Antibacterial activity and mechanisms of colonization of Pseudoalteromonas tunicata
Doralyn D. Saludes
A thesis in fulfillment 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
June 2004 U N S W 2 SEP 2005 LIBRARY Table of Contents Acknowledgments...... 7 Abstract...... 10 Certificate of Originality...... 11 List of Figures...... 12 List of Tables...... 15 List of Abbreviations...... 16
1. General introduction and review of related literature...... 18
1.1. Introduction...... 18
1.2. Bacterial secondary metabolites...... 19 1.2.1. Antibacterial agents: sources, classification, spectrum of activity and modes of action...... 21 1.2.2. Role of bacterial metabolites in the biofouling process...... 24
1.3. Bacterial colonization of nonliving and living surfaces...... 25 1.3.1. Bacterial attachment mechanisms...... 25 1.3.1.1. Cell surface structures that mediate attachment...... 25 1.3.2. Bacterial biofilm and microcolony formation...... 31 1.3.2.1. The negative impacts of biofilms...... 32 1.3.3. Biofouling in the marine environment...... 33 1.3.3.1. Settlement and colonization of macrofoulers...... 33 1.3.4. Survival of attached bacterial cells...... 34 1.3.4.1. Programmed cell-death (PCD)...... 35
1.4. The genus Pseudoalteromonas...... 38 1.4.1. Association with marine eukaryotes...... 39 1.4.2. Production of extracellular enzymes...... 39 1.4.3. Production of antibacterial compounds and toxins...... 40 1.4.4. Biological activities of Pseudoalteromonas tunicata...... 42
2 1.5. Aims of this study 43
2. Generation and analysis of Pseudoalteromonas tunicata transposon mutants with altered sensitivity to its autolytic protein, AlpP...... 45
2.1. Introduction...... 45
2.2. Materials and Methods...... 47 2.2.1. Bacterial strains and culture conditions...... 47 2.2.2. Preparation of P. tunicata cell-free concentrated supernatant...... 47 2.2.3. Fractionation of the AlpP protein from cell-free concentrated supernatant 48 2.2.4. Transposon mutagenesis...... 48 2.2.5. Phenotypic characterization of P. tunicata AlpP-sensitive and -resistant transposon mutants...... 49 2.2.5.1. Altered sensitivity to AlpP...... 49 2.2.5.2. Growth curves...... 50 2.2.6. Genotypic characterization of P. tunicata AlpP-sensitive and -resistant transposon mutants...... 50 2.2.6.1. Genomic DNA extractions...... 50 2.2.6.2. Panhandle PCR method for sequencing the regions flanking the inserted transposon...... 51 2.2.7. Sequence data analysis...... 53
2.3. Results...... 54 2.3.1. Generation of P. tunicata transposon mutants...... 54 2.3.2. Phenotypic characteristics of P. tunicata transposon mutants with altered sensitivity to its autolytic protein, AlpP...... 56 2.3.2.1. Analysis of the AlpP resistant and sensitive phenotypes...... 56 2.3.2.2. Growth of both AlpP-sensitive and -resistant transposon mutants...... 58 2.3.3. Genotypic characterization of P. tunicata transposon mutants...... 60 2.3.3.1. DNA sequence analysis of P. tunicata AlpP-sensitive mutants...... 60 2.3.3.2. DNA sequence analysis of the P. tunicata AlpP-resistant mutants .... 71
2.4. Discussion...... 88
3 3. Elucidation of the mode of action of AlpP and the response of Pseudoalteromonas tunicata to its autolytic protein product...... 94
3.1. Introduction...... 94
3.2. Materials and Methods...... 96 3.2.1. Bacterial strains and culture conditions...... 96 3.2.2. Preparation of cell-free concentrated supernatant from P. tunicata wild type, SMI and SM6 Alp-sensitive mutants...... 96 3.2.3. Fractionation of the AlpP protein from the cell-free concentrated supernatant...... 96 3.2.4. Determination of bacteriolytic or bacteriostatic activity of the AlpP protein ...... 97 3.2.5. Assay of inner-membrane permeability in the presence of AlpP protein ... 98 3.2.6. Transmission electron microscopy studies of P. tunicata AlpP protein.....98 3.2.7. Transposon mutagenesis and genetic analysis...... 100 3.2.8. Investigation of the toxin-antidote system in P. tunicata...... 100 3.2.8.1. Identification of a putative antidote activity in active cell-free concentrated P. tunicata supernatant...... 100 3.2.8.2. Fractionation of the cell-free concentrated supernatant to identify a putative antidote factor...... 101
3.3. Results...... 101 3.3.1. Chromogenic plate assay to evaluate bacteriolytic activity of P. tunicata AlpP protein...... 101 3.3.2. Inner-membrane permeability assay...... 103 3.3.3. Transmission electron microscopy studies of P. tunicata AlpP protein ... 104 3.3.4. P. tunicata genes associated with resistance and sensitivity to AlpP.107 3.3.5. Investigation of the toxin-antidote system in P. tunicata...... 109
3.4. Discussion...... 114 3.4.1. The AlpP is bacteriolytic to target cells...... 114 3.4.2. P. tunicata AlpP is not associated with phage or phage tail-like structures...... 115
4 3.4.3. P. tunicata genes encoding traits which mediate altered sensitivity to autolytic protein, AlpP...... 115 3.4.3.1. Transport and signal mechanisms are linked with AlpP activity.....116 3.4.3.2. Autolysis upon AlpP exposure may be mediated by a two-component hybrid sensor regulator...... 117 3.4.3.3. Expression of extracellular structures may be linked with resistance to AlpP activity...... 118 3.4.4. P. tunicata demonstrates mechanisms to prevent self-killing...... 119 3.4.5. Ecological roles of P. tunicata autolytic protein, AlpP...... 120
4. Identification and characterization of a putative mannose-sensitive hemagglutinin (MSHA) pilus biogenesis gene cluster and the role of MSHA pilus in the colonization of living marine surfaces...... 123
4.1. Introduction...... 123
4.2. Materials and Methods...... 125 4.2.1. Bacterial strains, plasmids and culture conditions...... 125 4.2.2. Panhandle PCR, DNA sequencing and sequence analysis...... 125 4.2.3. Transmission and scanning electron microscopy studies...... 126 4.2.4. Hemagglutination assay...... 128 4.2.5. Attachment assays...... 128 4.2.6. Preparation of axenic thallus of the green alga Ulva lactuca...... 129 4.2.7. Attachment assay with axenic Ulva lactuca...... 130
4.3. Results...... 130 4.3.1. DNA sequence analysis of the regions flanking the TnlO insert in the P. tunicata SM5 mutant...... 130 4.3.2. Detection and analysis of cell surface pili...... 141 4.3.3. Assay for agglutination of red blood cells...... 143 4.3.4. Pili promote the attachment of P. tunicata to abiotic and biotic surfaces. 144 4.3.5. Attachment of P. tunicata to the surface of axenic green alga U. lactuca 147
4.4. Discussion...... 151
5 5. The role of a putative cellulosome and its ecological importance in association of Pseudoalteromonas tunicata with higher marine organisms...... 156
5.1. Introduction...... 156
5.2. Materials and Methods...... 158 5.2.1. Bacterial isolates and culture conditions...... 158 5.2.2. Transmission and scanning electron microscopy studies...... 158 5.2.3. Attachment assay...... 159 5.2.4. Growth of P. tunicata in cellobiose or cellulose as the sole carbon source ...... 159 5.2.5. Isolation of bacterial fractions and preparation of cellulose-bound proteins ...... 159 5.2.6. Identification of cellulose-binding proteins by affinity based assay...... 160 5.2.7. Peptide sequencing...... 160 5.2.8. Cellulase activity assay...... 161 5.2.9. SDS-PAGE analysis and the zymogram assay...... 161
5.3. Results...... 162 5.3.1. P. tunicata produces cellulosome-like surface structures...... 162 5.3.2. P. tunicata attaches to cellulose...... 164 5.3.3. Growth of P. tunicata in different substrates...... 165 5.3.4. Cellulose-binding protein facilitates the attachment of P. tunicata to cellulose...... 167 5.3.5. Peptide sequencing of 46-48 kDa protein...... 171 5.3.6. Cellulase activity and zymogram assay...... 171
5.4. Discussion...... 174
6. Summary and General Discussion...... 179 Appendix 1...... 188 Appendix II...... 191 References...... 192
6 Acknowledgments
I owe a particular debt of gratitude to my supervisor Prof. Staffan Kjelleberg. No words of thanks can express my appreciation for all his support and guidance during my PhD studies. His endless encouragement and enthusiasm were heartwarming. Thanks for giving direction for the completion of my project. It is indeed a great privilege to have you as my supervisor and mentor!
I am grateful also for the supervision of Dr. Sally James. Her positive attitude and support towards the project were never failing. Thank you for being there since I started with the project and for being such a wonderful supervisor and friend. It is my pleasure to work with you over the years!
I deeply appreciate the support of Dr. Carola Holmstrom for reviewing the contents of my thesis. Her encouraging comments and suggestions helped me to keep on target. Thank you too for your enthusiasm about the project!
Thanks to Prof. Bill O’Sullivan for his last minute suggestions and thoughtful comments and for helping to provide extra substance to the content of the thesis.
My heartfelt thanks to the brilliant members of D2 group. To Carola, Sally, Su, Jeremy, Ashley, Sacha, Anne, Niina, Dhana, Andre and Flavia. Thanks for sharing your insights and for the technical advices. Thank you too for sharing the VNSS agar plates or NSS when I run out of my stock. It is great to work with you guys! We are the only people who can tolerate the malodorous smell of D2! What more can I say but all the best for D2!
My microscopy imaging skills would not have been developed without the technical expertise of the people from the Electron Microscope Unit, especially to Margaret Budanovic who was so patient in teaching me the skills and to Jenny Norman for her troubleshooting expertise. My thanks also to Luz Paje oif Biorad for teaching me to use the confocal laser scanning microscope. To Sohail aind Ayub of RC lab for their assistance in cellulose degradation project.
Thanks to Adam and Julie for all the administrative assistance and for conveying my messages to Staffan.
To all my friends in the university, who have been so supportive and for bringing joy during my PhD studies. Anne, thank you for being such a special friend and for being there always when I need someone to listen. Your friendship keeps my sanity during long hours of work in the lab. Wish we could have more time for fun and coffee together! Thanks also to Rene (Mr. Anne Mai) for his friendship and for all the help during the submission of this thesis. Johnny Q, you will always remain to be my nastiest friend! I mean it! Thank you for all the fun! You are the best “social organizer” I ever know! Keep up the good work. Joyce, as I always say don’t stop the networking...keep the ball rolling. I can always count on you. Thank you for granting all my favours and for the free coffee bean coated with chocolate! Thanks also to Dhana for being so nice and kind. I also appreciate the friendship of the members of SK and CMBB group especially to Dacre, Mike, Sharon, Megan, Maurice, Lyndal, Evi, Lan, Mathew, Krager, Wendy, Kin and Nidi.
Special thanks to my friends in KT lab, to Claus, Andrew and Jona, who have been so generous and kind for allowing me to use their refrigerated centrifuge and the “top of the line” PAGE device and accessories! Thank you too for the laughter and the friendship.
I deeply appreciate the friendship of my “Peenoy” friends Sarah and Fesca. To you Sarah, thanks for being there when I need your help. Thanks too for all the kindness and generosity you showed during my stay in Sydney. To my “matahum nga amiga” Fesca, truly you are an angel. Wish we could have met ages ago because it is only the two of us in Sydney that can fully understand our “Ilonggo” thing. Thanks for the Friday nights in Ritz cinema and for our weekend walks from Coogee to Bondi. I will surely miss that!
8 To the Australian Agency for International Development and Australian Development Scholarship (AusAID/ADS) for the scholarship grant to study in Australia.
I am grateful to my friends outside the uni who brought me joy through their company. To the wonderful couple, Toni and Edwin, my thanks for your understanding and thoughtfulness. To Jane and Rose, for being there to hear my “science thing” and for keeping me company during the frustrating times at uni.
I am deeply grateful to the Puyat Family of St. Clair for providing me a home in Australia and for making me feel that I belong to their family. Thank you Fr. Ray for being an instrument to know this family. To Tita Elvi and Tito Danny, I don’t know how to thank you for taking care of me when I was in Australia. Thank you for all the prayers and support in the completion of my studies. Thank you too to Daniel, Ronie, Rene, Ramil, Rielle and Bong for being my big brothers!
My heartfelt thanks to the Quinones family of Antelope, California for sharing their home during the writing of my thesis. To Manang Fe and Manong Boy, your generosity and kindness are so overwhelming. No words can express how much I appreciate your help. Thanks you to Billy J “Franchise” for granting all my favours pertaining to the writing of my thesis and for being such a good friend. To Stanley R, Vangie, Kristine, Adam and Gladys for their concern and support.
To my parents, brothers and their families. Thank you for all the prayers, concern, love and support. I know that you are always there for me.
To my wonderful husband Jonel. Having you forever is the best thing that ever happened to me during my PhD studies. Thank you for all your love, support and understanding. You are the fountain of my great joy and inspiration!
Finally, I would like to give thanks and praise to the God Almighty for being the ultimate source of my strength and intellect. Thank you Lord for your constant love and for all the many blessings and graces. What I am now is my gift to you!
9 Abstract
The marine bacterium Pseudoalteromonas tunicata is associated with the surfaces of the tunicate Ciona intestinalis and the green alga Ulva lactuca, where it is proposed to help protect the host organisms from biofouling. This is, in part, believed to be mediated by the production of a 190 kDa antibacterial (autolytic) protein, AlpP. This study is aimed at elucidating the mode of action of AlpP against P. tunicata cells and to investigate mechanisms of colonization and survival on surfaces by/3, tunicata.
The mode of action of the AlpP protein was found to be bacteriolytic and a series of P. tunicata transposon mutants were generated with altered resistance to AlpP exposure. Two membrane transport mechanisms, an ABC (type I) exporter and a type II secretory pathway, appear to be involved in the mode of action of AlpP without directly transporting AlpP. It was hypothesized that these transport mechanisms, in combination with a putative two-component hybrid sensor regulator, may allow AlpP to signal the production of secondary molecules resulting in the autolysis of P. tunicata. Evidence was also found that P. tunicata may protect itself from the effects of AlpP by producing a putative “antidote” molecule.
The colonization of P. tunicata on polystyrene and cellulose surfaces, as well as the green alga Ulva lactuca, is mediated by a putative mannose-sensitive hemagglutinin (MSHA) pilus. A gene cluster termed mshIlI2JKLM in P. tunicata was sequenced and showed a high similarity with the MSHA pilus biogenesis gene locus in Vibrio cholerae. This gene cluster is also most likely responsible for pilus production in this organism. Pilus production in P. tunicata was demonstrated to occur in response to exposure to both cellulose and cellobiose, indicating that this organism has a substrate as well as substratum sensing mechanism linked to pilus expression.
Ultrastructural studies revealed that P. tunicata also produces protuberant-like structures in response to cellulose exposure; similar to the multi-enzyme cellulosome complex reported for cellulolytic Clostridium species. It was demonstrated that the attachment of P. tunicata to cellulose, apart from the MSHA pili, was mediated by a 46-48 kDa membrane bound cellulose-binding protein, possibly associated with the cellulosome.
10 Certificate of Originality
I hereby declare that this submission is my own work and to the best of my knowledge it contains no matrial previously published or written by another person, nor material which to a substantial extent has been accepted for the award of any degree or diploma at UNSW or any other educational institution, except where due acknowledgment is made in the thesis. Any contribution made to the research by others, with whom I have worked at UNSW or elsewhere, is explicitly acknowledged in the thesis.
I also declare that the intellectual content of this thesis is the product of my own work, except to the extent that assistance from others in the project’s design and conception or in style, presentation and linguistic expression is acknowledged.
