidase (encoded by dspB) is necessary for biofilm dispersal of the organism. Similarly, in P. aeruginosa biofilms it was hypothesized that the phage induced cell death in the microcolony centre led to hollow colony formation and subsequent dispersal (Webb et al., 2003b). These findings suggest that biofilm dispersal may represent an adaptive response that facilitates the colonization of fresh surfaces under conditions of nutrient limitation within the environment.
27 Chapter 1 General introduction and literature review
1.1.4 The affect of cyclic di-GMP on biofilm development
Although first discovered two decades ago as an activator of cellulose synthase in
Gluconacetobacter xylinus (Ross et al., 1987), bis-(3’-5’)-cyclic dimeric guanosine monophosphate (c-di-GMP) has only recently emerged as a novel global secondary messenger molecule. C-di-GMP is synthesized by diguanylate cyclase (DGC) and hydrolysed by phosphodiesterase (PDE) activities, respectively (Tal et al., 1998; Chang et al., 2001; Simm et al., 2004). Proteins containing a GGDEF domain (responsible for
DGC activity) and/or an EAL domain (responsible for PDE activity) have been identified in almost every bacterial genome sequenced to date (Romling et al., 2005).
C-di-GMP signalling regulates complex biological processes, including virulence
(Tischler and Camilli, 2005), photosynthesis (Thomas et al., 2004) and biofilm formation (Simm et al., 2004; Tischler and Camilli, 2004). Simm et al. (2004) showed that in Salmonella enterica serovar Typhimurium, P. aeruginosa and E. coli, GGDEF and EAL domains mediate similar phenotypic changes related to the transition between sessility and motility. For example, GGDEF and EAL domain proteins had an opposite effect on motility function. Expression of the GGDEF domain containing protein
(AdrA) increases c-di-GMP levels in P. aeruginosa, S. typhimurium and E. coli leading to an inhibition of swimming, swarming and twitching. On the other hand the EAL domain containing protein YhjH, stimulated swarming. Thus it was suggested that
AdrA and YhjH inversely regulate sessility versus motility (Simm et al., 2004).
Several studies have shown that high c-di-GMP levels promote thick biofilms. In
Yersinia pestis a GGDEF domain containing protein, HmsT and an EAL domain containing protein HmsP, have been identified and studied for their role in c-di-GMP mediated biofilm formation. It was demonstrated that both a mutation in HmsP (when c-
28 Chapter 1 General introduction and literature review
di-GMP hydrolysis is diminished) and over-expression of HmsT (when c-di-GMP synthesis is increased) lead to an extremely thick biofilm in Y. pestis because of increased c-di-GMP levels (Kirillina et al., 2004). Similarly, mutations in wmsF encoding an EAL domain containing protein, also causes increased biofilm formation by P. aeruginosa (Hickman et al., 2005). In V. cholerae, VieA (containing an EAL domain) represses EPS synthesis genes involved in biofilm formation by reducing c-di-
GMP levels (Tischler and Camilli, 2004). As high c-di GMP levels promote thick biofilm formation, it was also demonstrated that induction of a gene (yhjH) encoding phosphodiesterase activity can induce biofilm detachment by decreasing c-di-GMP levels in Shewanella oneidensis (Thormann et al., 2006).
In summary, the regulation of intracellular c-di-GMP levels play a major role in the timing and amplitude of complex factors involved in biofilm formation in many (if not all) bacteria. Moreover, it was recently suggested that besides its role as an intracellular signalling molecule in bacteria, c-di-GMP also acts as an intercellular signalling molecule between prokaryotes and furthermore has effects in eukaryotes that could provide a perspective in cancer treatment (Romling and Amikam, 2006).
1.2 Genetic diversification of biofilm cells
A high degree of diversification is hypothesised to be beneficial to the population as the chances of thriving under different environmental conditions are enhanced. The occurrence of diverse phenotypes affects many traits in bacteria, including host- pathogen interactions (Craig and Scherf, 2003), production of exo-enzymes (Chabeaud et al., 2001) and secondary metabolites (van den Broek et al., 2003) as well as colonization behaviour such as biofilm formation (Deziel et al., 2001; Boles et al.,
2004). The process of generating diversity has a broad impact on the ecology of bacteria
29 Chapter 1 General introduction and literature review
and is particularly important for the adaptation to rapidly changing environmental conditions.
Biofilm variants can be detected when dispersal cells from mature biofilms are plated onto solid growth media and colonies display a different morphology (Figure 1.2) or other characteristic, such as increased antibiotic resistance. Most of these variants occur specifically during the biofilm mode of growth and arise at the later stages of biofilm development. Phenotypic variants have been detected in many biofilm forming organisms, including P. aeruginosa (Drenkard and Ausubel, 2002; Boles et al., 2004;
Haussler, 2004; Webb et al., 2004), Staphylococcus aureus (Proctor et al., 1995;
Sadowska et al., 2002), S. epidermidis (Conlon et al., 2004; Handke et al., 2004), V. cholerae (Ali et al., 2002; Matz et al., 2005) and Listeria monocytogenes (Monk et al.,
2004) suggesting that this is a common phenomenon among biofilm forming organisms.
Figure 1.2: Variant colonies on agar produced by a 5-day-old Pseudomonas aeruginosa biofilm. Modified from (Boles et al., 2004).
Random mutation and selection, gene regulation and environmental factors can all play a role in the emergence of variants from biofilms (Rainey and Travisano, 1998;
Drenkard and Ausubel, 2002; Haussler, 2004). In S. epidermidis, phenotypic variation is 30 Chapter 1 General introduction and literature review
regulated by mutation and transcriptional regulation of the ica operon encoding polysaccharide intracellular adhesion synthetases (Conlon et al., 2004; Handke et al.,
2004). Boles et al. (2004) showed that variation among biofilm dispersal cells of P. aeruginosa is recA mediated. However, environmental cues (such as those in the lung of patients with P. aeruginosa infections) were also shown to play a role in controlling the degree of antibiotic resistance in variants (Drenkard and Ausubel, 2002; O'Toole, 2002), suggesting that both intrinsic, regulatory elements and environmental factors are involved in the formation of population diversity among biofilm cells.
In the model organism P. aeruginosa, the emergence of small colony variants (SCVs) from biofilms is a well studied example (Drenkard and Ausubel, 2002; Haussler et al.,
2003; Webb et al., 2004). The variants have a rough colony appearance, show enhanced biofilm formation and type IV fimbriae production, enhanced antibiotic resistance and are defective or impaired in chemotaxis and different types of motility, including swimming, swarming and twitching (Deziel et al., 2001; Drenkard and Ausubel, 2002;
Kirisits et al., 2005). These properties might be important under certain environmental conditions favourable for biofilm growth. It has been suggested by Deziel et al. (2001) that SCVs emerge in anticipation to start biofilm formation when conditions are favourable. Moreover, Haussler et al. (2003) demonstrated that different SCVs derived from P. aeruginosa biofilms have an increased fitness and thus a selective advantage in late stationary phase compared to the wild-type strain, when both strains were grown in a co-culture. It was therefore speculated that the SCVs have an increased tolerance to nutrient and oxygen limitations (Haussler et al., 2003). In a study by Drenkard and
Ausubul (2002) a regulatory protein (pvrR) was identified which may regulate factors that control the emergence of SCVs in P. aeruginosa biofilms. It was also demonstrated that the expression of chaperone usher pathway (cup) genes, encoding putative fimbrial
31 Chapter 1 General introduction and literature review
adhesions are involved in the switch to SCVs in P. aeruginosa (Haussler, 2004).
Furthermore, Webb et al. (2004) showed that the emergence of SCVs in P. aeruginosa biofilm dispersal cells correlates with the appearance of plaque-forming filamentous phage Pf4. Thus it was concluded that phage expression can also mediate phenotypic variation in P. aeruginosa. In summary, phenotypic diversity is common during biofilm formation and can be mediated through a range of genetic and environmental factors.
1.3 Cell to cell communication within biofilms – quorum sensing
In bacteria, the regulation of many important changes in gene expression is mediated by systems of signalling between cells known as quorum sensing (Fuqua et al., 1994; Swift et al., 2001). In a population, cells can sense their density and number through the presence of small signalling molecules which diffuse freely across cell membranes and between cells. When a certain population density is reached and enough signalling molecules have been produced an auto-induced positive feedback mechanism is usually used to induce the appropriate phenotypes required for responding to a particular environmental condition or for proceeding with the differentiation process of the population (Kjelleberg and Molin, 2002). A number of QS systems, mainly the acylated homoserine lactone (AHL) systems of several Gram negative bacteria (Swift et al.,
1999), the auto-inducer 2 (AI2) system of a range of both Gram negative and Gram positive bacteria (Bassler, 1999), and peptide mediated quorum sensing signalling systems in several Gram positive species (Dunny, 1999), have been characterized.
Within a biofilm a high cell density is reached and thus quorum sensing systems are involved in several processes during biofilm development, including attachment (Huber et al., 2001; Tomlin et al., 2005), differentiation (Davies et al., 1998) and dispersal
(Allison et al., 1998; Rice et al., 2005). Furthermore, it was shown that QS signalling
32 Chapter 1 General introduction and literature review
systems play a role in both mixed and monoculture biofilms (McLean et al., 1997;
Charlton et al., 2000). QS systems are found to be involved in biofilm formation of an ever increasing number of organisms, including P. aeruginosa (Davies et al., 1998),
Aeromonas hydrophila (Lynch et al., 2002), Burkholderia cenocepacia (Huber et al.,
2001), S. marcescens (Labbate et al., 2004), Staphylococcus sp. (Vuong et al., 2003;
Kong et al., 2006; Xu et al., 2006), Streptococcus mutans (Li et al., 2002), V. cholerae
(Hammer and Bassler, 2003; Vance et al., 2003), Enterococcus faecalis (Carniol and
Gilmore, 2004), Klebsiella pneumoniae (Balestrino et al., 2005), Pseudomonas putida
(Arevalo-Ferro et al., 2005) and E. coli (Gonzalez Barrios et al., 2006). The involvement of QS systems in many biofilm forming organisms shows that this is a widespread mechanism used by bacteria to control different aspects of biofilm development. Understanding the role of these systems in biofilm formation can be an important tool in controlling biofilms.
1.3.1 The involvement of quorum sensing systems in the formation of a
differentiated biofilm architecture
AHL-mediated QS systems appear in many cases to control genes involved in the bacterial colonization of surfaces, such as biofilm formation. Through the use of mutants defective in AHL-systems it was shown that QS molecules are important for the differentiation of biofilms of B. cenocepacia (Huber et al., 2001), S. marcescens
(Labbate et al., 2004) and P. aeruginosa (Davies et al., 1998). The two cell-to-cell signalling systems identified in P. aeruginosa are the LasR-LasI and RhlR-RhlI systems
(Passador et al., 1993; Ochsner et al., 1994). It was observed that a specific signalling lasI mutant of P. aeruginosa formed a flat, undifferentiated, thin biofilm layer compared to the mushroom-like microcolonies separated by water-channels of the wild- type. Similarly, in S. marcescens a mutant deficient in the production of the 33 Chapter 1 General introduction and literature review
extracellular signal molecule formed a thin biofilm lacking the differentiation into cell aggregates and chains typical for the wild-type biofilm architecture (Labbate et al.,
2004). By adding back AHL to mutant biofilms of both P. aeruginosa and S. marcescens the differentiated wild-type phenotype was restored, demonstrating that
AHL molecules are necessary for the characteristic biofilm structure in these organisms.
1.4 Are biofilms multicellular communities?
An intensely discussed area in biofilm research is whether or not biofilm formation constitutes a developmental process in which cells commit to a change in lifestyle through a programmed cascade of changes in gene expression and thus shares features with development in higher organisms.
It is widely accepted that biofilm cells are different from their planktonic counterparts and that biofilm formation occurs in multiple steps (Costerton et al., 1995; Watnick and
Kolter, 1999). Some researchers have suggested that the formation of the characteristic biofilm architecture can be explained entirely by mathematical modelling, and that such structures are the result of individual cells responding to changes in their physicochemical environment where a biofilm is formed (Wimpenny et al., 2000; Kreft et al., 2001; Picioreanu et al., 2001; van Loosdrecht et al., 2002). More recently, others have suggested that biofilm formation may be viewed as a programmed developmental process with similarities to other prokaryotic developmental processes, such as sporulation of Gram positive bacteria (Dunny and Leonard, 1997), fruiting body formation in Myxococcus xanthus (Plamann et al., 1995; Shimkets, 1999) and stalked- cell formation in C. crescentus (Fukuda et al., 1977; Hecht and Newton, 1995) and thus also show properties of multicellularity. However, it is not entirely clear how many properties of biofilm bacteria are dictated by a developmental program of the organism
34 Chapter 1 General introduction and literature review
and how many are dictated by environmental conditions in which the biofilm is growing
(Kolter, 2005).
To regard biofilm formation as a programmed developmental process, biofilm cells must be able to sense their surroundings and adjust their behaviour to maximise survival success, such as: a) “decide” whether or not to form a biofilm, b) disintegrate an existing biofilm under unfavourable conditions or c) change metabolism according to nutrient availability within the small scale of a biofilm community. Several studies have demonstrated evidence for such behaviour in biofilm bacteria. It was shown that nutrient removal induces biofilm dispersal in E. coli and P. aeruginosa (Danese et al.,
2001; Sauer et al., 2004). Furthermore, different nutrient conditions lead to the formation of distinct biofilm structures and these structures can be interconverted
(Danese et al., 2001; Labbate et al., 2004), showing that biofilm cells can actively respond to environmental conditions. Bacteria in biofilms can also change gene expression to suit the biofilm lifestyle. For example, during the initial stages of biofilm formation P. aeruginosa increases the transcription of algC, the gene for the EPS alginate and at the same time decrease flagella synthesis (Garrett et al., 1999). A similar finding was made in E. coli biofilms where the production of the EPS colonic acid is increased while flagella synthesis decreased (Prigent-Combaret et al., 1999). Moreover, the fact that intercellular communication via signalling molecules (“quorum sensing”, see Section 1.3) has been demonstrated to play an important role in biofilm formation also suggests a form of multicellularity.
Another characteristic of multicellularity is the occurrence of altruistic behaviour, such as programmed cell death (PCD). It has been proposed that altruistic cell death events occur during developmental processes of prokaryotes, such as sporulation, fruiting body formation and mycelium differentiation (see below, Section 1.4.1). Cell death is often 35 Chapter 1 General introduction and literature review
observed during biofilm formation (see Section 1.1.2.1) and it could be speculated that this occurs as part of a developmental program and may show similar characteristics to eukaryotic PCD, such as altruism (Mai-Prochnow et al., 2006). Experiments to address this hypothesis are discussed in Chapter 4.
1.4.1 Cell death during differentiation processes in prokaryotes
In certain environmental conditions some bacteria form specialised structures, such as spores or fruiting bodies. During these processes cells differentiate and perform an assigned function, similar to cells in multicellular organisms. Cell death of subpopulations has been demonstrated to be a normal component during these differentiating processes, including sporulation of B. subtilis (Gonzalez-Pastor et al.,
2003), fruiting body formation of M. xanthus (Wireman and Dworkin, 1977; Rosenbluh and Rosenberg, 1990) and mycelium differentiation of S. antibioticus (Mendez et al.,
1985; Miguelez et al., 1999; Fernandez and Sanchez, 2002). Several implications for cell death events in these organisms have been investigated and are discussed below
(see Sections 1.4.1.1 to 1.4.1.3).
1.4.1.1 Cell death during fruiting body formation of Myxococcus xanthus
Myxococcus xanthus is a motile soil bacterium with a complex social behaviour. Under nutrient starvation conditions a developmental program is initiated where cells form aggregates resulting in the formation of fruiting bodies (Dworkin, 1972). Fruiting bodies contain a very high cell density to ensure a high local concentration of degrading enzymes for efficient hydrolysis of organic material and thus represent a form of cooperative feeding behaviour. Fruiting bodies differentiate into stalk and head, and cells accumulating in the head undergo morphogenesis to a resting cell type called
36 Chapter 1 General introduction and literature review
myxospores. Under favourable conditions myxospores can germinate and yield vegetative rods (Shimkets, 1990).
Early confocal laser scanning microscopy (CLSM) studies revealed that cell density in the centre of mature fruiting bodies is much lower than at the outer ring (Sager and
Kaiser, 1993). Recently, it was demonstrated that a characteristic spatial distribution of viable and dead cells occurs within mature fruiting bodies, with the majority of live cells localised in the outer layer and dead cells underneath (Lux et al., 2004). This phenomenon has also been observed in microcolonies of bacterial biofilms (Auschill et al., 2001; Webb et al., 2003b; Entcheva-Dimitrov and Spormann, 2004; Mai-Prochnow et al., 2004) (see Section 1.1.2.1), suggesting analogies between fruiting body and biofilm formation (Figure 1.3).
37 Chapter 1 General introduction and literature review
AB
CD
Figure 1.3: Structural similarities between microcolonies from bacterial biofilms and fruiting bodies formed by social Myxococcus spp. (A) Sharing of labour in P. aeruginosa microcolonies with caps formed by motile wild-type cells (yellow) and stalks formed by non-motile type-IV pilus mutant (pilA) cells (cyan). The image was taken from a 4-day biofilm inoculated with a 1:1 mixture of wild type and pilA cells. Bar = 20 µm (Klausen et al., 2003). (B) Mature fruiting bodies of M. xanthus viewed from the top and from the side in the insert. Bar = 0.2 mm (Jelsbak and Sogaard- Andersen, 2000). (C) 6-day old microcolony from P. tunicata biofilm stained with the LIVE/DEAD BacLight Bacterial Viability Kit. (D) Live versus dead cell distribution within fruiting body structures formed by M. xanthus at 48 h of development on MOPS 1.5% hard agar surfaces. Cells are stained with the LIVE/DEAD BacLight Bacterial Viability Kit (Lux et al., 2004).
38 Chapter 1 General introduction and literature review
A study by Wireman and Dworkin (1977) showed that the developmental program leading to fruiting body formation occurs in distinct steps: 1) growth; 2) aggregation; 3) formation of raised, darkened mounds of cells; 4) autolysis and 5) myxospore induction.
Autolysis was shown to be an essential requirement to complete the differentiation process. Up to 90 % of cells lyse by an autolytic mechanism independent of cell density or concentration of lysis products. In contrast, myxospore formation is dependent upon the concentration of lysis products, suggesting that autolysis is essential for spore induction (Wireman and Dworkin, 1977). Because fruiting body formation occurs normally under nutrient deprivation conditions, cells must have sufficient endogenous resources or must be provided with an exogenous nutrient source to complete the energy costly differentiation process. Thus it was suggested that lysis material may serve as an energy source for surviving cells undergoing myxospore formation (Wireman and
Dworkin, 1977).
1.4.1.2 Cell death during sporulation of Bacillus subtilis
The process of sporulation is particularly well demonstrated in the ubiquitous soil bacterium Bacillus subtilis. Under nutrient starvation and high cell density conditions B. subtilis cells start a morphological manifestation of the sporulation process: the asymmetrical position of the cell division septum or sporulation septum. This process leads to the formation of a large compartment, which develops into the mother cell and a small compartment which becomes the endospore (Piggot and Coote, 1976;
Sonenshein, 2000).
Sporulation in B. subtilis is a complex pathway, where the transcription factor Spo0A integrates signals from nutritional status, cell density and cell cycle to activate a cascade of interdependent sigma factors in both mother cell and spore (Burbulys et al., 1991). In
39 Chapter 1 General introduction and literature review
the last step of spore formation the mother cell is actively lysed to release the endospore. Three major autolysins responsible for mother cell lysis at different developmental stages have been identified, including CwlB, CwlC and CwlH. It was shown that only double cwlB cwlC and cwlC cwlH mutants are defective in lysis and single mutants still showed lysis (Smith and Foster, 1995; Nugroho et al., 1999).
Further, it was proposed that an additional unidentified factor is required to activate autolysis, because this process has to be tightly regulated and to specifically target the mother cell after the spore is formed (Lewis, 2000).
The obvious function of mother cell autolysis is to eliminate a barrier that could interfere with the outgrowth of a germinating spore. However, additional function(s) are also discussed. Similar to M. xanthus, it is proposed that nutrients released by the lysing mother cell are used by surviving sibling cells, because sporulation cannot occur under total nutrient deprivation. However, it is also possible that autolysis occurs simply because the mother cell cannot cope with the sudden decrease in energy and cell lysis is a consequence of a breakdown in the control of the cell wall machinery (Lewis, 2000).
Sporulation is an irreversible, energy costly process and only advantageous under growth limiting conditions. Thus the commitment to enter the sporulation cycle has to be tightly regulated. A study by González-Pastor et al. (2003) showed that B. subtilis cells that have entered the sporulation pathway produce and export a killing factor and a signalling protein that are together responsible for inhibiting sporulation in sister cells and causing them to lyse. Surviving cells, which have not yet committed to the sporulation process, feed on the released nutrients and delay sporulation. The operon skf has been identified to be responsible for the production of a killing factor. Skf is under the control of Spo0A and directs the production of a peptide antibiotic and a pump to export it. It was proposed that B. subtilis wild-type produces a mixed population where 40 Chapter 1 General introduction and literature review
Spo0A is active in some cells (and skf is transcribed) but not in others. Thus only the population with inactive Spo0A would be killed because of the inability to remove the peptide antibiotic through the action of the pump. Another operon, spd, was found to be responsible for the production of a signalling peptide, which turns on the transcriptional regulator YvbA. This regulator causes increased lipid oxidation and ATP synthesis, resulting in higher energy production and thus a delay in sporulation (which is triggered by nutrient depletion). It was speculated that this complex mechanism is in place to delay the energy costly sporulation process in case fresh nutrients become available
(Gonzalez-Pastor et al., 2003).
1.4.1.3 Cell death during mycelium differentiation of Streptomyces antibioticus
Streptomyces species are abundant soil bacteria that show an unusual and complex multicellular differentiation consisting of two distinct phases: vegetative mycelial growth (substrate mycelium) under optimal conditions and an aerial, reproductive mycelial growth (aerial mycelium) under nutrient limiting conditions. Exospores are produced from the aerial hyphae, which are relatively resistant to desiccation but sensitive to high temperatures (Chater, 1989; Champness and Chater, 1994).
During the differentiation process a large number of hyphae degenerate and die in two separate rounds of cell death, including hyphae of the substrate mycelium and aerial hyphae which do not differentiate into exospores (Wildermuth, 1970; Mendez et al.,
1985; Miguelez et al., 1999). Similar to cell death events within microcolonies the lytic processes in S. antibioticus start in the centre on the mycelial masses and extend progressively to the periphery, ending with the formation of the reproductive hyphae
(Figure 1.4) (Fernandez and Sanchez, 2001). Evidence was provided that cell death during differentiation in S. antibioticus is not a random process of autolysis
41 Chapter 1 General introduction and literature review
(degradation of the cell wall by uncontrolled, lytic action of murein hydrolases), but a highly organised process of PCD (Miguelez et al., 1999). It was demonstrated that hyphae undergo progressive disorganisation of internal cell constituents with extensive genome digestion followed by a loss of plasma membrane integrity. It was suggested that dead hyphae cells contribute to the mechanical support of the differentiated structure, because their cell walls stay intact after cell death has occurred (Miguelez et al., 1999). Moreover, it was proposed that this structure serves as a conducting system for translocating nutrients, as the aerial hyphae are completely dependent upon water and nutrients from the substrate mycelium (Mendez et al., 1985; Miguelez et al., 1999).
Some of these nutrients were shown to originate from the dead cells in the substrate mycelium (Mendez et al., 1985). As further evidence for this process, several different nucleases have been identified and proposed to play a role in the degeneration and recycling of DNA from dead cells, which then contributes to the nutrient support of surviving cells (Fernandez and Sanchez, 2002). These results suggest that cell death is an important part during differentiation of S. antibioticus and somehow included in the developmental process.
42 Chapter 1 General introduction and literature review
Figure 1.4: Cell death during differentiation of S. antibioticus. Laser confocal microscope observations of S. antibioticus mycelial pellets stained with the viability assay LIVE/DEAD BacLight Bacterial Viability Kit after 6 h of cultivation. Red Propidium Iodide-stained cells have a compromised cell membrane and are dead. Bar = 10 µm. Modified from (Fernandez and Sanchez, 2001).
In summary, prokaryotic cell death events show some characteristics of PCD. Cell death is clearly involved in differentiation processes and evidence suggests that it is an essential part during these developmental programs. It remains to be elucidated whether cell death in bacterial biofilms is also part of a developmental program and displays similar characteristics to PCD.
1.5 The genus Pseudoalteromonas
The Pseudoalteromonas genus resulted from the dividing of the genus Alteromonas into two genera after a comparison of the phylogenetic relationships among 17 isolates of the genera Alteromonas, Shewanella and Moritella by Gauthier et al. (1995) and today about 40 species have been described. Bacteria of the Pseudoalteromonas genus are
43 Chapter 1 General introduction and literature review
Gram-negative and belong to the Ȗ-Proteobacteria. Cells are heterotrophic and most are motile through the use of polar, bipolar or lateral flagella (Gauthier et al., 1995; Ivanova et al., 2002c).
One of the first and most extensively studied Pseudoalteromonas species is P. haloplanktis, which is a fast growing Antarctic bacterium with interesting cold adaptation properties. The recently released complete genome sequence revealed its complex genome structure, comprising two chromosomes as well as some unique cold adaptation features (Medigue et al., 2005). P. haloplanktis copes with the increased solubility of oxygen at low temperature by multiplying dioxygen scavenging genes while deleting whole pathways producing reactive oxygen species, thus protecting the organism from these species (Medigue et al., 2005).
Other well studied Pseudoalteromonas species include the halophilic P. carrageenovora
(Nandakumar et al., 2003; Silipo et al., 2005), the red pigmented antibiotic producing P. rubra (Ivanova et al., 2004) and the agar decomposing P. atlantica (Yaphe and Baxter,
1955) and P. citrea (Ivanova et al., 2002a; Nedashkovskaia et al., 2002; Alekseeva et al., 2004). Research in our laboratory has focused on the dark green pigmented P. tunicata (Holmström et al., 1998), which will be discussed in detail in Section 1.5.3.
1.5.1 Pseudoalteromonas species are associated with eukaryotic hosts
Bacteria belonging to the Pseudoalteromonas genus are commonly isolated from marine environments. A majority of Pseudoalteromonas species seem to be associated with eukaryotic hosts. Species have been isolated from various animals, such as mussels
(Ivanowa et al., 1996, 1998), pufferfish (Simudu et al., 1990), tunicates (Holmstöm et al., 1998) and sponges (Ivanowa et al., 1998, Lau 2005), as well as from a range of marine plants (Akagawa-Matsushita et al., 1992; Yoshikawa et al., 1997). 44 Chapter 1 General introduction and literature review
Some of the interactions between Pseudoalteromonas sp. and eukaryotes were shown to be pathogenic. For example, the pigmented P. bacteriolyticum was found to be the causative agent of the red spot disease of the red alga Laminaria japonica (Sawabe et al., 1998) and P. (previously Alteromonas) tetradonis produces a toxin which can cause poisoning of its pufferfish host (Simidu et al., 1990). Further, the agarolytic activity of
P. gracilis was demonstrated to be responsible for bleaching of the thalli from the red alga Gracilaria gracilis (Schroeder et al., 2003). However, relationships between
Pseudoalteromonas species and their hosts have also been described as symbiotic. For example, P. tunicata is hypothesized to be part of a biofouling defence mechanism for its host algae due to the production of a range of antifouling compounds (Holmström et al., 1998).
1.5.2 Production of secondary metabolites in Pseudoalteromonas species
The Pseudoalteromonas group has been of great interest to researchers, especially because it is known to produce a wide variety of biologically active compounds, including cytotoxins, hydrolytic enzymes and bio-controlling, antifouling compounds active against bacteria, invertebrate larvae, algal spores, fungi and diatoms (Holmström et al., 1998; Holmström and Kjelleberg, 1999; Isnansetyo and Kamei, 2003;
Kalinovskaya et al., 2004). It has been suggested that the diverse production of biologically active compounds results from the exploitation of a wide range of metabolic pathways when co-existing with eukaryotes (Jensen and Fenical, 1994).
The production of hydrolytic enzymes is often reported among Pseudoalteromonas species. They include proteinases, lipases, prominent laminarinases, agarases, amylases, alginases, ȕ-galactosidases, chitinases, alginases, fucoidan hydrolases, pustulanases, ȕ- galactosidases, ȕ-glucosidases and ȕ-xylosidases (Ivanova et al., 2003). These enzymes
45 Chapter 1 General introduction and literature review
break down material, such as host plant tissue (e.g. algae thallus) to low molecular weight compounds which can then be used as a nutrient source by the bacterium. The production of agarolytic enzymes which break down agar by species associated with red algae has been commonly observed. For example, P. gracilis and P. atlantica were found to be the primary pathogen of the red algae Gracilia gracilis due to their production of ȕ-agarase (Schroeder et al., 2003). The thallus of the brown alga Fucus evanescens is degraded by enzymes such as alginases and glycosidases produced by P. issachenkonii (Ivanova et al., 2002b). The cold-adapted P. haloplanktis was found to produce cellulase (Violot et al., 2003) and ȕ-galactosidase with high catalytic efficiency on natural and synthetic substrates (Hoyoux et al., 2001). Because of the high specificity and usability of the hydrolytic enzymes it has been suggested that their production is strain-specific and reflects the specific natural habitats of the strain
(Ivanova et al., 2003).
The competition for space and nutrients in the marine environment makes it advantageous for bacteria to produce inhibitory compounds against other organisms.