11 List of Figures
Figure 2-1. Selection of P. tunicata transposon mutant strains from the original template...... 55 Figure 2-2. P. tunicata wild type and AlpP-sensitive transposon mutants (logarithmic growth) displaying different responses to the different dilutions of the autolytic protein, AlpP...... 58 Figure 2-3. Growth curves of P. tunicata mutant strains. A) AlpP-sensitive mutant strains and B) AlpP- resistant mutant strains...... 59 Figure 2-4. Nucleotide sequence of the genomic-DNA regions flanking the transposon in the SM2 mutant...... 65 Figure 2-5. Nucleotide sequence of the genomic-DNA regions flanking the transposon in the SM4 mutant...... 70 Figure 2-6. Nucleotide sequence of the genomic-DNA region flanking the transposon in the RM1 mutant...... 75 Figure 2-7. Multiple amino acid sequence alignments of the deduced amino acid sequence of the signal transduction histidine kinase of the interrupted gene of P. tunicata RM1 with histidine kinase domain of other bacteria...... 75 Figure 2-8. Nucleotide sequence of the genomic-DNA regions flanking the transposon insert in the RM3 and RM4 mutants...... 81 Figure 2-9. Conserved domain identified in the RM3 and RM4 DNA regions disrupted by the mini-TnlO transposon...... 81 Figure 2-10. Multiple amino acid sequence alignments of the deduced amino acid sequence of the amino acid adenylating domains of the interrupted gene of RM3 and RM4 with AMP-binding domain of other bacteria...... 83 Figure 2-11. Multiple amino acid sequence alignments of the deduced amino acid sequence of the pp-binding domains of the interrupted gene of RM3 and RM4 with pp-binding domain of other bacteria...... 84 Figure 2-12. Nucleotide sequence of the genomic-DNA regions flanking the transposon inserted in the RM6 mutant...... 87 Figure 3-1. Chromogenic plate assay showing the bacteriolytic activity of P. tunicata AlpP protein...... 102
12 Figure 3-2. Effect of P. tunicata active cell-free concentrated supernatant on bacterial inner-membrane permeability...... 104 Figure 3-3. Transmission electron micrographs and immunolabeling studies of P. tunicata AlpP protein...... 106 Figure 3-4. Growth of P. tunicata cells in the presence of serial dilutions of the concentrated supernatant after 16 h...... 110 Figure 3-5. Hypothetical model for the mode of action of AlpP in P. tunicata...... 122 Figure 4-1. Panhandle PCR products of the digested and ligated genomic DNA of P. tunicata SM5 mutant...... 131 Figure 4-2. Nucleotide sequence of the genomic-DNA regions flanking the transposon in the SM5 mutant...... 136 Figure 4-3. Transmission electron micrographs of P. tunicata cells...... 142 Figure 4-4. Attachment of P. tunicata wild type and mshJ(SM5) mutant cells to polystyrene microtitre plate surfaces...... 145 Figure 4-5. Attachment of P. tunicata wild type and mshJ (SM5) mutant to microcrystalline cellulose (Avicel)...... 146 Figure 4-6. Transmission fluorescent micrographs using confocal laser scanning microscopy showing attachment of bacterial strains to microcrystalline cellulose (Avicel) after completion of the attachment assay...... 148 Figure 4-7. Scanning electron micrographs of P. tunicata cells grown in cellulose.... 149 Figure 4-8. Confocal laser scanning microscopy images of (A-l) GFP tagged P. tunicata wild type and (A-2) GFP tagged mshJ (SM5) mutant attached on the surface of treated U. lactuca...... 150 Figure 5-1. Transmission and scanning electron micrographs of P. tunicata cells grown in cellulose or cellobiose as the sole carbon source showing the cellulosome-like structures on their surfaces...... 163 Figure 5-2. Transmission electron micrographs of P. tunicata grown in cellulose as carbon source in different incubation conditions ...... 164 Figure 5-3. Attachment of P. tunicata wild type to microcrystalline cellulose (Avicel)...... 165 Figure 5-4. Growth curves of P. tunicata in different substrates...... 166 Figure 5-5. SDS-PAGE-Silver nitrate analysis of proteins from cellulose grown P. tunicata...... 168
13 Figure 5-6. SDS-PAGE-Coomassie blue analysis of membrane-bound proteins of P. tunicata grown in different substrates...... 169 Figure 5-7. SDS-PAGE-Silver nitrate analysis of the affinity-based assay of P. tunicata proteins eluted from the cellulose residues...... 170 Figure 5-8. Congo-red plate diffusion cellulase activity assay of membrane bound proteins of P. tunicata grown in cellulose as sole carbon source...... 172 Figure 5-9. SDS-PAGE and zymogram analysis of membrane-bound proteins of P. tunicata grown in cellulose as the sole carbon source...... 173 Figure 6-1. A hypothetical model of the occurrence and persistence of P. tunicata on surfaces of U. lactuca and C. intestinalis...... 187
14 List of Tables
Table 2-1. MID values to the AlpP protein of stable P. tunicata transposon mutant strains...... 57 Table 2-2. Oligonucleotides used in the primer walking strategy to sequence the DNA regions flanking the inserted transposon in the SM2 mutant...... 62 Table 2-3. Oligonucleotides used in the primer walking strategy to sequence DNA regions flanking the inserted transposon in the SM4 mutant...... 67 Table 2-4. Oligonucleotides used in the primer walking strategy to sequence DNA regions flanking the inserted transposon in theRM 1 mutant...... 72 Table 2-5. Oligonucleotides used in the primer walking strategy to sequence DNA regions flanking the inserted transposon in the RM3 and RM4 mutants...... 77 Table 3-1. Phenotypic characteristics of P. tunicata transposon mutants...... 108 Table 3-2. Relative activity of cell-free concentrated P. tunicata culture supernatant fraction mixes...... 111 Table 3-3. Drop test assay of the heat-treated (90l,C) inactive and active fractions of the P. tunicata cell-free concentrated supernatant after ion exchange chromatography...... 112 Table 3-4. Relative activity of cell-free concentrated P. tunicata culture supernatant fraction mixes...... 113 Table 4-1. Oligonucleotides used in the primer walking strategy to sequence DNA regions flanking the inserted transposon in the SM5 mutant...... 127 Table 4-2. Characteristics, location, size and predicted location of the gene products of the ORFs...... 137 Table 4-3. Hemagglutination of horse erythrocytes by bacteria at different growth stages3...... 143
15 List of Abbreviations
p: micro (10~6) aa: amino acid(s)
ABC: ATP-binding cassette alpP: autolytic protein Pseudo alt eromonas
Amp: ampicillin
ANGIS: Australian National Information Service
BLAST: Basic Local Alignment Search Tool bp: base pair(s)
BSA: bovine serum albumin
C: Celsius
CBP: cellulose binding protein
CLSM: confocal laser scanning microscope
CMCase: carboxymethyl cellulases
Da: Dalton
DNA: deoxyribonucleic acid dNTP: deoxyribonucleotide triphosphate
EPS: extracellular polymeric substances
ExPASy: Expert Protein Analysis System g: gram g- gravitational force GFP: green fluorescent protein h: hour(s)
IPTG: isopropyl-(3-D-thiogalactoside kb: kilobase(s), 1000 bp kDa: kilodalton(s), 1000 Da
Km: kanamycin
1: litre
LB: Luria Broth m: milli (10'3)
16 M: Molar (= molar per litre)
MIC: minimum inhibitory concentration
MID: maximum inhibitory dilution min: minute
MMM: Marine Minimal Media mol: mole (=6.022 x 1023)
MSHA: mannose sensitive hemagglutinin
MW: molecular weight
NCBI: National Center for Biotechnology Information
NSS: nine salts solution
OD: optical density
ORF: open reading frame
PAGE: polyacrylamide gel electrophoresis
PBS: phosphate buffer solution
PCD: programmed cell death
PCR: polymerase chain reaction pi: isoelectric point
PSORT: Prediction of Protein Localization Sites
RBS: ribosomal binding site
RFP: red fluorescent protein
RM: resistant mutant
RNA: ribonucleic acid
SDS: sodium dodecyl sulfate sec: second
SEM: scanning electron microscope
Sm: streptomycin
SM: sensitive mutant
SmR: streptomycin resistant sp.: species
SUPAMAC: Sydney University Prince Alfred Macromolecular Analysis
Centre
TEM: transmission electron microscope
VNSS: V-medium modified from vaatanen w/v: weight over volume
17 1. General introduction and review of related literature
1.1. Introduction
In marine surface environments, bacteria are usually the first species to colonize, after which the settlement of other surface colonizers occurs. Marine bacteria also express a wide array of secondary metabolites which exhibit significant biological activities and have been the target for the discovery and development of new therapeutic drugs (Fusetani, 2000). In marine surface environments, such bacterial metabolites may serve as chemical cues to induce or prevent the settlement and colonization of secondary fouling organisms (Steinberg et al., 2002). A group of bacteria that appear to be commonly associated with surfaces of marine plants and animals, and are known to produce a wide range of bioactive compounds, belong to the genus Pseudoalteromonas (Holmstrom and Kjelleberg, 1999). One member of this genus, P. tunicata, has become the focus of research into the process of marine biofouling due to its ability to inhibit the settlement and growth of a number of common biofouling organisms. This bacterium produces several bioactive agents, each of which targets a specific biofouling organism, including those of bacteria, fungi, algae, diatoms and invertebrate larvae (Holmstrom and Kjelleberg, 1999, James et al., 1996; Egan et al., 2001b).
The colonization of bacteria on surfaces is primarily influenced by the biological properties of the bacterium, such as surface located organelles (e.g. pili or fimbriae, flagella), membrane bound adhesins, and extracellular polymeric substances (EPS) (O’Toole and Kolter, 1998a; Mayer et al., 1999; Prigent-Combaret et al., 2000). Once attached to surfaces, bacteria often also express phenotypic characteristics that are profoundly distinct from those expressed when they are growing planktonically (Stoodley et al., 2002). Furthermore, in order to maintain the bacterial community on the surface, the attached bacterial cells undergo several adaptive survival strategies. One of these strategies involves a stationary phase-like behavior or non-growing state in order to adapt to the reduced nutrient levels on the interior of biofilm communities (Anderl et al., 2003; Pasmore and Costerton, 2003). Interestingly, another strategy involves the killing of a subpopulation of cells of the same species by the production of
18 autolytic substances, apparently in order to reduce the overall demand for nutrients, as well as to release nutrients from the lysed cells, a concept termed programmed-cell death (PCD), allows for the formation of dispersal cell s for colonization of new surfaces (Webb et al., 2003a).
The introductory section of this chapter reviews the production and mode of action of metabolic products of bacteria with an emphasis on antibacterial secondary metabolites of marine bacterial isolates. Some of the secondary metabolites produced by marine bacteria influence the settlement of fouling organisms and the roles of these compounds in biofouling formation and control will be discussed. The marine bacteria producing these bioactive metabolites are usually associated with surfaces of higher marine organisms (Jensen and Fenical, 2000). Mechanisms by which bacteria colonize surfaces as well as their adaptive survival strategies while living on different substrates will be reviewed. The ability of member species of the genus Pseudoalteromonas to produce several bioactive compounds as well as the association of these organisms with marine eukaryotes are presented with a particular focus on the species P. tunicata. This bacterium has been used as a model to understand the ecological role of surface associated bacteria as they influence and control surface associated microbial communities. Finally, this chapter outlines the aims of the thesis project with respect to antibacterial activities and colonization mechanisms of P. tunicata.
1.2. Bacterial secondary metabolites
The search for bioactive compounds produced by microorganisms has been the focus of intensive research due to their potential applications in the discovery and development of drugs. Enormous numbers of bioactive compounds have been isolated from bacteria and shown to exhibit several biological activities such as anticancer, antibacterial, anti inflammatory, antiviral and enzyme inhibition. For the past few decades, marine bacteria have been the new targets for the discovery of bioactive compounds. Marine bacterial metabolites represent diverse structure classes which include terpenes, peptides, polyketides, and compounds of mixed biosynthetic origin (Jensen and Fenical, 2000). The marine bacteria producing these bioactive compounds are usually associated with higher marine organisms such as algae, sponges, bryozoans, tunicates and ascidians, which themselves often are producers of bioactive compounds (Jensen and
19 Fenical, 2000). In fact, many researchers believe that the bioactive compounds from higher marine organisms are actually produced by the symbiotic bacteria (Fesutani, 2000; Jensen and Fenical, 2000). In this section, bacteriad secondary metabolites are presented according to their biomedical activities. Antibacterial compounds are discussed in section 1.2.1. Other bacterial metabolites with no known biomedical activity but which influence biofilm formation are discussed in section 1.2.2.
Several secondary metabolites produced by marine bacteria have been shown to exhibit anticancer activity. For example, alteramide A, a cytotoxic alkaloid isolated from Alteromonas sp. associated with the marine sponge Halichondria okadai, exhibits cytotoxic activity against leukemia, lymphoma, and human epidermal carcinoma cells (Shigemori et al., 1992). Another marine bacterium producing a cytotoxic agent is Pseudo alteromonas haloplanktis (Gauthier et al., 1995). This marine bacterium was isolated from marine sediments and has been shown to produce a cytotoxic compound, bisucaberin which sensitizes tumor cells to macrophage-mediated cytolysis (Takahashi et al., 1987). A strain of Streptomyces hygroscopicus isolated from the gastrointestinal tract of the marine fish Halichoeres bleekeri has been shown to produce the cytotoxic macrolide compound, halichomycin (Takahashi et al., 1994).
Marine bacteria also produce secondary metabolites with antiviral activity. Macrolactins (macrolide) and caprolactins (acylamino acid derivative) are produced by an unidentified Gram-positive deep-sea bacterium isolated from marine sediments and have been shown to inhibit several viruses including herpes simplex and human immunodeficiency virus (Gustafson et al., 1989; Davidson et al., 1993). Other biomedically active compounds act on specific enzymes involved in the pathogenesis of disease. Examples include flavocristamide, a DNA polymerase inhibitor produced by the marine bacterium Flavobacterium sp. (Kobayashi et al., 1995) and pyrostatin, an N-acetyl-beta-D-glucosaminidase inhibitor produced by Streptomyces sp. SA-3501 (Aoyama et al., 1995). A novel endothelin-converting enzyme (ECE), which has been shown to be effective in the treatment of myocardial ischemia, hypertension and renal failure, was isolated from a new marine species of the genus Blastobacter (Takaishi et al., 1998).
20 1.2.1. Antibacterial agents: sources, classification, spectrum of activity and modes of action
Prokaryotes have long been considered as a valuable resource for the identification and development of new antibacterial agents. Among the prokaryotes, both Gram-positive and -negative bacteria have been found to produce antibacterial compounds which inhibit or kill other bacteria. These antibacterial agents may be proteins, peptides, alkaloids, aromatics or terpenoids.
The Gram-positive bacteria are the best-studied group of microorganisms with respect to the production of antibacterial compounds. These bacteria usually produce ribosomally synthesized antibacterial proteins or peptides which are referred to as bacteriocins, and which target cells of the same or closely related species (Hechard and Sahl, 2002). Bacteriocins from lactic acid bacteria (LAB), particularly species of the genera Lactobacillus, Enterococcus and Lactococcus, have become the focus of intensive research because of their significant inhibitory activity against pathogenic food borne- bacteria, and their potential as biopreservatives (Jack et al., 1995). Bacteriocins primarily target the cytoplasmic membrane of the target cell and their mode of action is initiated by the formation of pores. Pore formation destroys the pH gradient resulting in the dissipation of the cell membrane proton motive force (PMF) (Bruno and Montville, 1993). The cationic and amphiphilic structural motif of most bacteriocins drives the formation of transmembrane pores via the interaction of the hydrophilic groups of antibiotic peptides with the phospholipid head groups of the membrane. In addition, hydrophobic helices are inserted into lipid bilayer membranes forming a rod like conformation (van den Hooven et al., 1996). Recently, it has been reported that bacteriocins inhibit target cells by binding to the membrane-bound cell wall precursor lipid II, resulting in inhibition of cell wall synthesis which consequently leads to cell death (Hechard and Shal, 2002).