Members of the genus Pseudoalteromonas are frequently found to produce toxic compounds, which could give them a competitive advantage (Holmström and
Kjelleberg, 1999). The production of antibacterial compounds is particularly well studied because of their potential application for new antibiotics in targeting human pathogens. Antibacterial activity has been demonstrated in P. aurantia (Gauthier and
Breittmayer, 1979), P. luteoviolacea (Gauthier and Flatau, 1976), P. rubra (Gauthier,
1979), P. citrea (Ivanova et al., 1998), P. maricaloris (Ivanova et al., 2002d), P. phenolica
(Isnansetyo and Kamei, 2003) and P. tunicata (James et al., 1996). These antibacterial compounds show a broad spectrum of activity against both Gram positive and Gram negative organisms. For example, it was demonstrated that two different classes of
46 Chapter 1 General introduction and literature review
compounds (cell-bound polyanionic macromolecules and small brominated compounds) were responsible for the antibacterial activity in different strains of P. luteoviolacea
(Gauthier and Flatau, 1976). The antibacterial compound of P. phenolica was found to be highly active against methicillin resistant S. aureus and different Enterococcus strains making it an interesting compound of potential clinical relevance. The novel antibacterial compound produced by P. tunicata is discussed in detail in Section 1.5.3.1.
Apart from antibacterial compounds, Pseudoalteromonas species also display a wide range of other inhibitory activity against higher organisms, including invertebrates, algae, diatoms, flagellates and fungi (Holmström and Kjelleberg, 1999). For example,
LPS of P. atlantica was found to be lethal for edible cancer crabs (Costa-Ramos and
Rowley, 2004). Further, a cytotoxic, yellow, chromo-peptide produced by P. maricalores can inhibit the development of sea urchin eggs (Ivanova et al., 2002d) and
Pseudoalteromonas strain Y causes rapid cell lysis of algal blooms (Lovejoy et al.,
1998). An extensively studied species in our laboratory is P. tunicata producing a range of biologically active compounds.
1.5.3 Pseudoalteromonas tunicata
P. tunicata was originally isolated from an adult tunicate, Ciona intestinales at a depth of 10 m off the Western coast of Sweden (Holmström et al., 1998) and has since been found associated with the green alga Ulva australis in Australian waters (Egan et al.,
2001). Furthermore, P. tunicata was detected in an algal bloom in Tasmania, Australia and on the surface of Ulva fusca, U. australis and C. intestinales in Aarhus Bay,
Denmark (Skovhus et al., 2004), suggesting that this species may be ubiquitous in the marine environment.
47 Chapter 1 General introduction and literature review
P. tunicata produces a range of inhibitory compounds against specific target organisms, including invertebrate larvae, algal spores, diatoms, heterotrophic flagellates, fungi and bacteria (Holmström et al., 1992; Holmström et al., 1998). Further, P. tunicata is dark green pigmented due to a yellow and a purple pigment and the production of these pigments has been linked to its antifouling activity (Egan et al., 2002a). A transposon library has been created and mutants with altered pigmentation were identified which had lost some or all antifouling properties. It was demonstrated that the production of inhibitory compounds is correlated with the production of the yellow pigment, because all yellow transposon mutants (which were disrupted in the purple pigment) retained full antifouling activity. In contrast, light purple, dark purple and white mutants have lost some, or all antifouling activities (Egan et al., 2002a). However, it was observed that only the white mutants lost the ability to inhibit bacterial growth along with the loss in ability to inhibit other target organisms. Sequence data indicated that in one of the white mutants, a gene involved in the regulation of pigment production and the expression of fouling inhibitors had been disrupted. The putative regulatory protein encoded in this disrupted gene (WmpR, white mutant phenotype R) is most similar to transcriptional activators CadC from E. coli and ToxR from V. cholerae (Egan et al., 2002a). Both
CadC and ToxR sense the environment and respond by regulating the transcription of specific genes. 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.
The different antifouling compounds produced by P. tunicata have been studied in detail. Inhibition of the larvae of Balanus amphitrite and C. intestinales was found to be due to an extracellular, heat stable, polar molecule of less than 500 kDa (Holmström et al., 1992). Algal spores of U. australis and Polysiphonia sp. were shown to be inhibited
48 Chapter 1 General introduction and literature review
by a small (3–10 kDa), extracellular, heat sensitive, polar compound (Egan et al., 2001).
A wide range of fungal and yeast isolates were also inhibited by P. tunicata and recently it was elucidated that the yellow pigment, which was classified as a tambjamine compound (Franks et al., 2005), is responsible for anti-fungal activity of P. tunicata.
Furthermore, preliminary evidence suggests that the purple pigment produced by P. tunicata is a violacein compound, which has been shown to be toxic to flagellates (Matz et al., 2004). So far the anti-diatom compound has not been further investigated.
1.5.3.1 The antibacterial and autolytic protein produced by P. tunicata
The antibacterial protein has been identified as a large extra-cellular protein of approximately 190 kDa in size. The protein contains at least two subunits of 60 and 80 kDa, linked together by noncovalent bonds (James et al., 1996). Analysis of the protein including Southern hybridisation and DNA sequencing from genomic libraries of P. tunicata indicate that it is encoded by a single open reading frame. Therefore a post- translational modification is required to retain the active protein of two subunits
(Stelzer, 1999). The antibacterial protein loses its activity when heated for 10 min at 80 ºC, however, it is stable across a pH range of 6.5 to 9.5 (James et al., 1996). A complete sequence analysis has been performed and the the protein was designated, AlpP (autolytic protein, Pseudoalteromonas). The primary nucleotide sequence data for the alpP gene has been deposited in the GenBank database under accession number AY 295768.
It has been proposed that AlpP confers an advantage for P. tunicata in the competition for space and nutrients on living marine surfaces (Holmström and Kjelleberg, 1999).
This protein inhibits the growth of both Gram+ and Gram- bacteria from a diverse range of environments including terrestrial, medical and marine isolates. Logarithmic phase growing cells of P. tunicata itself were found to be among the most sensitive of a broad
49 Chapter 1 General introduction and literature review
range of organisms tested. However, when cells reach the stationary phase they become resistant, which is seen as a possible explanation for their survival despite the production of an autotoxic factor (James et al., 1996). This observation suggests an important role for the antibacterial protein in the life cycle of P. tunicata. Investigations described in this thesis demonstrate that AlpP plays a role during biofilm formation of
P. tunicata (Chapters 2, 3 and 4). Further, elucidation of the mode of action is described in Chapter 3 and the roles of AlpP homologues in other organisms are discussed in
Chapter 5.
1.6 Aims and objectives of this study
This study uses the ubiquitous marine bacterium P. tunicata as a model organism to investigate biofilm development and a possible role for the antibacterial and autolytic protein AlpP. The overall hypothesis addressed is that the process of AlpP-mediated cell death benefits surviving cells of P. tunicata and may be common among other organisms containing an AlpP-homologue.
Specifically the aims of this study were:
1. To characterize and compare biofilm development of P. tunicata and a ǻAlpP
mutant in detail and determine a possible role for the antibacterial and autolytic
protein, AlpP (Chapter 2). Site directed mutagenesis of alpP was performed in
collaboration with Flavia Evans.
2. To determine the mode of action of AlpP in P. tunicata (Chapter 3)
3. To investigate whether the process of cell death benefits surviving dispersal cells
of P. tunicata by increasing their metabolic activity and rate of phenotypic
variation (Chapter 4). 50 Chapter 1 General introduction and literature review
4. To investigate whether AlpP has a common function during biofilm
development of organisms containing an AlpP homologue, including
Marinomonas mediterranea, Caulobacter crescentus, Chromobacterium
violaceum and Microbulbifer degradans. Biofilm experiments with M.
mediterranea and M. degradans were performed in collaboration with Dr.
Patricia Lucas-Elio.
51 Chapter 2: Biofilm development and cell death in P. tunicata. INTRODUCTION
2. Biofilm development and cell death in Pseudoalteromonas tunicata
2.1 Introduction
Pseudoalteromonas tunicata produces at least six novel extra-cellular compounds, each with inhibitory activity against a specific group of marine fouling organisms including bacteria, invertebrate larvae, algal spores, diatoms, heterotrophic flagellates and fungi
(Holmström et al., 1992; James et al., 1996; Holmström et al., 1998; Holmström and
Kjelleberg, 1999; Egan et al., 2000; Egan et al., 2001; Egan et al., 2002a; Egan et al.,
2002b). These inhibitory compounds are considered to provide an advantage to P. tunicata during the competitive colonization of living marine surfaces (Holmström et al., 1992; Holmström et al., 1998; Holmström and Kjelleberg, 1999; Egan et al., 2001;
Egan et al., 2002a).
One of the inhibitory compounds produced by P. tunicata is a 190 KDa antibacterial protein (AlpP) (James et al., 1996). AlpP inhibits growth of both Gram positive and
Gram negative bacteria including terrestrial, medical and marine isolates. Logarithmic phase growing cells of P. tunicata were found to be among the most sensitive to AlpP of the broad range of organisms tested, although stationary phase cells become resistant
(James et al., 1996).
There are examples where autolysis plays an important role in bacterial developmental processes (Lewis, 2000). For example, in Myxococcus xanthus a number of autocidal compounds are responsible for killing 80-90% of the cell population before fruiting body formation can occur (Wireman and Dworkin, 1977; Rosenbluh and Rosenberg,
1990). More recently a cell-cell signalling-mediated genetic mechanism was
52 Chapter 2: Biofilm development and cell death in P. tunicata. INTRODUCTION characterized whereby a subpopulation of Bacillus subtilis cells delay sporulation by killing sister cells and feeding on the nutrients that are released (Gonzalez-Pastor et al.,
2003). Thus, autolysis which appears undesirable to a single-cell organism may be advantageous to a bacterial population at the multicellular level.
Bacteria in biofilms are physiologically and morphologically different from their planktonic growing counterparts (Costerton et al., 1994). In laboratory systems biofilm bacteria often form highly differentiated 3-dimensional structures (microcolonies) which become surrounded by a network of water-channels. Water channels are important for delivering nutrients to the microcolony as well as removing toxic metabolites. Microcolony formation has been demonstrated for most model biofilm- forming bacteria, e.g. Escherichia coli, Vibrio cholerae and Pseudomonas aeruginosa
(Lawrence et al., 1991; Debeer et al., 1994; Watnick and Kolter, 1999; Davey and
O'Toole, 2000). Mature microcolonies can undergo complex differentiation. Dispersal of bacteria from the interior regions of microcolonies has been observed, resulting in the formation of transparent voids inside the microcolonies (Tolker-Nielsen et al., 2000;
Sauer et al., 2002; Kaplan et al., 2003a; Kaplan et al., 2003b).
It was recently shown that killing and lysis occurs reproducibly in localized regions in wild-type P. aeruginosa biofilms, inside microcolonies, by a mechanism that involves a prophage of P. aeruginosa (Webb et al., 2003b). The latter study proposed that cell death plays an important role in subsequent biofilm differentiation and dispersal, as surviving cells may benefit from the nutrients released during bacterial lysis (Webb et al., 2003b).
In this chapter a possible involvement of AlpP in P. tunicata biofilm formation is investigated. Biofilm development of wild-type P. tunicata and a ǻAlpP mutant was
53 Chapter 2: Biofilm development and cell death in P. tunicata. INTRODUCTION characterized using continuous culture flow-cells. It was shown that P. tunicata undergoes a highly reproducible pattern of cell death during normal development in a biofilm. The involvement of the autotoxic protein, AlpP, in the killing process was demonstrated through site-directed mutagenesis of the corresponding gene alpP.
54 Chapter 2: Biofilm development and cell death in P. tunicata. MATERIALS AND METHODS
2.2 Material and Methods
2.2.1 Bacterial strains and culture media
P. tunicata was routinely cultivated at room temperature in Väätänen nine salt solution
(VNSS) (Marden et al., 1985) (see Appendix). The ǻAlpP mutant was maintained on
VNSS medium containing the antibiotics streptomycin (100 µg ml-1) and kanamycin (50
µg ml-1). Biofilms were grown in marine minimal medium (MMM) (Neidhardt et al.,
1974) containing 0.01% trehalose (see Appendix).
2.2.2 Biofilm experiments
P. tunicata wild-type and 'AlpP mutant strains were grown in continuous-culture flow- cells (Figure 2.1) at room temperature as previously described (Moller et al., 1998).
Silicone tubing was connected to a peristaltic pump (model 323, Watson Marlow, UK).
A glass cover-slip was attached to a three channel autoclavable, poly-carbonate flow cell (channel dimensions 1 x 4 x 40 mm) using silicon glue (Selleys, Australia) and left to dry overnight. Before each run the flow cell was connected to silicon tubing on each side and autoclaved. The media was autoclaved separately and the glass-tubes connected to the flow cell inserted aseptically immediately before inoculation. The system was then filled with sterile media by turning on the flow. For inoculation the flow was stopped and metal screw down clamps positioned at each side of the flow cell.
Channels were inoculated with 0.5 ml of early stationary phase cultures containing approximately 1 x 109 cells ml-1 using a sterile syringe and needle. Flow cells were then inverted and incubated without flow for 1 h at room temperature allowing the cells to attach. After re-inverting the flow cells and removing the metal clamps flow was started with a mean flow velocity in the flow cells of 0.2 mm s-1, corresponding to laminar flow
55 Chapter 2: Biofilm development and cell death in P. tunicata. MATERIALS AND METHODS with a Reynolds number of 0.02.
media pump flow cell
waste
Figure 2.1: Flow cell setup. See text (section 2.2.2) for details.
2.2.3 Biofilm staining
To investigate cell death during biofilm development, biofilms were stained using the
Live/Dead Kit (Live/Dead® BacLight™ Bacterial Viability Kit for microscopy;
Molecular Probes, Eugene, Oregon USA). The Live/Dead Kit utilizes mixtures of
SYTO 9 green-fluorescent nucleic acid stain and the red-fluorescent nucleic acid stain, propidium iodide. These stains differ both in their spectral characteristics and in their ability to penetrate healthy bacterial cells. SYTO 9 stain generally labels all bacteria in a population; those with intact membranes and those with damaged membranes. In
56 Chapter 2: Biofilm development and cell death in P. tunicata. MATERIALS AND METHODS contrast, propidium iodide penetrates only bacteria with damaged membranes, causing a reduction in the SYTO 9 stain fluorescence when both dyes are present. Thus with an appropriate mixture of the SYTO 9 and propidium iodide stains, bacteria with intact cell membranes stain fluorescent green, whereas bacteria with damaged membranes stain fluorescent red. 500 µl of a 1:300 (vol/vol) dilution of the two components in NSS was injected into the flow cells. The flow was stopped for 15 min allowing the stain to penetrate into the cells and then washed away by flowing medium through the flow cell.
Microscopic observations were carried out immediately after staining.
2.2.3.1 Confocal laser scanning microscopy
Biofilms were examined using an Olympus LSMGB200 CSLM (Olympus Optical Co.
Ltd., Tokyo, Japan) equipped with 4X/0.13; 10X/0.4; 20X/0.7; 40X/0.95 and 60X/1.4 lenses. Image scanning was carried out with the 488-nm laser line of an argon laser.
2.2.3.2 Biofilm structure analysis with COMSTAT
The image analysis program, COMSTAT, was used to quantify potential differences between the P. tunicata wild-type strain and the ǻAlpP mutant (Heydorn et al., 2000).
This program quantifies three-dimensional biofilm image stacks. Images were acquired every 2 µm between the substratum layer and the tips of the microcolonies.
P. tunicata wild-type strain and the ǻAlpP mutant strain were grown in three different channels. From each channel, five image stacks were acquired at three different time points. Thus, 15 image stacks were analysed for each strain at each time point. Images were acquired at 2 µm intervals through the biofilm, and therefore the number of images in each stack varied according to the thickness of the biofilm. All images were acquired from random positions in the flow cell. Images were saved as TIFF files and
57 Chapter 2: Biofilm development and cell death in P. tunicata. MATERIALS AND METHODS thresholded before analysis. Thresholding an image stack results in a three-dimensional matrix with a value of ONE in positions where the pixel values in the original image are above or equal to the threshold value, and ZERO where the pixel values are below the threshold value. The value ONE represents positions containing biomass, while ZERO represents the background.
Four parameters were analysed for each time - point:
1. Maximum thickness of the biofilm [µm]
2. Average thickness of the biofilm [µm]: This provides a measure of the spatial size of the biofilm.
3. Total biomass [µm3 µm-2]: The bio-volume represents the overall volume of the biofilm, and provides an estimate of the biomass in the biofilm. The bio-volume is defined as the number of biomass pixels in all images of a stack multiplied by the voxel size [(pixel size)x x (pixel size)y x (pixel size)z] and divided by the substratum area of the image stack.
4. Roughness coefficient: This dimensionless coefficient is calculated from the thickness distribution of the biofilm. The thickness distribution locates the highest point
(µm) above each (x, y) pixel in the bottom layer containing biomass.
N L L * 1 fi f The roughness coefficient is defined as Ra ¦ N i 1 L f
in which Lfi is the ith individual thickness measurement, overlined Lf‘ is the mean thickness, and N is the number of thickness measurements. Biofilm roughness provides a measure of how much the thickness of the biofilm varies, and is an indicator of biofilm heterogeneity.
In order to compare biofilm structure between the wild-type strain and the mutant 15
58 Chapter 2: Biofilm development and cell death in P. tunicata. MATERIALS AND METHODS image stacks were analysed statistically using the student’s t-Test. An unpaired, two- sided t-Test was used to test the hypothesis that the difference between the means of the analysed parameters for both strains is equal to zero (null hypothesis). If the probability is less than the conventional 0.05, the null hypothesis is rejected and the conclusion is that the two means do indeed differ significantly.
2.2.4 Site-directed mutagenesis of alpP
To investigate the influence of AlpP on biofilm development, a ǻAlpP mutant derivative of P. tunicata was generated by allelic displacement. The alpP gene (2.3 kb)
(GenBank accession number AY 295768) was amplified using the following primers: forward (5’-gag aat tcc ata tga att taa aaa tcc atc c-3’) and reverse (5’-agt cta agc ata tgg gat cct gcg taa gtg ata tcc c-3’). A kanamycin (km) resistance cassette (1.2 kb) was amplified from the plasmid pUCR4K (Amersham) using the following primers: forward
(5’-tac tag atc tca cgt gcg tcg acc tgc agg g-3’) and reverse (5’-gtg aag atc tca cgt gcc gga tcc gtc gac c-3’). The alpP gene and km resistance cassette were each cloned separately into pGEM T-Easy vectors according to the manufactures’ instructions
(Invitrogen). The km resistance cassette was then inserted into alpP using the Pml I restriction enzyme to create the plasmid pGEM alpP::Kmr.
The alpP knockout plasmid, pAP704, was constructed by inserting the alpP::kmr construct into the Sma I site of the suicide vector pGP704 (Miller and Mekalanos,
1988), which was then transformed into E. coli SM10. The P. tunicata ǻAlpP mutant was constructed by site-directed mutagenesis using conjugation with P. tunicata Smr and E. coli SM10 containing the vector pAP704. Exconjugants with the alpP::kmr constructs inserted into the chromosome were selected using VNSS plates supplemented with streptomycin (200 µg ml-1) and kanamycin (85 µg ml-1), and confirmation that the
59 Chapter 2: Biofilm development and cell death in P. tunicata. MATERIALS AND METHODS alpP::kmr cassette has been inserted into the genome of P. tunicata was obtained by
PCR.
2.2.5 Characterization of ǻAlpP mutant
2.2.5.1 Testing ǻAlpP mutant supernatant for antibacterial activity
Loss of autocidal activity in the concentrated supernatants of P. tunicata exconjugants was confirmed by using the drop-plate assay for the detection of AlpP activity as previously described (James et al., 1996). Briefly, 20 µl drops were placed in a serial dilution on a lawn of P. tunicata cells. Plates were incubated at room-temperature and the zones of inhibition observed after 24 h.
2.2.5.2 Assessment of ǻAlpP mutant growth
A growth rate analysis was performed comparing the P. tunicata wild-type and ǻAlpP mutant. Overnight cultures (100 µl) were inoculated into 10 ml sterile VNSS and incubated with agitation (130 rpm) at room-temperature. The optical density was measured at 600 nm at 30 min intervals.
2.2.5.3 Acrylamide gel analysis of P. tunicata wild-type and ǻAlpP mutant
2.2.5.3.1 Preparation of concentrated supernatant and partial AlpP purification
Partially purified AlpP was prepared according to a method adapted from James et al.
(1996). Briefly, P. tunicata was grown in MMM (see Appendix) for 48 h with shaking at room-temperature. Cells were then harvested by centrifugation at 15,000 g for 15 minutes at 22 °C. The cell pellet was resuspended in fresh media (0.5 ml / 2g pellet) and incubated for a further 24 h under static conditions. Following centrifugation for 1.5 h at
23,700 g and 4 °C, the supernatant was filtered through a 0.22 µm membrane. Finally,
60 Chapter 2: Biofilm development and cell death in P. tunicata. MATERIALS AND METHODS the sample was partially purified by passing it through a 5-ml Econo-Pac Q anion- exchange cartridge (strongly basic anion exchanger; Bio-Rad Laboratories). The antibacterial fraction was eluted at between 0.1 and 0.2 M NaCl (Tris-HCl, pH 5) using a step gradient.
2.2.5.3.2 Native PAGE analysis and overlay assay
In order to assess the possible activity of AlpP in both P. tunicata wild-type and ǻAlpP mutant partially purified supernatants, non-denaturing, native polyacrylamide gel electrophoresis was 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) (Bio-Rad, Hercules, USA) 0.125 M
Tris-HCl (pH 6.8), 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), 0.06 % (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), 20 % (v/v) glycerol and a twentieth volume of
10 % (v/v) bromophenol blue solution. Broad range molecular weight markers (Bio-
Rad, Hercules, USA) were prepared by adding sample buffer. Partially purified supernatant (see Section 2.2.5.3.1) was loaded into duplicate gels and electrophoresis 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. One gel was stained with Coomassie brilliant blue solution (0.1% w/v, 50% v/v methanol, 7% v/v acetic acid) for 2 h, following de-staining in 50% v/v methanol and
7% v/v acetic acid. The second gel was washed for 30 min in MilliQ before overlaying with P. tunicata cells. Cells for the overlay were inoculated into VNSS broth and 61 Chapter 2: Biofilm development and cell death in P. tunicata. MATERIALS AND METHODS allowed to grow for 6 h to reach logarithmic growth phase. Immediately before the overlay the culture was mixed in a 1/10 dilution in VNSS at 40 °C containing 0.3% agar. The mixture was poured on top of the gel and incubated for 24 h before possible zones of inhibition were observed.
2.2.5.4 Liquid chromatography tandem mass spectrometry (LC-MSMS) analysis
In-gel digestion was performed as described by Nawrocki et al., (1998). The excised gel plugs were washed in 50 mM NH4HCO3/acetonitrile (60/40) and dried by vacuum
-1 centrifugation. Modified trypsin (8 ng Pl ), dissolved in 50 mM NH4HCO3, was added to the dry gel pieces and incubated on ice for 1 h for re-swelling. After removing the supernatant, additional digestion buffer was added to cover the gel pieces and the digestion continued at 37 oC for 4-18 h. For liquid chromatography tandem mass spectrometry (LC-MSMS), an aliquot of the peptide mixture was dried by vacuum centrifugation. The dried peptides were re-dissolved in 0.1 % trifluoroacetic acid (TFA) prior to LC-MSMS.
Nanoflow liquid chromatography tandem mass spectrometry analysis was performed using a QTOF Ultima mass spectrometer (Waters/Micromass UK Ltd., Manchester,
UK) employing automated data dependent acquisition. The mass spectrometer was operated in positive ion mode with a source temperature of 80 °C and a counter-current gas flow rate of 150 l h-1. Data dependent acquisition was employed. The peptides were bomb-loaded onto a custom-made capillary reversed-phase column (75 µm i.d.; 360 µm o.d.; Zorbax® SB-C18 3.5 µm (Agilent, Wilmington, DE)). A nanoflow-HPLC system
(Ultimate; Switchos2; Famos; LC Packings, Amsterdam, The Netherlands) was used for separation of the peptide mixture prior to MS detection. Peptides were eluted at 200 nl min-1 by an increasing concentration of acetonitrile (2 % min-1 gradient). A MS-TOF
62 Chapter 2: Biofilm development and cell death in P. tunicata. MATERIALS AND METHODS survey spectrum was recorded for 1 sec. The three most intense ions present in the MS-
TOF spectrum were selected and fragmented by collision-induced dissociation in the second quadruple (4 sec. per MS/MS spectrum).
The raw fragment data from each separation were converted to a Micromass pkl format using the MassLynx 3.5 ProteinLynx software, and the resulting pkl files were searched against protein sequence databases, using an in-house MASCOT server (v. 1.8) (Matrix
Sciences, London, UK).
2.2.5.5 Characterization of ǻAlpP mutant biofilm formation
Biofilm development of the ǻAlpP mutant was compared with P. tunicata wild-type.
Flow cell experiments were carried out and biofilms stained with the LIVE/DEAD
BacLight Bacterial Viability Kit as described above.
2.2.5.6 Add-back of purified AlpP to the ǻAlpP mutant and wild-type biofilms
To determine whether AlpP could restore killing in ǻAlpP mutant biofilms, purified
AlpP was added into flow cell channels containing ǻAlpP mutant biofilms. Purified
AlpP was also added to “young” P. tunicata wild-type biofilms, before the normal onset of cell death, in order to determine whether AlpP could induce early killing in the wild- type strain. AlpP was prepared and partially purified from the supernatant as described above (2.2.5.3.1). AlpP (10-12 µg in dialysis buffer; 20 mM Tris-HCl, 0.3 M NaCl, pH
5) was injected into the flow cells using a syringe needle. Silicone tubing either-side of the flow cell was then blocked-off by using tubing clamps. As a control, dialysis buffer was inoculated into separate flow cell channels. Biofilms were incubated at room temperature for 5 h without flow before staining with the LIVE/DEAD BacLight
Bacterial Viability Kit and visualizing with the CLSM.
63 Chapter 2: Biofilm development and cell death in P. tunicata. RESULTS
2.3 RESULTS
2.3.1 P. tunicata biofilm development and cell death
Biofilm development and possible cell death of P. tunicata wild-type was investigated in glass flow-cells by using the LIVE/DEAD BacLight Bacterial Viability Kit.
Immediately after inoculation single viable cells were observed attached to the substratum (Figure 2.2 A). After 24 h microcolonies had developed (Figure 2.2 B), and no dead cells were visible within the biofilm. Between 48 and 96 h post-inoculation, dead cells occurred in the interior portions of microcolonies and were surrounded by an outer layer of live cells (Figure 2.2 C, D). Dead cells and partially lysed cells were observed as well as amorphous red PI-stained material which was possibly DNA- containing debris from lysed cells. A subpopulation of cells in the region of killing remained viable. At this stage of biofilm development, the substratum was completely covered by bacteria. After 96 h, when killing had occurred inside all microcolonies, open voids within the regions of killing inside the microcolonies were observed (Figure
2.2 E). Regions of extensive killing subsequently occurred within and around microcolonies and the biofilm structure started to disrupt and detach. Once the biofilm had dispersed, no microcolonies could be observed while only single live cells remained attached within the flow cell (Figure 2.2 F). Similar results were observed in five additional sets of experiments.
64 Chapter 2: Biofilm development and cell death in P. tunicata. RESULTS
A B
C D
EF
Figure 2.2: Biofilm development and cell death of the P. tunicata wild-type strain. Biofilms stained with the BacLight LIVE/DEAD Bacterial Viability Kit. Red Propidium Iodide-stained cells have a compromised cell membrane and are dead. Time-points after inoculation are shown as follows: (A) 1 h (B) 24 h (C) 48 h (D) 72 h (E) 144 h (F) 168 h, Scale bar 50 µm.
65 Chapter 2: Biofilm development and cell death in P. tunicata. RESULTS
2.3.2 Characterization of the ǻAlpP mutant
2.3.2.1 Assessment of ǻAlpP mutant growth
To investigate the role of AlpP during P. tunicata biofilm growth a ǻAlpP mutant derivative of P. tunicata was generated. First, it was assessed whether the mutation in alpP causes an alteration in the growth ability. No significant difference in growth ability between P. tunicata wild-type and ǻAlpP mutant could be observed (Figure 2.3).
Both strains showed a lag phase of approximately 2 h followed by a logarithmic growth phase of 6 h before entering stationary phase 8 h post inoculation (Figure 2.3).
0.30
0.25
0.20
0.15 wild-type ǻAlpP 0.10
Absorbance 600 nm 0.05
0.00 0246810121416 Time (h)
Figure 2.3: Growth curve of P. tunicata wild-type (Ƈ) and ǻAlpP mutant (Ŷ). Cultures were inoculated into VNSS and the absorbance (600 nm) measured every 30 min. No significant difference in growth ability could be observed between the wild- type and the ǻAlpP mutant strain.
66 Chapter 2: Biofilm development and cell death in P. tunicata. RESULTS
2.3.2.2 Drop test assay for ǻAlpP mutant
To determine whether the ǻAlpP mutant has lost its autotoxic activity a drop test assay was performed. As previously reported, AlpP activity can be detected through the formation of an inhibition zone when droplets of P. tunicata wild-type supernatant are placed onto a lawn of bacteria (Figure 2.4 A) (James et al., 1996). However, no inhibition could be observed when concentrated supernatant of the ǻAlpP mutant was placed onto a lawn of P. tunicata cells, confirming that AlpP has been disrupted and made inactive (Figure 2.4 B).
A B
Figure 2.4: P. tunicata drop-plate assay. (A) P. tunicata wild-type concentrated supernatant on a lawn of P. tunicata cells. Each drop represents a concentration of AlpP protein from highest to lowest dilution in a clockwise direction. (B) ǻAlpP mutant concentrated supernatant on a lawn of P. tunicata cells. No inhibition could be observed.
67 Chapter 2: Biofilm development and cell death in P. tunicata. RESULTS
2.3.2.3 PAGE analysis of P. tunicata wild-type and ǻAlpP mutant supernatant
It has been previously reported that AlpP is approximately 190 kDa in size (James et al.,
1996). The results of the PAGE analysis showed that the wild-type partially purified supernatant showed a band of approximately 190 kDa in size under non-denaturing conditions (Figure 2.5 A; lane I, band a). This band correlated to a zone of inhibition observed in the overlay assay (Figure 2.5 B, lane I). A faint band of similar size to AlpP
(approximately 190 kDa) was also visible in the ǻAlpP mutant partially purified supernatant (Figure 2.5 A, lane II, band b), but no inhibition and thus antibacterial activity could be observed from this band (Figure 2.5 B, lane II).