Among the Gram-negative bacteria, species of the family Enterobacteriaceae produce effective antibacterial compounds, collectively known as colicins. Colicins are prototype bacteriocins which occur as large proteins of 30-70 kDa in molecular mass (Guder et al., 2000). Colicins have been observed to display a narrow range of inhibitory activity against closely related species (Jack et al., 1995). The mode of action of colicins is that
21 of forming pores in the cell membrane (Braun et al., 1994). Other colicins inhibit macromolecular biosyntheses, as is the case of colicin M, which was observed to inhibit peptidoglycan synthesis (Harkness and Braun, 1989). Colicin M prevents the regeneration of the lipid carrier, the third stage of peptidoglycan biosynthesis, in which the disaccharide-pentapeptide is transported to the acceptor site in the existing peptidoglycan layer. It was observed that the inhibition of this step subsequently led to cell lysis (Harkness and Braun, 1989). Protein synthesis can also be targeted, for example, colicin E3, which has ribonuclease activity, has been observed to inhibit protein synthesis by specific cleavage of the small subunit (16S) ribosome (Pugsley, 1984).
In the marine environment, several species of bacteria belonging to the genera Streptomyces, Bacillus, Pseudomonas, Vibrio, Alteromonas and Pseudo alt eromonas (see section 1.4.3) have been reported to produce antibacterial compounds (Gerard et al., 1997; Holmstrom and Kjelleberg, 1999; Isnansetyo et al., 2003; Shiozawa et al., 1993; Trischman et al., 1994).
Strains of Streptomyces produce antibacterial compounds that have a narrow spectrum of activity, inhibiting to only Gram-positive bacteria. For example, salinamides A and B are bicyclic depsipeptide compounds produced by a streptomycete isolated from the surface of the jellyfish Cassiopeia xamachana. These compounds exhibit antibacterial activity against Gram-positive bacteria (Trischman et al., 1994). The selective activity against Gram-positive bacteria is related to the mechanism of action of vancomycin, which inhibits bacterial cell wall synthesis by binding to D-alanyl-D-alanine, a cell wall precursor (Trischman et al., 1994). Bacteria associated with marine sediments have also been shown to produce antibacterial compounds. Bioxalomycin a2, a metabolite produced by Streptomyces viridodiastaticus sp., isolated from shallow-water sediment, exhibits antibacterial activity against Gram-positive bacteria by inhibiting DNA synthesis (Singh et al., 1994).
In addition, a marine bacterium Vibrio parahaemolyticus isolated from the toxic mucus of boxfish Ostracion cubicus, produces an antibacterial compound vibrindole A, which is inhibitory against Staphyloccus aureus, S. albus and Bacillus subtilis (Bell et al., 1994). Enterococcus faecium isolated from the tunicate Microcosmus australis
22 produces an alkaloid antibiotic identified as harman, which displays inhibitory activity against S. aureus and Vibrio anguillarum (Aassila et al., 2003). A broader range of antibacterial activity was also demonstrated by a marine Bacillus species isolated from a marine worm, which produces cyclic decapeptides, loloatins as the active compounds (Gerard et al., 1997; 1999).
Several species of the genus Pseudomonas have been reported as producing antibacterial compounds. Pseudomonas sp. AMSN, isolated from a red alga Ceratodiction spongiosum, produces a 2,4-diacetylphoroglucinol (DAPG) antibiotic with activity against vancomycin-resistant S. aureus (Isnansetyo et al., 2003). DAPG has been shown to be bactericidal against methicillin-resistant S. aureus and bacteriolytic against both Gram-negative and -positive bacteria (Kamei et al., 2003). Pseudomonas strains obtained from a marine alga and a tube worm produce eight cyclic depsipeptides (massetolides A-H) displaying antimycobacterial activity against Mycobacterium tuberculosis and M. avium-intracellulare (Gerard et al., 1997).
Species belonging to the genera of Alteromonas and Pseudo alter omonas have been reported to produce antibacterial compounds with a wide spectrum of activity. For example, “A. rava” sp. nov. SANK 73390 produces the metabolite thiomarinol, which shows in vitro antibacterial activity against Gram-negative and Gram-positive bacteria and particularly against S. aureus, including methicillin-resistant strains of the species (Shiozawa et al., 1993). Several Pseudoalteromonas species demonstrate antibacterial activities, including P. aurantia, P. luteoviolacea, P. rubra, P. citrea, P. tunicata, P. maricaloris and P. phenolica (Gauthier and Breittmayer, 1979; Gauthier and Flatau, 1976; Gauthier, 1979; James et al., 1996; Ivanova et al., 2002e; Isnansetyo and Kamei, 2003). In particular the antibacterial agents produced by these species are discussed in section 1.4.3. P. tunicata has been reported to produce an extracellular 190 kDa antibacterial protein (James et al., 1996). A detailed discussion of the antibacterial activity of P. tunicata will be presented in section 1.4.4 and the investigation of the mode of action of the antibacterial protein is presented in chapters 2 and 3.
23 1.2.2. Role of bacterial metabolites in the biofouling process
Bacteria produce chemical metabolites that can influence the development of biofouling communities (see section 1.3.3). Bacterial metabolites either induce or prevent the settlement and colonization of eukaryotic fouling organisms (reviewed in Steinberg et al., 2002). Bacterial water-borne metabolites and cell surface-associated signals are known to induce larval settlement (Fitt et al., 1990; Maki et al., 1990; Rodriguez and Epifanio, 2000). Bacterially derived settlement signals include: lipids, which induce stolon settlement of the cnidaria Aurelia aurita (Schmahl, 1985), oligopeptides, which induce larval settlement and metamorphosis of the cnidaria Cassiopea andromeda (Neumann, 1979), glycoconjugates, which induce settlement and metamorphosis of the polychaete Janua brasiliensis (Kirchman et al., 1982), and L-dihyroxyphenylalanine (L- DOPA), which induces the settlement of larvae of the oyster Crassostrea gigas (Weiner et al., 1989).
Marine bacterial isolates have been shown to prevent the colonization of fouling organisms. However, many bioactive compounds from these bacteria responsible for inhibiting settlement of the invertebrate larvae have not been isolated and the nature of their activities have not been determined. It has been shown that epibiotic bacteria of the green alga Ulva reticulata belonging to genera Alteromonas, Pseudo alt eromonas and Vibrio inhibit the larval settlement of polychaete Hydroides elegans (Dobretsov and Qian, 2002). The anti-larval activity of these bacteria has been proposed to protect the alga from fouling (Dobretsov and Qian, 2002). It has also been shown that the metamorphosis of the polychaete H. elegans is inhibited by Vibrio strains (Unabia and Hadfield, 1999). Extracts of Pseudomonas sp. NUDMB50-1 have been shown to inhibit the larval settlement of the barnacle Balanus amphitrite and algal spores of U. lactuca (Burgess et al., 2003). The antifouling compounds responsible for the activity of the extract have not been fully elucidated. One well-known and studied antifouling bacterium is P. tunicata. This bacterium has been shown to inhibit the settlement and colonization of a variety of biofouling organisms as it produces a wide range of antifouling compounds. These compounds will be described and discussed in section 1.4.4.
24 1.3. Bacterial colonization of nonliving and living surfaces
The ability of bacteria to colonize surfaces offers important advantages, including increased access to nutrients, protection against toxins (e.g., biocides), maintenance of extracellular enzyme activities and shelter from predation (Dang and Lovell, 2000). All surfaces submerged in marine waters, both nonliving and living, are colonized by bacteria. The association of bacteria with living surfaces (e.g. marine plants and animals) may reflect different survival strategies of bacteria in the marine environment. The surfaces of marine plants and animals may serve as a nutritional niche for colonizing bacteria as well as shelter from predation. However, bacterial colonization may be detrimental to host organisms as this leads to tissue degradation and disease due to the production of toxins, enzymes and other harmful metabolites. Moreover, bacterial colonization can influence the subsequent colonization of eukaryotic fouling organisms. They can induce settlement and colonization of macrofoulers and the development of biofouling (Johnson et al., 1997). Other surface associated bacteria, such as P. tunicata prevent the settlement and colonization of macrofoulers (Holmstrdm and Kjelleberg, 1999).
1.3.1. Bacterial attachment mechanisms
Attachment to surfaces is an important event in bacterial colonization. This process is believed to commence when bacteria sense environmental signals that initiate the expression of their attachment phenotypes (reviewed in Costerton et al., 1995). The environmental cues include nutrient availability, temperature, osmolarity, pH, iron and oxygen (O’Toole et al., 1998a; 1998b; 2000). The membrane bound adhesins and the structural polymers of polysaccharides and proteins present on the cell surface mediate the attachment of bacteria to surfaces.
1.3.1.1. Cell surface structures that mediate attachment
Bacteria have a well-developed attachment strategy, where they for example can use surface structures such as flagella and pili to attach themselves to surfaces. In
25 Pseudomonas aeruginosa, flagella and type 4 pili play an important role in the early stages of attachment of the bacterium to a surface (O’Toole and Kolter, 1998). In particular, fimbriae or pili have attachment moieties or binding domains positioned at their distal ends that recognize different binding receptors in host tissues or other surfaces. Flagellar motility has also been demonstrated to mediate cell-to-surface interactions. For example, it has been shown that non-motile P. aeruginosa flgK mutant strains attach to polyvinyl chloride (PVC) less effectively than the wild type strain (O’Toole and Kolter, 1998).
In Vibrio cholerae, attachment to surfaces requires three components: flagella, MSHA (mannose-sensitive hemagglutinin) type 4 pili and EPS (O’Toole and Kolter, 1998a; Watnick et al., 1999). Flagella and type 4 pili accelerate the attachment of V. cholerae to abiotic surfaces and EPS is involved in the formation of the three-dimensional biofilm architecture (Watnick and Kolter, 1999). For example, flagellar mutants of V. cholerae are inefficient colonizers of the intestinal epithelium (Watnick et al., 2001). V. cholerae also expresses different attachment structures depending on the surrounding environment. Firstly, attachment to human host cells requires the involvement of tcp (toxin-coregulated pilus), a type 4 pilus, and a virulence factor (Herrington et al., 1988). Secondly, the colonization of V. cholerae in the aquatic environment, particularly with respect to attachment to phytoplankton, is mediated by the MSHA pilus (Watnick et al., 1999). It has also been reported that there is an additional attachment factor present in V. cholerae for attachment to chitin, since neither of the two pili - the MSHA and the tcp pili - appear to be involved in attachment to chitin (Chiavelli et al., 2001).
The screening of a library of Escherichia coli mutants to identify genes involved in attachment to surfaces resulted in the isolation of mutants defective in flagellum- mediated motility (Genevaux et al., 1996). Flagella were demonstrated to initiate the early stages of the attachment of E. coli to surfaces. Another research team carried out a similar screening and demonstrated that type-I pili mutants are defective in their abilities to attach to surfaces (Pratt and Kolter, 1998). The type-I pili retract and mediate surface- dependent motility (Pratt and Kolter, 1998). Furthermore, type-I pili retraction is important for close associations with human cells (Mulvey et al., 1998).
26 Another surface structure that mediates the attachment of E. coli to surfaces, are fimbriae-like binding proteins termed curb. Curli have been associated with the binding of E. coli to proteins including fibronectin and plasminogen (Olsen et al., 1989; Sjobring et al., 1994). Pringet-Combaret et al (2000) have demonstrated that curli synthesis is required for the initial attachment of E. coli to inert surfaces. Curli are believed to interact directly with the substratum and form interbacterial bundles, resulting in a cohesive and stable association of the cell and the substratum.
The attachment of bacteria to living surfaces is also mediated by the aforementioned cell surface structures. Flagella have been demonstrated to mediate the attachment of plant pathogens such as Pseudomonas species to surfaces of their host organisms (Panopoulos and Schroth, 1974, Haefele and Lindow, 1987). Well-investigated surface structures known to be involved in the attachment of bacteria to plant surfaces are the type 4 pili. For example, in P. syringae pv. phaseolicola, attachment to the host’s stomata is mediated by type 4 pili (Romantschuk, and Bamford, 1986). It was also reported that type 4 pili contribute to the ability of the plant growth-stimulating P. putida WCS358 to colonize plant roots (de Groot et al., 1994). Similarly, the attachment of Azoarcus sp. to grass roots as well as to fungal mycelia is also mediated by type 4 pili, which are expressed in response to plant recognition (Dorr et al., 1998). The survival of P. syringae pv. tomato DC3000 on the plant leaf surface is mediated by the expression of type 4 pili (Roine et al., 1998). In V. cholerae El Tor, the attachment of this bacterium to phytoplankton and zooplankton surfaces is mediated by the MSHA pilus, which belongs to the type 4 pili family (Chiavelli et al., 2001). Recently, it was reported that the MSHA pilus promotes an interaction between V. cholerae El Tor and the hemolymph of the mussel Mytilus galloprovincialis (Zampini et al., 2003). The characteristics of the type 4 pili will be discussed in section 1.3.1.1.1.
Bacteria also interact with living surfaces via proteinaceous adhesins which are directly associated with the cell wall or outer membrane. Examples include chitin-binding proteins, which bind specifically to chitin or chitinous polymers, such as the chitin binding proteins of V. alginolyticus (Tarsi and Pruzzo, 1999), V. harveyi (Soto-Gill and Zyskind, 1984), Serratia marcescens (Brurberg et al., 1994), Aeromonas caviae (Sitrit et al., 1995) and Pseudoalteromonas sp. strain S91 (Techkamjanaruk et al., 1997). In cellulolytic clostridia, attachment to surfaces that contain cellulose is mediated by
27 protuberant-like structures called cellulosomes. These cell surface structures bind closely to the cellulose through the cellulose-binding domains of the scaffolding protein and the catalytic enzymes (Bayer et al., 1998). Although the binding of these proteinaceous adhesins is specific, it establishes an important advantage for the bacterium especially if they degrade and utilize these polymers (chitin and cellulose) as a source of carbon and energy. A detailed discussion of the cellulosome will be presented in section 1.3.1.1.2.
1.3.1.1.1. Type 4 pili: structure and biogenesis
Type 4 pili are flexible, filamentous surface appendages that are found in a wide variety of Gram-negative bacteria (Strom and Lory, 1993). The type 4 pili are characterized by conserved features of the structural pilin subunits and are classified as either type 4a or 4b based on amino acid sequence similarities within the amino terminus region of the subunit polypeptide. The majority of the type 4 pili family belongs to the type 4a subclass. This subclass is composed of prepilin subunits which are characterized by a short, positively charged leader sequence of six or seven amino acids which, upon removal, result in a mature pilin subunit with a modified amino acid N- methylphenylalanine as the first residue, and a highly conserved hydrophobic amino terminal domain (Dalrymple and Mattick, 1987). The type 4a pili are usually distributed either peritrichously or in polar positions on the bacterial cell surface. The type 4b prepilins have longer leader peptides and the N-terminal amino acid of the mature pilin subunit is variable, being either methionine or leucine (Donnenberg et al., 1992; Giron et al., 1994). The type 4b pili form large bundles of laterally associated fibers on the bacterial cell surface. The type 4 pili of either subclass mediate bacterial attachment, surface motility and aggregation or biofilm formation (O’Toole and Kolter, 1998).
The biogenesis of type 4 pili requires numerous gene products, including a structural prepilin subunit, ancillary proteins with prepilin-like leader sequences, inner and outer membrane proteins and nucleotide binding proteins (Aim and Mattick, 1997). It has been reported that a mutation in one these genes prevents the assembly of the functional pili. For example, mutation in P. aeruginosa pilO, pilP, or pilQ results in a phenotype devoid of pili (Martin et al., 1995). The pilO,pilP and pilQ belong to a secretory operon that includes two other genes (pilM and pilN) required in the biogenesis of fimbriae in
28 P. aeruginosa (Martin et al., 1995). Another example illustrating the effect of mutation in the pili biogenesis proteins is observed in N. gonorrhoeae. It was reported that neisserial pilC and pilP mutants showed a reduction in the expression of pili (Jonsson et al., 1991; Drake et al., 1997). These genes encode proteins necessary for neisserial pilus biogenesis. The interdependence of gene products required for pilus biogenesis is also observed in cyanobacteria. In Synechocystis sp. PCC6803, a unicellular cyanobacterium, it was found that inactivation of pilC, which encodes for a protein for pilus biogenesis, resulted in the absence of pili on the cell surface (Bhaya et al., 2000). In V. cholerae, biogenesis of MSHA pilus is dependent on sixteen gene products of the MSHA gene locus, which is made up of secretory and structural operons (Marsh and Taylor, 1999). A deletion in any of the putative promoter regions upstream of mshl, a secretory gene, or mshB, a stmctural gene, abolished MSHA pilus assembly, secretion and expression (Marsh and Taylor, 1999). Additionally, mutation in mshE, a secretory gene in the MSHA biogenesis locus, was shown to abolish hemagglutination (Hase et al., 1994).
Many Gram-negative bacteria bind to surfaces via the tip adhesins of the pili (Strom and Lory, 1993). The pili are incorporated with minor pilin-like subunits that are responsible for the recognition and binding of different receptors. In P. aeruginosa, binding to epithelial tissue is mediated by pili containing the cysteine-cysteine bridge structure (Irvin et al., 1989). The ability of N. meningitis to adhere to epithelial cells is mediated by the adhesin protein, PilC which is found on the tip of the pilus fibers (Rudel et al., 1995). The adhesion-promoting region is located in thepilCl allele which is specifically located in the amino terminal part of the molecule (Morand et al., 2001). In V. cholerae, El Tor strains, a hemagglutinin pilus with preference for mannose receptors is expressed (Jonson et al., 1991). A hemagglutinin pilus with preference for both mannose and fucose is expressed by the V. cholerae 01 strain Bgdl7 (Ehara et al., 1991).
1.3.1.1.2. Characteristics of cellulosome complexes
Cellulosomes are large, stable, multienzyme complexes specialized in the adhesion to and degradation of cellulose, that reside within protuberances visible on the cell surface (Bayer et al., 1994). The cellulosome organization, as supported by molecular evidence, has been described in detail only for the cellulolytic Clostridium species, including Clostridum thermocellum (Bayer and Lamed, 1986), C cellulovorans (Doi and Tamaru,
29 2001), C. josiu (Kakiuchi et al., 1998), C. cellulolyticum (Belaich et al., 1997) and C. acetobutylicum (Sabathe et al., 2002). Among these Clostridium species, C. thermocellum has been extensively investigated. The cellulosome complex in this bacterium contains different types of glycosyl hydrolases such as cellulases, which are bound to a scaffolding protein (also known as cellulose integrating protein, CipA) (Bayer et al., 1998). This scaffolding protein is the characteristic signature of cellulosomes. It possesses a series of functional domains involved either in enzyme adhesion (via the cohesin), cellulose binding (via the cellulose binding domain, CBD) or anchoring to the bacterial surface (via the dockerin domain) (Bayer et al., 1998). In addition to a definitive catalytic domain, the cellulosomal enzymes (glycosyl hydrolases) possess an additional domain called dockerin, which binds tightly with the cohesins of the scaffolding protein (Shoham et al., 1999). The assembly of the enzymatic subunits into a cellulosome complex is dependent on the cohesin-dockerin interactions and the interaction of the cellulosome complex with cellulose which is mediated by the cellulose-binding domains (CBDs) of the scaffolding protein and the catalytic enzymes (Bayer et al., 1994). The attachment of the cellulosome to the cell surface is mediated by a unique type of cohesin-dockerin interaction, which involves the binding of type-II dockerin of the scaffolding protein and the type-II cohesins of the cell surface anchoring protein (Leibovitz and Begui, 1996), which contain a SLH protein (S- layer homology) (Sleytr and Beveridge, 1999), believed to be associated with the cell surface of Gram-positive bacteria. The cellulosomal enzymes are relatively large proteins ranging from 40 to 180 kDa and belonging to different families of glycosyl hydrolases (Shoham et al., 1999). Many of the enzymes are cellulases, including both endo and exo-acting p-glucanases. Not all the cellulosomal enzymes are cellulases, as xylanases, mannanases, lichenases, and chitinases can also be found within cellulosome complexes (Fontes et al., 1995; Zverlov et al., 2002a; 2002b).
The cellulosomes of C. thermocellum were observed in cells grown in cellulose and cellobiose (Mayer et al., 1987). These structures undergo dynamic structural transformations during different stages of growth. It has been reported that in early logarithmic phase, cellulosomes are closely bound to the cell surface (Bayer et al., 1986) and begin to disengage in late exponential phase, finally detaching from the cell surface during the stationary phase of growth. Loosely bound cellulosomes were found to be attached to the residual substrate (Lamed and Bayer, 1983). Cellulosomes are also
30 observed in other nonclostridial anaerobic bacteria including Ruminococcus flavefaciens, R. albus, Acetivibrio cellulolyticus and B'acteroides cellulosolvens (Ohara et al., 2000; Ding et al., 1999; 2000; 2001).
1.3.2. Bacterial biofilm and microcolony formation
Once attached to a surface, bacteria continue to move in order to spread laterally. It has been proposed that the retraction and extension of pili, a behavior termed twitching motility, can cause bacteria to move across a surface (Bradley, 1980). Consequently, twitching motility causes the formation of microcolonies as individual cells move across the surface towards each other (O’Toole and Kolter, 1998). The type 4 pili of P. aeruginosa mediate twitching motility on surfaces. Studies have shown that P. aeruginosa type 4 pili mutants are defective in successive development events that are necessary to the formation of a biofilm (O’Toole and Kolter, 1998). Although type 4 pili have also been found in V. cholerae, its potential type 4 pili-mediated motility has not been reported or explored. In the enteropathogenic E. coli (EPEC), microcolony formation depends on the expression of a type 4 pilus called the bundle forming pilus, BFP (Giron et al., 1991; Donnenberg et al., 1992).
The binary division of the attached cells also contributes to the formation and development of bacterial biofilms. Cell clusters are formed as the cells divide upward and outward from the attached surface (Heydom et al., 2000). As biofilms expand and grow, they form pillars or mushroom-like structures with fluid-filled channels, which are believed to facilitate the diffusion of nutrients, as well as the transport of toxic waste products away from the cell (Costerton et al., 1995).
Important for biofilm formation is the increase in synthesis of extracellular polymeric substances (EPS) that act as binders providing mechanical adhesive (surface-related) and cohesive stability between the cells (Mayer et al., 1999). This structural polymer matrix is made up of not only polysaccharides, but also of proteins, nucleic acids and phospholipids (Wingender et al., 1999). The production of EPS in biofilms has been demonstrated in several bacteria such as P. aeruginosa (Gacesa, 1998), V. cholerae (Watnick et al., 1999) and S. aureus (Baselga et al., 1993). More structurally
31 differentiated biofilms were found to be formed by mucoid strains of P. aeruginosa (Nivens et al., 2001). These mucoid P. aeruginosa strains have been shown to have an overexpression of alginate, the main component of EPS (Hentzer et al., 2001), which results in the formation of differentiated biofilm structures such as mound- and mushroom-shaped cell clusters. In E. coli, the formation and differentiation of biofilms requires colonic acid, a main component of its EPS (Danese et al., 2000). EPS has also been shown to affect the formation and differentiation of V. cholerae biofilms (Watnick et al., 1999).
1.3.2.1. The negative impacts of biofilms
Biofilms are ubiquitous as they are formed on a wide range of biotic and abiotic surfaces (O’Toole, Kaplan and Kolter, 2000). Most environments are predominated by mixed- species biofilms, whereas single-species biofilms are more likely to exist under specific conditions such as on the surfaces of implanted medical devices causing persistent disease and infections (Stewart and Costerton, 2001). Biofilms are a serious problem in many industrial and medical settings. For example, the cooling water systems that are used as a cooling medium in many industries for power generation, chemical manufacture and other manufacturing processes, are prone to biofilm development. The deposition of undesirable bacteria and other microorganisms has detrimental effects such as mechanical blockages, reduced heat transfer efficiency and biodeterioration of polymeric and metallic systems, causing the loss of millions of dollars in terms of lost productivity (Boot, 1999). In medical setting, biofilms associated with implanted medical devices or damaged tissues are a major cause of antibiotic resistance. The high antibiotic resistance of biofilms is believed to be due to slow or incomplete penetration of antibiotics into the biofilm, as bacteria are encased in a matrix of extracellular polymer substances (EPS) (Stewart and Costerton, 2001), as well as stationary phase induced resistance by the cells in the interior of the biofilm (Drenkard and Ausubel,
2002).
32 1.3.3. Biofouling in the marine en vironment
Biofouling in the marine environment takes place according to a sequence of specific events consisting of 1) bacterial attachment; 2) development of biofilm formation; and 3) colonization and settlement of macrofoulers (Wahl, 1989). The mechanisms by which bacteria attach to surfaces and develop into biofilms have been discussed in section 1.3.1 and 1.3.2.
The presence of biofouling communities on the surface of living marine organisms can be beneficial or detrimental to the host. The attached microorganisms may serve as protective barriers that protect the host from invading predators. Alternatively, the presence of biofouling can be unfavorable, if bacterial colonization causes tissue degradation and disease. Two examples are coral bleaching (Torren et al., 1998) and Coralline Lethal Orange Disease (CLOD), both of which are processes of microalgal decay that can destroy tropical coral or algae (Littler and Littler, 1995). In addition, the biofouling of marine macroalgae can cause a decrease in photosynthesis, because of the shading effect of the fouling organisms (Sand-Jensen, 1977). Biofouling poses not only detrimental effects to fouled marine organisms, but is also a costly problem, particularly with respect to the fouling of artificial or man-made substrata such as ship hulls, docks, pipelines and marine aquaculture facilities. Detrimental effects of biofouling upon such structures include 1) increased load, 2) mechanical damage to protective coatings, 3) blockage of sea-water inlets, and 4) corrosion (Wahl, 1997). For example, the biofouling of ship hulls in the U.S. caused a loss of more than $500 million in additional fuel costs per annum as well as significantly reduction in speed (Flemming, 2002).
1.3.3.1. Settlement and colonization of macrofoulers
The final event in the formation of a biofouling community is the settlement and colonization of macrofoulers such as algae and invertebrates. The life cycle of many of these macrofoulers includes a free-living settlement stage (Chia, 1978). The dispersal of invertebrate larvae and algal spores is a critical phase during the life cycle because this is when they are able to select, attach to and subsequently settle on a surface (Pawlik, 1992). There are several factors (both physical and chemical) known to influence the settlement of macrofoulers. Physical factors include surface topography and
33 hydrophobicity, and light. Algal spores prefer to settle; on and colonize rough surfaces, and the strength of their attachment to such surfaces is stronger than their attachment to smooth surfaces (Woods and Fletcher, 1991). Surface hydrophobicity has also been shown to influence the settlement preferences of invertebrate larvae. The larvae of the marine tunicate C. intestinalis prefer to settle on hydrophobic surfaces (Szewzyk et al., 1991) while larvae of the barnacle B. amphiphrite prefer to settle on hydrophilic surfaces (Rittschof et al., 1989). Light also appears to influence the settlement of macrofoulers. It has been reported that algal spores prefer to settle under illuminated surfaces (Leadbeater and Callow, 1992).
Diatoms are the most common and abundant of the early algal colonizers. Only a few cells are needed to attach to a surface in order for cell division to occur and give rise to colonies that eventually coalesce to form a compact biofilm. The most common macroalgae found attached to marine surfaces are the green algae (e. g. Enteromorpha) and the brown algae (e. g. Ectocarpus). In some species, germination gives rise to a thallus with a spreading behavior which allow the algae to grow along the substrate and through the existing biofilm (Leadbeater and Callow, 1992).
1.3.4. Survival of attached bacterial cells
It has been proposed that the bacteria develop several adaptive survival strategies as they age on surfaces (Stoodley et al., 2002). In these conditions, bacteria employ survival strategies as they grow and expand; they require an increasing amount of nutrients. However, the community of attached bacteria may experience nutrient poor conditions, particularly in cells that are in close contact with the solid surface. It is proposed that the presence of an EPS matrix prevents the continuous diffusion of nutrients (Stewart and Costerton, 2001). In these conditions, it has been proposed that bacteria employ survival strategies typical of a stationary phase-like response to reduced nutrient levels (Pasmore and Costerton, 2003). It has been demonstrated that the expression of stationary phase sigma factor-regulated genes (rpoS) and genes encoding for putative transport proteins associated with starvation response were upregulated during the transition of E. coll from planktonic to biofilm growth (Schembri et al., 2003). Moreover, gene clusters (cydAB and B2991-hybABC) involved in survival under
34 oxygen-limiting conditions were also upregulated. The identity of the induced gene clusters indicates that the biofilm employs mechanisms for maintaining the cells in the community during oxygen and nutrient limitation (Schembri et al., 2003).
It has been demonstrated that bacterial cells detach from surfaces as they age and it has been suggested that this detachment mechanism is nutrient dependent (Stoodley et al., 2002). Nutrient starvation triggers the release of small cell aggregates and individual cells that can more easily locate and use dissolved nutrients (Peyton and Characklis, 1993). Moreover, these detachment mechanisms have developed in mature bacterial biofilms to release planktonic cells in order for the propagules to colonize surfaces (Pasmore and Costerton, 2003). In P. aeruginosa, overexpression of alginate lyase contributes to cell detachment and sloughing from biofilms, as this enzyme dissolves the EPS matrix (Boyd and Chakrabarty, 1994). The possibility of an EPS dissolution detachment mechanism has also been explored in P. fluorescens biofilm (Allison et al., 1998). However, the regulatory pathway is unclear. The detachment of Streptococcus mutans biofilm has been shown to be mediated by a surface protein releasing enzyme (SPRE) that dissolves EPS to facilitate the release of the cells from the biofilm (Lee et al., 1996).
1.3.4.1. Programmed cell-death (PCD)
Another adaptive survival strategy, which is now generally accepted, is the concept of programmed-cell death (PCD). PCD is defined as a genetically regulated process of cellular suicide that is activated by several signals, including the metabolic status of the cells, harsh environmental conditions and developmental history of the cells (Lewis, 2002). A recent article proposes that bacterial biofilms undergo PCD (Webb et al., 2003a) and that PCD may serve as a key role in the multicellular differentiation of microcolonies during the formation of mature biofilms. It has also been reported that for many bacterial species, the development of mature biofilms includes stages where hollow microcolonies form as result of localized cellular differentiation, where dissolution and dispersal of the cells occurred inside the microcolonies (Sauer et al., 2002; Tolker-Nielsen et al., 2000). It was proposed that the dispersal of cells from inside the microcolonies is triggered by nutrient limitation (Webb et al., 2003a). Cell death or autolysis has been observed in mono-species biofilms of P. aeruginosa (Webb et al.,
35 2003b), and the marine bacterium P. tunicata (Mai-Prochnow et al., 2004) as well as mixed-species biofilms (Auschill et al., 2001). In these biofilms, cell death occurred inside microcolony structures, leaving behind a sub-population of alive cells. In P. aeruginosa biofilms, cell death is linked to the expression of a Pfl-like prophage, which encodes for proteins with similarity to a PCD operon in E. coli (Thompson, 2003) . Cell death during biofilm development in the biofilms of P. tunicata is believed to be mediated by the production of the autolytic protein, AlpP (Mai-Prochnow et al., 2004) .