A B III III
190 kDa a b
Figure 2.5: Native PAGE analysis (A) and overlay of P. tunicata wild-type and ǻAlpP mutant (B). (A) Coomassie stained gel with lane I, partially purified supernatant of P. tunicata wild-type; lane II, partially purified supernatant of ǻAlpP mutant. Band a and b correlate to protein(s) of approximately 190 kDa in size from wild-type and ǻAlpP mutant, respectively. (B) Corresponding gel with overlay of P. tunicata cells. The 190 kDa band of wild-type partially purified supernatant has autotoxic activity as determined by a zone of inhibition in the overlay (black arrow). 68 Chapter 2: Biofilm development and cell death in P. tunicata. RESULTS
2.3.2.3.1 LC-MSMS analysis
Because a faint band of similar size to AlpP was also present in the supernatant of the
ǻAlpP mutant (Figure 2.5 A, lane II), bands from the partially purified supernatant of both wild-type (“band a”) and ǻAlpP mutant (“band b”) were excised from the gel and mass spectrometry analysis was performed to confirm their identity. As expected “band a” excised from the wild-type gel corresponding to the zone of inhibition (Figure 2.5 B, lane I) was identical to AlpP (score 911). However, “band b” excised from the ǻAlpP mutant gel of similar size (190 kDa, Figure 2.5 A, lane II) matched with a very low score (74) to a putative cytosol aminopeptidase, further confirming that the mutant does not produce AlpP.
2.3.3 Biofilm development of the P. tunicata ǻAlpP mutant
The biofilm development of the ǻAlpP mutant was investigated compared to the P. tunicata wild-type. In a similar fashion to the wild-type, the ǻAlpP mutant formed a differentiated biofilm with a 3-dimensional structure consisting of microcolonies
(Figure 2.6). However, unlike P. tunicata wild-type, the ǻAlpP mutant did not undergo cell death at any stage of biofilm development, and no hollow cavities within microcolonies were observed. Moreover, detachment of the biofilm was considerably delayed, with the majority of microcolonies persisting after 9 days (wild-type biofilms were completely dispersed after 7 days) (Figure 2.6 F). These results indicate that AlpP- mediated killing may affect dispersal from within the interior of attached microcolonies as well as large scale detachment of the whole biofilm.
69 Chapter 2: Biofilm development and cell death in P. tunicata. RESULTS
A B
C D
EF
Figure 2.6: Biofilm development of the P. tunicata ǻAlpP mutant. Biofilms stained with the LIVE/DEAD BacLight Bacterial Viability Kit. Time-points after inoculation are shown as follows: (A) 1 h (B) 24 h (C) 48 h (D) 72 h (E) 144 h (F) 168 h, Scale bar 50 µm.
2.3.4 Characterization of the biofilm structure with COMSTAT
Biofilm structures of both P. tunicata wild-type and ǻAlpP mutant were also characterized with the biofilm quantification software COMSTAT (Heydorn et al., 70 Chapter 2: Biofilm development and cell death in P. tunicata. RESULTS
2000). The results are summarized in Table 2.1. The following parameters were assessed: Average and maximum thickness, total biomass and roughness. No significant differences between the wild-type and the ǻAlpP mutant strain were observed for all parameters after 24, 72 and 120 h of biofilm development. Biofilms of both strains increased in thickness and biomass over time, whereas roughness decreased, indicating that the substratum coverage was higher and a denser, even layer of cells was formed over the time-course of the experiment.
The t-Test analysis revealed that there was no significant difference between wild-type and ǻAlpP mutant biofilms in maximum thickness, average thickness, total biomass and roughness coefficient for all three time-points. These results show that despite the lack of cell death within ǻAlpP mutant biofilms it is still capable of forming the normal biofilm architecture characteristic of wild-type P. tunicata.
71 Chapter 2: Biofilm development and cell death in P. tunicata. RESULTS
Table 2.1: Characterization of P. tunicata wild-type and ǻAlpP mutant biofilm structure using the biofilm image analysis software COMSTAT.
Parameter Hours Wild-type ¨AlpP t-Test Maximum thickness 24 12.00 ± 2.45 16.67 ± 4.68 0.07 (µm) 72 28.29 ± 5.59 24.00 ± 4.62 0.11 120 38.00 ± 5.29 37.60 ± 2.19 0.91
Average thickness 24 2.01 ± 0.48 2.15 ± 1.98 0.86 (µm) 72 14.45 ± 1.96 10.63 ± 3.62 0.06 120 31.64 ± 6.7 24.95 ± 1.44 0.22
Total biomass 24 2.12 ± 0.46 2.00 ± 1.78 0.87 (µm3 µm-2) 72 11.77 ± 1.41 10.49 ± 4.87 0.39 120 24.79 ± 5.64 27.38 ± 1.35 0.51
Roughness coefficient 24 1.21 ± 0.20 1.38 ± 0.48 0.07 72 0.25 ± 0.07 0.49 ± 0.36 0.07 120 0.11 ± 0.04 0.14 ± 0.02 0.35
2.3.5 Add-back of purified AlpP to P. tunicata biofilms
To provide direct evidence that AlpP is involved in cell death in P. tunicata biofilms, purified AlpP was added to the ǻAlpP mutant strain after 48 h of biofilm development in an attempt to restore AlpP-mediated killing in the biofilm. Similar to the P. tunicata wild-type, the autotoxic protein caused cell death in the centre of microcolonies, and also killed other cells throughout the biofilm (Figure 2.7 A).
In addition, purified AlpP was also added to young wild-type biofilms (30 h) before the normal onset of killing within the microcolonies. Add back of AlpP induced early killing in P. tunicata wild-type biofilms (Figure 2.7 C). Cell death occurred within microcolonies as well as in other regions of the biofilm.
72 Chapter 2: Biofilm development and cell death in P. tunicata. RESULTS
A B
CD
Figure 2.7: Addition of purified of AlpP to P. tunicata biofilms. Biofilms stained with the LIVE/DEAD BacLight Bacterial Viability Kit. (A) Add back of AlpP to ǻAlpP mutant 48 h biofilms, (B) ǻAlpP mutant biofilms plus buffer control (20 mM Tris, 0.3 M NaCl), (C) AlpP add back to young (30 h) P. tunicata wild-type biofilms before the normal onset of killing within the biofilm, (D) P. tunicata wild-type biofilm plus buffer control (20 mM Tris, 0.3 M NaCl). Scale bar 50 µm. Similar results were obtained in four replicate experiments.
73 Chapter 2: Biofilm development and cell death in P. tunicata. DISCUSSION
2.4 Discussion
It was demonstrated that P. tunicata forms a microcolony based biofilm and a repeatable pattern of cell death occurs during normal biofilm development mediated by the autotoxic protein AlpP. Although the experiments were carried out in an artificial flow cell environment, differentiated microcolony structures, similar to those reported here in P. tunicata biofilms, are commonly observed in natural and medical situations
(Costerton et al., 1999; Singh et al., 2000; Anderson et al., 2003; Webb et al., 2003a).
This study showed that in P. tunicata, regions of lysis lead to the formation of hollow microcolonies presumably aiding dispersal. Similar cell death events and the formation of hollow microcolonies were shown to occur in several other biofilm forming bacteria, including V. cholerae (D. McDougald, J. S. Webb and S. Kjelleberg, unpublished),
Serratia marcescens (K.W. Lam, S.A. Rice and S. Kjelleberg, unpublished),
Caulobacter crescentus (Entcheva-Dimitrov and Spormann, 2004) and the opportunistic human pathogen P. aeruginosa (Tolker-Nielsen et al., 2000; Sauer et al., 2002; Webb et al., 2003b). Other observations have shown that loss of viability also occurs inside microcolonies in mixed species biofilms formed by oral bacteria (Auschill et al., 2001;
Hope et al., 2002) and within the biofilm flocs of waste-water treatment processes
(Meyer et al., 2003). Thus cell death inside microcolonies may be widespread among biofilm-forming bacteria and it is possible that these killing events play an important role in biofilm development and dispersal.
With the exception of P. aeruginosa, where loss of viability was demonstrated to be due to the activity of a prophage (Webb et al., 2003b), the mechanisms leading to biofilm cell death have not been elucidated in other organisms. Results presented in this chapter provide evidence that cell death in P. tunicata is mediated by the autolytic protein AlpP. 74 Chapter 2: Biofilm development and cell death in P. tunicata. DISCUSSION
A ǻAlpP mutant derivate was generated which did not undergo cell death during any stage of biofilm development. Further, the ǻAlpP mutant did not form hollow microcolonies and was deficient in biofilm dispersal. It was shown that the ǻAlpP mutant had lost its autotoxic activity as no inhibition could be observed when concentrated supernatant was placed on a lawn of susceptible P. tunicata cells.
Moreover, a non-denaturing PAGE analysis revealed that the protein band previously identified as AlpP only occurred in the P. tunicata wild-type and this band correlated to a zone of inhibition when the gel was overlaid with P. tunicata cells.
COMSTAT analysis revealed that P. tunicata wild-type and ǻAlpP mutant biofilms have a similar structure. No significant differences were measured in the four parameters assessed, despite the lack of cell death during ǻAlpP mutant biofilm formation, suggesting a role for this process in biofilm dispersal rather than biofilm architecture. It was observed that cell death occurred in localized regions within microcolonies of P. tunicata biofilms, but that a subpopulation of cells remained viable in these regions. It is possible that killing and lysis benefits this subpopulation of cells
(through the release of nutrients and DNA) which then undergo continued differentiation and dispersal. However, little is known about the mechanism/s by which hollow microcolonies are formed, or by which cells disperse from the internal regions inside microcolonies. Generally, large-scale biofilm detachment and sloughing are known to be influenced by both physical and physiological properties. Hydrodynamic conditions such as shear stress and velocity of fluids can greatly affect the biofilm matrix and detachment of biofilms (Cowan et al., 2000; Purevdorj et al., 2002). In addition to hydrodynamics, biofilm detachment is also influenced by environmental sensors and nutritional conditions. For example, it was shown that the global regulatory protein CsrA (carbon storage regulator) serves as an activator of biofilm dispersal
75 Chapter 2: Biofilm development and cell death in P. tunicata. DISCUSSION through the regulation of intracellular glycogen biosynthesis and catabolism in E. coli
(Jackson et al., 2002).
Another possible role for AlpP-mediated cell death in P. tunicata is that it may be involved in regulating biofilm growth on the surface of living hosts in the marine environment. It is known that many sessile algae and animals have evolved defence mechanisms against fouling by producing metabolites that can influence the settlement, growth and survival of other organisms e.g. (Davis and Wright, 1990; de Nys et al.,
1995). However, one hypothesis is that algae and animals lacking chemical and non- chemical defences rely on secondary metabolites produced by associated surface bacteria such as P. tunicata as their defence against fouling (Thomas and Allsopp, 1983;
Holmström et al., 1992; Kon-ya et al., 1995). Cell death and dispersal of a subpopulation of cells within P. tunicata biofilms may protect its host against uncontrolled biofilm formation and fouling by P. tunicata itself.
It could also be speculated that the process of cell death during biofilm formation is an evolved capacity and may benefit the bacteria at the multicellular level in a similar manner to that previously described for autolysis in Myxococcus xanthus (Wireman and
Dworkin, 1977; Rosenbluh and Rosenberg, 1990), Bacillus subtilis (Gonzalez-Pastor et al., 2003) and Streptomyces antibioticus (Mendez et al., 1985; Miguelez et al., 1999;
Fernandez and Sanchez, 2002). In these organisms, death of a subpopulation is required to complete a developmental process such as fruiting body formation and sporulation, respectively. Possible benefits of cell death during biofilm development of P. tunicata are described in Chapter 4.
The mechanism of action of AlpP-mediated cell death and biofilm killing is elucidated in Chapter 3.
76 Chapter 3 The mode of action of AlpP in P. tunicata INTRODUCTION
3. The mode of action of AlpP in P. tunicata
3.1 Introduction
The production of antibacterial compounds from marine bacteria has long been well documented in the literature. Already at the beginning of the last century it was discovered that natural seawater is bactericidal for many non-marine bacteria (Korinek,
1927; Rosenfeld and Zobell, 1947) and thus suggested that the water contains toxic compounds, some possibly produced by marine bacteria. More recently, a diverse range of biologically active compounds have been isolated and characterized from marine bacteria, including brominated, low molecular weight compounds from the genera
Alteromonas (Gauthier and Flatau, 1976) and Pseudoalteromonas (Kalinovskaya et al.,
2004; Sobolevskaya et al., 2004) as well as compounds from the genera Pseudomonas
(Burkholder et al., 1966; Wratten et al., 1976) and Chromobacterium (Andersen et al.,
1974). It has become clear that a diverse range of antibiotic substances is produced by different marine bacteria.
Since the first description of antibacterial properties of P. tunicata by James (1996), the mode of action of AlpP has been the focus of several research projects. AlpP was found to be bacteriolytic and a series of P. tunicata transposon mutants were generated with altered resistance to AlpP exposure (Dalisay-Saludes, 2004). Two membrane transport mechanisms, an ABC (type I) exporter and a type II secretion pathway, appeared to be involved in the mode of action of AlpP without directly transporting AlpP.
Recently the mode of action of an antibacterial protein (LodA, previously marinocine) from the marine bacterium Marinomonas mediterranea was discovered (Lucas-Elío et al., 2006).
This protein was found to have homology to AlpP of P. tunicata (Lucas-Elio et al., 2005) and was shown to generate hydrogen peroxide from L-lysine which subsequently led to 77 Chapter 3 The mode of action of AlpP in P. tunicata INTRODUCTION cell death. This lysine oxidase activity could be inhibited by the presence of catalase
(Lucas-Elío et al., 2006).
In this chapter it is demonstrated that the antibacterial activity of AlpP is also due to its lysine oxidase activity. AlpP produced hydrogen peroxide from the substrate L-lysine and its activity could be inhibited by catalase. It is also shown that hydrogen peroxide production occurs within P. tunicata wild-type biofilms. The hydrogen peroxide is likely a result of the AlpP activity, as hydrogen peroxide is only detected at the time of killing localised to microcolonies. Further, hydrogen peroxide could not be detected within ǻAlpP mutant biofilms during any stage of biofilm development. This study also revealed that removal of hydrogen peroxide from the biofilm through the addition of catalase leads to less microcolony formation, further demonstrating that AlpP mediated cell death, through hydrogen peroxide production, plays an important role in biofilm differentiation of P. tunicata.
.
78 Chapter 3 The mode of action of AlpP in P. tunicata MATERIALS AND METHODS
3.2 Materials and Methods
3.2.1 Bacterial strains and culture conditions
P. tunicata was routinely cultivated at room temperature in Väätänen nine salt solution
(VNSS) (Marden et al., 1985) (see Appendix). The ǻAlpP mutant was maintained on
VNSS medium containing the antibiotics streptomycin (100 µg ml-1) and kanamycin (50
µg ml-1). Biofilms were grown in marine minimal medium (MMM) (Neidhardt et al.,
1974) containing 0.01 % trehalose (see Appendix).
3.2.2 AlpP purification
An improved method for AlpP purification adapted from James et al. (1996) was developed. AlpP was purified from P. tunicata culture supernatant using dialysis, ion- exchange chromatography and ultra-filtration. Concentrated supernatant was prepared as described in Section 2.2.5.3.1. The concentrated supernatant was then dialyzed
(12,000 kDa cut off; Sigma, Castle Hill, Australia) overnight against 2 l 20 mM Tris
(pH 7.4). A strong anionic ion-exchange matrix (High Q strong anion-exchanger, Bio-
Rad, Hercules, USA) was used to purify AlpP from the P. tunicata supernatant. The protocol was performed on a BioLogic chromatography unit (BioRad, USA). After washing the column with 50 ml of 20 mM Tris-HCl (pH 7.4) buffer, the dialysed supernatant was loaded onto the column (1.5 ml x min-1). Following 2 rinsing steps of the column with 100 ml of 20 mM Tris-HCl buffer and 150 ml of 150 mM NaCl, a linear gradient ranging from 150 to 500 mM NaCl concentration was applied. AlpP was eluted between 250 mM and 350 mM NaCl in 20 mM Tris-HCl (pH 7.4). The eluate was further purified by ultrafiltration with a 100 kDa molecular cut-off filter (YM100;
Diaflo, Amicon, Lexington, USA).
79 Chapter 3 The mode of action of AlpP in P. tunicata MATERIALS AND METHODS
3.2.2.1 Sodium dodecyl sulphate polyacrylamide gel (SDS PAGE) analysis of
purified AlpP
Electrophoretic analysis of samples of the different purification steps (crude supernatant, dialysis, ion-exchange chromatography and ultra-filtration) was performed using a similar protocol as described in Section 2.2.5.3.2. To achieve denaturing conditions sodium dodecyl sulphate (SDS) was added to the stacking gels (0.3 % (w/v) and the separating gels (0.1 % (w/v). 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 (Bio-
Rad, Hercules, USA) were prepared by adding sample buffer. Both samples and markers were boiled at 100 °C for 90 sec. The samples were loaded on the gel and electrophoresis 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 and 0.1 % (w/v) SDS.
3.2.2.2 Determination of protein concentration
Total protein concentration was determined using the bicinchoninic acid (BCA) method on a microtitre plate based assay. This system combines the reaction of proteins with
Cu2+ ions to yield cuprous ions (Cu1+) which in the presence of BCA leads to a colour reaction. Ten microliters 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, Castle Hill,
Australia) and 1 part 4 % (w/v) CuSO4 solution (Sigma, Castle Hill, Australia), was added. The plate was incubated at 37 °C for 30 min. The colour reaction was measured
80 Chapter 3 The mode of action of AlpP in P. tunicata MATERIALS AND METHODS by absorbance at 600 nm in a Wallac 1420 multilabel counter. Samples were compared to a standard curve consisting of known concentrations of bovine serum albumin (BSA)
(Sigma, Castle Hill, Australia) ranging from 0.2 to 1.2 mg ml-1. A blank consisting of only the diluents and the working reagents was used as a base line reaction.
3.2.3 Drop test assay for AlpP activity testing
Following the ultra-filtration the purified fractions were tested for activity against P. tunicata. Twenty µl drops were placed in a serial dilution on a lawn of P. tunicata cells.
Plates were incubated at room-temperature and the zones of inhibition observed after
24 h.
3.2.4 Amplex Red assay
The Amplex Red Hydrogen Peroxide/Peroxidase Assay Kit (Molecular Probes A22188,
Eugene, Oregon, USA) assay was performed according to the manufacturer’s instruction. In the presence of peroxidase the Amplex Red reagent reacts with hydrogen peroxide in a 1:1 stoichiometry to produce the red fluorescent oxidation product resuforin. Resuforin fluorescence was measured at excitation of 550 nm and an emission of 590 nm in a Wallac 1420 multilabel counter. All reactions were prepared in generic 8 x 12 microtiter plates. Hydrogen peroxide (Univar, Kirkland, USA) concentrations ranging from 2 µm to 12 µM were used as a standard. L-lysine (50 mM,
Sigma-Aldrich, Castle Hill, Australia) was used as a substrate for AlpP. The ability of
AlpP to generate hydrogen peroxide from L-lysine was tested with AlpP concentrations ranging from 0.4 to 4 ng. Separate reactions were prepared by adding catalase (0.1 mg ml-1) to the reaction of AlpP and L-lysine.
81 Chapter 3 The mode of action of AlpP in P. tunicata MATERIALS AND METHODS
3.2.5 Determination of the minimum inhibitory catalase concentration
Catalase reduces H2O2 to H2O and O2 and thus counteracts the activity of AlpP by removing the produced hydrogen peroxide. AlpP was purified as described above ( see
Section 3.2.2) and 1.3 ng were mixed with catalase (Sigma, Saint-Louis, USA) in concentrations ranging from 0 to 1 mg. Droplets of the mixtures (20 µl) were placed on a lawn of P. tunicata cells to determine the minimum catalase concentration necessary to abolish AlpP activity.
3.2.6 Assessment of storage conditions on the stability of AlpP
AlpP was routinely stored in glycerol (50 % vol/vol) at -20 °C. However, to investigate the stability of AlpP, storage temperatures of 4 °C and 25 °C were also tested. The activity of AlpP was tested after 1 month using the drop test assay (see Section 3.2.3).
3.2.7 Biofilm experiments
P. tunicata wild-type and ǻAlpP mutant strains were grown in continuous-culture flow- cells as described in Chapter 2 (see Section 2.2.2). Briefly, channels were inoculated with 0.5 ml of early stationary phase cultures containing approximately 1 x 109 cells ml-
1 and incubated without flow for 1 h at room temperature. Flow was then started with a mean flow velocity in the flow cells of 0.2 mm s-1, corresponding to laminar flow with a
Reynolds number of 0.02.
3.2.7.1 Detecting hydrogen peroxide in biofilms
To investigate whether hydrogen peroxide production occurs during biofilm formation, biofilms were stained with the Amplex Red reagent. Amplex Red stock solution (25
µM) and horseradish peroxidase (HPR) (0.14 units ml-1) were diluted into 1 ml biofilm
82 Chapter 3 The mode of action of AlpP in P. tunicata MATERIALS AND METHODS media (MMM). The stain was injected into the flow cells using a syringe needle.
Silicone tubing at either-side of the flow cell, was then blocked-off by using tubing clamps. Immediately after staining, biofilms were visualized with an epifluorescence microscope (Leica Microsystems, Wetzlar, Germany). The microscope is equipped with x10/0.25; x40/0.65 and x100/1.25 objectives. Amplex Red fluorescence was observed at an excitation of 515-560 nm and an emission of 590 nm. Bright-field images were taken from the same field of view.
3.2.7.2 Removing hydrogen peroxide from biofilms
To remove hydrogen peroxide from biofilms, catalase was added to the biofilm media in a non-growth inhibitory concentration (see Section 3.2.8). First, biofilms were allowed to establish for 24 h. A stock concentration of catalase was sterile filtered and added to the biofilm media at a final concentration of 200 µM. After 3 days of incubation, the biofilm was stained with Amplex Red and observed with an epifluorescence microscope
(see Section 3.2.7.1).
3.2.8 Assessment of catalase on growth of P. tunicata
A growth rate analysis of P. tunicata wild-type was performed in the presence of different catalase concentrations (0, 3.4, 53.6, 107.3, 429.1 µg ml-1). Overnight cultures
(100 µl) were inoculated into 10 ml sterile VNSS with the appropriate catalase concentration and incubated with agitation (130 rpm) at room-temperature. The optical density was measured at 600 nm at 30 min intervals.
83 Chapter 3 The mode of action of AlpP in P. tunicata RESULTS
3.3 Results
3.3.1 AlpP purification
AlpP was purified using dialysis, ion-exchange chromatography and ultra-filtration. The resulting SDS PAGE protein profile after each purification step is shown in Figure 3.1.
After the last purification step (ultra-filtration), two major bands (60 and 80 kDa) which had been previously identified as AlpP (James et al., 1996) were observed (Figure 3.1).
ABCDE
80 kDa 60 kDa
Figure 3.1: SDS PAGE analysis of AlpP purification steps. Molecular weight markers (A), crude supernatant (B), after dialysis (C), after ion-exchange (D), and after ultra-filtration (E)
3.3.1.1 Determination of AlpP concentration and activity
Total protein indicated a yield of 0.13 mg ml-1. The antibacterial activity was confirmed using the drop plate assay. The minimum inhibitory concentration was determined as
84 Chapter 3 The mode of action of AlpP in P. tunicata RESULTS
32.7 ng ml-1 (Figure 3.2).
3.3.1.1.1 AlpP storage conditions
The purified AlpP was stored at 25 °C, 4 °C and -20 °C to investigate whether storage temperature influences activity of the protein. The different storage temperatures did not affect the activity of AlpP after one month of incubation. All samples showed zones of inhibition for the same minimum inhibitory concentration (32.7 ng ml-1) when placed upon a lawn of P. tunicata cells (Figure 3.2).
ABC
65.4 ng ml -1 AlpP
32.7 ng ml-1 AlpP
………..
Figure 3.2: Drop plate assay for purified AlpP. Twenty µl of AlpP, stored at 25 °C (A), 4 °C (B) and -20 °C (C) were placed onto a lawn of P. tunicata cells. Zones of inhibition demonstrate AlpP antibacterial activity.
85 Chapter 3 The mode of action of AlpP in P. tunicata RESULTS
3.3.2 AlpP mode of action investigation
The antibacterial activity of the AlpP homologue, LodA, produced by Marinomonas mediterranea, was recently shown to be due to the generation of hydrogen peroxide from L-lysine (Lucas-Elío et al., 2006). To investigate whether AlpP from P. tunicata also produces hydrogen peroxide from L-lysine the fluorometric Amplex Red assay was used. High fluorescence intensity (390000 counts) was detected after 6 min when AlpP
(0.13 mg ml-1) was incubated with L-lysine and Amplex Red (Figure 3.3). The same fluorescence was measured in the positive control containing Amplex Red and hydrogen peroxide. However, no fluorescence was detected when catalase was added to the reaction of AlpP and L-lysine, to remove the hydrogen peroxide produced (Figure
3.3). The results suggest that AlpP acts as a lysine oxidase and that catalase can abolish the activity of AlpP by removing hydrogen peroxide.
86 Chapter 3 The mode of action of AlpP in P. tunicata RESULTS
500000
400000
300000
200000 Fluorescence units 100000
0 12345678910 Time (min)
Figure 3.3: Amplex Red assay for purified AlpP. The Amplex Red reagent reacts with H2O2 in the presence of peroxidase (HPR) to produce the red fluorescent oxidation product resuforin. High fluorescence (390000 counts) was detected in the presence of the H2O2 standard (12 µM) (Ÿ). A similar fluorescence intensity was detected when AlpP (0.13 mg ml-1) was added to the substrate L-lysine (250 mM) (Ŷ) in the presence of Amplex Red. However, no fluorescence was detected in the presence of catalase (1 -1 -1 mg ml ), AlpP (0.13 mg ml ) and L-lysine (250 mM) () and when no AlpP was added to the reaction (¨).
3.3.2.1 Minimum catalase concentration to abolish AlpP activity
A range of different catalase concentrations (0; 0.01; 0.1; 1; 50; 100; 500 µg and 1 mg ml-1) were mixed with purified AlpP and the activity of the mixture tested by the drop test assay. Catalase concentrations below 50 µg ml-1 did not abolish AlpP activity as the same zone of inhibition was observed (Figure 3.4) for concentrations below that threshold. However, catalase concentration greater than 50 µg ml-1 completely abolished AlpP activity and no zones of inhibition could be observed (Figure 3.4).
87 Chapter 3 The mode of action of AlpP in P. tunicata RESULTS
0 µg 50 µg
0.01 µg 100 µg
0.1 µg 500 µg
1 µg 1 mg
Figure 3.4: Catalase effect on AlpP activity. Ten µl AlpP (0.13 mg ml-1) were mixed with 10 µl catalase at concentrations 0; 0.01; 0.1; 1; 50; 100; 500 µg and 1 mg ml-1 and placed onto a lawn of P. tunicata cells. AlpP activity was abolished at a catalase concentration 50 µg as no zone of inhibition can be observed.
3.3.3 Hydrogen peroxide detection in P. tunicata biofilms
The results described in Chapter 2 showed that AlpP leads to cell death during P. tunicata wild-type biofilm formation. Because of the findings that AlpP mediates cell death through the oxidation of L-lysine and subsequent hydrogen peroxide production, it was speculated that hydrogen peroxide could be detected in biofilms at the onset of killing inside microcolonies. Amplex Red staining was used to visualize hydrogen peroxide in biofilms of P. tunicata. High red fluorescence was observed in P. tunicata wild-type biofilms associated with microcolonies (Figure 3.5 D). Further, the detection of hydrogen peroxide correlated with the onset of cell death at 3 days of biofilm
88 Chapter 3 The mode of action of AlpP in P. tunicata RESULTS development, as no Amplex Red fluorescence could be detected prior to that time
(Figure 3.5 B).
A B
C D
Figure 3.5: Hydrogen peroxide detection in P. tunicata wild-type biofilms. At 24 h of biofilm development small microcolonies could be observed (A, bright field image).
However, no H2O2 was detected at this stage (B, epifluorescent image from the same field of view). After 72 h of biofilm development, large microcolonies had formed (C) and H2O2 can be detected associated with the microcolony (D). Scale bar 50 µm.
3.3.3.1 Hydrogen peroxide detection in ǻAlpP mutant biofilms
To confirm that the hydrogen peroxide detected in P. tunicata wild-type biofilms is due to AlpP activity, biofilms of the ǻAlpP mutant were also stained with Amplex Red at 24 h and 72 h during biofilm formation. No Amplex Red fluorescence was detected at any
89 Chapter 3 The mode of action of AlpP in P. tunicata RESULTS time-point during biofilm development of the ǻAlpP mutant, even when large microcolonies had developed and a very thick biomass covered the substratum (Figure
3.6).
A B
C D
Figure 3.6: Hydrogen peroxide detection in ǻAlpP mutant biofilms. At 24 h biofilm development small microcolonies could be observed (A, bright field image) and no
H2O2 was detected (B, epifluorescent image from the same field of view). However, even at 72 h of biofilm development, when large microcolonies had formed (C), no
H2O2 was detected (D). Scale bar 50 µm.
90 Chapter 3 The mode of action of AlpP in P. tunicata RESULTS
3.3.4 The effect of catalase on microcolony formation in P. tunicata wild-type
biofilms
To investigate the effect of hydrogen peroxide on P. tunicata biofilms, catalase was added to the biofilm media to remove the hydrogen peroxide produced by the biofilm.
Biofilms were allowed to establish for 24 h before adding catalase. At this time small microcolonies had formed and all cells were viable (Figure 3.7 A). In the presence of catalase at 48 h the substratum was completely covered but no microcolonies could be observed (Figure 3.7 B). Furthermore, the addition of catalase affected cell death in P. tunicata wild-type biofilms (Figure 3.7 C). A layer of dead cells was observed covering the substratum, unlike in biofilms without catalase where dead cells were observed throughout the biofilm. The thickness of the layer of dead cells was extended after 72 h of biofilm development. However the top layer of the biofilm always remained completely viable. The data further support the hypothesis that the AlpP mediated killing process through hydrogen peroxide production plays a role in differentiation of the biofilm architecture.
91 Chapter 3 The mode of action of AlpP in P. tunicata RESULTS
A B Substratum
C D Substratum
E F Substratum Catalase added to the biofilm
Figure 3.7: Catalase effect on microcolony formation in P. tunicata wild-type. Biofilms stained with the BacLight LIVE/DEAD Bacterial Viability Kit. Red Propidium Iodide-stained cells have a compromised cell membrane and are dead. The left panel pictures are horizontal scans and the right panel images are vertical scans of the same field of view. Time-points after inoculation are shown as follows: (A, B) 24 hours (before the addition of catalase), (C, D) 48 hours and (E, F) 72 hours, Scale bar 50 µm.
92 Chapter 3 The mode of action of AlpP in P. tunicata RESULTS
3.3.5 Assessment of catalase on growth of P. tunicata
A growth curve experiment was performed to assess whether the addition of catalase affects the growth ability of P. tunicata. No significant difference in growth ability under conditions of different catalase concentrations, compared to the control, could be observed (Figure 3.8).
0.30
0.25
0 0.20 3.4
0.15 catalase 53.6 -1
107.3
0.10 µg ml 429.1 Absorbance 600 nm 0.05
0.00 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 Time (h)
Figure 3.8: Growth curve P. tunicata wild-type in the presence of catalase. Cultures were inoculated into VNSS and the absorbance (600 nm) measured every 30 min. No significant difference in growth ability was observed with the different catalase concentrations.
93 Chapter 3 The mode of action of AlpP in P. tunicata DISCUSSION
3.4 Discussion
The antibacterial and autolytic protein (AlpP) produced by P. tunicata was shown to lead to cell death through the generation of hydrogen peroxide from the amino acid L- lysine. Catalase can remove hydrogen peroxide and thus counteracts the activity of
AlpP. The minimum catalase concentration necessary to abolish the activity of AlpP was determined to be 50 µg ml-1. As shown in Chapter 2, AlpP leads to cell death, differentiation and dispersal during biofilm development. Here it is observed that hydrogen peroxide can be detected in P. tunicata wild-type biofilms at the onset of cell death and hydrogen peroxide is associated with microcolonies. Further, it was found that the addition of catalase to biofilm media abolishes microcolony formation, presumably through the removal of hydrogen peroxide.
AlpP was successfully purified to an 80 and a 60 kDa band previously identified as the two subunits of AlpP (Figure 3.1) (James et al., 1996). The resulting purified fraction had a minimum inhibitory activity against P. tunicata of 32.7 ng ml-1 AlpP. Further, purified AlpP was found to be very stable and remained active independent of the storage temperature (Figure 3.2).
Several members of the genus Pseudoalteromonas have been found to produce antibacterial and autotoxic compounds, including Pseudoalteromonas luteoviolacea
(previously Alteromonas luteoviolacea CH130) (Gauthier and Flatau, 1976; McCarthy et al., 1994), Pseudoalteromonas phenolica (Isnansetyo and Kamei, 2003) and
Pseudoalteromonas sp. F-420 (Yoshikawa et al., 2003). However, the mode of action of these compounds has not been fully established. The strain P. luteoviolacea was found to produce a macromolecular antibiotic which was able to modify aerobic respiration in the target bacterial strain. Furthermore, the activity of the antibiotic produced by P.
94 Chapter 3 The mode of action of AlpP in P. tunicata DISCUSSION luteoviolacea was inhibited by the addition of catalase (Gauthier and Flatau, 1976), suggesting that the mode of action of this compound could be similar to the lysine oxidase activity of AlpP produced by P. tunicata.
L-lysine oxidases were first isolated from the fungus Trichoderma sp. and catalyse the following reaction:
L-lysine + O2 + H2O Æ 6-amino-2-oxo-hexanoate + NH3 + H2O2
Lysine oxidases are of particular interest because of their unique properties, including cytotoxic, antitumor, antimetastatic, antiinvasive, antiviral and antibacterial activities
(Lukasheva and Berezov, 2002). These characteristics are suggested to be due to a decrease in the essential amino acid L-lysine in target cells as well the production of damaging hydrogen peroxide. The antibacterial activity of lysine oxidases was first demonstrated using a rec- strain of Bacillus subtilis as a target strain. The rec- strain was more sensitive to the antibacterial effect of the lysine oxidase than the B. subtilis wild- type (Kusakabe et al., 1979), suggesting that the damaging effect of hydrogen peroxide is responsible for the antibacterial activity, rather than the decreasing L-lysine concentration. Further evidence for this correlation derived from the ability of catalase to protect the cells by removal of hydrogen peroxide (Kusakabe et al., 1979).
The AlpP homologue produced by M. mediterranea was the first L-lysine oxidase to be described in bacteria (Lucas-Elío et al., 2006). Because of its homology to AlpP it was speculated that both proteins share similar characteristics. Indeed, the Amplex Red assay showed that AlpP can also generate hydrogen peroxide from the amino acid L- lysine. It is likely that the antibacterial activity of AlpP in P. tunicata is due to the hydrogen peroxide generated, rather than the decrease of the essential amino acid L- lysine, because the addition of catalase completely abolishes AlpP activity by removing 95 Chapter 3 The mode of action of AlpP in P. tunicata DISCUSSION hydrogen peroxide (Figure 3.3 and 3.4).
The addition of catalase to the biofilm media had a significant effect upon the biofilm structure as the formation of microcolonies was prevented (Figure 3.7). Because catalase removes hydrogen peroxide produced by AlpP, it could be expected that cell death would not occur in P. tunicata wild-type biofilms after the addition of catalase and its biofilm structure would be similar to ǻAlpP mutant biofilms were no AlpP is produced. However, a layer of dead cells covering the substratum was still observed after the addition of catalase to wild-type biofilms (Figure 3.7 D, E). Catalase is a large macromolecule (diameter ~ 10.5 nm, 248 kDa) and it is feasible that penetration to the bottom of the biofilm through a thick layer of cells as well as the biofilm matrix would be hindered. Furthermore, it remains to be established whether the observed effect of catalase on the biofilm structure is directly mediated through catalase or indirectly through the ability of catalase to remove the hydrogen peroxide produced by AlpP. The fact that the ǻAlpP mutant is still capable of forming microcolonies suggests a direct involvement of catalase. However, a more complex effect involving AlpP is also possible. The effect of catalase on biofilms of other organisms is described in Chapter 5.
Following the elucidation of the mode of action of AlpP-mediated cell death, experiments were designed to address a possible role of cell death in P. tunicata biofilms as reported in Chapter 4.
96 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata INTRODUCTION
4. Ecological advantages of autolysis during biofilm development and dispersal of P. tunicata
4.1 Introduction
Models for biofilm formation include that of a developmental lifecycle (De Kievit et al.,
2001; Sauer et al., 2002; Southey-Pillig et al., 2005), or of biofilm behaviours resulting simply from the cumulative effect of single cells responding to changes within their environment (Kjelleberg and Molin, 2002). Biofilm formation has been proposed as a developmental process (Davies and Geesey, 1995; Davey and O'Toole, 2000; Sauer et al., 2002) because several studies indicate that it is a regulated process where large- scale temporal and spatial changes in gene expression occur during different stages.
Different gene expression as a response to biofilm formation has been demonstrated in model biofilm forming bacteria, including Pseudomonas aeruginosa (Southey-Pillig et al., 2005), Escherichia coli (Prigent-Combaret et al., 1999; Schembri et al., 2003) and
B. subtilis (Stanley et al., 2003). On the other hand it has been hypothesized that environmental factors (e.g. hydrodynamics or nutrient availability) and conditions simply resulting from the biofilm lifestyle (e.g. intracellular carbon flux or oxidative stress) have a major impact on biofilm formation (Kjelleberg and Molin, 2002). These factors may lead to the alteration of expression of cellular traits necessary for the adaptation to the biofilm lifestyle (Kjelleberg and Molin, 2002).
Biofilm dispersal and phenotypic variation are two behavioural aspects of biofilm cells which may represent an adaptive response or are part of a regulated, developmental process. These two aspects have been a focus of attention because of their ability to greatly enhance survival and spread of bacteria within the environment. Biofilms are known to generate a differentiated population of phenotypic and genotypic variants of
97 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata INTRODUCTION bacteria that enhance survival in the face of changing environmental conditions (Boles et al., 2004; Haussler, 2004; Kirisits et al., 2005). This has been shown for several organisms, including Staphylococcus aureus (Sadowska et al., 2002), Vibrio cholerae
(Ali et al., 2002), Listeria monocytogenes (Monk et al., 2004) and P. aeruginosa
(Oliver et al., 2000; Deziel et al., 2001; Webb et al., 2004). Diverse mechanisms have been investigated that may lead to an increased genetic and phenotypic variation in biofilms. These mechanisms include phase variation (Drenkard and Ausubel, 2002), adaptive mutation (Oliver et al., 2000; Bjedov et al., 2003) and enhanced gene transfer through conjugation, transformation (Hausner and Wuertz, 1999; Hendrickx et al.,
2003; Molin and Tolker-Nielsen, 2003) and phage transduction (Webb et al., 2004). In summary, high levels of genetic diversity are common among cells that disperse from biofilms and may benefit bacteria in their ability to survive and colonize diverse new environments.
In addition, cells within biofilms can at times disperse in order to colonize new surfaces
(Boyd and Chakrabarty, 1994; Kaplan et al., 2003a; Fux et al., 2004; Sauer et al.,
2004). Active processes are often used by bacteria to promote dispersal, for example enzyme-mediated breakdown of the biofilm matrix (Boyd and Chakrabarty, 1994; Lee et al., 1996; Kaplan et al., 2003a; Kaplan et al., 2003b), or the production of surfactants which loosen cells from the biofilm (Davey et al., 2003). Dispersal processes may also be under the control of cell-cell communication systems that facilitate coordinated group behaviour of bacteria in biofilms (Rice et al., 2005). Biofilm development is thus a dynamic process, in which phenotypic variation and dispersal are key factors that influence biofilm function within the environment.
Another important feature of biofilm formation is the occurrence of cell death of subpopulations within microcolonies. This has been recently demonstrated for several 98 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata INTRODUCTION organisms, such as P. aeruginosa (Webb et al., 2003b) and P. tunicata (Mai-Prochnow et al., 2004) (Chapter 2). Notably, similar cell death events were observed during developmental processes of differentiating organisms, including fruiting body formation of Myxococcus xanthus (Wireman and Dworkin, 1977; Rosenbluh and Rosenberg,
1990), sporulation of Bacillus subtilis (Gonzalez-Pastor et al., 2003) and mycelium differentiation of Streptomyces sp. (Mendez et al., 1985; Fernandez and Sanchez, 2002).
In M. xanthus death of a cell subpopulation is necessary to complete the developmental process of fruiting body formation (Wireman and Dworkin, 1977). Surviving cells of B. subtilis use nutrients released from their lysed sister cells (Gonzalez-Pastor et al., 2003).
Similarly, in Streptomyces sp. dead mycelium cells serve as a nutrient source for perpendicular hyphae cells (Mendez et al., 1985; Fernandez and Sanchez, 2002). In summary, cell death of cell subpopulations is proposed to be beneficial to the population as a whole and important for differentiation processes in these organisms. It could be hypothesized that cell death events during bacterial biofilm development have similar implications and may be advantageous to the bacterial community.
Results in this chapter provide evidence that AlpP-mediated cell death confers ecological advantages to P. tunicata by ensuring a metabolically active and phenotypically diverse dispersal population. Furthermore, it is shown that cell death of a subpopulation can increase the growth activity of surviving cells during nutrient starvation.
99 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata MATERIALS AND METHODS
4.2 Materials and Methods
4.2.1 Bacterial strains and culture media
P. tunicata was routinely cultivated at room temperature in Väätänen nine salt solution
(VNSS) (Marden et al., 1985) supplemented with streptomycin (100 µg ml-1) and kanamycin (50 µg ml-1) for the P. tunicata ǻAlpP mutant (Mai-Prochnow et al., 2004).
Biofilms were grown in marine minimal medium (MMM) (Neidhardt et al., 1974) containing 0.01 % trehalose. For starvation experiments the MMM base nine salt solution (NSS) was used without a carbon source.
4.2.2 Biofilm experiments
P. tunicata wild-type and ǻAlpP mutant strains were grown in continuous-culture flow- cells (channel dimensions 1 x 4 x 40 mm) at room temperature as described in Chapter 2
(see Section 2.2.2). Briefly, channels were inoculated with 0.5 ml of early stationary phase cultures containing approximately 1 x 109 cells ml-1 and incubated without flow for 1 h at room temperature. Flow was then started with a mean flow velocity in the flow cells of 0.2 mm s-1, corresponding to laminar flow with a Reynolds number of
0.02.
4.2.3 Phenotypic characterization of biofilm dispersal cells
To characterize biofilm dispersal, the number of viable bacteria in the effluent of both
P. tunicata wild-type and ǻAlpP mutant biofilms was determined by serial plate counts on VNSS over 192 h of biofilm formation. To investigate the hypothesis that cell lysis within microcolonies promotes phenotypic variation in biofilms, effluent was spread plated onto VNSS, LB10, Marine Agar (Difco, Becton Dickenson, USA) and Davis
100 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata MATERIALS AND METHODS
Minimal Medium (Difco, Becton Dickenson, USA) in order to detect colony morphology variation. In addition, 20 colonies derived from wild-type and ǻAlpP mutant biofilms were randomly picked from VNSS agar and screened for growth, biofilm formation and motility. Three time-points were chosen: 24 h (before the onset of cell death), 72 h (shortly after the onset of cell death) and 144 h (when cell death had extended throughout the biofilm). Overnight cultures (15 µl) of the 20 colonies were inoculated into 1.5 ml fresh VNSS in 24-well tissue culture plates. Plates were incubated at room temperature with agitation (130 rpm). OD 600 nm was measured after 24 h as an indicator for growth ability. To measure biofilm forming ability wells of the tissue culture plates were stained with crystal violet for 20 min. After washing the wells twice with NSS, crystal violet was extracted in 95 % ethanol and the absorbance read at 600 nm. As an indicator for swimming-motility, the variants were stab- inoculated into 0.4 % VNSS agar and the growth radius measured after 24 h.
4.2.4 Phenotypic variation of batch culture cells
To provide evidence that the measured variation is a biofilm specific trait the wild-type and ǻAlpP mutant strains were inoculated into 500 ml planktonic cultures in MMM with agitation (130 rpm) and also screened for variation in motility, growth and biofilm formation. Aliquots were taken at 24 h, 72 h and 144 h and spread onto VNSS. Ten colonies were randomly picked and subjected to the same characterization process as described above (see Section 4.2.3).
4.2.5 Calculating the variation coefficient
To determine the relative variation among the wild-type and ǻAlpP mutant strains for each time-point, a statistical coefficient of variation (CV) was calculated. The coefficient of variation is a dimensionless number that allows comparison of the 101 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata MATERIALS AND METHODS variation of populations that have significantly different mean values. It is defined as a
V ratio of the standard deviation (ı) to the mean (µ): Cv P
CV is reported here on a scale of 0 to 100% by multiplying the above calculation by
100 %.
A variance analysis (F test) was performed to determine whether a difference in variation is statistically significant. The F test can be used to determine whether variances of two samples are different. For a sample size of 20 and a 95 % confidence level the F value should be >2.1 to reject the hypothesis that the two variances are equal.
4.2.6 Add back experiments
Purified AlpP was added to the 144 h ǻAlpP mutant biofilms to determine whether
AlpP mediated killing induces dispersal and increase phenotypic variation among dispersal cells. AlpP was prepared as described in Chapter 3 (see Section 3.2.2) and the add back procedure was described in Chapter 2 (see Section 2.2.5.6). Biofilms were incubated at room temperature for 5 h before turning on the flow and collecting effluent at 30 min intervals. Effluent of both control and AlpP add back channels were spread onto VNSS plates in serial dilutions to quantify biofilm dispersal of viable cells.
Phenotypic variation was measured in motility, growth and biofilm formation as described above (see Section 4.2.3).
102 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata MATERIALS AND METHODS
4.2.7 Analysing metabolic activity of biofilm effluent
4.2.7.1 Fluorescent staining of biofilm effluent
The membrane potential probe bis-(1,3-dibutylbarbituric acid) trimethine oxonol
(DiBAC4(3)) (Molecular Probes, Inc., Oregon, USA) was used to investigate whether cell death influences the metabolic activity of dispersal cells. DiBAC4(3) is a green fluorescent anionic dye and commonly used in flow cytometry studies as an indicator of membrane potential. It enters the cell as a result of membrane depolarization and binds to intracellular proteins or membranes exhibiting green fluorescence. Increased depolarization results in a greater influx of the anionic dye and thus results in an increased green fluorescence. Viable, active cells with a charged membrane largely exclude the dye and remain less fluorescent (Brauner et al., 1984). Effluent of both wild-type and ǻAlpP mutant biofilms at 72 h (shortly after the onset of cell death) and
144 h (with extensive cell death throughout the wild-type biofilm) were stained according to the manufacturer’s description. As controls, P. tunicata cells in logarithmic growth phase were stained. One half of the control cells were not treated (viable, active cells) and the other half heat killed for 10 min at 70 °C (non-viable, dead cells) before staining.
4.2.7.2 Flow cytometry
Flow cytometric analysis was performed using a BD FACSCalibur-sort flow cytometer
(BD Biosciences, Sydney, Australia) equipped with an air-cooled 100 mW argon ion laser (488 nm) for excitation. The positive and negative controls were analysed first and the threshold was set to just below the population of bacteria on a bivariate dot-plot of side scatter (SSC) versus forward scatter (FSC). A gate was defined around this control population and the fluorescence monitored on a histogram of green florescence (FL1). 103 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata MATERIALS AND METHODS
All samples were run in triplicate with 10,000 cells analysed on a histogram of green fluorescence. Data analysis was carried out using Summit offline software (Cytomation
Inc). Positive and negative controls were analysed first to ensure both populations were separated on the FL1 histogram (green fluorescence detector). For all samples a region was defined around the main bacterial population in the bivariate dot-plot of SSC versus
FSC and only cells within this region were analysed in the univariate FL1 histogram.
Regions were then defined around active (R1) and depolarized (R2) cells on the univariate FL1 histogram. The percentage of cells in each region was recorded and the mean value and standard deviation calculated from triplicate samples. An unpaired, two-sided t-Test was performed to test the hypothesis that the analysed samples differed significantly.
4.2.8 Starvation experiments
Biofilms were allowed to pre-establish in the flow system (see Chapter 2, Section 2.2.2) for 72 h in MMM, before changing to nutrient starvation media (NSS). They were stained with the LIVE/DEAD BacLight Bacterial Viability Kit (Molecular Probes Inc.,
Eugene, Oreg.). The two stock solutions of the stain (SYTO 9 and propidium iodide
(PI)) were diluted to 3 µl ml-1 in biofilm medium and injected into the flow channels.
Actively respiring bacteria were localized within the biofilm by the reduction of 5- cyano-2, 3-ditolyl tetrazolium chloride (CTC) (Polyscience, Inc., Warrington, Pa.). CTC was diluted to a final concentration of 5 mmol before injection into the flow channels.
The resulting fluorescence was visualized with a confocal laser scanning microscope
(CLSM) (Olympus) equipped with fluorescein isothiocyanate and tetramethyl rhodamine isocyanate optical filters, respectively.
104 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
4.3 Results
4.3.1 Quantification of biofilm dispersal cells
Death and lysis of a subpopulation of cells was shown to occur in the centre of microcolonies after 48 h of P. tunicata biofilm development leading to the formation of hollow microcolonies (Mai-Prochnow et al., 2004) (Chapter 2). Here it was found that this process also plays a role in P. tunicata biofilm dispersal. Numbers of viable dispersal cells in the biofilm effluent from both P. tunicata wild-type and ǻAlpP mutant strains were quantified and statistically compared over a period of 192 h (8 days) of biofilm development. Both strains showed little dispersal for the first 48 h of biofilm development with 1.51 x 106 wild-type colony forming units (CFU ml-1) and 1.86 x 106
ǻAlpP mutant CFU ml-1 in the biofilm effluent (Figure 4.1). The P. tunicata wild-type strain showed a steady increase in viable cells dispersing from the biofilm over the time- course of the experiment; there was a major detachment event at 192 h where most of the biofilm had been removed from the substratum. This detachment event correlates with extensive AlpP mediated killing within the biofilm. At this time-point the effluent contained as much as 4.43 x 107 CFU ml-1. In contrast, the ǻAlpP mutant did not show a major biofilm detachment event and dispersal remained low over the time-course of the experiment, with only 2.28 x 106 CFU ml-1 dispersing at 192 h (Figure 4.1). Several replicate experiments over longer periods of time (up to 240 h) failed to show a large dispersal event in the ǻAlpP mutant strain.
105 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
50 45 40 35 6 30 25 20 CFU x 10 15 10 5 0 5 27 48 54 73 98 120 144 168 192 Biofilm age (h)
Figure 4.1: Dispersal of viable cells from P. tunicata wild-type and ǻAlpP mutant biofilms. Biofilm dispersal is shown in effluent viable counts (CFU). P. tunicata wild- type (Ŷ) shows a significant sloughing event at 192 h. No major increase in dispersal can be detected in the ǻAlpP mutant (Ÿ).Error bars represent the standard deviation of three independent experiments.
4.3.1.1 AlpP add back to induce dispersal of ǻAlpP mutant biofilms
To show that AlpP mediated cell death is linked to the dispersal event observed in the P. tunicata wild-type, purified AlpP was added to ǻAlpP mutant biofilms. It was previously shown that addition of AlpP to ǻAlpP mutant biofilms induces killing within the biofilm (Chapter 2). After the AlpP add back, the ǻAlpP mutant showed a 10 fold increase in the number of dispersal cells compared to the Tris buffer control (Figure
4.2). The reduction in CFU subsequent to a major dispersal event is due to the complete loss of the biofilm similar to what occurs after the main dispersal event of the wild-type biofilm past 192 h (Figure 4.1). The nature of the add back experiment, i.e. stopping the flow for 5 h led to a considerably higher number of CFU in the effluent of both control
106 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS and add back channels compared to the experiment shown in Figure 4.1. Three independent experiments revealed similar results.
3500
3000
2500
6 2000
1500 Cfu x 10
1000
500
0 5 h 5.5 h 6 h 6.5 h 7 h 18 h Time after AlpP add back
Figure 4.2: Add back of purified AlpP (Ŷ) and Tris (20 mmol) buffer control (Ÿ). Dispersal is shown in effluent viable counts 5 to 18 h after inoculating AlpP and Tris, respectively. Error bars represent the standard deviation of three independent experiments.
4.3.2 Metabolic activity of biofilm dispersal cells
The metabolic activity of the dispersal population of both P. tunicata wild-type and
ǻAlpP mutant strains was examined. Biofilm effluent was stained at two different time- points during biofilm formation (72 h and 144 h) using the fluorescent dye DiBAC4(3) in conjunction with flow-cytometry. DiBAC4(3) exhibits enhanced fluorescence when it
107 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS enters cells with depolarized membranes. At 72 h, dispersal cells of both wild-type and the ǻAlpP mutant showed two distinct cell populations (Figure 4.3 A, B). The sub- population with lower fluorescence corresponded to viable, active cells with polarized membranes, and the sub-population with higher fluorescence corresponds to cells with reduced activity (depolarized membranes). At 72 h in both strains a large proportion of dispersal cells were active and a smaller fraction of the population had depolarized membranes (Figure 4.3 A, B). At 144 h the ǻAlpP mutant dispersal population consisted mainly of cells with depolarized membranes and only a very small population of active cells could be detected (Figure 4.3 D). In contrast, at 144 h of biofilm development when large regions of lysis occurred, wild-type dispersal cells still demonstrated two distinct fluorescent peaks, showing a large active cell population as well as a population of cells with depolarized membranes (Figure 4.3 C).
A t-Test was performed to establish whether the variability between subpopulation phenotypes observed at 72 h and 144 h was significant. The results demonstrated that the difference between the wild-type and ǻAlpP mutant active subpopulations was highly significant (P = 4.9 x 10-5). Similarly, the difference of the depolarized subpopulations between both time-points also revealed a highly significant P value of
5.2 x 10-5. It confirmed that the subpopulations observed at 144 h were significantly different from those observed at the 72 h time-point.
108 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
Figure 4.3: Flow cytometry analysis of biofilm effluent. Cells were stained with the green membrane potential probe DiBAC4(3) at 72 h (wild-type (A) and ǻAlpP mutant (B)) and at 144 h (wild-type (C) and ǻAlpP mutant (D)) of biofilm development. The histograms show green fluorescence (FL1-H in relative fluorescence units) with R1 corresponding to active cells and R2 corresponding to cells with depolarized membranes. Mean counts and standard deviations for each region as a percentage of total event counts were calculated from triplicate samples.
4.3.3 Phenotypic variation of biofilm dispersal cells
Many organisms show enhanced phenotypic variation during biofilm growth (Oliver et al., 2000; Deziel et al., 2001; Ali et al., 2002; Sadowska et al., 2002; Monk et al., 2004;
Webb et al., 2004). The occurrence of morphological colony variants during biofilm formation was investigated by plating biofilm effluent onto several different media types over 10 days of biofilm formation. Unexpectedly, no biofilm specific morphological colony variants could be detected in the wild-type or the ǻAlpP mutant as all colonies had normal wild-type appearance at all times investigated (Figure 4.4).
However, it was also investigated whether variation among dispersal cells could occur 109 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS in phenotypic traits other than colony morphology; in particular traits that are relevant for colonization of new surfaces. Therefore 20 randomly picked colonies from VNSS agar plates, derived from biofilms of both strains were screened for their growth ability, biofilm formation and motility. It was observed that colonies from the wild-type showed a very high variation in their biofilm forming ability (Figure 4.5) and growth ability
(Figure 4.6). The highest variation was detected at the latest time-point of biofilm development (144 h) when biofilm killing was well established (as determined in
Chapter 2). At this time-point, wild-type phenotypic variation ranked as high as 62 % for biofilm formation (Figure 4.5 C) and 57 % for growth (Figure 4.6 C). Colonies derived from ǻAlpP mutant biofilms did not differ greatly in all three investigated traits and variation at 144 h was only 21.02 % for biofilm formation (Figure 4.5 B, C) and
7.56 % for growth (Figure 4.6 B, C). Variation in motility of biofilm dispersal cells was comparably low for both strains (Figure 4.7). However, a small number (0.03 %) of wild-type colonies showed an interesting phenotype with significantly decreased biofilm formation as well as increased motility, as is likely to be beneficial for dispersal away from the mother colony in times of poor resources. Despite repeat experiments, this phenotype was not detected in the ǻAlpP mutant colonies.
An F test analysis showed that differences in variance between wild-type and ǻAlpP mutant were significant, i.e., with an F value >2.1 for biofilm formation at 144 h (F =
14.2), for growth at 24 h (F = 11.4) and at 144 h (F = 46.0) and for motility at 24 h (F =
12.4), 72 h (F = 4.9) and 144 h (F = 9.3).
110 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
Figure 4.4: Biofilm effluent of a 7-day old P. tunicata wild-type biofilm spread onto VNSS agar. No variation in colony morphology could be detected
111 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
A 0.4 Biofilm formation of wild-type biofilm dispersal cells AlpP
24 h 72 h 144 h ǻ Biofilm 0.3 ) and A
0.2
0.1 Absorbance 600 nm
0 wild-type biofilms ( 1 3 5 7 9 11 13 15 17 19 21 23 25 27 29 31 33 35 37 39 41 43 45 47 49 51 53 55 57 59 Variant number mut coefficient for each strain and time-point is coefficient for each strain and time-point
B Biofilm formation of ǻAlpP mutant biofilm dispersal cells P. tunicata 2.0 24 h 72 h 144 h 144 h AlpP ant dispersal cells. biofilm 1.8 ǻ 1.6 add back 1.4 1.2 1.0 0.8
0.6 wild-type and 0.4 Absorbance 600 nm 0.2 0.0 M1 M3 M5 M7 M9 P. tunicata M11 M13 M15 M17 M19 M21 M23 M25 M27 M29 M31 M33 M35 M37 M39 M41 M43 M45 M47 M49 M51 M53 M55 M57 M59 M61a M63a M65a M67a M69a M71a M73a M75a M77a M79a Variant number
C Biofilm formation variation comparison 100 ) 24 h 72 h 144 h
% 90 ( 80 70 e e e
60 p yp mutant yp y t P mutant P mutant 50 P mutant p p p ) at 3 time-points during biofilm formation. The calculated variation The calculated formation. ) at 3 time-points during biofilm B Al AlpP Al
40 Al Wild-t ǻ Wild-t add back ǻ ǻ ǻ 30 Wild- ). C
Variation coefficient 20 10 0 shown in ( mutant biofilms ( Figure 4.5: Variation in biofilm formation of formation quantified with crystal violet staining for individual colonies derived from with crystal violet formation quantified
112 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
Growth of wild-type biofilm dispersal cells A 2 1.8 24 h 72 h 144 h 1.6
1.4 ) at 3 time- B 1.2 1 ).