PCD or autolysis has been known to be a part of the developmental process of a number of bacterial species. For example, PCD has been observed during cell development in sporulating mother cells of Bacillus subtilis, where lysis occurs prior to the release of the spores (Smith and Foster, 1995). It is suggested that in this case, autolysis is an altruistic behavior of the sporulating mother cell, as it releases nutrients for the kin cells to complete sporulation. Autolysis has also been observed during the fruiting body formation and sporulation of Myxococcus xanthus (Rosenbluh and Rosenberg, 1990). The lysis of these vegetative cells is also believed to represent altruistic death, where nutrients are released in order to enhance the survival of the developing cells. Compelling similarities between fruiting body formation in myxobacteria and microcolony biofilm formation in non-differentiated bacteria such as P. aeruginosa have been demonstrated by Webb et al (2003a).
1.3.4.1.1. Mechanisms of controlling cell death and survival
The expression of autolysins has been observed during the development of bacteria. Cell death in sporulating B. subtilis is mediated by three autolysins, namely CwlB, CwlC and CwlH. CwlB is a major autolysin released at the end of the logarithmic phase of growth (Smith and Foster, 1995). The autolysins CwlC and CwlH are produced during the late sporulation stage (Nugroho et al., 1999). It is unclear how these autolysins are activated during the sporulation process. Autolysins have also been shown to play an active part in fruiting body formation and sporulation in M. xanthus. The pathway by which these autolysins lead to cell death is unknown.
36 In Streptococcus pneumoniae, autolysis is known to occur in response to cellular damage caused by antibiotics and other toxic molecules (Tomasz et al., 1970). Bacterial autolysis in S. pneumoniae is mediated by the autolysin LytA. It was reported that mutation of the lytA gene causes resistance to inhibitors of cell wall synthesis, indicating that these antibiotics may operate via the activity of this gene (Tomasz et al., 1970). In E. coli, SulA activates cell death in response to DNA damage, similar to induced cell death after exposure to quinolone antibiotics (Piddock and Walters, 1992). The expression of the sulA gene triggers the elimination of cells with serious DNA damage. While SulA activates cell death in response to antibiotic exposure, inhibition of antibiotic-induced autolysis in E. coli is controlled by the expression of mar genes. The expression of these genes is responsible for the development of multiple drug resistance (MDR) to antimicrobials in E. coli (Goldman et al., 1996).
One well-researched genetic mechanism for controlling cell death and survival in bacteria, is the toxin-antidote (TA) system, which is controlled via a set of genes encoded on plasmids, bacteriophages (Jensen and Gerdes, 1995) or as part of the genomic DNA (Aizenman et al., 1996). The TA loci are organized into operons, which encode for a stable toxin that mediates cell death and an unstable antitoxin that counteracts the effect of toxin resulting in the survival of the cells.
In E. coli, the plasmid-encoded TA loci function during the postsegregational killing (PSK) of the plasmid-free segregants, which assures the stability and maintenance of plasmids among the general E. coli population. The toxin-antidote plasmid encodes for a stable protein toxin and an unstable protein antitoxin. Alternatively, the toxin is a protein synthesized from a stable mRNA and the antidote is a small unstable antisense RNA molecule (Engelberg-Kulka and Glaser, 1999). In both cases, the toxin and antitoxin are normally produced at the same time. One of the best-investigated toxin- antidote systems is ccd from plasmid F of E. coli (Jensen and Gerdes, 1995). This system involves the expression of two small proteins CcdA (antitoxin) and CcdB (toxin). When the F plasmid is lost, bacterial killing occurs because CcdA is degraded which subsequently allows the CcdB toxin to exert its lethal effect on the cell (Maki et al., 1996). The E. coli cells harboring the F plasmids are resistant to the lethal effects of CcdB because of the formation of a tight complex between CcdA and CcdB that binds
37 to the ccd promoter-operator, which in turn represses the synthesis of toxin and antitoxin proteins (Tam and Kline, 1989).
Other toxin-antidote systems are encoded on the bacterial chromosomes. An example is chpA, a TA system involved in controlling cell death and survival in E. coli K-12 (Masuda et al., 1993; Metzger et al., 1988). The chpA locus has also been called mazEF, which has been shown to be responsible for cell death in E. coli (Aizenman et al., 1996). This system expresses MazF which is toxic and MazE which is anti-toxic and short lived. The mazEF genes form an operon upstream of the relA gene, encoding an ATP:GTP 3’-pyrophosphotransferase, ppGpp, during amino acid starvation (Aizenman et al., 1996). It has been demonstrated that mazEF mediated cell death is induced by the overproduction of ppGpp, as this compound inhibits the coexpression of mazE and mazF. Under these conditions, MazF exerts its toxic effect and causes cell death in the absence of MazE, which is an unstable protein and is thus more rapidly depleted than MazF (Aizenman et al., 1996). This finding strongly suggests that mazEF elicits PCD during nutrient deprivation (Aizenman et al., 1996).
1.4. The genus Pseudoalteromonas
The genus Pseudoalteromonas belongs to the class of Gammaproteobacteria of the recently proposed phylum Proteobacteria (Garrity et al., 2002). Members of this genus are motile, rod-shaped Gram-negative bacteria with an oxidative metabolism. Some member species are pigmented and able to produce a range of extracellular compounds. Most of the Pseudoalteromonas species are isolated exclusively from marine waters in particular in association with higher organisms. The majority of Pseudoalteromonas species produce bioactive compounds such as exoenzymes, toxins and secondary metabolites.
38 1.4.1. Association with marine eukaryotes
The association of Pseudoalteromonas species w ith a range of marine eukaryotes indicates a diverse distribution of the species in the marine environment. The Pseudoalteromonas species can be classified as a pathogen or symbiont based on their effect on the host and their interaction with higher marine organisms. Pathogenic Pseudoalteromonas include P. bacteriolytica (currently known as Algicola bacteriolytica, (Ivanova et al., 2004)), which causes red spot disease in the red algae, Laminaria japonica, (Sawabe et al., 1998) and P. tetraodonis, which releases toxin precursors causing poisoning by pufferfish (Simidu et al., 1990). Additionally, P. gracilis strain B9, elicits disease symptoms in the red alga Gracilaria gracilis (Schroeder et al., 2003). It has been suggested that the interaction between P. tunicata and higher marine organisms, for example with the tunicate C. intestinalis or the green alga U. lactuca, is symbiotic because P. tunicata is believed to provide these marine organisms with protection against surface colonizers (Homstrom and Kjelleberg, 1999). The nature of the interaction between other Pseudoalteromonas species and other higher marine organisms with which they have been found in association with, has not been elucidated. This includes for example P. maricaloris and P. translucida which have been isolated from marine sponges (Ivanova et al., 2002c; 2002e), and P. ruthenica sp. nov., isolated from marine invertebrates (Ivanova et al., 2002d).
1.4.2. Production of extracellular enzymes
Some members of the genus Pseudoalteromonas demonstrate specific physiological responses in order to survive in a particular habitat or ecological niche. These bacterial species produce extracellular enzymes, which hydrolyze material to form soluble substrates that can be taken up by the cells. The enzymes are of fundamental importance in the degradation of polymer substrates and organic matter in the marine environment (Martinez et al., 1996). They include proteinases, lipase, chitinase, agarase, cellulase, amylase, [3-galactosidase and p-glucosidase (Ivanova et al., 2003). Several species associated with red algae derive nutrients from agar, a polysaccharide present in the cell walls of the algae. The breakdown of agar is based on the specific activity of a-agarase and [3-agarase. Pseudoalteromonas species which produce agarase include
39 P. agarolyticus, P. antarctica strain N-l, P carrageenovora, P. atlantica, P. agarivorans and P. gracilis (Akagawa-Matsushita et al., 1992; Bozal et al., 1997; Vera et al., 1998; Romanenko et al., 2003; Schroeder et al., 2003). P. gracilis strain B9 produces p-agarase, which degrades the mucilaginous component of the cell wall of the bleached thallus of the red alga Gracilaria gracilis (Schroeder et al., 2003). Another agarase producing Pseudoalteromonas species is P. antarctica which was isolated from the Chilean coast. This bacterium has been shown to produce p-agarase which hydrolyzes agar yielding neoagarotetraose and neoagarohexaose (Vera et al., 1998). Enzymes such as alginases and glycosidases have been reported in P. issachenkonii, a bacterium isolated from the degraded thallus of the brown alga Fucus evanescens (Ivanova et al., 2002b). Chitinases are also reported to be produced by a Pseudoalteromonas sp. strain S91. The cluster gene of this enzyme has been identified and well investigated (Techkamjanaruk and Goodman, 1999). A cold-active cellulase with endocellulase activity has recently been isolated and identified in the antarctic psychrophile P. haloplanktis (Violot et al., 2003). Moreover, P. haloplanktis has been shown to produce a cold-active p-galactosidase with a high catalytic efficiency on natural and synthetic substrates (Hoyoux et al., 2001). A marine bacterium, P. sagamiensis sp. nov. has been reported to produce three types of protease inhibitors, namely, marinostatin, which mediated inhibitory activity against serine proteases; monastatin, a thiolprotease inhibitor against the protease produced by fish bacterial pathogens; and leupeptin, which has inhibitory activity against both thiol and serine proteases (Kobayashi et al., 2003).
1.4.3. Production of antibacterial compounds and toxins
Member species of the genus Pseudoalteromonas produce potent toxins presumably designed to kill competitors in the same ecological niche. Several species demonstrate antibacterial activities, including P. aurantia, P. luteoviolacea, P. rubra, P. citrea, P. tunicata, P. maricaloris and P. phenolica (Gauthier and Breittmayer, 1979; Gauthier and Flatau, 1976; Gauthier, 1979; James et al., 1996; Ivanova et al., 2002e; Isnansetyo and Kamei, 2003a). These Pseudoalteromonas species have been found to be active against both Gram-positive and Gram-negative bacteria. It has been proposed that the antibacterial agents released by the producing strains may provide them with a
40 competitive advantage in the competition for living spa^ce and thus in the maintenance of their population (Holmstrom and Kjelleberg, 1999). These antibacterial agents occur as both small and large molecules. P. bacteriolytica (currently known as Algicola bcicteriolytica, (Ivanova et al., 2004)), has been reported to produce at least six bacteriolytic substances showing a broad spectrum of activity (Takamoto et al., 1994). A polyanionic antibiotic molecule is produced by P. citrea (Gauthier, 1977). P. phenolica sp. nov. has been reported to produce an antibacterial agent, specifically a phenolic compound, against methicillin-resistant S. aureus (MC21-A, a biphenyldiol derivative antibiotic) (Isnansetyo and Kamei, 2003). An extracellular 190 kDa protein has been reported to be responsible for the antibacterial activity of P. tunicata (James et al., 1996). Two pale-orange-pigmented strains of P. ruthenica sp. nov (KMM 3001 and KMM 290), isolated respectively from a mussel, Crenomytilus grayanus, and a scallop, Patinopecten yessoensis, produce a number of antimicrobial compounds (Ivanova et al., 2002d).
The toxic products of Pseudoalteromonas species target higher organisms as well as bacteria. Other toxins have been shown to be effective against invertebrates, algae and mammals. Extracellular products (ECP) of P. atlantica have been shown to cause rapid death in crabs (Cancer pagurus). The ECP target hemocytes, resulting in a rapid decline of the number of circulating blood cells. This damages the nervous system of the crab, ultimately resulting in limb paralysis and a lack of antennal sensitivity (Costa-Ramos and Rowley, 2004). P. maricaloris species isolated from the Australian sponge, Fascaplysinopsis reticulata have been reported to produce cytotoxic lemon-yellow, chromopeptide pigments that prevent the development of sea urchin eggs (Ivanova et al., 2002e). The yellow-pigmented Pseudoalteromonas strain Y has been shown to cause rapid cell lysis of algal blooms including gymnodinoids (Gymnodinium catenatum) and raphidophytes (Chattonella marina and Heterosigma akashiwo) (Lovejoy et al., 1998). The hemolysin of P. issachenkonii is a low-molecular weight, red-brown compound with the formula (C9H7N30S3Na), produced in stationary phase. This compound causes the hemolysis of human, rabbit and mouse red blood cells (Alexeeva et al., 2003). P. tunicata has been shown to inhibit the settlement and colonization of invertebrate larvae, alga and diatoms by producing low molecular weight bioactive molecules (Egan et al., 2001b).
41 1.4.4. Biological activities of Pseudoalteromonas tunicata
The Pseudoalteromonas species that is currently being investigated in our laboratory is P. tunicata. It is a dark green-pigmented bacterium that was first isolated from the surface of a marine tunicate, Ciona intestinalis, from the west coast of Sweden (Holmstrom et al., 1992). The same species was later isolated from Australian waters on the surface of the green alga Ulva lactuca (Egan et al., 2001a). Recently, it has been reported that the same species was isolated from Aarhus Bay, Denmark on the surfaces of U. lactuca, C. intestinalis and Ulvaria fusca (Shovhus et al., 2004), indicating that P. tunicata is widely distributed in different marine waters. It has been reported that this bacterium exhibits biocontrol activity, as it produces a number of bioactive compounds that are inhibitory against algal spores, fungi, invertebrate larvae and bacteria (Holmstrom et al., 1992, 1996; Holmstrom and Kjelleberg, 1999; James et al., 1996; Egan, 2001).
The dark green pigmentation of P. tunicata is due to the production of yellow and purple pigments. The link between the production of pigmentation and antifouling activities (eg. antibacterial, antilarval) was investigated using transposon mutagenesis (Egan et al., 2002a). Four different classes of mutants were produced: yellow, dark purple, light purple and white-pigmented. It was found that the yellow mutants retained full antifouling activities while the purple and white mutants lost some or all of their antifouling activities. Based on the results of the antifouling activity assays and genotypic characterization of the transposon mutants, it was proposed that the production of inhibitory products is linked to the synthesis of the yellow pigment. However, this does not apply to the antibacterial protein since all the transposon mutants with the exception of the white mutants displayed antibacterial activity. Among the different transposon strains, the light purple mutant showed reduced antibacterial activity suggesting that the antibacterial protein and production of purple pigmentation are jointly regulated (Egan, 2001). A wmpR gene similar to toxR of V. cholerae and cadC of E. coli was found to be interrupted by a transposon insertion in the white mutant. It has been suggested that WmpR is essential for the expression of the antifouling agents in P. tunicata (Egan et al., 2002b).
42 The identity of the different antifouling compounds has been elucidated. Inhibition of larvae of B. amphitrite and C. intestinalis is due to an extracellular inhibitory molecule of less than 500 Da, which is heat stable, polar and uncharged (Holmstrom et al., 1992). An extracellular antialgal compound was observed to display inhibition against algal spores of the algae U. lactuca and Polysiphonia sp. The antialgal compound is heat sensitive and polar with a molecular size somewhere between 3-10 kDa (Egan et al., 2001b). P. tunicata was also observed to have antifungal activity against a wide range of fungal and yeast isolates. The yellow pigment was found to be the anti-fungal compound (Franks et al., submitted for publication).
The 190 kDa antibacterial protein produced by P. tunicata has been identified and characterized (James et al., 1996). The production of the antibacterial protein was demonstrated to occur during the stationary phase of growth. Interestingly, P. tunicata cells are sensitive to the antibacterial protein when the cells are in exponential phase. However, it was observed that P. tunicata cells could clearly survive at high bacterial densities even in the presence of the self-inhibitory protein. The sensitivity and resistance of P. tunicata to its own antibacterial protein has been investigated and the results are presented in chapters 2 and 3 of this thesis. The antibacterial protein has been shown to inhibit marine, soil and clinical bacterial isolates. It is very effective against Staphylococcus aureus, Bacillus subtilis, B. cereus, B. globi, B. primulus, Enterococcus faecalis, E. coli, Proteus mirabilis, P. aeruginosa, P. putida and Alcaligenes faecalis (James et al., 1996; Stelzer, 1999).