0.8 C 0.6
Absorbance 600 nm 0.4 Growth ability quantified by quantified Growth ability 0.2 0 1 3 5 7 9 11 13 15 17 19 21 23 25 27 29 31 33 35 37 39 41 43 45 47 49 51 53 55 57 59 AlpP mutant biofilms ( Variant number ǻ ) and B Growth of ǻAlpP mutant biofilm dispersal cells A 1 24 h 72 h 144 h 144 h add back 0.8
0.6
0.4 wild-type biofilms ( AlpP mutant biofilm dispersal cells. AlpP mutant biofilm ent for each strain and time-point is shown in ( time-point ent for each strain and ǻ 0.2 Absorbance 600 nm
0 P. tunicata M1 M3 M5 M7 M9 M11 M13 M15 M17 M19 M21 M23 M25 M27 M29 M31 M33 M35 M37 M39 M41 M43 M45 M47 M49 M51 M53 M55 M57 M59 M61a M63a M65a M67a M69a M71a M73a M75a M77a M79a Variant number wild-type and
C Growth variation comparison 100 P. tunicata ) 90 24 h 72 h 144 h % ( 80 70
60 e e utant e yp yp
50 yp P mutant P mutant P mutant p p 40 p AlpP m Al Al Al Wild-t Wild-t ǻ ǻ add back ǻ ǻ 30 Wild-t
Variation coefficient 20 10 0 Figure 4.6: Variation in growth of OD measurements for individual colonies derived from points during biofilm formation. The calculated variation coeffici variation points during biofilm formation. The calculated
113 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
A Motility of wild-type biofilm dispersal cells 2.0 1.8 24 h 72 h 144 h 1.6
) 1.4 1.2 mm ( 1.0 Motility radius for 0.8
Radius 0.6 0.4 0.2
0.0 ) at 3 time-points during biofilm B 1 3 5 7 9 11 13 15 17 19 21 23 25 27 29 31 33 35 37 39 41 43 45 47 49 51 53 55 57 59 Variant number . ) C ( B Motility of ǻAlpP mutant biofilm dispersal cells 2 1.8 24 h 72 h 144 h 144 h 1.6 add back ) 1.4 AlpP mutant biofilms ( ǻ oint is shown in
mm 1.2 ( AlpP mutant biofilm dispersal cells. AlpP mutant biofilm p
1 ǻ
0.8 ) and A
Radius 0.6 0.4 0.2 0 wild-type and M1 M3 M5 M7 M9 M11 M13 M15 M17 M19 M21 M23 M25 M27 M29 M31 M33 M35 M37 M39 M41 M43 M45 M47 M49 M51 M53 M55 M57 M59 M61a M63a M65a M67a M69a M71a M73a M75a M77a M79a Variant number wild-type biofilms (
C Motility variation comparison P. tunicata 100
) 90 24 h 72 h 144 h P. tunicata %
( 80 70 60 e e e
50 yp yp yp P mutant P mutant P mutant
40 p p p Al Al Al AlpP mutant ǻ ǻ ǻ Wild-t ǻ add back 30 Wild-t Wild-t 20 Variation coefficient 10 0 formation. The calculated variation coefficient for each strain and time- each strain for variation coefficient calculated formation. The Figure 4.7: Variation in motility of individual colonies derived from
114 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
4.3.4 AlpP add back to induce phenotypic variation in ǻAlpP mutant dispersal
cells
In order to show that the generation of phenotypic variation among dispersal cells is linked to AlpP-mediated cell death events, purified AlpP was added back to ǻAlpP mutant biofilms. Add back of purified AlpP was shown to induce cell death in ǻAlpP mutant biofilms (Chapter 2) (Mai-Prochnow et al., 2004) as well as dispersal (see
4.3.1.1). Here it was observed that add back of AlpP also leads to a significant increase in variation among ǻAlpP mutant dispersal cells. Variation in growth increased from
7.6 to 31 % (Figure 4.6 C), variation in biofilm formation from 21 to 49.6 % (Figure 4.5
C) and variation in motility from 8.2 to 23.9 % (Figure 4.7 C) after AlpP add back.
4.3.5 Phenotypic variation in batch culture
To investigate whether the high variation among wild-type dispersal cells occurs specifically in biofilms (and not planktonic cells) wild-type and ǻAlpP mutant colonies deriving from non-biofilm, batch cultures were also tested for variation in the same traits. Variation in wild-type planktonic cells was found to be significantly lower than in wild-type biofilm cells and likewise variation in the ǻAlpP mutant planktonic cells was lower than ǻAlpP mutant biofilm cells in motility (Figure 4.8), biofilm formation
(Figure 4.9) as well as growth (Figure 4.10). In contrast to biofilm dispersal cells where variation was highest at the latest time-point (144 h), batch culture cells of the wild-type showed highest variation at the early time-point (24 h after inoculation) for all three traits. Cells derived from ǻAlpP mutant batch culture showed very small variation
(below 7 %) in motility (Figure 4.8) as well as growth (Figure 4.10) and biofilm formation (Figure 4.9). These data suggest that the phenotypic variation in motility, growth and biofilm formation are biofilm specific properties.
115 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
Motility of wild-type batch culture cells A 1.2 24 h 72 h 144 h 1
) 0.8 mm ( 0.6
0.4 Radius 0.2 Motility radius for individual for Motility radius
0 The calculated ) at 3 time-points. 1 2 3 4 5 6 7 8 9 B 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Variant number
Motility of ǻAlpP mutant batch culture cells B 1.2 24 h 72 h 144 h 1 ) 0.8 mm ( 0.6 ( cultures batch AlpP mutant AlpP mutant batch culture cells. ǻ ǻ
0.4 ). C Radius ) and
0.2 A
0 M1 M2 M3 M4 M5 M6 M7 M8 M9 wild-type and M10 M11 M12 M13 M14 M15 M16 M17 M18 M19 M20 M21 M22 M23 M24 M25 M26 M27 M28 M29 M30 Variant number
C
Motility variation comparison P. tunicata 100 )
90 24 h 72 h 144 h ( batch cultures wild-type % ( 80 70 e e 60 e P. tunicata P. yp yp 50 yp P mutant P mutant P mutant p p 40 p Al Al Al Wild-t Wild-t ǻ Wild-t ǻ 30 ǻ 20 Variation coefficient 10 0 Figure 4.8: Variation in motility of variation coefficient for each strain and time-point is shown in ( for variation coefficient colonies derived from
116 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
A Biofilm formation of wild-type batch culture cells 0.5 24 h 72 h 144 h 0.4
0.3 Biofilm formation AlpP mutant batch ǻ 0.2 ). C ) and
0.1 A Absorbance 600 nm 0.0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Variant number batch cultures ( Biofilm formation of ǻAlpP mutant batch culture cells B 0.5 24 h 72 h 144 h
0.4 culture cells. AlpP mutant batch ǻ P. tunicata
0.3
0.2
0.1 wild-type and
Absorbance 600 nm 0.0 M1 M2 M3 M4 M5 M6 M7 M8 M9 M10 M11 M12 M13 M14 M15 M16 M17 M18 M19 M20 M21 M22 M23 M24 M25 M26 M27 M28 M29 M30 Variant number P. tunicata
C Biofilm formation variation comparison 100 ) 90 24 h 72 h 144 h % ( 80 70 e e 60 e yp yp yp P mutant P mutant 50 P mutant p p p 40 Al Al Al Wild-t ǻ Wild-t Wild-t ǻ 30 ǻ ) at 3 time-points. The calculated variation coefficient for each strain and time-point is shown in ( variation ) at 3 time-points. The calculated
Variation coefficient 20 B 10 0 Figure 4.9: Variation in biofilm formation of cultures ( cultures quantified with crystal violet staining for individual colonies derived from
117 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
A Growth ability of wild-type batch culture cells 1.2 24 h 72 h 144 h 1.0
0.8
0.6
0.4 Absorbance 600 nm 0.2 batch cultures AlpP mutant Growth ability quantified by Growth ability quantified ǻ ).
0.0 C 1 2 3 4 5 6 7 8 9 ) and 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 Variant number A
B Growth ability of ǻAlpP mutant batch culture cells 1.2 24 h 72 h 144 h 1.0
0.8
0.6 wild-type batch cultures ( wild-type AlpP mutant batch culture cells.
0.4 ǻ
0.2 Absorbance 600 nm
0.0 P. tunicata M1 M2 M3 M4 M5 M6 M7 M8 M9 M10 M11 M12 M13 M14 M15 M16 M17 M18 M19 M20 M21 M22 M23 M24 M25 M26 M27 M28 M29 M30 Variant number wild-type and
C Growth ability variation comparison
100 P. tunicata
) 90 24 h 72 h 144 h % ( 80 70 e e 60 e yp yp 50 yp P mutant P mutant P mutant 40 p p p Al Al Al Wild-t Wild-t ǻ ǻ ǻ 30 Wild-t
Variation coefficient 20 10 0 ) at 3 time-points. The calculated variation coefficient for each strain and time-point is shown in ( for each strain coefficient variation calculated ) at 3 time-points. The B Figure 4.10: Variation in growth of OD measurements for individual colonies derived from (
118 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
4.3.6 Influence of nutrient starvation on metabolic activity within biofilms
In differentiating bacteria, one effect of programmed cell lysis is that surviving bacteria can utilize nutrients released from dead cells in order to facilitate continued development under starvation conditions (Mendez et al., 1985; Fernandez and Sanchez,
2002; Gonzalez-Pastor et al., 2003). As a first step towards understanding whether similar processes may occur in bacterial biofilm development, the role of biofilm cell death on the activity and viability of remaining cells within the biofilm under starvation conditions was examined. As previously observed (Chapter 2) (Mai-Prochnow et al.,
2004), 72 h P. tunicata wild-type biofilms under non-starvation conditions showed dead cells localized in the centre of microcolonies (Figure 4.11 B). At this time-point, CTC staining of wild-type biofilms revealed that metabolic activity was highest at the outside of microcolonies, with low activity in the microcolony centre (Figure 4.11 A). However, after switching to nutrient starvation conditions, a reduction in the amount of PI stained material was observed (Figure 4.11 D) and the highest CTC fluorescence was then detected in the centre of microcolonies (Figure 1.1 C), suggesting a shift of metabolic activity to the microcolony centre. The ǻAlpP mutant did not show cell death during any stage of biofilm development (Chapter 2) (Mai-Prochnow et al., 2004). Despite the fact that all cells are viable during non-starvation conditions, their metabolic activity was generally lower than that of wild-type biofilms (Figure 4.11 E). Metabolic activity was evenly distributed within the ǻAlpP mutant biofilm and no specific pattern of localized activity was noted. Upon starvation, an overall loss of biomass was observed with a further decrease in metabolic activity (Figure 4.11 G, H).
119 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
Metabolic activity staining (CTC) Live/Dead staining A Wild-type B Wild-type
C Wild-type starved D Wild-type starved
EFǻAlpP mutant ǻAlpP mutant
G ǻAlpP mutant starved H ǻAlpP mutant starved
120 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata RESULTS
Figure 4.11 (previous page): Starvation influence on 72 h P. tunicata biofilms. P. tunicata wild-type in MMM (control) (A, B); P. tunicata wild-type in NSS (starvation) (C, D); ǻAlpP mutant in MMM (control) (E, F); ǻAlpP mutant in NSS (starvation) (G, H). Left panels visualised with metabolic activity stain CTC and right panels visualised with LIVE/DEAD stain. Scale bar 50 µm.
121 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata DISCUSSION
4.4 Discussion
It was established that self-induced cell lysis plays an important role during biofilm formation and dispersal of P. tunicata. In particular, cell death within microcolonies affected numbers, metabolic activity and the degree of phenotypic variation of dispersal cells as well as the metabolic activity of surviving attached cells. Only the P. tunicata wild-type showed a major detachment event after cell death had occurred. In contrast, the ǻAlpP mutant biofilm dispersal remained low over the time-course of the experiment. Furthermore, a large, metabolically active dispersal population with high phenotypic variation was found to be generated after cell death had occurred (Figure
4.3, Figure 4.5 - Figure 4.7). Cell death was also demonstrated to influence the spatial distribution of metabolically active cells within biofilms under nutrient starvation conditions (Figure 4.11).
Biofilm formation in microorganisms is generally linked to development and differentiation processes leading to the formation of multicellular 3-dimensional structures. This community-like behaviour gives rise to many benefits for biofilm dwelling cells, including sharing of labour, cell to cell interactions and resistance to different physical stresses. However, there are specialized needs for growth in crowded
‘multicellular’ communities of cells, primarily imposed by the development of strong gradients of nutrients and a range of physical and chemical conditions (Debeer et al.,
1994; Sternberg et al., 1999). Bacterial behaviours under these conditions include loss of viability of a subpopulation of cells and adaptive responses that allow for dispersal and the generation of phenotypic variation among dispersal cells to enhance chances of survival and successful re-colonization (McCann, 2000; Webb et al., 2003a; Boles et al., 2004; Mai-Prochnow et al., 2004).
122 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata DISCUSSION
Multicellular eukaryotes use apoptosis or PCD to remove surplus or damaged cells during development, an adaptation to the specialized needs of multicellular life. In some specialized bacteria, cell death also occurs as an essential part of development during the formation of certain stress induced structures (e.g. spores or fruiting bodies)
(Wireman and Dworkin, 1977; Fernandez and Sanchez, 2002; Gonzalez-Pastor et al.,
2003). Evidence has been presented that the benefits of PCD, such as removal of surplus or damaged cells, not only apply to eukaryotes but also to bacteria during biofilm development, under conditions of cellular crowding and physical and chemical gradients through inducing dispersal of metabolically active and diverse cells.
Specifically, it would appear that generation of dispersal cells with high phenotypic variation provide ecological advantages to bacteria during development.
It is generally recognized that a high diversity within a community protects against unfavourable conditions by increasing the range of conditions in which a community as a whole can thrive (McCann, 2000; Singh et al., 2000). As seen in this study, an increase in diversity of cells within P. tunicata biofilms affected three different traits important for survival and colonization of new surfaces; motility, growth ability and biofilm formation. Variation in the three traits appears to be relatively stable in the dispersal cells as three culturing steps did not allow for reversion of the mutations. The highest variation was detected in the wild-type after cell death had occurred, with some variants showing high growth rates and rapid biofilm formation and some showing slow growth rates and being mostly deficient in biofilm formation (Figure 4.5 - Figure 4.7).
Further, some variants derived from wild-type biofilms showed increased motility
(Figure 4.7). Each of these phenotypes may play a role in the colonization of surfaces under different environmental conditions. For example, a higher growth and biofilm formation rate may be beneficial under high nutrient availability and conversely a slow
123 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata DISCUSSION growth rate and decreased biofilm formation may be advantageous under nutrient limited conditions. A phenotype with increased motility could be vital for settlement at more distant surfaces, ensuring a wider distribution of the organism. Similar events for the generation of dispersal propagules have been observed for many sessile colonizing eukaryotes (Moran and Emlet, 2001; Marshall et al., 2003; Marshall and Keough,
2003). It is clear that variation in a range of phenotypes within propagule populations is a strategy used to ensure successful colonization at new surfaces in different habitats.
For example, differences in the swimming ability due to larvae size and nutritional status of larvae of the bryozoans Bugula neritina and Watersipora subtorquata and the ascidian Diplosoma listerianum leads to variation in settlement distances (Marshall and
Keough, 2003).
Mechanisms by which biofilms regulate dispersal are only beginning to be explored and include quorum sensing signals (Rice et al., 2005), EPS-degrading enzymes (Boyd and
Chakrabarty, 1994; Davey et al., 2003), nutrient levels (Hunt et al., 2004; Sauer et al.,
2004), cell division cycle (Allison et al., 1990), the carbon storage regulator (Jackson et al., 2002) and the global secondary messenger c-di-GMP (Simm et al., 2004). Recently, it was proposed that the generation of phenotypically different and stable dispersal cells in the model biofilm forming bacterium P. aeruginosa follows a phage-mediated death of a subpopulation of cells (Webb et al., 2003b; Webb et al., 2004). Although the mechanisms for inducing cell death and dispersal variants in P. tunicata biofilms are yet to be fully investigated, the antibacterial and autolytic protein, AlpP, is clearly involved as add back of purified AlpP induced cell death and dispersal in mature ǻAlpP mutant biofilms as well as increased phenotypic variation among dispersal cells. It could be suggested that the AlpP-mediated development of large areas of cell death contributes to weakening of the biofilm architecture and facilitates dispersal of cells displaying a wide
124 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata DISCUSSION range of phenotypic characteristics. Furthermore, it may be hypothesized that lysis of neighbouring cells releases nutrients which provide energy for active dispersal of surviving cells.
Bacteria in biofilms are subject to rapid changes in nutrient availability. Because autolysis is known to be important during starvation induced differentiation processes in other bacteria, the possibility that cell death in P. tunicata also influences metabolic activity of surviving cells within the biofilm during nutrient starvation was explored.
The results showed an increased metabolic activity in the region of lysis, as well as decreased PI stained material within microcolonies of P. tunicata wild-type, when the supply of external nutrients was removed. In contrast, the ǻAlpP mutant did not show increased metabolic activity, but an overall loss of biomass was observed upon the removal of external nutrients. A possible explanation for this phenomenon is that cell lysis material which has accumulated in the centre of wild-type microcolonies during non-starvation conditions is metabolized by the cells when no other nutrients are available. However, no lysis material is available to ǻAlpP mutant biofilm cells and thus the biofilm may not be able to withstand nutrient starvation conditions to the same extent as the wild-type. Cell lysis material was shown to be used as a nutrient source for the sporulating organism B. subtilis (Gonzalez-Pastor et al., 2003). In B. subtilis, sporulating cells export a killing factor and a signalling factor which cooperatively prevent neighbouring cells from sporulating and cause them to lyse. The sporulating cells feed on the released nutrients, which then allow them to grow and delay sporulation morphogenesis. Another example is Streptomyces antibioticus where the mycelium, during the life cycle of the organism, undergoes a highly regulated process of PCD, which then contributes to the nutrient support of remaining cells (Miguelez et al., 1999; Fernandez and Sanchez, 2002). Similarly, it may be speculated that cell lysis
125 Chapter 4: Ecological advantages of autolysis during biofilm development of P. tunicata DISCUSSION material in P. tunicata biofilms serves as a nutrient source for other cells in the absence of external nutrients.
This chapter demonstrates that lysis of a subpopulation of cells in P. tunicata biofilms can influence several traits important for dispersal, colonization and survival within the environment and therefore may confer certain social advantages on the organism during development. It was observed that AlpP is required for a major detachment and dispersal event in mature P. tunicata wild-type biofilms and that the wild-type dispersal population consisted of more active cells compared to the ǻAlpP mutant without lysis.
Furthermore, it was demonstrated that cell death of a sub-population is linked to enhanced phenotypic variation in biofilm dispersal cells and enhanced metabolic activity of biofilm cells under nutrient starvation conditions.
Studies on the occurrence of AlpP-homologues in other organisms and their possible role during biofilm formation are reported in Chapter 5.
126 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms INTRODUCTION
5. AlpP homologues appear to have a conserved function during biofilm development and dispersal of several Gram negative organisms
5.1 Introduction
After a role for AlpP mediated cell death in P. tunicata biofilms had been established
(Chapter 4) it was of interest whether this process is specific to P. tunicata. In a recent study by Lucas-Elio et al. (2006) AlpP homologues were identified in a range of Gram negative organisms, including Marinomonas mediterranea MMB-1, Chromobacterium violaceum ATCC 12472, Magnetococcus sp. MC-1, Caulobacter crescentus CB15,
Microbulbifer degradans 2-40, Rhodopseudomonas palustris CGA009 and
Rhodopirellula baltica SH-1, suggesting that it is a common protein and may play an important role in the lifestyle of these organisms.
M. mediterranea is a marine bacterium which was originally isolated from the water column in the Mediterranean Sea (Solano et al., 1997) but was recently also found associated with the sea grass Posidonia oceanica in different Mediterranean areas (E.
Marco-Noales, personal communication; D. Gómez and A. Sánchez-Amat, unpublished data). It has been studied extensively because of its production of secondary metabolites, including melanin. Melanins are dark-brown polyphenolic pigments which are synthesized by both eukaryotes and prokaryotes and are involved in the defence against oxidants, free radicals and UV radiation. M. mediterranea has a multifunctional polyphenol oxidase system for the production of melanin (Solano et al., 1997). M. mediterranea is taxonomically similar to P. tunicata and was in fact originally placed in the genus Alteromonas. Furthermore, M. mediterranea was also found to produce a
127 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms INTRODUCTION large, antibacterial protein (LodA, previously marinocine) with activity against both
Gram positive and Gram negative organisms (Lucas-Elio et al., 2005). LodA has lysine oxidase activity and its antibacterial effect is due to the hydrogen peroxide generated
(Lucas-Elío et al., 2006). LodA has high homology to AlpP and both proteins display the same mode of action (Chapter 3).
C. violaceum is a ȕ-Proteobacterium ubiquitous in soil and water and is found in a variety of ecosystems in tropical and subtropical regions. It was first described in 1882 and has since been the focus of research mainly because of the production of a deep- violet pigment, called violacein (Boisbaudran, 1882). Violacein is reported to have antimicrobial activity against tropical pathogens, including Mycobacterium tuberculosis
(de Souza et al., 1999), Trypanosoma cruzi (Duran et al., 1994), and Leishmania sp.
(Leon et al., 2001) as well as displaying other bactericidal (Lichstein and Van de Sand,
1945), antiviral (Duran and Menck, 2001), and anticancer (Ueda et al., 1994) activities.
Violacein is part of a series of compounds released by C. violaceum presumably to oppose competitors and predators in soil and water environments (Duran and Menck,
2001). The complete genome sequence of C. violaceum has been made available and revealed many interesting and novel properties, including extensive alternative pathways for energy generation, 500 ORFs for transport-related proteins, complex and extensive systems for stress adaptation and motility, and widespread utilization of quorum sensing for control of inducible systems, demonstrating the versatility and adaptability of the organism (Vasconcelos et al., 2003).
C. crescentus is a Gram negative aquatic bacterium and belongs to the group of Į-
Proteobacteria. C. crescentus differentiates and divides asymmetrically at each cell cycle. Furthermore, it has two distinct stages during its lifecycle, an adhesive stage
128 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms INTRODUCTION where the stalk cell attaches via a holdfast to solid surfaces and a motile swarmer cell stage (Stove and Stanier, 1962). An obligate motile phase during its lifecycle was considered to give C. crescentus a competitive advantage in nutrient poor environments by ensuring new settlement at a distance with fresh nutrients (Brun and Janakiraman,
2000). The complete genome sequence revealed that the organism contains a very high number of two component signal transduction proteins which are known to play a significant role in cell cycle progression. Moreover, multiple clusters of genes encoding proteins essential for survival in a nutrient poor habitat were identified, providing C. crescentus with the ability to respond to a wide range of environmental fluctuations
(Nierman et al., 2001).
M. degradans is a Ȗ-Proteobacterium originally isolated from a decaying salt marsh grass in Chesapeake Bay watershed in coastal Virginia (Andrykovitch and Marx, 1988).
It has the ability to degrade at least 10 complex insoluble polysaccharides, including agar, alginic acid, carrageenan, cellulose, chitin, glucan, laminarin, pectin, pullulan, starch, and xylan (Ensor et al., 1999). Because of this characteristic M. degradans may have an important role in carbon and nitrogen recycling in the marine environment.
This chapter describes the detection of AlpP homologues in 13 Gram negative organisms. It was found that similar to P. tunicata death of a subpopulation of cells within microcolonies is displayed by at least 4 species containing an AlpP-homologue, including M. mediterranea, C. violaceum, C. crescentus and M. degradans. The AlpP- homologues are implicated in these cell death events as hydrogen peroxide is detected at the time of killing and catalase can prevent cell death. Further, similar to P. tunicata, this process is linked to the generation of a phenotypically diverse dispersal population in M. mediterranea. Because AlpP-homologues have been identified within several
129 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms INTRODUCTION species it is hypothesised that AlpP-mediated autotoxic events occur across a range of
Gram negative bacterial groups and play an important role in biofilm development and differentiation.
130 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms MATERIALS AND METHODS
5.2 Materials and Methods
5.2.1 Strains and culture conditions
All bacterial strains and the respective culture media used are shown in Table 5.1. P. tunicata and M. mediterranea were grown at 25ºC and C. violaceum, C. crescentus and
M. degradans were grown at 30 °C.
131 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms MATERIALS AND METHODS
Table 5.1: Bacterial strains and culture media
Minimal biofilm Strain Reference Culture media media 3M supplemented with Pseudoalteromonas (Holmström et al., VNSS (Marden et 0.01% trehalose tunicata 1998) al., 1985) (Neidhardt et al., 1974) VNSS + Sm (100 3M supplemented with P. tunicata ǻAlpP (Mai-Prochnow et -1 µg x ml ) + Km 0.01% trehalose mutant SmR, KmR al., 2004) (50 µg x ml-1) (Neidhardt et al., 1974) Marinomonas Marine minimal media Marine media mediterranea (Solano et al., 1997) MN (Hernandez- 2216 (Difco) MMB-1 Romero et al., 2003) Marine media M. mediterranea Marine minimal media (Lucas-Elío et al., 2216 SB1 (lodA mutant) MN (Hernandez- 2006) + Km (50 µg x KmR Romero et al., 2003) ml-1) (Difco) UNSW culture Chromobacterium Luria Bertani collection (Acc. M9 (Miller, 1972) violaceum broth No.: 040100) Caulobacter Australian medium collection of Caulobacter 2 g x l-1 peptone, M2 supplemented with microorganisms crescentus CB15 1 g x l-1 yeast 0.2% xylose (Ely, 1991) (Acc. No.: ACM extract, 0.2 g x l-1 5171 MgSO4 x 7 H2O MN (Hernandez- Marine media Microbulbifer (Andrykovitch and Romero et al., 2003) + 2216 degradans 2-40 Marx, 1988) 0.2% low melting Agarose Escherichia coli Luria Bertani (Hanahan, 1983) - DH5Į broth
5.2.2 Purification of LodA from Marinomonas mediterranea
LodA purification from M. mediterranea cultures was performed as previously described (Lucas-Elio et al., 2005). Briefly, M. mediterranea was inoculated at OD600 of
0.05 in MN media (Hernandez-Romero et al., 2003). After 48 h of culturing, the supernatant was separated by centrifugation and 2 volumes of ethanol were added and kept overnight at 4º C in order to precipitate LodA. The suspension obtained was then
132 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms MATERIALS AND METHODS centrifuged at 19,000 g at 4º C for 20 min. The pellet was allowed to air dry and re- suspended in 1/15 volume of 0.1 M sodium phosphate buffer, pH7. Insoluble material was removed by centrifugation (13,000 g at 4 ºC for 20 min). The activity of the extract obtained was calculated by measuring the halos formed in the antibiogram inhibition test against E. coli DH5Į (Lucas-Elio et al., 2005).
5.2.3 Biofilm experiments
All strains were grown in continuous-culture flow-cells (channel dimensions 1 x 4 x 40 mm) as previously described (Moller et al., 1998). For a detailed description of the method see Chapter 2, Section 2.2.2. In brief, channels were inoculated with 0.5 ml of early stationary phase cultures containing approximately 1 x 109 cells ml-1 and incubated without flow for 1 h at 25 ºC. Flow was then started with a mean flow velocity in the flow cells of 0.2 mm s-1, corresponding to laminar flow with a Reynolds number of
0.02. Biofilms were run at 25 ºC for P. tunicata and M. mediterranea, and at 30 °C for
C. violaceum, C. crescentus and M. degradans.
5.2.3.1 Biofilm staining
To investigate cell death during biofilm development, biofilms were stained using the
LIVE/DEAD BacLight Bacterial Viability Kit (Molecular Probes Inc., Eugene, Oregon,
USA) as described in Chapter 2 (see Section 2.2.3).
To visualize hydrogen peroxide production during biofilm development, biofilms were stained with Amplex Red as described in Chapter 3 (see Section 3.2.7.1).
133 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms MATERIALS AND METHODS 5.2.3.2 Removal of hydrogen peroxide from biofilms
To remove hydrogen peroxide from biofilms, catalase was added to the biofilm media.
All biofilms were allowed to establish for 24 h before catalase was added at a final concentration of 100 µM. This catalase concentration did not affect the planktonic growth rate of the strains. After 3 days of incubation, the biofilm was stained with
Amplex Red and observed with an epifluorescence microscope (see Section 5.2.3.1).
5.2.3.3 Add back of LodA protein to SB1 mutant biofilms
M. mediterranea LodA was prepared as described above (see Section 5.2.2) and approximately 60 µg were added back to each flow cell containing M. mediterranea
SB1 mutant biofilms. The add back experiment was performed as described for P. tunicata in Chapter 2 (see Section 2.2.5.6). As a control, 0.1 M pH 7.0 phosphate buffer was inoculated into separate flow cell channels. Biofilms were incubated at 25 ºC for
3 h before staining with the LIVE/DEAD kit and visualizing with the CLSM.
5.2.4 Phenotypic variation of M. mediterranea dispersal cells
To investigate the hypothesis that cell lysis within microcolonies correlates with phenotypic variation in M. mediterranea, effluent was spread plated onto marine agar
(Difco, Becton Dickenson, USA) at three time-points during biofilm formation: 24 h
(before the onset of cell death), 72 h (shortly after the onset of cell death) and 144 h
(when cell death was more extended throughout the biofilm). Twenty colonies derived from M. mediterranea wild-type and SB1 biofilms were randomly picked from marine agar and screened for growth and biofilm formation as described for P. tunicata in
Chapter 4 (see Section 4.2.3).
134 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms RESULTS
5.3 Results
5.3.1 AlpP homologues in Gram negative organisms
The complete AlpP nucleotide sequence was compared with sequences in the GenBank- database available through the National Centre for Biotechnology Information (NCBI) web site (http://www.ncbi.nlm.nih.gov). This search revealed that AlpP has homology to proteins in at least 13 Gram negative organisms (Table 5.2). Most organisms containing AlpP homologues belong to the group of Proteobacteria, except for one cyanobacterium (Synechococcus sp.) and one planctomycete (R. baltica). While the majority of proteins with similarity to AlpP are hypothetical proteins, the AlpP homologue in M. mediterranea MMB-1 is known to be a lysine oxidase. AlpP homologues are also annotated as a catalase in R. solanacearum, a 3-isopropylmalate dehydrogenase in R. baltica and a Valyl-tRNA synthetase in M. degradans.
135 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms RESULTS
Table 5.2 AlpP homologues in Gram negative organisms
Identity/ Similarity Organism Protein Accession Number Protein identity to AlpP Classification Habitat Autolytic and Pseudoalteromonas tunicata AY 295768 100% (747/747) -Proteobacteria Marine antibacterial protein Ȗ Marinomonas mediterranea 53% (411/769) AY968053 Lysine oxidase -Proteobacteria Marine MMB-1 67% (520/769) Ȗ Chromobacterium violaceum Conserved 36% (191/529) NP_902938 -Proteobacteria Soil/water ATCC 12472 hypothetical protein 52% (277/529) Ȗ 27% (164/594) Caulobacter crescentus CB15 NP_419374 Hypothetical protein -Proteobacteria Soil/water 42% (252/594) Į Magnetococcus sp. Conserved 32% (172/524) ZP_00607515 -Proteobacteria Water MC-1 hypothetical protein 47%(250/524) Į 29% (155/529) Hahella chejuensis KCTC 2396 YP_434329 Hypothetical protein -Proteobacteria Marine 43% (232/529) Ȗ Marinomonas mediterranea 27% (155/574) EAQ66745 Hypothetical protein -Proteobacteria Marine MED121 43% (248/574) Ȗ Rhodopseudomonas palustris 28% (169/588) NP_947813 Hypothetical protein -Proteobacteria Soil/water CGA009 42% (249/588) Į 29% (164/560) Synechococcus sp. WH 7805 ZP_01125241 Hypothetical protein Cyanobacteria Marine 45% (252/560) 3-isopropylmalate 29% (159/533) Rhodopirellula baltica SH 1 NP_866009 Planctomycetes Marine dehydrogenase 43% (232/533) Ralstonia solanacearum 28% (153/542) ZP_00942644 Catalase -Proteobacteria Soil UW551 42% (228/542) ȕ 34% (120/375) Nitrobacter hamburgensis X 14 ZP_00626223 Hypothetical protein -Proteobacteria Soil 46% (173/375) Į Valyl-tRNA 29% (111/377) Microbulbifer degradans 2-40 ZP_00318137 -Proteobacteria Marine synthetase 46% (174/377) Ȗ
Out of these organisms four were chosen for further studies, namely M. mediterranea, C. violaceum, C. crescentus and M. degradans.
136 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms RESULTS
5.3.2 Biofilm development and cell death in M. mediterranea
The lysine oxidase LodA in M. mediterranea has been well characterized and shows high similarity to AlpP (Lucas-Elío et al., 2006). It was demonstrated that AlpP shares the same mode of action and also acts as a lysine oxidase (Chapter 3). Here the possibility that LodA is also implicated in cell death events during M. mediterranea biofilm formation similar to P. tunicata was explored. Biofilms were allowed to form in continuous culture flow cells and stained with the BacLight Live/Dead kit before visualizing with a CLSM. Single cells attached to the substratum and small microcolonies were observed 24 h after inoculation (Figure 5.1 A). After 48 h, larger microcolonies were formed, consisting only of viable cells at this stage (Figure 5.1 C).
However, at three days after inoculation, cell death occurred within microcolonies.
Similar to P. tunicata, dead cells were mainly localized in the centre of microcolonies.
Four days after inoculation subpopulations of dead cells were observed in almost all microcolonies (Figure 5.1 E). Cell death extended throughout the biofilms, before the biofilm structure started to disrupt and detach, suggesting that, similar to P. tunicata, cell death plays a role during biofilm development and subsequent dispersal in M. mediterranea.
5.3.3 Biofilm development of the SB1 mutant strain
To show that biofilm cell death in M. mediterranea is mediated by LodA, biofilm development of a lodA mutant strain (SB1) (Lucas-Elío et al., 2006) was investigated.
The SB1 mutant also formed a biofilm with microcolony based architecture. However, similar to the P. tunicata ǻAlpP mutant major cell death events did not occur during any stage of biofilm development (Figure 5.1 B, D, F) and only few single, dead cells were observed in the SB1 mutant biofilm. To further support the hypothesis that LodA
137 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms RESULTS causes cell death in M. mediterranea, purified LodA was added back to SB1 mutant biofilms. Add back of purified LodA to mature SB1 mutant biofilms induced cell death
(Figure 5.2 A), while the buffer control still showed only viable cells, suggesting that
LodA mediates cell death in M. mediterranea biofilms (Figure 5.2 B).
138 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms RESULTS
A B
C D
EF
Figure 5.1: Biofilm development of M. mediterranea wild-type and SB1 mutant. Biofilms were stained with the BacLight Live/dead viability kit. (A) wild-type 24 h, (B) SB1 mutant 24 h, (C) wild-type 48 h, (D) SB1 mutant 48 h, (E) wild-type 72 h and (F) SB1 mutant 72 h. A corresponding vertical scan is shown below each image.
139 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms RESULTS
A B
Figure 5.2: Add back of the LodA to M. mediterranea SB1 mutant. Biofilms were stained with the BacLight Live/Dead kit. (A) Add back of LodA to 72 h SB1 mutant biofilms (B) 72 h SB1 mutant biofilm plus buffer control. A corresponding vertical scan is shown below each image.
5.3.4 Phenotypic variation of M. mediterranea biofilm dispersal cells
In P. tunicata, AlpP mediated cell death has been linked to the generation of a metabolically active and phenotypically diverse dispersal population (Mai-Prochnow et al., 2006) (Chapter 4). Dispersal cells of the ǻAlpP mutant showed significantly lower variation in motility, growth and biofilm formation. Here it was investigated whether
LodA-mediated cell death in M. mediterranea is also implicated in the generation of phenotypic variation among biofilm dispersal cells. M. mediterranea wild-type and SB1 mutant (which does not show cell death during biofilm formation) dispersal cells were tested for variation in growth and biofilm formation at three different time-points during biofilm development. The M. mediterranea wild-type showed higher variation in growth at all time-points investigated. Variation in growth among the 20 randomly picked colonies was 19 % in the wild-type compared to only 5 % in the SB1 mutant strain (Figure 5.3 A). Variation in biofilm formation was highest in the wild-type at the 140 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms RESULTS 144 h time-point when cell death had occurred extensively throughout the biofilm. At this time-point variation was 45 % in the wild-type compared to only 25 % in the mutant strain (Figure 5.3 B). These results suggest that similar to P. tunicata, LodA mediated biofilm cell death in M. mediterranea is linked to the generation of a phenotypically diverse dispersal population.
A 50
40
30
20
10
Variation coefficient 0 24 h 72 h 144 h Biofilm age
B 50
40
30
20
10
Variation coefficient 0 24 h 72 h 144 h Biofilm age
Figure 5.3: Phenotypic variation in M. mediterranea wild-type and SB1 mutant biofilm dispersal cells. Variation coefficient of M. mediterranea wild-type (Ŷ) and SB1 mutant (Ŷ) biofilm dispersal cells in growth (A) and biofilm formation (B). Variation coefficient (%) calculated for 20 colonies for time-points 24 h, 72 h and 144 h after biofilm inoculation.
141 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms RESULTS 5.3.5 Biofilm formation and cell death in C. violaceum, C. crescentus and M.
degradans
Because C. violaceum, C. crescentus and M. degradans also contain AlpP homologues
(see Section 5.3.1) it was hypothesized that these homologous proteins mediate similar biofilm cell death events and thus play a role in the lifestyle of these organisms. Biofilm formation and possible cell death events in these organisms were investigated using the flow cell system and Live/Dead staining. In all three strains, single cells attached to the substratum followed by microcolony formation at 48 h. Initially microcolonies consisted only of viable cells. However, similar to the observation made for P. tunicata and M. mediterranea, cell death started to occur within the centre of microcolonies after three days of biofilm formation (Figure 5.4). After four days of biofilm formation, subpopulations of dead cells were observed in almost all microcolonies, before the biofilm structure started to disrupt and detach, suggesting that cell death plays a role during biofilm development and subsequent dispersal in these organisms.
A B C
Figure 5.4: Mature biofilms of other organisms containing the AlpP homologue. Biofilms were stained with the BacLight Live/Dead kit at 4 days post inoculation. (A) Chromobacterium violaceum, (B) Caulobacter crescentus and (C) Microbulbifer degradans. A corresponding vertical scan is shown below each picture.
142 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms RESULTS 5.3.6 Detection of hydrogen peroxide in M. mediterranea, C. violaceum, C.
crescentus and M. degradans biofilms
In P. tunicata hydrogen peroxide can be detected at the time of biofilm killing, presumably from the lysine oxidase activity of AlpP (Chapter 3). To show that AlpP homologues are implicated in cell death events of other organisms, the presence of hydrogen peroxide was investigated in these biofilms. Amplex red staining was used to visualize hydrogen peroxide in biofilms of, M. mediterranea, C. violaceum, C. crescentus and M. degradans. Indeed, high red fluorescence was observed in all biofilms investigated associated with microcolonies at the onset of cell death (Table
5.3), indicating the presence of hydrogen peroxide in the biofilms. To give further evidence for the hypothesis that the activity of the AlpP homologues is responsible for cell death events via the production of hydrogen peroxide in these organisms, catalase was added to the biofilm media to remove hydrogen peroxide. Biofilm cell death was almost entirely prevented by the addition of catalase, and little or no hydrogen peroxide was detected within the biofilms after the catalase treatment (Table 5.3).
143 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms RESULTS
Table 5.3: Detection of hydrogen peroxide and cell death in biofilms with and without the addition of catalase.
M. mediterranea M. mediterranea C. violaceum C. crescentus M. degradans wild-type SB1 mutant Biofilms in PI fluorescence +++ - +++ ++ +++ minimal media a (dead cells) b
AR fluorescence b +++ + +++ ++ ++ (H2O2)
Biofilms in PI fluorescence ---+- minimal media (dead cells) b plus catalase a AR fluorescence b ++--+ (H2O2)
a Two sets of triplicate biofilms were inoculated for each strain, one set in minimal media and the other set in minimal media with the addition of catalase (100 µg ml-1) after 24 h. b Biofilms were allowed to establish for 4 days before staining with the BacLight Live/Dead viability kit to detect cell death and the Amplex Red reagent to localize hydrogen peroxide, respectively. Fluorescence intensity was noted as +++ high ++ medium + low - no
144 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms DISCUSSION
5.4 Discussion
The data presented in this chapter suggest that homologues of the autolytic protein AlpP play a similar role during biofilm development of M. mediterranea, C. violaceum, C. crescentus and M. degradans. Similar to AlpP in P. tunicata biofilms, AlpP homologue production caused death of a subpopulation of cells in the centre of microcolonies in these organisms. Moreover, this process was also linked to the production of hydrogen peroxide and is suggested to be important for biofilm differentiation and dispersal.
Most AlpP homologues in the GenBank database are currently annotated as hypothetical proteins and three proteins with lower homology to AlpP show similarity to proteins with a known function, including a catalase, a 3-isopropylmalate dehydrogenase and a Valyl-tRNA synthetase. However, these protein identities are derived from whole genome automatic annotation and thus do not necessarily represent a correct identity of the proteins. Therefore it is feasible that these proteins have a similar mode of action to AlpP and may be autotoxic due to lysine oxidase activity.
M mediterranea strain MMB-1 is a melanogenic marine bacterium originally isolated from a water column sample from the Mediterranean Sea (Solano et al., 1997).
Recently, however new strains have been isolated from the microflora associated with the sea grass Posidonia oceanica in different Mediterranean areas (E. Marco-Noales, personal communication; D. Gómez and A. Sánchez-Amat, unpublished data), suggesting that this could be the microhabitat it occupies. The results described in this chapter demonstrated, for the first time, that M. mediterranea is able to form biofilms which may contribute to its colonization ability and establishment of M. mediterranea on the plant surface. Similarly to P. tunicata, subpopulations of cells die during its biofilm formation. Furthermore, it was demonstrated that cell death in M. mediterranea 145 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms DISCUSSION biofilms is mediated by its lysine oxidase LodA and that this process plays a role in the dispersal of the organisms.
Similar to the P. tunicata biofilm life cycle, cell death seems to play a role in generating a phenotypically diverse dispersal population from M. mediterranea biofilms. This process is hypothesised to be beneficial to the population as the chances of thriving under different environmental conditions are enhanced. Variation in growth among biofilm dispersal cells was highest for the M. mediterranea wild-type cells released after cell death had occurred in the biofilm and the SB1 mutant showed less variation at all time-points investigated (Figure 5.3). Variation in biofilm formation increased for wild type cells released after cell death had occurred in the biofilm. Because variation in biofilm formation among dispersal cells of the SB1 mutant can also increase with biofilm age, although to a lesser extend than the wild-type, other factors for inducing variation may play a role in M. mediterranea biofilms. Variation in cells growing in biofilms has been shown for many organisms including Pseudomonas aeruginosa
(Drenkard and Ausubel, 2002; Boles et al., 2004; Haussler, 2004; Webb et al., 2004),
Staphylococcus aureus (Proctor et al., 1995; Sadowska et al., 2002), Staphylococcus epidermidis (Conlon et al., 2004; Handke et al., 2004), Vibrio cholerae (Ali et al., 2002;
Matz et al., 2005) Listeria monocytogenes (Monk et al., 2004) and P. tunicata (Chapter
3). Diverse mechanisms that may lead to an increased genetic and phenotypic variation in biofilm forming bacteria have been investigated. These mechanisms include phase variation (Drenkard and Ausubel, 2002), adaptive mutation (Oliver et al., 2000; Bjedov et al., 2003), enhanced gene transfer through conjugation, transformation (Hausner and
Wuertz, 1999; Hendrickx et al., 2003; Molin and Tolker-Nielsen, 2003), phage induction (Webb et al., 2004) and self-induced lysis (Chapter 3). Here it is shown that the onset of LodA-mediated killing correlates with the generation of high variation
146 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms DISCUSSION among dispersal cells of M. mediterranea biofilms. It suggests that similar to AlpP in P. tunicata, LodA has an important effect on the biofilm life-style of the organism.
Biofilm formation of C. crescentus has previously been described (Entcheva-Dimitrov and Spormann, 2004). Interestingly, the latter authors observed the occurrence of cell death within microcolonies which was hypothesized to be due to senescence. Here it is suggested that hydrogen peroxide production as a result of the activity of an AlpP homologue is responsible for cell death in C. crescentus biofilms. Biofilm formation by
C. violaceum and M. degradans had not been previously characterized. Results presented in this chapter show that these organisms form a differentiated, microcolony based biofilm. Moreover, cell death occurs in the centre of microcolonies due to the production of hydrogen peroxide during the normal course of biofilm development leading to subsequent dispersal of the biofilms.
Hydrogen peroxide is a low molecular weight compound that diffuses rapidly and acts on a wide range of molecules. These properties are likely to facilitate cell death and dispersal even in complex microbial communities such as those on living marine surfaces. It was found that in biofilms of M. mediterranea, C. violaceum, C. crescentus and M. degradans, hydrogen peroxide is mainly detected at the outer, viable cell layers of microcolonies and is not restricted to the microcolony centre where cell death occurs
(Table 5.3). This could suggest that hydrogen peroxide is actively generated by viable cells rather than released upon cell death in the centre of microcolonies. Furthermore, it may be hypothesised that cells in the centre of microcolonies are more susceptible to hydrogen peroxide due to senescence. Senescence involves increased levels of intracellular reactive oxygen species and thus a small additional exogenous amount of hydrogen peroxide could lead to the death of these cells.
147 Chapter 5 AlpP homologues appear to have a conserved function during biofilm development and dispersal of several gram negative organisms DISCUSSION Because hydrogen peroxide is a by-product of aerobic respiration it could be expected to occur in aerobic areas of all biofilms and not only biofilms of organisms investigated in this study. However, hydrogen peroxide cannot be detected with Amplex Red staining in Pseudomonas aeruginosa biofilms (Barraud, N., J. S. Webb and S.
Kjelleberg, unpublished). Moreover, the fact that Amplex Red staining revealed no or very little hydrogen peroxide in the ǻAlpP mutant (Chapter 3) and SB1 mutant biofilms
(Table 5.3) suggests that hydrogen peroxide is actively produced as result of AlpP or its homologues activity by cells in wild-type biofilms of M. mediterranea, C. violaceum, C. crescentus and M. degradans.
148 Chapter 6 General discussion and future outlooks
6. General discussion and future outlooks
Apoptosis in eukaryotes is known as the deliberate life relinquishment of an unwanted cell and generally confers advantages during the lifecycle of an organism (Kerr et al.,
1972; Ellis et al., 1991). The work described in this thesis has provided evidence for analogous processes during the formation of biofilms by the marine surface associated bacterium Pseudoalteromonas tunicata.
Almost every natural and man-made surface in an aqueous environment can become colonized by biofilm forming bacteria (Costerton et al., 1994; Costerton et al., 1999).
Biofilms are extensively studied today, in part because of the realization that there is a systematic developmental sequence displayed by biofilm forming bacterial communities
(Southey-Pillig et al., 2005), and because the colonization of surfaces by bacteria can cause significant industrial and medical problems (Lappin-Scott and Costerton, 1989;
Zottola and Sasahara, 1994; Tenke et al., 2006).
6.1 Cell death promotes biofilm dispersal and phenotypic variation
During P. tunicata biofilm development an apoptosis-like behaviour was found to occur, leading to the production of a metabolically active and phenotypically diverse dispersal population. It was demonstrated that the involvement of cell death in these parameters plays a significant role in the spread and survival of P. tunicata in the environment. Thus, the results obtained in this thesis showed for the first time that developmental cell death within bacterial biofilms benefits surviving cells similar to programmed cell death in higher organisms.
After cell death occurred within the P. tunicata wild-type biofilm a major dispersal event was induced and the dispersal population showed increased metabolic activity. In
149 Chapter 6 General discussion and future outlooks
contrast, no major dispersal event was observed from ǻAlpP mutant biofilms and only low numbers of viable cells dispersed. The metabolic activity of the mutant dispersal cells remained low compared to the wild-type dispersal population. A sudden increase in the number of viable cells dispersing from a biofilm such as in P. tunicata wild-type may be beneficial when the local conditions become unfavourable and the spread of the organism to new colonisation sites can increase chances of survival.
Biofilm cell death also influenced phenotypic traits of dispersal cells. P. tunicata wild- type dispersal cells showed a higher phenotypic variation than dispersal cells from the
ǻAlpP mutant biofilms. An analogous link between differentiation, phenotypic variation and cell lysis occurs in the opportunistic pathogen P. aeruginosa. In this organism, the activity of a prophage (Pf4) causes death of a subpopulation of cells and also generates a subpopulation of small colony phenotypic variants that over-express the phage (Webb et al., 2004).
A high diversity in different phenotypic traits would be beneficial to eukaryotic and prokaryotic communities by increasing the range of conditions under which a population can thrive (McCann, 2000; Boles et al., 2004). In P. tunicata, within population diversification appeared to occur as a response to AlpP-mediated death of a subpopopulation of cells. One explanation for the link between cell death and phenotypic variation may be that AlpP-mediated cell death triggers stress responses within the cell which induce high mutation rates (e.g. by the SOS response). Increased mutation rates in this manner could explain the increases in phenotypic variation observed in P. tunicata, producing cell subpopulations with a range of different characteristics. Cells with beneficial mutations would have an ecological benefit so that evolutionary selection can take place.
150 Chapter 6 General discussion and future outlooks
In addition to the importance of regulating biofilm dispersal for P. tunicata itself, cell death may also play a role for the living hosts (e.g. algae or tunicates) in the marine environment. Many sessile algae and animals have evolved defence mechanisms against fouling by producing metabolites that can influence the settlement, growth and survival of other organisms (Davis and Wright, 1990; de Nys et al., 1995). However, algae and animals lacking chemical and non-chemical defences are thought to rely on secondary metabolites produced by associated surface bacteria such as P. tunicata, as their defence against fouling (Thomas and Allsopp, 1983; Holmström et al., 1992; Kon-ya et al.,
1995). Thus cell death and dispersal of a subpopulation of cells within P. tunicata biofilms may protect its host against uncontrolled biofilm formation and fouling by P. tunicata itself.
6.2 Biofilm dispersal displays similarities to dispersal of sessile
invertebrates
For most organisms in the aquatic environment, successful colonisation of a surface by dispersive propagules (e.g. invertebrate larvae, bacterial cells) is a fundamental constraint on their life histories. Benthic organisms have developed a two phase lifestyle, with a sessile stage alternating with a planktonic dispersal stage (Caley et al.,
1996; Kisdi, 2002). It is possible that bacterial biofilms display a similar lifecycle, where the surface attached mode of growth is followed by a dispersal phase. Other similarities between microbial biofilms and traditional multicellular organisms include communication between cells (Davies et al., 1998; Huber et al., 2001), differentiation of cells within the biofilm (Sauer et al., 2002) and the development of predictable multicellular structures (microcolonies) (Costerton et al., 1999).
151 Chapter 6 General discussion and future outlooks
Variation among dispersive propagules occurs in prokaryotes as well as eukaryotes, where it has been widely studied and shown to have profound effects on dispersal, colonisation and post colonisation success. For example in benthic aquatic, eukaryotic organisms, variation in larval/offspring size affects subsequent juvenile growth of snails and colony growth of bryozoans, both crucial to the post settlement success of these organisms (Moran and Emlet, 2001; Marshall et al., 2003; Marshall and Keough, 2003).
For marine invertebrates, differences in the swimming ability of larvae leads to variation in settlement distances (Marshall and Keough, 2003), ensuring optimal spread and survival chances of the species. In bacterial biofilms, dispersal cells show a similar level of variation in key traits. Biofilms release dispersal cells of varying types and quantities, for example, different sized cells, flagellated versus non-flagellated cells
(Deziel et al., 2001; Haussler et al., 2003; Boles et al., 2004). In P. tunicata variation affected motility, growth and biofilm formation in dispersal cells (Mai-Prochnow et al.,
2006) (Chapter 4). It can be concluded that benthic, aquatic eukaryotes and bacterial biofilms, such as those formed by P. tunicata, use similar strategies to ensure optimal spread, colonisation success and survival of the species. Moreover, the evolutionary and ecological pressures acting on dispersal and colonisation are fundamentally the same and thus dispersal strategies used by bacterial biofilms may represent an evolutionary precursor to the dispersal strategies from higher eukaryotes.
6.3 A novel mechanism mediates biofilm cell death in P. tunicata and
other Gram negative organisms
Biofilm cell death was previously observed in several mono species biofilms and mixed species biofilm communities (Auschill et al., 2001; Hope et al., 2002; Webb et al.,
2003b; Entcheva-Dimitrov and Spormann, 2004). However, the current study is the first
152 Chapter 6 General discussion and future outlooks
to fully elucidate a mechanism of biofilm killing. Notably, biofilm cell death only occurred in P. tunicata wild-type and not the ǻAlpP mutant. Other mutants of P. tunicata have also been generated, including D2W2 (disrupted in wmpR) and D2W3
(disrupted in wmpD, which encodes the general secretion pathway protein) (Egan et al.,
2002a; Egan et al., 2002b), both of which are deficient in the production of AlpP and show no cell death during biofilm development despite the formation of a similar biofilm architecture (Mai-Prochnow, 2002). Thus, it was concluded that AlpP is involved in biofilm killing in P. tunicata.
In order to gain mechanistic insight into AlpP-mediated killing, the mode of action of the protein was determined. The autolytic effect of AlpP was demonstrated to be due to the production of hydrogen peroxide from the oxidation of lysine (Chapter 3). Hydrogen peroxide is a small, rapidly diffusing molecule, likely to affect many organisms and thus can play an important role in complex biofilm communities in the marine environment.
Catalase breaks down hydrogen peroxide and could therefore act as an antidote to AlpP.
It is naturally produced by most aerobic microorganism and is possibly involved in the protection of cells from the effect of AlpP.
Lysine oxidases have been described for the fungus Trichoderma sp. (Lukasheva and
Berezov, 2002), but were only recently found in a prokaryote, the marine bacterium
Marinomonas mediterranea (Lucas-Elío et al., 2006). The lysine oxidase of M. mediterranea, LodA, shows high sequence homology to AlpP and several other AlpP homologues were identified in a range of Gram negative organisms (Chapter 5).
Therefore, it seems possible that AlpP homologues are common and may have a conserved function during biofilm development of several bacteria. Indeed, it was shown that similar to P. tunicata, cell death occurs within the centre of microcolonies in at least 4 organisms containing an AlpP homologue, including M. mediterranea, C. 153 Chapter 6 General discussion and future outlooks
violaceum, C. crescentus and M. degradans. Furthermore, cell death in M. mediterranea biofilms was linked to the generation of a phenotypically diverse dispersal population
(Chapter 5), demonstrating that cell death events also influence traits of dispersal cells in these organisms.
In summary, the results of this study demonstrate that self-induced lysis of a subpopulation of cells mediated by hydrogen peroxide production through AlpP or
AlpP-like proteins occurs across several bacterial taxa and may be a common process during biofilm differentiation. It is proposed that this process is an important mechanism for facilitating dispersal and generating diversity in a range of bacteria. For the first time it is shown that self induced lysis, while appearing undesirable for single cell prokaryotes can benefit a species during the biofilm mode of growth. The process of biofilm cell death shows similarities to apoptosis in eukaryotes and thus it may be speculated that it represents the prokaryotic precursor of programmed cell death.
6.4 Future directions
Several questions worthy of further investigations have emerged from this work:
1. Given the importance of AlpP in P. tunicata and its homologues in several other
organisms, it is important to investigate how AlpP is regulated and expressed.
More specifically:
a. How is AlpP expression regulated within the cell? A global regulator,
WmpR, has already been identified as involved in AlpP expression
(Stelzer et al., 2006) and it appears likely that a complex regulatory
network is in place.
154 Chapter 6 General discussion and future outlooks
b. Do external signals (e.g. nutrient availability, such as the substrate lysine
for the production of hydrogen peroxide) or cell density dependent
signals (such as quorum sensing) trigger AlpP expression? Following
annotation of the newly available sequence of the genome of P. tunicata
it would be possible to identify regulators or possible signalling systems
involved in such a response.
c. How do P. tunicata cells protect themselves from the autolytic action of
AlpP? The role of catalase should be investigated, as catalase is likely to
break down the released hydrogen peroxide and thus protects the cell.
Three different catalase genes have been identified within the P. tunicata
genome (unpublished). Studying their regulation and expression within a
biofilm, for example using GFP tagged promoter constructs could
provide insights into mechanisms of resistance by P. tunicata cells to
AlpP.
2. What is the genetic basis of diversity in P. tunicata biofilm dispersal cells? The
occurrence of diversity within phenotypes has been identified, but not the
genomic level changes. Because high phenotypic variation occurs only among P.
tunicata wild-type biofilm cells and not among ǻAlpP mutant biofilm cells,
experiments could be designed to study real-time evolution in the very important
ecological context of biofilms. Sequencing of dispersal populations could
identify possible mutations responsible for various phenotypes.
155 Appendix
Solutions and Buffers
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 x 6H2O,
0.41 g CaCl2 x 2H2O,
0.008 g SrCl2 x 6H2O,
0.008 g H3BO3,
- Adjust to pH 7
VNSS (per litre NSS) (Marden et al., 1985)
1.0 g peptone,
0.5 g yeast extract,
0.5 g glucose,
0.01 g FeSO4 x 7H2O,
0.01 g Na2HPO4,
- For agar plates add 15 g agar before autoclaving
156 Luria Broth (LB) medium (per litre)
10 g NaCl,
10 g tryptone,
5 g yeast extract
- Adjust to pH 7.5
- For agar plates add 15 g agar before autoclaving
Peptone Yeast extract (per litre) for C. crescentus
2 g peptone
1 g yeast extract
1 ml of 1M MgSO4
0.5 ml of 1 M CaCl2
Marine Minimal Medium (MMM) for P. tunicata
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 + FeSO4 x 7H2O (pH 7.8) (sterile filtered)
10 ml 132 mM K2HPO4 (autoclaved, add slowly while stirring)
10 ml 952 mM NH4Cl (pH 7.8) (autoclaved)
10 ml 20% Trehalose (sterile filtered)
157 Marine Minimal Medium (MN) per litre for M. mediterranea and M. degradans
20 g NaCl
7 g MgSO4 7H2O
5.3 g MgCl2 5H2O
0.7 g KCl
1.25 g CaCl2
6.1 g Tris base
- Adjust to pH 7.4 before autoclaving
- After autoclaving add:
2 ml of 2.5 g/l FeSO4 7H2O (autoclaved)
427.5 µl 1 M K2HPO4 (autoclaved)
40 ml of 10% sodium glutamate (sterile filtered)
Marine Minimal Media (M2) per litre for C. crescentus
2.19 g Na2HPO4
0.53 g KH2PO4
0.5 g NH4Cl
3.82 mg FeSO4
- After autoclaving add:
0.5 ml of 1M MgSO4 (autoclaved)
0.5 ml of 1M CaCl2 (autoclaved)
10 ml of 20 % glucose (sterile filtered)
158 Minimal Media (M9) per litre for C. violaceum
6.78 g Na2HPO4
3 g KH2PO4
0.5 g NaCl
1 g NH4Cl
- After autoclaving add
2 ml of 1M MgSO4 (autoclaved)
0.1 ml of 1M CaCl2 (autoclaved)
5 ml of 20% glucose (sterile filtered)
Phosphate Buffer Solution (PBS), pH 7.4 (per litre)
8 g NaCl
0.2 g KCl
1.44 g Na2HPO4
0.24 g KH2PO4
159 REFERENCES
References
Agladze, K., D. Jackson and T. Romeo (2003). "Periodicity of cell attachment patterns during Escherichia coli biofilm development." J Bacteriol 185: 5632-8.
Agladze, K., X. Wang and T. Romeo (2005). "Spatial periodicity of Escherichia coli K-
12 biofilm microstructure initiates during a reversible, polar attachment phase of development and requires the polysaccharide adhesin PGA." J Bacteriol 187: 8237-46.
Alekseeva, S. A., I. Y. Bakunina, O. I. Nedashkovskaya, V. V. Isakov, V. V. Mikhailov and T. N. Zvyagintseva (2004). "Intracellular alginolytic enzymes of the marine bacterium Pseudoalteromonas citrea KMM 3297." Biochemistry (Mosc) 69: 262-9.