1.5. Aims of this study
The distinctive features of the marine bacterium P. tunicata suggest that this bacterium plays an active role in surface associated communities. The ability of this organism to produce an array of bioactive compounds which target different biofouling organisms indicates a competing role in controlling the establishment of marine biofouling communities. Moreover, the association P. tunicata with higher marine organisms suggests unique adaptation traits in the marine ecosystem.
43 Studies to date aimed at the isolation, purification and characterization of the antibacterial protein produced by P. tunicata revealed that there is still much to be learned about this protein, particularly its mode of action. While most of the current works on P. tunicata focuses on the production and regulation of antifouling agents, no investigation has been carried out to identify mechanisms for colonization and adaptation to surfaces. Molecular, genetic and ecological studies will contribute to the further understanding of the biology and physiology of this bacterium and how it successfully colonizes higher marine organisms. The findings of this thesis will also potentially contribute to the development of techniques and methods for the study of the ecology of surface associated bacteria as well as developing strategies to control biofilm formation in the marine environment.
The work described in this thesis aims to provide a better understanding of the mode of action of the antibacterial protein, AlpP and the role of this protein with respect to the colonization mechanisms and physiological responses of P. tunicata on living surfaces. The study focuses on the following specific goals.
a. To elucidate the mode of action of AlpP toxicity towards the producing organism P. tunicata, by identifying genes responsible for resistance and sensitivity using transposon mutagenesis, and to determine whether the primary event of killing is bacteriolytic or bacteriostatic (Chapters 2 and 3).
b. To investigate the mechanisms of colonization of P. tunicata to surfaces (biotic and abiotic), specifically on the surface of the green alga Ulva lactuca (Chapter4).
c. To investigate the mechanisms by which P. tunicata is able to adhere to and utilize cellulose as source of carbon and energy (Chapter 5).
44 2. Generation and analysis of Pseudoalteromonas tunicata transposon mutants with altered sensitivity to its autolytic protein, AlpP
2.1. Introduction
The marine bacterium Pseudoalteromonas tunicata produces a range of bioactive agents that specifically target different classes of fouling organisms (Holmstrom et al., 1996; James et al., 1996; Holmstrom and Kjelleberg, 1999; Egan et al., 2001b). One of these bioactive compounds is a novel 190 kDa antibacterial protein, an extracellular product produced during the stationary phase of growth (James et al., 1996). N-terminal sequence analysis of the purified protein showed no similarities in its protein sequence to other proteins in the Swiss Protein data base, which indicates that it represents a novel antibacterial agent (James et al., 1996). SDS-PAGE analysis indicated that the 190 kDa antibacterial protein is made up of two subunits of 60 and 80 kDa, linked together by noncovalent bonds. N-terminal amino acid sequencing of these two protein bands showed that the first 27 amino acids of both subunits are identical raising the question as to whether these protein subunits are encoded on one or two genes. Oligonucleotide probes targeting the N-terminal region of the antibacterial protein, were used to probe Southern blots of P. tunicata genomic DNA digested with several different restriction enzymes. In all cases, a single band was observed, indicating that both subunits originate from the same gene (James, 1998). The entire antibacterial protein gene, termed alpP (autolytic protein Pseudoalteromonas) has been sequenced and the translated amino acid sequence showed identity with only two hypothetical proteins, more recently entered into GenBank. These matches were found within the genomes of Magnetococcus sp. MC-1 (GenBank accession number: ZP_00044814) and Caulobacter crescentus CB15 (GenBank accession number NP_ 419374), with identities of 32% and 27% respectively, both being hypothetical proteins of no known function (Mai- Prochnow et al., 2004). The AlpP protein was found to be auto-toxic during the logarithmic phase of growth (James et al., 1996). However, it was observed that P.
45 tunicata cells survived at high bacterial densities in the presence of the AlpP protein (for example when grown as colonies on agar plates) during the stationary phase of growth (James et al., 1996). Cells that are sensitive or resistant to AlpP are also present in P. tunicata biofilms. Autolysis mediated by AlpP occurs inside biofilm microcolonies, resulting in the formation of hollow voids within these structures and leaving behind a subpopulation of alive and persistent cells (Mai-Prochnow et al., 2004).
Microorganisms exhibit autolysis or “self-killing” as an adaptive and regulatory response to adverse environmental conditions, such as nutrient limitation, antibiotics and toxic chemicals, extreme changes in temperature and variations in osmolarity and pH (Gerdes, 2000). Autolysis has also been shown to occur during bacterial development and processes such as cell division and cell separation (Lewis, 2000). Recently, autolysis has been linked to programmed cell death (PCD), a concept of “altruistic suicide” whereby microbial communities eliminate cells programmatically during development as well as eradicating defective cells during exposure to harsh conditions, thus ensuring survival of the genome (Frolich and Madeo, 2000; Lewis, 2000). In P. tunicata, its autolytic behavior may play an important part in its role as an antifouling bacterium. For example, autolysis may suggest a mechanism for controlling population density, and prevent overgrowth on surfaces of host organisms, at which P. tunicata may produce antifouling agents. Autolysis by P. tunicata may also suggest an important survival mechanism during nutrient limitation (as AlpP is produced during stationary phase), where autolysis may be an altruistic process that would result in the release of nutrients from the lysed cells for maintenance and survival of the remaining P. tunicata population. The mechanism of the autolytic activity of AlpP in P. tunicata is unclear and further investigation is needed. Understanding the function of the antibacterial protein AlpP produced by P. tunicata will provide information on its mechanism of action and understanding its ecological role.
This chapter focuses on the identification of P. tunicata genes encoding traits mediating resistance and sensitivity to AlpP protein. Mutants from a P. tunicata transposon library were screened for increased resistance and sensitivity to AlpP. The AlpP-sensitive and - resistant mutants were selected and identified based on their phenotypic and genotypic characteristics. The genetic information obtained from this study will provide an understanding of the mechanisms of resistance and sensitivity of P. tunicata to its AlpP.
46 2.2. Materials and Methods
2.2.1. Bacterial strains and culture conditions
Transposon donor E. coli SmlO (containing the pLOF mini-TnlO system, Alexeyev et al., 1995) was grown and maintained at 37°C on LB 10 agar plates (Appendix I) containing 85 pg/ml kanamycin (Km) and 100 pg/ml ampicillin (Amp). Transposon recipient P. tunicata (SmR) (Egan, 2001) was grown and maintained at room temperature on VNSS agar plates (Appendix I) containing 100 pg/ml streptomycin (Sm). Wild type P. tunicata was grown and maintained at room temperature on VNSS agar plates.
2.2.2. Preparation of P. tunicata cell-free concentrated supernatant
A cell-free concentrated supernatant of P. tunicata was prepared using the method developed by Mai-Prochnow et al (2004). Briefly, a 20 ml overnight culture of wild- type P. tunicata was added to two litres of Marine Minimal Media (MMM) with 0.2 % trehalose as the sole carbon source (MMM-trehalose) (Appendix I). The mixture was grown shaking for 48 h at room temperature and cells were harvested at 12, 000 x g for 20 min at 20°C. The supernatant was removed and the cells were resuspended in 1 ml of MMM-trehalose for every 2 grams of the wet weight of the cells. This mixture was incubated at room temperature for 24 h without shaking. The concentrated supernatant containing the AlpP protein was separated from the cells by centrifugation at 14, 000 x g for 1.5 h at 4°C. Prior to use, the supernatant was filter sterilized using a 0.22 pm filter. Aliquots of the cell-free concentrated supernatant were stored at -4°C. Bio-Rad Protein Quantification KIT™ was used with bovine serum albumin as the standard. A drop test assay (James et al., 1996) was performed to check the autolytic activity of the cell-free concentrated supernatant.
47 2.2.3. Fractionation of the AlpP protein from cell-free concentrated supernatant
Fractionation of the cell-free concentrated supernatant was performed using the ion- exchange method described in James (1998). Filter sterilized cell-free concentrated supernatant was dialyzed overnight in 20 mM Tris-HCl, pH 7.5 prior to ion-exchange fractionation using a 5 ml Eco-Pac hydroxyapatite cartridge (Bio-Rad). The dialyzed cell-free concentrated supernatant was diluted with 20 mM Tris-HCl, pH 7.5 (1/10 of its contained total volume). The diluted supernatant was loaded on the cartridge at a rate of 30 drops per min. The column was eluted with 10 ml of 0.1 M NaCl in 20 mM Tris- HCl buffer, pH 7.5 followed by 0.3 M NaCl in 20 mM Tris-HCl, pH 7.5. The eluate from the wash solution contains the AlpP protein and the purified fraction was stored at -20°C. BioRad Protein Quantification KIT™ was used with bovine serum albumin as the standard. A drop test assay (James et al., 1996) was performed to check the autolytic activity of the collected fractions of the cell-free concentrated supernatant.
2.2.4. Transposon mutagenesis
The transposon mutagenesis protocol established by James (1998) was used to generate mutants of P. tunicata with altered sensitivity to AlpP. Ten millilitre overnight cultures of both donor E. coli SmlO and recipient P. tunicata (SmR) were prepared as described in section 2.2.1. The E. coli strain was grown in LB medium at 37°C with shaking (Appendix I). P. tunicata (SmR) was grown in VNSS medium at room temperature with shaking. All media were supplemented with antibiotics as described in section 2.2.1.
Overnight cultures of both donor and recipient cells were mixed at a volume ratio of 1:3 (50 pi E. coli + 150 pi P. tunicata) in 5 ml of wash solution (50 % NSS: 50 % 10 mM MgSCL) and mixed by gently inverting the tube five times. The mixture of donor and recipient cells was suction-filtered through a 0.22 pm filter (2.5 cm diameter) and washed with 5 ml of wash solution. The filters were then placed cell side up onto LB 15 agar plates (Appendix I) containing 3 mM isopropyl-fl-D-thiogalactoside (IPTG) and incubated for 4 to 5 h at 30°C. After incubation the filters were placed into Eppendorf tubes with 1 ml of NSS (Appendix I) and vortexed. The cell suspension was spread on
48 to selective VNSS plates containing Km (85 pg/ml) and Sm (100 pg/ml) and incubated at room temperature for 24 h.
Recipient P. tunicata strains carrying the mini-TnlO transposon were picked from the selective VNSS plates and transferred to fresh VNSS plates containing Km (85 pg/ml) and Sm (100 pg/ml). The mutant strains were inoculated using a sterile toothpick, according to a template pattern onto the VNSS plates. Each template contained 48 mutant strains. The plates were incubated at room temperature for 24 h.
Screening of mutants for altered resistance or sensitivity to AlpP was done by transferring mutant strains from the original template, using a sterile metal grid corresponding to the template, onto VNSS plates which contained serial dilutions of P. tunicata cell-free concentrated supernatant. The serial dilutions were carried out at the following total protein concentrations in the cell-free concentrated supernatant: 1.64 mg/ml, 0.890 mg/ml, 0.410 mg/ml, 0.205 mg/ml and 0.102 mg/ml. The selection of the mutants was based on their sensitivity to the AlpP protein (within the cell-free concentrated supernatant) relative to that of the P. tunicata wild type. The mutant strains growing at all protein concentrations were classified as resistant mutants, and mutants that were inhibited in growth were classified as sensitive mutants. The selected mutant strains were tested in the drop test assay (James et al., 1986) and compared to P. tunicata wild type to confirm their sensitivity to AlpP. Selected AlpP-sensitive and -resistant mutant strains were grown in VNSS medium containing Km (85 pg/ml) and Sm (100 pg/ml) and stored in 30 % (v/v) glycerol at -80°C.
2.2.5. Phenotypic characterization of P. tunicata AlpP-sensitive and -resistant transposon mutants
2.2.5.1. Altered sensitivity to AlpP
Overnight cultures of mutant strains were exposed to different concentrations of the supernatant (section 2.2.4.) containing the AlpP protein. To further test their resistance or sensitivity to AlpP, selected mutant strains were assayed in a microtitre plate
49 containing 180 pi fresh VNSS, 10 jlxI overnight culture of selected mutant strain, 10 pi of AlpP protein containing a specified concentration which was the same as the concentrations used in the drop test assay. This mixture was incubated at room temperature and the absorbance was measured at 610 nm after 12 h of incubation. To determine if the AlpP-sensitive and -resistant mutant strains retained their level of response to AlpP at different stages of the growth curve, drop test assays were performed in early logarithmic and mid-stationary phase of growth. A clearing zone observed around the drop of AlpP protein added on the lawn of mutant strains was scored as a sensitive response. Sensitivity to the AlpP protein was expressed as a Maximum Inhibitory Dilution (MID), which was scored as the reciprocal of the last serial dilution showing a visible clearing zone on the lawn of mutant strains. All of these experiments were carried out in triplicate.
2.2.5.2. Growth curves
The growth rates of the AlpP-sensitive and -resistant transposon mutants and wild type P. tunicata were compared. The growth curve assay was performed in 50 ml side arm flasks containing 10 ml of VNSS medium for wild type and VNSS medium with antibiotics, Km (85 pg/ml) and Sm (100 pg/ml), for the AlpP-sensitive and -resistant transposon mutants. Ten microlitres of an overnight culture was inoculated into each appropriate flask and incubated with shaking at room temperature. The growth was monitored by measuring the optical density of the cultures at 610 nm over a 24 h period. The experiments were carried out in triplicate.
2.2.6. Genotypic characterization of P. tunicata AlpP-sensitive and -resistant transposon mutants
2.2.6.1. Genomic DNA extractions
The genomic DNA of the AlpP-sensitive and -resistant transposon mutants was extracted using the XS-buffer (Xanthogenate-SDS) extraction protocol (Tillet and Neilan, 2000). Three to four ml of an overnight cultures of transposon mutant strains
50 were centrifuged for 5 min at 6, 000 x g. The cells were resuspended in 1 ml of XS- buffer (Appendix I). The suspension was incubated at 65°C for 2 h or 70°C for 1 h then vortexed for 10 sec and incubated on ice for 30 min. After incubation, the suspension was centrifuged at high speed (10, 000 x g) for 10 min at 4°C. The clear supernatant was transferred to a new tube, to which 1 ml of isopropyl alcohol was added to precipitate the genomic DNA. The mixture was incubated on ice for 5 min and the precipitated DNA was removed by spooling using a glass hook. The spooled DNA was washed twice in 70 % EtOH and excess EtOH was allowed to drain before suspending in 400 pi of Molecular Grade Water™ (Brinkmann). Following extraction, genomic DNA was visualized on a 1 % agarose gel alongside a molecular standard (7-DNA digested with EcoRUHindlll) to check for integrity, concentration and RNA contamination. If there was a large amount of RNA contamination detected on the gel, the extracted DNA was treated with RNAse, then extracted with phenol/chloroform/isoamylalcohol and precipitated with ethanol (Appendix II).
2.2.6.2. Panhandle PCR method for sequencing the regions flanking the inserted transposon
The panhandle polymerase chain reaction (panhandle-PCR) method, which was modified by Tillet (2000) from Siebert et al (1995), was used to obtain sequence information from the genes disrupted by the mini-TnlO transposon. This is possible because the DNA sequence of this transposon is known. This PCR approach employs the concept of "suppression PCR" using adaptor molecules which contain inverted terminal repeats. Generated PCR products containing adaptor sequences at both sides form "panhandle" structures following every denaturation step. Further amplification is suppressed because these structures are more stable than the primer-template hybrid. Moreover, PCR products generated by specific primers and the primer for the adaptor contain the adaptor sequence only in one end and therefore do not form the "panhandle" structure, permitting further PCR amplification.