Ali, A., M. H. Rashid and D. K. Karaolis (2002). "High-frequency rugose exopolysaccharide production by Vibrio cholerae." Appl Environ Microbiol 68: 5773-8.
Allison, D. G., D. J. Evans, M. R. Brown and P. Gilbert (1990). "Possible involvement of the division cycle in dispersal of Escherichia coli from biofilms." J Bacteriol 172:
1667-9.
Allison, D. G., B. Ruiz, C. SanJose, A. Jaspe and P. Gilbert (1998). "Extracellular products as mediators of the formation and detachment of Pseudomonas fluorescens biofilms." FEMS Microbiol Lett 167: 179-84.
Andersen, R. J., M. S. Wolfe and D. J. Faulkner (1974). "Autotoxic production by a marine Chromombacterium." Marine Biology 27: 281-285.
160 REFERENCES
Anderson, G. G., J. J. Palermo, J. D. Schilling, R. Roth, J. Heuser and S. J. Hultgren
(2003). "Intracellular bacterial biofilm-like pods in urinary tract infections." Science
301: 105-7.
Andrykovitch, G. and I. Marx (1988). "Isolation of a New Polysaccharide-Digesting
Bacterium from a Salt Marsh." Appl Environ Microbiol 54: 1061-1062.
Arevalo-Ferro, C., G. Reil, A. Gorg, L. Eberl and K. Riedel (2005). "Biofilm formation of Pseudomonas putida IsoF: the role of quorum sensing as assessed by proteomics."
Syst Appl Microbiol 28: 87-114.
Auschill, T. M., N. B. Arweiler, L. Netuschil, M. Brecx, E. Reich, A. Sculean and N. B.
Artweiler (2001). "Spatial distribution of vital and dead microorganisms in dental biofilms." Arch Oral Biol 46: 471-6.
Balestrino, D., J. A. Haagensen, C. Rich and C. Forestier (2005). "Characterization of type 2 quorum sensing in Klebsiella pneumoniae and relationship with biofilm formation." J Bacteriol 187: 2870-80.
Bjedov, I., O. Tenaillon, B. Gerard, V. Souza, E. Denamur, M. Radman, F. Taddei and
I. Matic (2003). "Stress-induced mutagenesis in bacteria." Science 300: 1404-9.
Boisbaudran, L. D. (1882). "Matiere colorante se formant dans la cole de farine." Comp
Rend Acad Sci 94: 562.
Boles, B. R., M. Thoendel and P. K. Singh (2004). "Self-generated diversity produces
"insurance effects" in biofilm communities." Proc Natl Acad Sci U S A 101: 16630-5.
Boles, B. R., M. Thoendel and P. K. Singh (2005). "Rhamnolipids mediate detachment of Pseudomonas aeruginosa from biofilms." Mol Microbiol 57: 1210-23.
161 REFERENCES
Bomchil, N., P. Watnick and R. Kolter (2003). "Identification and characterization of a
Vibrio cholerae gene, mbaA, involved in maintenance of biofilm architecture." J
Bacteriol 185: 1384-90.
Boyd, A. and A. M. Chakrabarty (1994). "Role of alginate lyase in cell detachment of
Pseudomonas aeruginosa." Appl Environ Microbiol 60: 2355-9.
Brauner, T., D. F. Hulser and R. J. Strasser (1984). "Comparative measurements of membrane potentials with microelectrodes and voltage-sensitive dyes." Biochim
Biophys Acta 771: 208-16.
Brun, Y. V. and R. Janakiraman (2000). The dimorphic life cycle of Caulobacter and stalked bacteria, p. 297–318. In Y. V. Brun and L. J. Shimkets (ed.), Prokaryotic
Development. ASM Press Washington DC, Washington DC
Burbulys, D., K. A. Trach and J. A. Hoch (1991). "Initiation of sporulation in B. subtilis is controlled by a multicomponent phosphorelay." Cell 64: 545-52.
Burkholder, P. R., R. M. Pfister and F. H. Leitz (1966). "Production of a pyrrole antibiotic by a marine bacterium." Appl Microbiol 14: 649-53.
Busscher, H. J. and A. H. Weerkamp (1987). "Specific and non-specific interactions in bacterial adhesion to solid substrata." FEMS Microbiol Review 46: 165-173.
Caley, M. J., M. H. Carr, M. A. Hixon, T. P. Hughes, G. P. Jones and B. A. Menge
(1996). "Recruitment and the local dynamics of open marine populations." Annu Rev
Ecol Syst 27: 477-500.
162 REFERENCES
Carniol, K. and M. S. Gilmore (2004). "Signal transduction, quorum-sensing, and extracellular protease activity in Enterococcus faecalis biofilm formation." J Bacteriol
186: 8161-3.
Chabeaud, P., A. de Groot, W. Bitter, J. Tommassen, T. Heulin and W. Achouak
(2001). "Phase-variable expression of an operon encoding extracellular alkaline protease, a serine protease homolog, and lipase in Pseudomonas brassicacearum." J
Bacteriol 183: 2117-20.
Champness, W. C. and K. F. Chater (1994). Regulation and integration of antibiotic production and morphological differentiation in Streptomyces spp., p. 61–93. In P.
Piggot, C. P. Moran and P. Youngman (ed.), Regulation of bacterial differentiation.
American Society for Microbiology, Washington, D.C
Chang, A. L., J. R. Tuckerman, G. Gonzalez, R. Mayer, H. Weinhouse, G. Volman, D.
Amikam, M. Benziman and M. A. Gilles-Gonzalez (2001). "Phosphodiesterase A1, a regulator of cellulose synthesis in Acetobacter xylinum, is a heme-based sensor."
Biochemistry 40: 3420-6.
Characklis, W. G. (1981). "Fouling biofilm development: a process analysis." Biotech
Bioeng 23: 1923-1960.
Charlton, T. S., R. de Nys, A. Netting, N. Kumar, M. Hentzer, M. Givskov and S.
Kjelleberg (2000). "A novel and sensitive method for the quantification of N-3-oxoacyl homoserine lactones using gas chromatography-mass spectrometry: application to a model bacterial biofilm." Environ Microbiol 2: 530-41.
163 REFERENCES
Chater, K. F. (1989). Sporulation in Streptomyces p. 277–299. In I. Smith, R. A.
Slepecky and P. Setlow (ed.), Regulation of prokaryotic development: structural and functional analysis of bacterial sporulation and germination. American Society for
Microbiology, Washington D. C.
Chen, C. (2001). "Periodontitis as a biofilm infection." J Calif Dent Assoc 29: 362-9.
Conlon, K. M., H. Humphreys and J. P. O'Gara (2004). "Inactivations of rsbU and sarA by IS256 represent novel mechanisms of biofilm phenotypic variation in
Staphylococcus epidermidis." J Bacteriol 186: 6208-19.
Costa-Ramos, C. and A. F. Rowley (2004). "Effect of extracellular products of
Pseudoalteromonas atlantica on the edible crab Cancer pagurus." Appl Environ
Microbiol 70: 729-35.
Costerton, J. W., Z. Lewandowski, D. E. Caldwell, D. R. Korber and H. M. Lappin-
Scott (1995). "Microbial biofilms." Annu Rev Microbiol 49: 711-45.
Costerton, J. W., Z. Lewandowski, D. DeBeer, D. Caldwell, D. Korber and G. James
(1994). "Biofilms, the customized microniche." J Bacteriol 176: 2137-42.
Costerton, J. W., P. S. Stewart and E. P. Greenberg (1999). "Bacterial biofilms: a common cause of persistent infections." Science 284: 1318-22.
Cowan, S. E., E. Gilbert, D. Liepmann and J. D. Keasling (2000). "Commensal interactions in a dual-species biofilm exposed to mixed organic compounds." Appl
Environ Microbiol 66: 4481-5.
164 REFERENCES
Craig, A. and A. Scherf (2003) Antigenic variation. Academic Press Elsevier,
Amsterdam.
Crossman, L. and J. M. Dow (2004). "Biofilm formation and dispersal in Xanthomonas campestris." Microbes Infect 6: 623-9.
Dalisay-Saludes, D. (2004). Antibacterial activity and mechanisms of colonization of
Pseudoalteromonas tunicata. School of Biotechnology and Biomolecular Sciences,
Faculty of Science. Sydney, University of New South Wales. PhD thesis.
Danese, P. N., L. A. Pratt and R. Kolter (2000). "Exopolysaccharide production is required for development of Escherichia coli K-12 biofilm architecture." J Bacteriol
182: 3593-6.
Danese, P. N., L. A. Pratt and R. Kolter (2001). "Biofilm formation as a developmental process." Methods Enzymol 336: 19-26.
Davey, M. E., N. C. Caiazza and G. A. O'Toole (2003). "Rhamnolipid surfactant production affects biofilm architecture in Pseudomonas aeruginosa PAO1." J Bacteriol
185: 1027-36.
Davey, M. E. and G. A. O'Toole (2000). "Microbial biofilms: from ecology to molecular genetics." Microbiol Mol Biol Rev 64: 847-67.
Davies, D. G. and G. G. Geesey (1995). "Regulation of the alginate biosynthesis gene algC in Pseudomonas aeruginosa during biofilm development in continuous culture."
Appl Environ Microbiol 61: 860-7.
165 REFERENCES
Davies, D. G., M. R. Parsek, J. P. Pearson, B. H. Iglewski, J. W. Costerton and E. P.
Greenberg (1998). "The involvement of cell-to-cell signals in the development of a bacterial biofilm." Science 280: 295-8.
Davis, A. R. and A. E. Wright (1990). "Inhibtion of larval settlement by natural products from ascidian, Eudistoma olivaceum (Van Name)." Journal of Chemical
Ecology 16: 1349-1357.
De Kievit, T. R., R. Gillis, S. Marx, C. Brown and B. H. Iglewski (2001). "Quorum- sensing genes in Pseudomonas aeruginosa biofilms: their role and expression patterns."
Appl Environ Microbiol 67: 1865-73. de Nys, R., P. D. Steinberg, P. Willemsen, S. A. Dworjanyn, C. L. Gabelish and R. J.
King (1995). "Broad spectrum affects of secundary metabolites from the red algae
Delisea pulchra in antifouling assays." Biofouling 8: 259-271. de Souza, A. O., D. C. G. Aily, D. N. Sato and N. Durán (1999). "Atividade da violaceína in vitro sobre o Mycobacterium turbeculosis H37RA." Rev Inst Adolfo Lutz
58: 59–62.
Debeer, D., P. Stoodley, F. Roe and Z. Lewandowski (1994). "Effects of Biofilm
Structures on Oxygen Distribution and Mass- Transport." Biotechnol. Bioeng. 43: 1131-
1138.
Deziel, E., Y. Comeau and R. Villemur (2001). "Initiation of biofilm formation by
Pseudomonas aeruginosa 57RP correlates with emergence of hyperpiliated and highly adherent phenotypic variants deficient in swimming, swarming, and twitching motilities." J Bacteriol 183: 1195-204.
166 REFERENCES
Dow, J. M., L. Crossman, K. Findlay, Y. Q. He, J. X. Feng and J. L. Tang (2003).
"Biofilm dispersal in Xanthomonas campestris is controlled by cell-cell signaling and is required for full virulence to plants." Proc Natl Acad Sci U S A 100: 10995-1000.
Drenkard, E. and F. M. Ausubel (2002). "Pseudomonas biofilm formation and antibiotic resistance are linked to phenotypic variation." Nature 416: 740-3.
Dunny, G. M. and B. A. Leonard (1997). "Cell-cell communication in gram-positive bacteria." Annu Rev Microbiol 51: 527-64.
Duran, N., R. V. Antonio, M. Haun and R. A. Pili (1994). "Biosynthesis of a
Trypanocide by Chromobacterium violaceum." World J Microbiol Biotechnol 10: 686-
690.
Duran, N. and C. F. Menck (2001). "Chromobacterium violaceum: a review of pharmacological and industiral perspectives." Crit Rev Microbiol 27: 201-22.
Dworkin, M. (1972). The myxobacteria: new directions in studies of procaryotic development, p. 435-452. In A. I. Laskin and H. Lechevalier (ed.), CRC critical reviews in microbiology, vol. 1. CRC press, Cleveland
Egan, S., S. James, C. Holmstrom and S. Kjelleberg (2002a). "Correlation between pigmentation and antifouling compounds produced by Pseudoalteromonas tunicata."
Environ Microbiol 4: 433-42.
Egan, S., S. James, C. Holmström and S. Kjelleberg (2001). "Inhibition of algal spore germination by the marine bacterium Pseudoalteromonas tunicata." FEMS Microbiol
Ecol 35: 67-73.
167 REFERENCES
Egan, S., S. James and S. Kjelleberg (2002b). "Identification and characterization of a putative transcriptional regulator controlling the expression of fouling inhibitors in
Pseudoalteromonas tunicata." Appl Environ Microbiol 68: 372-8.
Egan, S., T. Thomas, C. Holmström and S. Kjelleberg (2000). "Phylogenetic relationship and antifouling activity of bacterial epiphytes from the marine alga Ulva lactuca." Environ Microbiol 2: 343-7.
Ellis, R. E., J. Y. Yuan and H. R. Horvitz (1991). "Mechanisms and functions of cell death." Annu Rev Cell Biol 7: 663-98.
Ely, B. (1991). "Genetics of Caulobacter crescentus." Methods Enzymol 204: 372-84.
Ensor, L., S. Stosz and R. Weiner (1999). "Expression of multiple complex polysaccharide-degrading enzyme systems by marine bacterium strain 2-40." J Ind
Microbiol Biotechnol 23: 123-126.
Entcheva-Dimitrov, P. and A. M. Spormann (2004). "Dynamics and control of biofilms of the oligotrophic bacterium Caulobacter crescentus." J Bacteriol 186: 8254-66.
Fernandez, M. and J. Sanchez (2001). "Viability staining and terminal deoxyribonucleotide transferase-mediated dUTP nick end labelling of the mycelium in submerged cultures of Streptomyces antibioticus ETH7451." J Microbiol Methods 47:
293-8.
Fernandez, M. and J. Sanchez (2002). "Nuclease activities and cell death processes associated with the development of surface cultures of Streptomyces antibioticus ETH
7451." Microbiology 148: 405-12.
168 REFERENCES
Franks, A., P. Haywood, C. Holmström, S. Egan, S. Kjelleberg and N. Kumar (2005).
"Isolation and structure elucidation of a novel yellow pigment from the marine bacterium Pseudoalteromonas tunicata." Molecules 10: 1286 -1291.
Fukuda, A., H. Iba and Y. Okada (1977). "Stalkless mutants of Caulobacter crescentus." J Bacteriol 131: 280-7.
Fuqua, W. C., S. C. Winans and E. P. Greenberg (1994). "Quorum sensing in bacteria: the LuxR-LuxI family of cell density-responsive transcriptional regulators." J Bacteriol
176: 269-75.
Fux, C. A., S. Wilson and P. Stoodley (2004). "Detachment characteristics and oxacillin resistance of Staphyloccocus aureus biofilm emboli in an in vitro catheter infection model." J Bacteriol 186: 4486-91.
Garrett, E. S., D. Perlegas and D. J. Wozniak (1999). "Negative control of flagellum synthesis in Pseudomonas aeruginosa is modulated by the alternative sigma factor AlgT
(AlgU)." J Bacteriol 181: 7401-4.
Gauthier, G. (1979). "Alteromonas rubra sp. nov. marine antibiotic-producing bacterium." Int J Syst Bacteriol 26: 459-466.
Gauthier, G. and V. Breittmayer (1979). "A new antibiotic-producing bacterium from seawater: Alteromonas aurantia sp. nov. ." Int J Syst Bacteriol 29: 366-372.
169 REFERENCES
Gauthier, G., M. Gauthier and R. Christen (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: 755-61.
Gauthier, M. J. and G. N. Flatau (1976). "Antibacterial activity of marine violet- pigmented Alteromonas with special reference to the production of brominated compounds." Can J Microbiol 22: 1612-9.
Genevaux, P., P. Bauda, M. S. DuBow and B. Oudega (1999). "Identification of Tn10 insertions in the rfaG, rfaP, and galU genes involved in lipopolysaccharide core biosynthesis that affect Escherichia coli adhesion." Arch Microbiol 172: 1-8.
Genevaux, P., S. Muller and P. Bauda (1996). "A rapid screening procedure to identify mini-Tn10 insertion mutants of Escherichia coli K-12 with altered adhesion properties."
FEMS Microbiol Lett 142: 27-30.
Gonzalez-Pastor, J. E., E. C. Hobbs and R. Losick (2003). "Cannibalism by sporulating bacteria." Science 301: 510-3.
Gonzalez Barrios, A. F., R. Zuo, Y. Hashimoto, L. Yang, W. E. Bentley and T. K.
Wood (2006). "Autoinducer 2 controls biofilm formation in Escherichia coli through a novel motility quorum-sensing regulator (MqsR, B3022)." J Bacteriol 188: 305-16.
Hammer, B. K. and B. L. Bassler (2003). "Quorum sensing controls biofilm formation in Vibrio cholerae." Mol Microbiol 50: 101-4.
170 REFERENCES
Hanahan, D. (1983). "Studies on transformation of Escherichia coli with plasmids." J
Mol Biol 166: 557-80.
Handke, L. D., K. M. Conlon, S. R. Slater, S. Elbaruni, F. Fitzpatrick, H. Humphreys,
W. P. Giles, M. E. Rupp, P. D. Fey and J. P. O'Gara (2004). "Genetic and phenotypic analysis of biofilm phenotypic variation in multiple Staphylococcus epidermidis isolates." J Med Microbiol 53: 367-74.
Hausner, M. and S. Wuertz (1999). "High rates of conjugation in bacterial biofilms as determined by quantitative in situ analysis." Appl Environ Microbiol 65: 3710-3.
Haussler, S. (2004). "Biofilm formation by the small colony variant phenotype of
Pseudomonas aeruginosa." Environ Microbiol 6: 546-51.
Haussler, S., I. Ziegler, A. Lottel, F. von Gotz, M. Rohde, D. Wehmhohner, S.
Saravanamuthu, B. Tummler and I. Steinmetz (2003). "Highly adherent small-colony variants of Pseudomonas aeruginosa in cystic fibrosis lung infection." J Med Microbiol
52: 295-301.
Hecht, G. B. and A. Newton (1995). "Identification of a novel response regulator required for the swarmer-to-stalked-cell transition in Caulobacter crescentus." J
Bacteriol 177: 6223-9.
Hendrickx, L., M. Hausner and S. Wuertz (2003). "Natural genetic transformation in monoculture Acinetobacter sp. strain BD413 biofilms." Appl Environ Microbiol 69:
1721-7.
171 REFERENCES
Hernandez-Romero, D., P. Lucas-Elio, D. Lopez-Serrano, F. Solano and A. Sanchez-
Amat (2003). "Marinomonas mediterranea is a lysogenic bacterium that synthesizes R- bodies." Microbiology 149: 2679-86.
Heydorn, A., A. T. Nielsen, M. Hentzer, C. Sternberg, M. Givskov, B. K. Ersboll and S.
Molin (2000). "Quantification of biofilm structures by the novel computer program
COMSTAT." Microbiology 146: 2395-407.
Hickman, J. W., D. F. Tifrea and C. S. Harwood (2005). "A chemosensory system that regulates biofilm formation through modulation of cyclic diguanylate levels." Proc Natl
Acad Sci U S A 102: 14422-7.
Hinsa, S. M., M. Espinosa-Urgel, J. L. Ramos and G. A. O'Toole (2003). "Transition from reversible to irreversible attachment during biofilm formation by Pseudomonas fluorescens WCS365 requires an ABC transporter and a large secreted protein." Mol
Microbiol 49: 905-18.
Holmström, C., S. James, B. A. Neilan, D. C. White and S. Kjelleberg (1998).
"Pseudoalteromonas tunicata sp. nov., a bacterium that produces antifouling agents."
Int J Syst Bacteriol 48: 1205-12.
Holmström, C. and S. Kjelleberg (1999). "Marine Pseudoalteromonas species are associated with higher organisms and produce biologically active extracellular agents."
FEMS Microbiol Ecol 30: 285-293.
Holmström, C., D. Rittschoff and S. Kjelleberg (1992). "Inhibition of attachment of larval barnacles, Balanus amphitrite and Ciona intestinales by a surface colonizing marine bacterium." Appl Environ Microbiol 58: 2111-2115.
172 REFERENCES
Hope, C. K., D. Clements and M. Wilson (2002). "Determining the spatial distribution of viable and nonviable bacteria in hydrated microcosm dental plaques by viability profiling." J Appl Microbiol 93: 448-55.
Hoyoux, A., I. Jennes, P. Dubois, S. Genicot, F. Dubail, J. M. Francois, E. Baise, G.
Feller and C. Gerday (2001). "Cold-adapted beta-galactosidase from the Antarctic psychrophile Pseudoalteromonas haloplanktis." Appl Environ Microbiol 67: 1529-35.
Huber, B., K. Riedel, M. Hentzer, A. Heydorn, A. Gotschlich, M. Givskov, S. Molin and L. Eberl (2001). "The cep quorum-sensing system of Burkholderia cepacia H111 controls biofilm formation and swarming motility." Microbiology 147: 2517-28.
Hunt, S. M., E. M. Werner, B. Huang, M. A. Hamilton and P. S. Stewart (2004).
"Hypothesis for the role of nutrient starvation in biofilm detachment." Appl Environ
Microbiol 70: 7418-25.
Inglis, T. J. (1993). "Evidence for dynamic phenomena in residual tracheal tube biofilm." Br J Anaesth 70: 22-4.
Isnansetyo, A. and Y. Kamei (2003). "MC21-A, a bactericidal antibiotic produced by a new marine bacterium, Pseudoalteromonas phenolica sp. nov. O-BC30(T), against methicillin-resistant Staphylococcus aureus." Antimicrob Agents Chemother 47: 480-8.
Ivanova, E. P., I. Y. Bakunina, O. I. Nedashkovskaya, N. M. Gorshkova, Y. V.
Alexeeva, E. A. Zelepuga, T. N. Zvaygintseva, D. V. Nicolau and V. V. Mikhailov
(2003). "Ecophysiological variabilities in ectohydrolytic enzyme activities of some
Pseudoalteromonas species, P. citrea, P. issachenkonii, and P. nigrifaciens." Curr
Microbiol 46: 6-10.
173 REFERENCES
Ivanova, E. P., S. Flavier and C. Richard (2004). "Phylogenetic relationships among marine Alteromonas-like proteobacteria: emended description of the family
Alteromonadaceae and proposal of Pseudoalteromonadaceae fam. nov., Colwelliaceae fam. nov., Shewanellaceae fam. nov., Moritellaceae fam. nov., Ferrimonadaceae fam. nov., Idiomarinaceae fam. nov. and Psychromonadaceae fam. nov.
." Int J Syst Evol Microbiol 54: 1773-1788.
Ivanova, E. P., E. A. Kiprianova, V. V. Mikhailov, G. F. Levanova, A. D. Garagulya, N.
M. Gorshkova, M. V. Vysotskii, D. V. Nicolau, N. Yumoto, T. Taguchi and S.
Yoshikawa (1998). "Phenotypic diversity of Pseudoalteromonas citrea from different marine habitats and emendation of the description." Int J Syst Bacteriol 48 247-56.
Ivanova, E. P., G. R. Matte, M. H. Matte, T. Coenye, A. Huq and R. R. Colwell
(2002a). "Characterization of Pseudoalteromonas citrea and P. nigrifaciens isolated from different ecological habitats based on REP-PCR genomic fingerprints." Syst Appl
Microbiol 25: 275-83.
Ivanova, E. P., T. Sawabe, Y. V. Alexeeva, A. M. Lysenko, N. M. Gorshkova, K.
Hayashi, N. V. Zukova, R. Christen and V. V. Mikhailov (2002b). "Pseudoalteromonas issachenkonii sp. nov., a bacterium that degrades the thallus of the brown alga Fucus evanescens." Int J Syst Evol Microbiol 52: 229-34.
Ivanova, E. P., T. Sawabe, A. M. Lysenko, N. M. Gorshkova, K. Hayashi, N. V.
Zhukova, D. V. Nicolau, R. Christen and V. V. Mikhailov (2002c).
"Pseudoalteromonas translucida sp. nov. and Pseudoalteromonas paragorgicola sp. nov., and emended description of the genus." Int J Syst Evol Microbiol 52: 1759-66.
174 REFERENCES
Ivanova, E. P., L. S. Shevchenko, T. Sawabe, A. M. Lysenko, V. I. Svetashev, N. M.
Gorshkova, M. Satomi, R. Christen and V. V. Mikhailov (2002d). "Pseudoalteromonas maricaloris sp. nov., isolated from an Australian sponge, and reclassification of
[Pseudoalteromonas aurantia] NCIMB 2033 as Pseudoalteromonas flavipulchra sp. nov." Int J Syst Evol Microbiol 52: 263-71.
Jackson, D. W., K. Suzuki, L. Oakford, J. W. Simecka, M. E. Hart and T. Romeo
(2002). "Biofilm formation and dispersal under the influence of the global regulator
CsrA of Escherichia coli." J Bacteriol 184: 290-301.
James, S. G., C. Holmström and S. Kjelleberg (1996). "Purification and characterization of a novel antibacterial protein from the marine bacterium D2." Appl Environ Microbiol
62: 2783-8.
Jelsbak, L. and L. Sogaard-Andersen (2000). "Pattern formation: fruiting body morphogenesis in Myxococcus xanthus." Curr Opin Microbiol 3: 637-42.
Jensen, P. R. and W. Fenical (1994). "Strategies for the discovery of secondary metabolites from marine bacteria: ecological perspectives." Annu Rev Microbiol 48:
559-84.
Kalinovskaya, N. I., E. P. Ivanova, Y. V. Alexeeva, N. M. Gorshkova, T. A.
Kuznetsova, A. S. Dmitrenok and D. V. Nicolau (2004). "Low-molecular-weight, biologically active compounds from marine Pseudoalteromonas species." Curr
Microbiol 48: 441-6.
Kaplan, J. B., M. F. Meyenhofer and D. H. Fine (2003a). "Biofilm growth and detachment of Actinobacillus actinomycetemcomitans." J Bacteriol 185: 1399-404.
175 REFERENCES
Kaplan, J. B., C. Ragunath, N. Ramasubbu and D. H. Fine (2003b). "Detachment of
Actinobacillus actinomycetemcomitans biofilm cells by an endogenous beta- hexosaminidase activity." J Bacteriol 185: 4693-8.
Kaplan, J. B., C. Ragunath, K. Velliyagounder, D. H. Fine and N. Ramasubbu (2004).
"Enzymatic detachment of Staphylococcus epidermidis biofilms." Antimicrob Agents
Chemother 48: 2633-6.
Kerr, J. F., A. H. Wyllie and A. R. Currie (1972). "Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics." Br J Cancer 26: 239-57.
Kirillina, O., J. D. Fetherston, A. G. Bobrov, J. Abney and R. D. Perry (2004). "HmsP, a putative phosphodiesterase, and HmsT, a putative diguanylate cyclase, control Hms- dependent biofilm formation in Yersinia pestis." Mol Microbiol 54: 75-88.
Kirisits, M. J., L. Prost, M. Starkey and M. R. Parsek (2005). "Characterization of colony morphology variants isolated from Pseudomonas aeruginosa biofilms." Appl
Environ Microbiol 71: 4809-21.
Kisdi, É. (2002). "Dispersal: Risk spreading versus local adaptation." Am Nat 159: 579-
596.
Kjelleberg, S. and S. Molin (2002). "Is there a role for quorum sensing signals in bacterial biofilms?" Curr Opin Microbiol 5: 254-8.
Klausen, M., A. Aaes-Jorgensen, S. Molin and T. Tolker-Nielsen (2003). "Involvement of bacterial migration in the development of complex multicellular structures in
Pseudomonas aeruginosa biofilms." Mol Microbiol 50: 61-8.
Kolter, R. (2005). "Surfacing views of biofilm biology." Trends Microbiol 13: 1-2.
176 REFERENCES
Kon-ya, K., N. Shimidzu, N. Otaki, A. Yokoyama, K. Adachi and W. Miki (1995).
"Inhibitory effect of bacterial ubiquinones on the settling of barnacle, Balanus amphitrite." Experientia 51: 153-155.
Kong, K. F., C. Vuong and M. Otto (2006). "Staphylococcus quorum sensing in biofilm formation and infection." Int J Med Microbiol 296: 133-9.
Korinek, J. (1927). "Ein Beitrag zur Mikrobiologie des Meeres." Zentr Bakt Parasitenk
71: 73-79.
Kreft, J. U., C. Picioreanu, J. W. Wimpenny and M. C. van Loosdrecht (2001).
"Individual-based modelling of biofilms." Microbiology 147: 2897-912.
Kusakabe, H., K. Kodama, A. Kuninaka, H. Yoshino and K. Soda (1979).
"Antibacterial activity of L-Lysine Į-Oxidase against rec+ and rec- Strains of Bacillus subtilis." Agr Biol Chem 43: 1371-1373.
Labbate, M., S. Y. Queck, K. S. Koh, S. A. Rice, M. Givskov and S. Kjelleberg (2004).
"Quorum sensing-controlled biofilm development in Serratia liquefaciens MG1." J
Bacteriol 186: 692-8.
Laemmli, U. K. (1970). "Cleavage of structural proteins during the assembly of the head of bacteriophage T4." Nature 227: 680-5.
Lappin-Scott, H. M. and J. W. Costerton (1989). "Bacterial biofilms and surface fouling
" Biofouling 1: 323-342.
177 REFERENCES
Lawrence, J. R., M. R. Chenier, R. Roy, D. Beaumier, N. Fortin, G. D. Swerhone, T. R.
Neu and C. W. Greer (2004). "Microscale and molecular assessment of impacts of nickel, nutrients, and oxygen level on structure and function of river biofilm communities." Appl Environ Microbiol 70: 4326-39.
Lawrence, J. R., D. R. Korber, B. D. Hoyle, J. W. Costerton and D. E. Caldwell (1991).