51 2.2.6.2.1. Preparation of adaptor ligated DNA
The genomic DNA extracted from the AlpP-sensitive and -resistant transposon mutant strains, was used for restriction digestion and the ligation of adaptor molecules in a one- step process. One microgram or less (ca. 500 ng) of the genomic DNA, 1 pi of adaptor 1,5’CTAATACGACTCACTATAGGGCTCGAGCGGCCGCCCGGGCAGGT 3’(10 pmol/pl stock), 1 pi of adaptor 2, 5’P-ACCTGCCC-NH23’ (10 pmol/pl stock), 40 mM ATP, 2.5 units T4 ligase, 10 units of blunt-end restriction enzyme (Dral, EcoRV, Hindi, Hpal, Pvull, Rsal, Seal, Sspl or Xmnl), 1 pi lOx One-Phor-All buffer PLUS and milli-Q water to make 20 pi were mixed and incubated at 20°C for 16 h. The reaction was deactivated at 68°C for 10 min. The DNA was precipitated using ethanol (Appendix II) and resuspended in 50 pi of milli-Q water. This solution was used as template DNA for the panhandle-PCR reaction.
2.2.6.2.2. Panhandle-PCR
The 20 pi volume of PCR mix contained 1 pi of template DNA, 2 pi Taq lOx reaction buffer containing 25 mM MgCl2, 1 pi 10 mM dNTP mix, 10 pmol of adaptor primer 1, 10 pmol of gene specific primer (Tnl0C-5’GCTGACTTGACGGGACGGCG3’ or TnlOD-5’ CCTCGAGCAAGACGTTTCCCG3’), 1 unit Taq and PFU in 10:1 ratio and milli-Q water. The cycle parameters used were as follows: 95°C for 30 sec for the denaturation step and 68°C for 7 min for annealing/extension. The cycles varied between 25 and 30 depending on the template.
2.2.6.2.3. Preparation of PCR templates and DNA sequencing
The generated PCR products were visualized on a 1 % (w/v) agarose gel and compared to molecular weight standards to estimate the size and concentration of the product. The single band products were excised from the gel and purified using QIAquick Gel Extraction Kit ™ according to the manufacturer's instruction. The final concentration of the purified panhandle-PCR products was determined by UV-spectroscopy using DNA/Oligo Quantitation program in a Beckman DU-600 spectrometer.
52 Fifty to one hundred nanograms of double stranded template DNA, 4 pi of CSA buffer, 15-20 pmol of sequencing primer (SI, 5’GGGTATTCAGGCTGACCC3’ for panhandle PCR products of TnlOC or TnlOD and various primers, see Table 2-2 to Table 2-5 for primer walking strategy) and 4 pi of BigDye™ terminator cycle sequencing reaction mix (Applied Biosystems) in a final volume of 20 pi was used in a standard DNA sequencing reaction. Amplification was conducted using the following parameters: 94°C for the initial denaturation step followed by 25 cycles of 94°C for 10 sec, 55°C for 5 sec and 60°C for 4 min. After the PCR amplification, the sequencing mixture was cleaned and purified using Butanol Purification of Dye Terminator Sequencing protocol outlined below (Tillet, 2001). Ninety microlitres of Molecular Grade Water™ (Brinkmann) was added to each of the sequencing reaction tubes and transferred to 0.5 ml Eppendorf tubes containing 100 pi of phenol. The tubes were vortexed for 5 sec and centrifuged (—12, 000 x g) for 4 min. The aqueous phase was removed and transferred to a 1.5 ml Eppendorf tube containing 900 pi of n-butanol. The mixture was vortexed for 10 sec until the aqueous phase was completely mixed and then centrifuged at 12, 000 x g for 10 min. The supernatant was removed and the tubes were dried in a speedivac for 5 min to remove the remaining n-butanol. Separation of the sequencing products was carried out on an ABI 377 DNA sequencing system at the Sydney University Prince Alfred Macromolecular Analysis Centre (SUPAMAC).
2.2.7. Sequence data analysis
The DNA-sequence-electropherograms were analyzed using ABI-PRISM software. The results were processed and multiple sequence alignments were performed using a Staden Package System (Medical Research Council-Laboratory of Molecular Biology, Cambridge, England). The complete consensus DNA sequence was compared with sequences in the GenBank-database available through the National Center for Biotechnology Information (NCBI) web site (http://www.ncbi.nlm.nih.gov). The GCG- software package provided by the Australian National Genomic Information Service (ANGIS) web site (http://www.angis.org.au/WebANGIS/) and the ExPASy (Expert Protein Analysis System) site (http://expasy.proteom.org.au/index/html) of the Swiss Institute of Bioinformatics (SIB) were also used for sequence analysis.
53 2.3. Results
2.3.1. Generation of P. tunicata transposon mutants
To identify the genes in P. tunicata which encode for traits that mediate resistance and sensitivity, respectively, to AlpP, transposon insertion mutants of P. tunicata were generated. The transposon mini-TnlO system was used for mutagenesis and this generated a library of 10,080 transposon mutant strains of P. tunicata. Transposon mutagenesis produced several mutant strains with altered sensitivity to AlpP (Figure 2-1). The genes mutated in AlpP-sensitive and -resistant transposon mutant strains, provided information on how P. tunicata responds to its autolytic protein, AlpP. Out of 10,080 transposon mutants screened for altered sensitivity to AlpP, 54 AlpP- resistant mutants and 23 AlpP-sensitive mutants were selected after the first initial screening using a drop test assay (Table 2-1). A transposon mutant strain with a MID value < 4 was scored as resistant whereas those with a MID value > 4 were scored as sensitive.
54 B » « • • « » c • ♦ • • f f • • * » Ji ♦ <♦'»»* a « to to i> J * • »
<1 • . J > j «. to. > * V «> «• a* to* ^ ^ b A f *> V'
• • i « » •
Figure 2-1. Selection of P. tunicata transposon mutant strains from the original template. The template was made up of 48 mutant strains grown on VNSS agar plates containing Km (85 pg/ml) and Sm (100 pg/ml) for 24 h (A). The template was transferred to each of the VNSS agar plates containing dilutions of P. tunicata active concentrated supernatant from the highest to the lowest concentration (B, C, D, E and F). Note that some mutants failed to grow on VNSS agar plates containing the undiluted concentrated supernatant (B). Some mutants were showed to be sensitive up to a 4x dilution of the concentrated supernatant (D).
55 2.3.2. Phenotypic characteristics of P. tunicata transposon mutants with altered sensitivity to its autolytic protein, AlpP
2.3.2.1. Analysis of the AlpP resistant and sensitive phenotypes
After initial screening using the drop test assay, 54 AlpP-resistant and 23 AlpP-sensitive mutants were selected from the transposon library. The selected mutants varied in their sensitivity to AlpP protein showing a MID value of 0 for the most resistant mutant and a MID value of 16 for the most sensitive mutant. Only one mutant (RM1) showed complete resistance to the AlpP protein. This AlpP-resistant mutant consistently demonstrated complete resistance after repeated subculturing in different concentrations of AlpP. In order to confirm the phenotypic characteristics of the selected strains, a series of drop tests and microtitre plate assays were carried out. Out of 54 AlpP-resistant mutant strains initially selected from the transposon library, only 4 AlpP-resistant mutant strains consistently showed resistance to the AlpP protein. Similar to the AlpP- resistant mutants, many AlpP-sensitive mutant strains were not phenotypically stable. Only 5 AlpP-sensitive mutant strains were consistently sensitive out of the 23 AlpP- sensitive mutant strains. The AlpP-resistant mutants were designated as RM1, RM3, RM4 and RM5. The AlpP-resistant mutants RM3, RM4 and RM5, were found to display the same degree of resistance. The 5 AlpP-sensitive mutant strains were designated as SMI, SM2, SM4, SM5 and SM6 (Table 2-1). Three AlpP-sensitive mutant strains (SMI, SM2 and SM5) displayed the same degree of sensitivity to AlpP. Two AlpP-sensitive mutant strains (SM4 and SM6) were very sensitive to the AlpP protein, exhibiting a MID value of 16. The AlpP-sensitive mutant strains showed a distinct clearing zone in the drop test assay (Figure 2-2).
In order to determine if the selected AlpP-sensitive and -resistant transposon mutant strains retain their phenotypic characteristics at different stages of growth, drop test assays were performed in early logarithmic and mid-stationary phase (Table 2-1). Four AlpP-sensitive mutant strains showed altered sensitivity when exposed to the AlpP protein during mid-stationary phase. There was a 2-fold decrease in sensitivity observed in three AlpP-resistant mutant strains when exposed to AlpP protein during the mid-
56 stationary phase of growth. The AlpP-sensitive mutant SM4 showed the highest decrease in sensitivity, with a 4-fold reduction in sensitivity in log phase compared to its sensitivity to AlpP in mid-stationary phase of growth. No changes in MID value were observed in AlpP-resistant mutant strains when assayed both in log and mid-stationary phase of growth.
Table 2-1. MID values to the AlpP protein of stable P. tunicata transposon mutant strains.
MIDa (drop test assay) Mutant strains Logarithmic Phase Mid-stationary Phase P. tunicata wild type 4 0 SMI 8 8 SM2 8 4 SM4 16 4 SM5 8 8 SM6 16 8 RM1 0 0 RM3 2 2 RM4 2 2 RM5 2 2 d MID — Maximum Inhibitory Dilution. Values represent the last serial dilution showing a visible clearing zone of the bacterial lawn grown on VNSS agar plates.
57 Bacterial P. tunicata SMI SM2 SM4 SMS strains wild-type
MID 4 8 8 16 8
Drop test assay • 9•
Figure 2-2. P. tunicata wild type and AlpP-sensitive transposon mutants (logarithmic growth) displaying different responses to the different dilutions of the autolytic protein, AlpP. Each drop represents a concentration of AlpP protein from highest to lowest concentration in a counter clockwise direction. MID values (maximum inhibitory dilution) represent the last serial dilution showing a visible clearing zone of the bacterial lawn grown on VNSS agar plates. The SMI mutant showed dark purple pigmentation identical to the dark purple transposon mutant generated by Egan (2001) (see section 2.3.3.1.1).
2.3.2.2. Growth of both AlpP-sensitive and -resistant transposon mutants
The growth of the AlpP-resistant and -sensitive mutant strains was observed over 24 h by measuring the optical density of the cultures at 610 nm. All of the AlpP-sensitive mutant strains, except SM5 and SM6, showed the same growth pattern as that of the wild type. Strain SM5 and SM6 showed growth inhibition after 4 h of incubation and later showed slower logarithmic growth. This suggests that these mutants possibly undergo autolysis but eventually build up an antidote, which counteracts the effect of AlpP, thus allowing them to grow slowly after several hours. The logarithmic phase of growth of SM5 and SM6 started 8 h after incubation, which is 4 h later than the onset of logarithmic phase of growth for the other sensitive mutant strains (Figure 2-3A). This result suggests that genes mutated in SM5 and SM6 are important for growth in the logarithmic phase.
58 P. tunicata wild-type
Time (h)
P. tunicata wild-type RM1 -*-RM4
Time (h)
Figure 2-3. Growth curves of P. tunicata mutant strains. A) AlpP-sensitive mutant strains and B) AlpP- resistant mutant strains. The growth assay was performed in 50 ml side arm flasks containing 10 ml of VNSS medium for wild type and VNSS medium with antibiotics, Km (85 pg/ml) and Sm (100 pg/ml), for the AlpP-sensitive and -resistant transposon mutants. The optical density was measured at 610 nm.
59 All of the AlpP-resistant mutant strains showed the same pattern of growth compared to the wild type. The onset of the logarithmic phase of growth occurred 3 h after incubation (Figure 2-3B).
2.3.3. Genotypic characterization of P. tunicata transposon mutants
2.3.3.1. DNA sequence analysis of P. tunicata AlpP-sensitive mutants
The five AlpP-sensitive mutants were further analyzed for their genotypic characteristics. Using the pan-handle PCR method, the initial sequence analysis of the regions flanking the transposon in each of the 5 AlpP-sensitive mutants showed that they had been disrupted in 5 different DNA regions. Using primer walking and additional pan-handle PCR reactions on the known sequenced region from both sides of the transposon in the mutant, substantial DNA sequence information was obtained for analysis. The DNA sequences were assembled, aligned and analyzed using the Staden package software (Medical Research Council-Laboratory of Molecular Biology, Cambridge, England). The resulting consensus nucleotide sequence was submitted to the ORF-finder and BLASTX and BLASTP programs of NCBI. The ExPASy (Expert Protein Analysis System) program packages of the Swiss Institute of Bioinformatics (SIB) were used for the analysis of deduced amino acid sequence of the identified open reading frames.
2.3.3.1.1. Sequence analysis of the SMI mutant
The sequence analysis showed that the transposon had disrupted a 702 bp open reading frame (ORF) with 100 % identity to the dppB gene of P. tunicata (GenBank accession number AF441247) (Egan, 2001). This gene was identified in a dark purple transposon mutant generated by Egan (2001). The DppB protein is similar to the YvrO, ABC-transporter protein of Bacillus subtilis (GenBank accession number AJ223978) and the TptC, ATP-binding protein of Streptococcus cristatus (GenBank accession
60 number AAB97961). The genotypic characteristics of this mutant have been reported by Egan et al (2002a).
2.3.3.1.2. Sequence analysis of the SM2 mutant
A total of 2600 bp of nucleotide sequence flanking the transposon inserted in the SM2 mutant was determined by pan-handle PCR and primer walking using primers shown in Table 2-2. Several primers were designed to ensure the fidelity of the sequence data. Sequence analysis revealed that the mutation occurred in a 1455 bp ORF which encodes for 484 amino acids (aa) (Figure 2-4). The DNA sequence generated has been entered in the GenBank databases under accession number AY751752. The ATG codon starts at base position 103, which is preceded by a putative ribosomal binding sequence (RBS) (5’GCAGAT3’) 4 bp upstream. The putative promoter region, based on homology with consensus sequences for -10 and -35 promoter elements of E. coli, was identified 25 bp upstream of the start codon. The putative sequence terminator was identified 189 bp down stream of the stop codon. Only one ORF was identified from this DNA region.
The deduced aa sequence of this ORF indicates that the protein has a predicted isoelectric point (pi) of 4.20 and a molecular weight of 52.8 kDa. The ORF gene product has 43 % similarity to Chi A, a chitinase precursor of Vibrio harveyi (GenBank accession number AF323180). Other proteins with homology to the ORF1 gene product are ChiA, a chitinase of V. cholerae with 44 % similarity (GenBank accession number AAF95941), and ChiA, an extracellular chitinase A of Aeromonas hydrophila with 44 % similarity (GenBank accession number AAF70180). The PSORT (Prediction of Protein Localization Sites) program (Nakai and Kanehisa, 1991) predicted no transmembrane region on the ORF1 gene product and that the gene product is located in the bacterial cytoplasm.
61 Table 2-2. Oligonucleotides used in the primer walking strategy to sequence the DN> regions flanking the inserted transposon in the SM2 mutant.