"Optical sectioning of microbial biofilms." J Bacteriol 173: 6558-67.
Lee, S. F., Y. H. Li and G. H. Bowden (1996). "Detachment of Streptococcus mutans biofilm cells by an endogenous enzymatic activity." Infect Immun 64: 1035-8.
Leon, L. L., C. C. Miranda, A. O. De Souza and N. Duran (2001). "Antileishmanial activity of the violacein extracted from Chromobacterium violaceum." J Antimicrob
Chemother 48: 449-50.
Lewis, K. (2000). "Programmed death in bacteria." Microbiol Mol Biol Rev 64: 503-14.
Li, Y. H., N. Tang, M. B. Aspiras, P. C. Lau, J. H. Lee, R. P. Ellen and D. G.
Cvitkovitch (2002). "A quorum-sensing signaling system essential for genetic competence in Streptococcus mutans is involved in biofilm formation." J Bacteriol 184:
2699-708.
Lichstein, H. C. and V. F. Van de Sand (1945). "Violacein, an antibiotic pigment produced by Chromobacterium violaceum." J Infect Dis 76.
Lovejoy, C., J. P. Bowman and G. M. Hallegraeff (1998). "Algicidal effects of a novel marine Pseudoalteromonas isolate (class Proteobacteria, gamma subdivision) on harmful algal bloom species of the genera Chattonella, Gymnodinium, and
Heterosigma." Appl Environ Microbiol 64: 2806-13.
178 REFERENCES
Lucas-Elío, P., D. Gómez, F. Solano and A. Sanchez-Amat (2006). "The antimicrobial activity of marinocine, synthesized by Marinomonas mediterranea, is due to the hydrogen peroxide generated by its lysine oxidase activity." J Bacteriol 188: 2493-2501.
Lucas-Elio, P., P. Hernandez, A. Sanchez-Amat and F. Solano (2005). "Purification and partial characterization of marinocine, a new broad-spectrum antibacterial protein produced by Marinomonas mediterranea." Biochim Biophys Acta 1721: 193-203.
Lukasheva, E. V. and T. T. Berezov (2002). "L-Lysine alpha-oxidase: physicochemical and biological properties." Biochemistry (Mosc) 67: 1152-8.
Lux, R., Y. Li, A. Lu and W. Shi (2004). "Detailed three-dimensional analysis of structural features of Myxococcus xanthus fruiting bodies using confocal laser scanning microscopy." Biofilms 1: 293-303.
Lynch, M. J., S. Swift, D. F. Kirke, C. W. Keevil, C. E. Dodd and P. Williams (2002).
"The regulation of biofilm development by quorum sensing in Aeromonas hydrophila."
Environ Microbiol 4: 18-28.
Mai-Prochnow, A. (2002). Biofilm development of Pseudoalteromonas tunicata - role of the antibacterial protein. School of Biotechnology and Bio-Molecular Sciences.
Sydney, University of New South Wales and Universitaet Rostock. Diplom Biologie:
78.
Mai-Prochnow, A., F. Evans, D. Dalisay-Saludes, S. Stelzer, S. Egan, S. James, J. S.
Webb and S. Kjelleberg (2004). "Biofilm development and cell death in the marine bacterium Pseudoalteromonas tunicata." Appl Environ Microbiol 70: 3232-8.
179 REFERENCES
Mai-Prochnow, A., J. S. Webb, B. C. Ferrari and S. Kjelleberg (2006). "Ecological advantages of autolysis during the development and dispersal of Pseudoalteromonas tunicata biofilms." Appl Environ Microbiol 72: in press.
Marden, P., A. Tunlid, K. Malmcrona-Friberg, G. Odham and S. Kjelleberg (1985).
"Physiological and morphological changes during short term starvation of marine bacterial isolates." Arch Microbiol 142: 326-332.
Marshall, D. J., T. F. Bolton and M. J. Keough (2003). "Offspring size affects the post- metamorphic performance of a colonial marine invertebrate." Ecology 84: 3131-3137.
Marshall, D. J. and M. J. Keough (2003). "Variation in the dispersal potential of non- feeding larvae: the desperate larva hypothesis and larval size." Marine Ecology Progress
Series 255: 145-153.
Matz, C., P. Deines, J. Boenigk, H. Arndt, L. Eberl, S. Kjelleberg and K. Jurgens
(2004). "Impact of violacein-producing bacteria on survival and feeding of bacterivorous nanoflagellates." Appl Environ Microbiol 70: 1593-9.
Matz, C., D. McDougald, A. M. Moreno, P. Y. Yung, F. H. Yildiz and S. Kjelleberg
(2005). "Biofilm formation and phenotypic variation enhance predation-driven persistence of Vibrio cholerae." Proc Natl Acad Sci U S A 102: 16819-24.
McCann, K. S. (2000). "The diversity-stability debate." Nature 405: 228-33.
McCarthy, S. A., R. M. Johnson and D. Kakimoto (1994). "Characterization of an antibiotic produced by Alteromonas luteoviolacea Gauthier 1982, 85 isolated from
Kinko Bay, Japan." J Appl Bacteriol 77: 426-32.
180 REFERENCES
McLean, R. J., M. Whiteley, D. J. Stickler and W. C. Fuqua (1997). "Evidence of autoinducer activity in naturally occurring biofilms." FEMS Microbiol Lett 154: 259-
63.
Medigue, C., E. Krin, G. Pascal, V. Barbe, A. Bernsel, P. N. Bertin, F. Cheung, S.
Cruveiller, S. D'Amico, A. Duilio, G. Fang, G. Feller, C. Ho, S. Mangenot, G. Marino,
J. Nilsson, E. Parrilli, E. P. Rocha, Z. Rouy, A. Sekowska, M. L. Tutino, D. Vallenet, G. von Heijne and A. Danchin (2005). "Coping with cold: the genome of the versatile marine Antarctica bacterium Pseudoalteromonas haloplanktis TAC125." Genome Res
15: 1325-35.
Mendez, C., A. F. Brana, M. B. Manzanal and C. Hardisson (1985). "Role of substrate mycelium in colony development in Streptomyces." Can J Microbiol 31: 446-50.
Meyer, R. L., A. M. Saunders, R. J. Zeng, J. Keller and L. L. Blackall (2003).
"Microscale structure and function of anaerobic-aerobic granules containing glycogen accumulating organisms." FEMS Microbiol Ecol 45: 253-261.
Miguelez, E. M., C. Hardisson and M. B. Manzanal (1999). "Hyphal death during colony development in Streptomyces antibioticus: morphological evidence for the existence of a process of cell deletion in a multicellular prokaryote." J Cell Biol 145:
515-25.
Miller, J. H. (1972) Experiments in Molecular Genetics. Cold Spring Habour
Laboratory Press, New York.
Miller, V. L. and J. J. Mekalanos (1988). "A novel suicide vector and its use in construction of insertion mutations: osmoregulation of outer membrane proteins and virulence determinants in Vibrio cholerae requires toxR." J Bacteriol 170: 2575-83.
181 REFERENCES
Mitrakul, K., C. Y. Loo, C. V. Hughes and N. Ganeshkumar (2004). "Role of a
Streptococcus gordonii copper-transport operon, copYAZ, in biofilm detachment." Oral
Microbiol Immunol 19: 395-402.
Molin, S. and T. Tolker-Nielsen (2003). "Gene transfer occurs with enhanced efficiency in biofilms and induces enhanced stabilisation of the biofilm structure." Curr Opin
Biotechnol 14: 255-61.
Moller, S., C. Sternberg, J. B. Andersen, B. B. Christensen, J. L. Ramos, M. Givskov and S. Molin (1998). "In situ gene expression in mixed-culture biofilms: evidence of metabolic interactions between community members." Appl Environ Microbiol 64: 721-
32.
Monk, I. R., G. M. Cook, B. C. Monk and P. J. Bremer (2004). "Morphotypic conversion in Listeria monocytogenes biofilm formation: biological significance of rough colony isolates." Appl Environ Microbiol 70: 6686-94.
Moran, A. L. and R. B. Emlet (2001). "Offspring size and performance in variable environments: field studies on a marine snail." Ecology 82: 1597-1612.
Nandakumar, K., H. Obika, T. Shinozaki, T. Ooie, A. Utsumi and T. Yano (2003).
"Laser impact assessment in a biofilm-forming bacterium Pseudoalteromonas carrageenovora using a flow cytometric system." Biotechnol Bioeng 82: 399-402.
Nawrocki, A., M. R. Larsen, A. V. Podtelejnikov, O. N. Jensen, M. Mann, P.
Roepstorff, A. Gorg, S. J. Fey and P. M. Larsen (1998). "Correlation of acidic and basic carrier ampholyte and immobilized pH gradient two-dimensional gel electrophoresis patterns based on mass spectrometric protein identification." Electrophoresis 19: 1024-
35.
182 REFERENCES
Nedashkovskaia, O. I., E. P. Ivanova, I. Bakunina, V. I. Svetashev, T. N. Zviagintseva and V. V. Mikhailov (2002). "[Characterization of marine bacteria Pseudoalteromonas citrea degradating fucoidan]." Mikrobiol Z 64: 3-10.
Neidhardt, F. C., P. L. Bloch and D. F. Smith (1974). "Culture medium for enterobacteria." J Bacteriol 119: 736-747.
Nierman, W. C., T. V. Feldblyum, M. T. Laub, I. T. Paulsen, K. E. Nelson, J. A. Eisen,
J. F. Heidelberg, M. R. Alley, N. Ohta, J. R. Maddock, I. Potocka, W. C. Nelson, A.
Newton, C. Stephens, N. D. Phadke, B. Ely, R. T. DeBoy, R. J. Dodson, A. S. Durkin,
M. L. Gwinn, D. H. Haft, J. F. Kolonay, J. Smit, M. B. Craven, H. Khouri, J. Shetty, K.
Berry, T. Utterback, K. Tran, A. Wolf, J. Vamathevan, M. Ermolaeva, O. White, S. L.
Salzberg, J. C. Venter, L. Shapiro and C. M. Fraser (2001). "Complete genome sequence of Caulobacter crescentus." Proc Natl Acad Sci U S A 98: 4136-41.
Nugroho, F. A., H. Yamamoto, Y. Kobayashi and J. Sekiguchi (1999).
"Characterization of a new sigma-K-dependent peptidoglycan hydrolase gene that plays a role in Bacillus subtilis mother cell lysis." J Bacteriol 181: 6230-7.
O'Toole, G. (2002). "A resistance switch." Nature 416: 695-696.
O'Toole, G., H. B. Kaplan and R. Kolter (2000). "Biofilm formation as microbial development." Annu Rev Microbiol 54: 49-79.
O'Toole, G. A. and R. Kolter (1998). "Flagellar and twitching motility are necessary for
Pseudomonas aeruginosa biofilm development." Mol Microbiol 30: 295-304.
183 REFERENCES
Ochsner, U. A., A. K. Koch, A. Fiechter and J. Reiser (1994). "Isolation and characterization of a regulatory gene affecting rhamnolipid biosurfactant synthesis in
Pseudomonas aeruginosa." J Bacteriol 176: 2044-54.
Oliveira, D. R. (1992). Physico-chemical aspects of adhesion, p. 45-48. In L. F. Melo,
T. R. Bott, M. Fletcher and B. Capdeville (ed.), Biofilms – Science and Technology.
Kluwer Academic Publishers, London
Oliver, A., R. Canton, P. Campo, F. Baquero and J. Blazquez (2000). "High frequency of hypermutable Pseudomonas aeruginosa in cystic fibrosis lung infection." Science
288: 1251-4.
Passador, L., J. M. Cook, M. J. Gambello, L. Rust and B. H. Iglewski (1993).
"Expression of Pseudomonas aeruginosa virulence genes requires cell-to-cell communication." Science 260: 1127-30.
Picioreanu, C., M. C. van Loosdrecht and J. J. Heijnen (2001). "Two-dimensional model of biofilm detachment caused by internal stress from liquid flow." Biotechnol
Bioeng 72: 205-18.
Piggot, P. J. and J. G. Coote (1976). "Genetic aspects of bacterial endospore formation."
Bacteriol Rev 40: 908-62.
Piriou, P., S. Dukan, Y. Levi and P. A. Jarrige (1997). "Prevention of bacterial growth in drinking water distribution systems." Water Sci Technol 35: 283-287.
Plamann, L., Y. Li, B. Cantwell and J. Mayor (1995). "The Myxococcus xanthus asgA gene encodes a novel signal transduction protein required for multicellular development." J Bacteriol 177: 2014-20.
184 REFERENCES
Prigent-Combaret, C., O. Vidal, C. Dorel and P. Lejeune (1999). "Abiotic surface sensing and biofilm-dependent regulation of gene expression in Escherichia coli." J
Bacteriol 181: 5993-6002.
Proctor, R. A., P. van Langevelde, M. Kristjansson, J. N. Maslow and R. D. Arbeit
(1995). "Persistent and relapsing infections associated with small-colony variants of
Staphylococcus aureus." Clin Infect Dis 20: 95-102.
Purevdorj, B., J. W. Costerton and P. Stoodley (2002). "Influence of hydrodynamics and cell signaling on the structure and behavior of Pseudomonas aeruginosa biofilms." Appl
Environ Microbiol 68: 4457-64.
Putnins, E. E., D. Di Giovanni and A. S. Bhullar (2001). "Dental unit waterline contamination and its possible implications during periodontal surgery." J Periodontol
72: 393-400.
Rainey, P. B. and M. Travisano (1998). "Adaptive radiation in a heterogeneous environment." Nature 394: 69-72.
Ramsey, M. M. and M. Whiteley (2004). "Pseudomonas aeruginosa attachment and biofilm development in dynamic environments." Mol Microbiol 53: 1075-87.
Reisner, A., J. A. Haagensen, M. A. Schembri, E. L. Zechner and S. Molin (2003).
"Development and maturation of Escherichia coli K-12 biofilms." Mol Microbiol 48:
933-46.
Rice, S. A., K. S. Koh, S. Y. Queck, M. Labbate, K. W. Lam and S. Kjelleberg (2005).
"Biofilm formation and sloughing in Serratia marcescens are controlled by quorum sensing and nutrient cues." J Bacteriol 187: 3477-85.
185 REFERENCES
Romling, U. and D. Amikam (2006). "Cyclic di-GMP as a second messenger." Curr
Opin Microbiol 9: 218-28.
Romling, U., M. Gomelsky and M. Y. Galperin (2005). "C-di-GMP: the dawning of a novel bacterial signalling system." Mol Microbiol 57: 629-39.
Rosenbluh, A. and E. Rosenberg (1990). "Role of autocide AMI in development of
Myxococcus xanthus." J Bacteriol 172: 4307-14.
Rosenfeld, D. W. and C. E. Zobell (1947). "Antibiotic production by marine microorganisms." J Bacteriol 54: 393-398.
Ross, P., H. Weinhouse, Y. Aloni, D. Michaeli, P. Weinberger-Ohana, R. Mayer, S.
Brann, E. De Vroom, G. Van der Marel, J. Van Boom and M. Benziman (1987).
"Regulation of cellulose synthesis in Acetobacter xylinum by cyclic diguanylic acid."
Nature 325: 279–281.
Ryu, J. H., H. Kim and L. R. Beuchat (2004). "Attachment and biofilm formation by
Escherichia coli O157:H7 on stainless steel as influenced by exopolysaccharide production, nutrient availability, and temperature." J Food Prot 67: 2123-31.
Sadowska, B., A. Bonar, C. von Eiff, R. A. Proctor, M. Chmiela, W. Rudnicka and B.
Rozalska (2002). "Characteristics of Staphylococcus aureus, isolated from airways of cystic fibrosis patients, and their small colony variants." FEMS Immunol Med
Microbiol 32: 191-7.
Sager, B. and D. Kaiser (1993). "Two cell-density domains within the Myxococcus xanthus fruiting body." Proc Natl Acad Sci U S A 90: 3690-4.
186 REFERENCES
Sauer, K., A. K. Camper, G. D. Ehrlich, J. W. Costerton and D. G. Davies (2002).
"Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm." J Bacteriol 184: 1140-54.
Sauer, K., M. C. Cullen, A. H. Rickard, L. A. Zeef, D. G. Davies and P. Gilbert (2004).
"Characterization of nutrient-induced dispersion in Pseudomonas aeruginosa PAO1 biofilm." J Bacteriol 186: 7312-26.
Sawabe, T., H. Makino, M. Tatsumi, K. Nakano, K. Tajima, M. M. Iqbal, I. Yumoto, Y.
Ezura and R. Christen (1998). "Pseudoalteromonas bacteriolytica sp. nov., a marine bacterium that is the causative agent of red spot disease of Laminaria japonica." Int J
Syst Bacteriol 48 769-74.
Schembri, M. A., K. Kjaergaard and P. Klemm (2003). "Global gene expression in
Escherichia coli biofilms." Mol Microbiol 48: 253-67.
Schroeder, D. C., M. A. Jaffer and V. E. Coyne (2003). "Investigation of the role of a beta(1-4) agarase produced by Pseudoalteromonas gracilis B9 in eliciting disease symptoms in the red alga Gracilaria gracilis." Microbiology 149: 2919-29.
Shimkets, L. J. (1990). "Social and developmental biology of the myxobacteria."
Microbiol Rev 54: 473-501.
Shimkets, L. J. (1999). "Intercellular signaling during fruiting-body development of
Myxococcus xanthus." Annu Rev Microbiol 53: 525-49.
187 REFERENCES
Silipo, A., R. Lanzetta, M. Parrilli, L. Sturiale, D. Garozzo, E. L. Nazarenko, R. P.
Gorshkova, E. P. Ivanova and A. Molinaro (2005). "The complete structure of the core carbohydrate backbone from the LPS of marine halophilic bacterium
Pseudoalteromonas carrageenovora type strain IAM 12662T." Carbohydr Res 340:
1475-82.
Simidu, U., K. Kita-Tsukamoto, T. Yasumoto and M. Yotsu (1990). "Taxonomy of four marine bacterial strains that produce tetrodotoxin." Int J Syst Bacteriol 40: 331-6.
Simm, R., M. Morr, A. Kader, M. Nimtz and U. Romling (2004). "GGDEF and EAL domains inversely regulate cyclic di-GMP levels and transition from sessility to motility." Mol Microbiol 53: 1123-34.
Singh, P. K., A. L. Schaefer, M. R. Parsek, T. O. Moninger, M. J. Welsh and E. P.
Greenberg (2000). "Quorum-sensing signals indicate that cystic fibrosis lungs are infected with bacterial biofilms." Nature 407: 762-4.
Skovhus, T. L., N. B. Ramsing, C. Holmström, S. Kjelleberg and I. Dahllof (2004).
"Real-time quantitative PCR for assessment of abundance of Pseudoalteromonas species in marine samples." Appl Environ Microbiol 70: 2373-82.
Smith, T. J. and S. J. Foster (1995). "Characterization of the involvement of two compensatory autolysins in mother cell lysis during sporulation of Bacillus subtilis
168." J Bacteriol 177: 3855-62.
188 REFERENCES
Sobolevskaya, M. P., O. F. Smetanina, M. Speitling, L. S. Shevchenko, P. S.
Dmitrenok, H. Laatsch, T. A. Kuznetsova, E. P. Ivanova and G. B. Elyakov (2004).
"Controlling production of brominated cyclic Depsipeptides by Pseudoalteromonas maricaloris KMM 636T." Letters in Applied Microbiology 40: 243-248.
Solano, F., E. Garcia, E. Perez De Egea and A. Sanchez-Amat (1997). "Isolation and
Characterization of Strain MMB-1 (CECT 4803), a Novel Melanogenic Marine
Bacterium " Appl Environ Microbiol 63: 3499-3506.
Sonenshein, A. (2000). Endospore-forming bacteria: an overview, p. 133-150. In Y. V.
Brun and L. J. Shimkets (ed.), Prokaryotic Development. ASM Press, Washington DC
Southey-Pillig, C. J., D. G. Davies and K. Sauer (2005). "Characterization of temporal protein production in Pseudomonas aeruginosa biofilms." J Bacteriol 187: 8114-26.
Stanley, N. R., R. A. Britton, A. D. Grossman and B. A. Lazazzera (2003).
"Identification of catabolite repression as a physiological regulator of biofilm formation by Bacillus subtilis by use of DNA microarrays." J Bacteriol 185: 1951-7.
Stapper, A. P., G. Narasimhan, D. E. Ohman, J. Barakat, M. Hentzer, S. Molin, A.
Kharazmi, N. Hoiby and K. Mathee (2004). "Alginate production affects Pseudomonas aeruginosa biofilm development and architecture, but is not essential for biofilm formation." J Med Microbiol 53: 679-90.
Stelzer, S. (1999). Investigation of the antibacterial protein produced by
Pseudoalteromonas tunicata. School of Microbiology. Sydney, University of New
South Wales. Bachelor of Science (Honours).
189 REFERENCES
Stelzer, S., S. Egan, M. R. Larsen, D. H. Bartlett and S. Kjelleberg (2006). "Unravelling the role of the ToxR-like transcriptional regulator WmpR in the marine antifouling bacterium Pseudoalteromonas tunicata." Microbiology 152: 1385-94.
Sternberg, C., B. B. Christensen, T. Johansen, A. Toftgaard Nielsen, J. B. Andersen, M.
Givskov and S. Molin (1999). "Distribution of bacterial growth activity in flow- chamber biofilms." Appl Environ Microbiol 65: 4108-17.
Stoodley, P., R. Cargo, C. J. Rupp, S. Wilson and I. Klapper (2002). "Biofilm material properties as related to shear-induced deformation and detachment phenomena." J Ind
Microbiol Biotechnol 29: 361-7.
Stoodley, P., I. Dodds, J. D. Boyle and H. M. Lappin-Scott (1999). "Influence of hydrodynamics and nutrients on biofilm structure." J Appl Microbiol 85 19S-28S.
Stoodley, P., S. Wilson, L. Hall-Stoodley, J. D. Boyle, H. M. Lappin-Scott and J. W.
Costerton (2001). "Growth and detachment of cell clusters from mature mixed-species biofilms." Appl Environ Microbiol 67: 5608-13.
Stove, J. L. and R. Y. Stanier (1962). "Cellular differentiation in stalked bacteria."
Nature 196: 1189-1192.
Sutherland, I. W. (1985). "Biosynthesis and composition of gram-negative bacterial extracellular and wall polysaccharides." Annu Rev Microbiol 39: 243-70.
Swift, S., J. A. Downie, N. A. Whitehead, A. M. Barnard, G. P. Salmond and P.
Williams (2001). "Quorum sensing as a population-density-dependent determinant of bacterial physiology." Adv Microb Physiol 45: 199-270.
190 REFERENCES
Tal, R., H. C. Wong, R. Calhoon, D. Gelfand, A. L. Fear, G. Volman, R. Mayer, P.
Ross, D. Amikam, H. Weinhouse, A. Cohen, S. Sapir, P. Ohana and M. Benziman
(1998). "Three cdg operons control cellular turnover of cyclic di-GMP in Acetobacter xylinum: genetic organization and occurrence of conserved domains in isoenzymes." J
Bacteriol 180: 4416-25.
Tenke, P., B. Kovacs, M. Jackel and E. Nagy (2006). "The role of biofilm infection in urology." World J Urol 24: 13-20.
Thomas, C., C. R. Andersson, S. R. Canales and S. S. Golden (2004). "PsfR, a factor that stimulates psbAI expression in the cyanobacterium Synechococcus elongatus PCC
7942." Microbiology 150: 1031-40.
Thomas, R. W. S. P. and D. Allsopp (1983). "The effects of certain periphytic marine bacteria upon the settlement and growth of Enteromorpha, a fouling alga."
Biodeterioration 5: 348-357.
Thormann, K. M., S. Duttler, R. M. Saville, M. Hyodo, S. Shukla, Y. Hayakawa and A.
M. Spormann (2006). "Control of formation and cellular detachment from Shewanella oneidensis MR-1 biofilms by cyclic di-GMP." J Bacteriol 188: 2681-91.
Tischler, A. D. and A. Camilli (2004). "Cyclic diguanylate (c-di-GMP) regulates Vibrio cholerae biofilm formation." Mol Microbiol 53: 857-69.
Tischler, A. D. and A. Camilli (2005). "Cyclic diguanylate regulates Vibrio cholerae virulence gene expression." Infect Immun 73: 5873-82.
191 REFERENCES
Tolker-Nielsen, T., U. C. Brinch, P. C. Ragas, J. B. Andersen, C. S. Jacobsen and S.
Molin (2000). "Development and dynamics of Pseudomonas sp. biofilms." J Bacteriol
182: 6482-9.
Tomlin, K. L., R. J. Malott, G. Ramage, D. G. Storey, P. A. Sokol and H. Ceri (2005).
"Quorum-sensing mutations affect attachment and stability of Burkholderia cenocepacia biofilms." Appl Environ Microbiol 71: 5208-18.
Ueda, H., H. Nakajima, Y. Hori, T. Goto and M. Okuhara (1994). "Action of
FR901228, a novel antitumor bicyclic depsipeptide produced by Chromobacterium violaceum no. 968, on Ha-ras transformed NIH3T3 cells." Biosci Biotechnol Biochem
58: 1579-83. van den Broek, D., A. W. T. F. Chin, K. Eijkemans, I. H. Mulders, G. V. Bloemberg and B. J. Lugtenberg (2003). "Biocontrol traits of Pseudomonas spp. are regulated by phase variation." Mol Plant Microbe Interact 16: 1003-12. van Loodsrecht, M. C. M., J. Lyklema, W. Norde and A. J. B. Zehnder (1990).
"Influence of interfaces on microbial activity." Microbiol Review 54: 75-87. van Loosdrecht, M. C., J. J. Heijnen, H. Eberl, J. Kreft and C. Picioreanu (2002).
"Mathematical modelling of biofilm structures." Antonie Van Leeuwenhoek - Int J of
General and Molec Microbiol 81: 245-56.
Vance, R. E., J. Zhu and J. J. Mekalanos (2003). "A constitutively active variant of the quorum-sensing regulator LuxO affects protease production and biofilm formation in
Vibrio cholerae." Infect Immun 71: 2571-6.
192 REFERENCES
Vasconcelos, A. T. R., D. F. Almeida, M. Hungria, C. T. Guimaraes, R. V. Antonio, F.
C. Almeida, L. G. P. Almeida and R. Almeida (2003). "The complete genome sequence of Chromobacterium violaceum reveals remarkable and exploitable bacterial adaptability." Proc Natl Acad Sci U S A 100: 11660-5.
Violot, S., R. Haser, G. Sonan, D. Georlette, G. Feller and N. Aghajari (2003).
"Expression, purification, crystallization and preliminary X-ray crystallographic studies of a psychrophilic cellulase from Pseudoalteromonas haloplanktis." Acta Crystallogr D
Biol Crystallogr 59: 1256-8.
Vuong, C., C. Gerke, G. A. Somerville, E. R. Fischer and M. Otto (2003). "Quorum- sensing control of biofilm factors in Staphylococcus epidermidis." J Infect Dis 188: 706-
18.
Walker, J. T., C. W. Mackerness, D. Mallon, T. Makin, T. Williets and C. W. Keevil
(1995). "Control of Legionella pneumophila in a hospital water system by chlorine dioxide." J Ind Microbiol 15: 384-90.
Watnick, P. I. and R. Kolter (1999). "Steps in the development of a Vibrio cholerae El
Tor biofilm." Mol Microbiol 34: 586-95.
Watnick, P. I., C. M. Lauriano, K. E. Klose, L. Croal and R. Kolter (2001). "The absence of a flagellum leads to altered colony morphology, biofilm development and virulence in Vibrio cholerae O139." Mol Microbiol 39: 223-35.
Webb, J. S., M. Givskov and S. Kjelleberg (2003a). "Bacterial biofilms: prokaryotic adventures in multicellularity." Curr Opin Microbiol 6: 578-85.
193 REFERENCES
Webb, J. S., M. Lau and S. Kjelleberg (2004). "Bacteriophage and phenotypic variation in Pseudomonas aeruginosa biofilm development." J Bacteriol 186: 8066-73.
Webb, J. S., L. S. Thompson, S. James, T. Charlton, T. Tolker-Nielsen, B. Koch, M.
Givskov and S. Kjelleberg (2003b). "Cell death in Pseudomonas aeruginosa biofilm development." J Bacteriol 185: 4585-92.
Wildermuth, H. (1970). "Development and organization of the aerial mycelium in
Streptomyces coelicolor." J Gen Microbiol 60: 43-50.
Wimpenny, J., W. Manz and U. Szewzyk (2000). "Heterogeneity in biofilms." FEMS
Microbiol Rev 24: 661-71.
Wireman, J. W. and M. Dworkin (1977). "Developmentally induced autolysis during fruiting body formation by Myxococcus xanthus." J Bacteriol 129: 798-802.
Wratten, S. J., M. S. Wolfe, R. J. Andersen and D. J. Faulkner (1976). "Antibiotic
Metabolites from a Marine Pseudomonad." Antimicrob Agents Chemother 11: 411-414.
Xu, L., H. Li, C. Vuong, V. Vadyvaloo, J. Wang, Y. Yao, M. Otto and Q. Gao (2006).
"Role of the luxS quorum-sensing system in biofilm formation and virulence of
Staphylococcus epidermidis." Infect Immun 74: 488-96.
Yaphe, W. and B. Baxter (1955). "The enzymic hydrolysis of carrageenin." Appl.
Microbiol 3: 380-383.
Yoshikawa, K., K. Adachi, F. Nishida and K. Mochida (2003). "Planar structure and antibacterial activity of korormicin derivatives isolated from Pseudoalteromonas sp. F-
420." J Antibiot (Tokyo) 56: 866-70.
194 REFERENCES
Zobell, C. E. and D. Q. Anderson (1936). "Observations on the multiplication of bacteria in different volumes of stored seawater and the influence of oxygen tension and solid surfaces." Biol Bull Woods Hole 71: 324-342.
Zottola, E. A. and K. C. Sasahara (1994). "Microbial biofilms in the food processing industry--should they be a concern?" Int J Food Microbiol 23: 125-48.
195