Oligonucleotide Sequence (5’-3’) Target region
SM2TnlOC-Dl CATGATTGATACCTTCCCTGC 1335-1355
SM2TnlOC-Dl A GTCGACCTATTTATGTTGGCG 1632-1652
SM2TnlOC-DlB T GT G AAATCC AAGCGAT GC 2213-2231
SM2TnlOC-DlC TTAACAGCTTCTTGCGGTACG 2502-2522
SM2TnlOD-D5 C GGT A AT GGTT AGGGT AGC 928-946
SM2TnlOD-D5A CGTTTAGGTCGACATTCTCCC 765-785
SM2TnlOD-D5B C GTT GT ATTT G A AGT G AC CTCGG 392-414
SM2TnlOD-D5C TAGGTCGACCGCATTTACTCC 1620-1640
SM2TnlOD-D5D CATGCTGCGCCTTTACTAAACG 1177-1197
62 1 CTGAAAGTGACGCTACATTAAGCTCACTGATAAGCTCAAGCACTAGTTTT 50
-35 region -10 region RBS 51 TCGGCATCAGTAAATGGGGAATATGTTGTTGGATTACGACTAGCAGATTC 100
101 TCatggcgcctatagtgatggaatctatcaagtcaggtgtcgcttcaagt 150 MAPIVMES I KSGVASS
151 aatcaagctccaattgcacaattaggtaacgacatgagcacagtacttgg 200 NQAPIAQLGNDMSTVLG
201 gcagtcagttactttatcctgcgagtattgttttgatcctgaaggggacg 250 QSVTLSCEYCFDPEGDE
251 aattgagctacaactggcaattattagggcaacccgacaccagtactagc 300 LSYNWQLLGQPDTSTS
301 gtgcttgaatccaccaccagcccaattgcaactcttaacccagatatgat 350 VLESTTS PIATLNPDMI
351 aggcgaatatttagttgcagtgataatttctgatggcgaaaccgaggtca 400 GEYLVAVI ISDGETEVT
401 cttcaaatacacagataatcgatgtaagaaataaccaaaaaccaatctca 450 SNTQI IDVRNNQKPIS
451 aaaatatttgctcctttaacagcttcattaggagagagagtaattctaga 500 K I FAPLTASLGERVILD
501 tggttcacacagctttgatcctgaaggtgctgaacttaactatgaatgga 550 GSHS FDPEGAELNYEWK
551 aaataattgatcagccaaatcatgatgagatatatgaagactcttctgct 600 I IDQPNHDEIYEDSSA
601 ttagcatatttcacccctatgagtatcggcgactatattgtctccttgcg 650 LAYFTPMSIGDYIVSLR
651 aaccaatgatggaactcaatattcagatccagaaacaacggctatttctg 700 TNDGTQYSDPETTAISV
701 tcaaagaaaatcaagcacctgtcattgtaattcaaggagatcttaatcga 750 KENQAPVIVIQGDLNR
751 actatttgcattaggggagaatgtcgacctaaacgcagcggcaaagtcta 800 TICIRGECRPKRSGKVY
801 cgacccagaaggtgaagcgattacgttcctttggacattgcacaaaccag 850 DPEGEAITFLWTLHKPD
851 ataactctaccgtaaccctacaagatgatagtgccaaaatactacagttc 900 NSTVTLQDDSAKILQF
901 actcctgatattgcaggtatttatatcgctaccctaaccattaccgatcc 950 TPDIAGIYIATLTITDP
63 951 agcaaataacttatctagtgaatcaataacagttgaagttactgatctat 1000 ANNLSSESITVEVTDLS
1001 ctgatgttctaactggaacagttaaagggcgcattgtcgacacgactttg 1050 DVLTGTVKGRIVDTTL
1051 agcggtgtagaaaatgcactacttagtatcaacggtaaagaatatagctc 1100 SGVENALLSINGKEYSS
1101 tgatcaagaaggcttttttaatattactctggatatagaagaaggaaacg 1150 DQEGFFNITLDIEEGNA
1151 ctataaagattcttaccgcagacagccgtttagtaaaggcgcagcatgtc 1200 IKILTADSRLVKAQHV
1201 actgcagtaattgctcagcaagattttattatcgatttaggcgaatcttt 1250 TAVIAQQDFI I DLGESF
1251 cgttccagttaaacaggaagtagatacttacttatggacctgtggcgact 1300 VPVKQEVDTYLWTCGDY
1301 attcgggcccagaaaatattaatttacgctttaacatgattgataccttc 1350 SGPENINLRFNMIDTF
1351 cctgcgacaaacaagttcactactcactttgatgaaacatttagtttcgc 1400 PATNKFTTHFDETFSFA
1401 cattgatagcaataaaagtatagcgcttccttcaacggcttcttatgaat 1450 IDSNKSIALPSTASYEL
1451 tattcgtagatgatgaagtaacaattacagcgccaggacctaaagtcacc 1500 FVDDEVTITAPGPKVT
1501 atatattatgcacctattttaggggcaatgaacattgtcactatttgtaa 1550 IYYAPILGAMNIVTICN
1551 taaataaATTAACAGGAAACTATAAAAATGAAATTAATCAGCACCTTAAC 1600 K *
1601 AGCTCTCGCATTTTCTTTTGGAGTAAATGCGGTCGACCTATTTATGTTGG 1650
1651 CGGAACCCAGTATAGCTATACCGAAGCTGATTTTAATGCCATTCATCTTG 1700
1701 ATGGTAACGTTATCGGTTATGAAACAGAAATAGATGTCTTACTAGATGAA 1750
1751 TACGATGAATATCTAGGCGGATATCCAGAGCTTATTTTYGATGTGAGCGG 1800
1801 CCACGTGGTAGGAACTATTGGTCGCCTGCCTATGAAAGTAGAAATGGATG 1850
1851 TTACCTGCTCGGGCAAAGACATAGGTTACGACGCAGATCTGATAATCAAT 1900
1901 AGAAGGGGCCAATTAACCCCTGCCCGTTTTTCTATCTATAACAGACTTAC 1950
1951 AACCGAAGGCCAATGTAAGAAACTAAAGCTAAAAATAAAAAAAATGGATT 2000
64 2001 CATCAACTTCTGATACTGAAGCAACCATCGACGCAT'TCAACGTTAATCTA 2050
2051 ACTATTTATCTCAACTTAATGTAAGGAACACCCCCATGAAAACACCTATT 2100
2101 AATGTTATTCATTTTTATCACACTCAATATTTTTTCATTTAGTGCATTCT 2150
2151 CTTCTGAAGAACAGGATGTAAATCCATTAAGACGGCAATGTTGCGATGAC 2200
2201 C G AG AC AG AAT C T GT G AAAT C C AAC G C AT G C AAG AT T C AGT T T C GAT AG A 2250
2251 TCAAATGTTTGCTTGCAGCAAAACCTATGATTTTAAGCACGCCAATTTAC 2300
2301 GTTGGCGGAGCAAGATACACCTTAACTAAATCAGACTTTAATCCTGTTTA 2350
2351 TACCACTAGAGTGATTTATGACCCATGTTTTATGAGTGAGCCTCGAGTAA 2400
2401 TCAATATCTTAGTTGGATATGAAGTAGTTTTCCCAGTCGATGGTGATTAT 24 50
2451 GGACAAAACCTCTTTTTTGACATTAATGGTCGTTCAAATAAATCAATGAG 2500
2501 TTTAACAGCTTCTTGCGGTACGTTTAGCGCATCTGATTCAGGGAGTAAAT 2550
2551 TCATAATATCTCAACGAAAAAATACAGGTGGGAGCTGTAAAAATATGCAG 2 600
Figure 2-4. Nucleotide sequence of the genomic-DNA regions flanking the transposon in the SM2 mutant. The nucleotide sequence with the translated amino acid sequence in one-letter code is shown. The arrow ( ) indicates where the mini-TnlO transposon insertion occurred. The potential promoter region and putative ribosomomal binding site (RBS) are underlined. The putative sequence terminator is underlined, italicized and shown in red.
2.3.3.1.3. Sequence analysis of the SM4 mutant
The pan-handle PCR and primer walking strategy identified 1961 bp of DNA region flanking the transposon inserted in SM4 mutant. The primers used for primer walking are summarized in Table 2-3. Three ORFs were identified after submitting the consensus DNA sequence to the ORF-finder program in NCBI (Figure 2-5). Sequence analysis showed that the transposon had disrupted a 987 bp ORF (designated as ORF2) which encodes a sequence of 328 aa. The ORF2 region ranged between base positions 552 and 1538. A putative RBS was identified 5 bp upstream of the start codon, which is preceded by a putative promoter region 12 bp upstream.
65 The deduced aa sequence of ORF2 was analyzed and the product was predicted to have a molecular weight of 37.28 kDa and a pi of 5.13. The ORF2 gene product is of unknown function and is 38 % similar to a hypothetical protein of Magnetococcus sp. MC-1 (GenBank accession number ZP_00043311). The open reading frame contains a tetratricopeptide repeat domain (TPR). The TPR region was identified in aa positions 71 to 138 with a TPR repeat in aa positions 105-138. The TPR domain is a degenerate sequence of about 34 amino acids with a consensus residue of (L(X2)IA(X2)L(X8)S(X3)Y(X2)S(X4)K) (Blatch and Lassie, 1999). PSORT analysis of the gene product of ORF2 predicted its location in the inner bacterial membrane.
Two ORFs flanking the ORF2 were identified. Downstream from ORF2 is a 183 bp ORF (ORF1) encoding 60 aa. This ORF is 324 bp down stream of ORF2. The ATG start site is at base position 145, which is preceded by a putative RBS (5’GACAATC3’) 11 bp upstream. The putative promoter region was identified 16 bp upstream of the RBS. The predicted pi and molecular weight of the deduced aa sequences are 9.52 and 6.34 kDa, respectively. Analysis using the BlastX program of NCBI showed that the gene product of ORF1 has no known function and has a 61 % identity and 81 % similarity to a hypothetical protein of Shexvanella oneidensis MR-1 (GenBank accession number NP_715948.1). The ORF1 appears to have an uncleavable N-terminal signal. The predicted location of the gene product is in the inner membrane of bacteria, which is indicated by the presence of the transmembrane protein signal.
The third ORF (ORF3) was identified to begin at base position 1669 upstream of ORF2 and encodes a sequence of 76 aa. The deduced aa sequence has a predicted pi and molecular weight of 9.64 and 9.10 kDa, respectively. The putative RBS was located 11 bp upstream of the start codon, which is preceded by a putative promoter region 10 bp upstream. The gene product has an unknown function and no transmembrane proteins were identified, revealing the location of the protein in the bacterial cytoplasm.
66 Table 2-3. Oligonucleotides used in the primer walking strategy to sequence DNA regions flanking the inserted transposon in the SM4 mutant.
Oligonucleotide Sequence (5’-3’) Target region
SM4TnlOF CTT AT GGCGCG AGTTT AT GGG 14-34
SM4TnlOJ TTTGCTGACCAATGTGATCGGC 229-250
SM4TnlOK TCCGCCCGCTAAGGCACAGC 956-976
SM4TnlOC-D2 T AAGTCGGT AG AT GC ATTT GC 1079-1097
SM4TnlOC-D2A A A ATTT GC ACCGCT AGATT GG 811-831
SM4TnlOC-D2B AC AAGGAGAAAACC AC ACT GC 570-590
SM4TnlOC-D2C AGTCGAAACGGGCAATAAAGC 339-359
SM4TnlOD-D4 GCGCTAGAAACTCACATG GG 1377-1396
67 1 TCTGGTTTTATCGCTTATGGCGCGAGTTTATGGGGTGCGTTGTACTTTGA 50
-35 region 51 TTATGCATTAGGCGTATTTGTTGGTGCCTTTGCCGTAGGCGTTTACAGTA 100
-10 region RBS 101 ATTTATTTACCCGTTTGGCAAATGCGCCTGCGGTTATTGTTGCGatgcaa 150 M Q
151 gggctcatagttttggtacccggtagtaaaatttatattggattaaactc 200 GLIVLVPGSKIYIGLNS
201 aatcatttcaggtcaaaactttgtagctgccgatcacattggtcagcaaa 250 I I SGQNFVAADHIGQQT
251 catttttgatattaatgtccctagtggctggtttaatctttgcaaacgtt 300 FLILMSLVAGLIFANV 301 gctctgccgccaaagaaaattctctagCTTATTATTGAGCTTTATTGCCC 350 ALPPKKIL*
351 GTTTCGACTCAATCGTTACGGGCTGTTATTTAACATTTACTTCACTACCA 400
4 01 CCTTGTACCCATTTATGAATCTAAACGTTCGTTTTAATTCATAAAATGCC 4 50
4 51 TAACGAACTTTTCTGCAAAATATCTCATTTTCAAAATTTTGGACTAGCAT 50 0
-35 region -10 region RBS 501 TTAATCAATAAACTCAGTTTGATAATAAGTTTACTTTTTAAAGGCAAAAG 550
551 Catgaccaataataacattgcagtgtggttttctccttgtcggtatttta 600 MTNNNIAVWFSPCRYFI
601 tttttagcttagcttttttatttatatctcctatagttgcaagcccaacc 650 FSLAFLFISPIVASPT
651 actcctaattacattcaaattctgaataatgccgatccttttctaaaaga 700 TPNYIQILNNADPFLKD
701 caaaaactttcagggtttattcgattatttaattagttacgaattcgatt 750 KNFQGLFDYLISYEFDF
751 ttgcaggacaagctgagtatgactatgtacttggacttgcagcgttagag 800 AGQAEYDYVLGLAALE
801 tcgaatcaacccaatctagcggtgcaaattttacaacgagcagtggacgt 850 SNQPNLAVQILQRAVDV
851 cgatgctcttttttctggggctagaatggccttagcacgcgcttattttt 900 DALFSGARMALARAYFS
901 ctaccggagatttagagcgggcaaaatttcatttcactttattgcttgag 950 tgdlerakfhftllle'
951 caaaatccgcccgctaaggcacagcctgttatcaaacaatatttagcgaa 1000 QNPPAKAQPVIKQYLAN
68 1001 catagaacagttatcaaaaaactatagtgccagtttgcacacttatcttg 1050 IEQLSKNYSASLHTYLE
1051 aattaggcatgggctttgatagtaatgcaaatgcatctaccgacttagat 1100 LGMGFDSNANAST DLD
1101 ttattttatggctttcaattagatccaaaaaatatagaaactcaatctgt 1150 LFYGFQLDPKNIETQSV
1151 ttttagtactttattaggtgcattaaaatataactatcccattgatgctt 1200 FSTLLGALKYNYPI DAY
1201 atcaaaaactaaccgctaatatgatgcttggtaacagaaccaatgcaagc 1250 QKLTANMMLGNRTNAS 1 1251 gcacattatgtcgacatgaatatgctaagcatggatttaaaatatagtta 1300 AHYVDMNMLSMDLKYSY
1301 tgtattctcgtcttttagtgcctatggattagttcgtgaatatcacaatc 1350 VFSSFSAYGLVREYHNQ
1351 aaatcgaagataaattcaatcaaaatgcgctagaaactcacatgggatta 1400 IEDKFNQNALETHMGL
1401 gactttaaccaaagtgataattccttttttaatattgatttaagtgctgc 1450 DFNQSDNSFFNIDLSAA
1451 caaacaacgttttaataatcagcctgagtattcaagatgctacaagttac 1500 KQRFNNQPEYSRCYKLL
1501 tccgcttggctgacatcaacatctcggttggaaaataaTTGGTTAATCGG 1550 RLADINISVGK*
1551 AGCGTCTTTTGGCGTCGCTAAAGATGATGCAAGCGAAGCATTAAGCCCTT 1600
-35 region -10 region 1601 ATTCGAACAATAAAATAAGTGCGAGACTCTTTTCATTTACACCAGTCACA 1650
RBS 1651 GACAATCTCACATTTAATatgcagcttggggcatataaaacagattacag 1700 MQLGAYKTDYS
1701 taaagagcagctattttttggtgaacaacgtaaagataaacgctataacc 1750 KEQLFFGEQRKDKRYNL
1751 taatgacaagcttaagttacaacaacttcttagcaaaaagttggcaatta 1800 MTSLSYNNFLAKSWQL
1801 acgggtcggcttatcctgacaaaacaccaatctaacatctctatttatca 1850 TGRLILTKHQSNISIYQ
1851 atttgaccgcggtgaaattggcatttatttacgtaaaacatttgattaaA 1900 FDRGEIGIYLRKTFD*
69 1901 GG AC C T AAC AT G AAT AT T AAC t T T T T C AAAT GGGCCTGTATTT GC AT T AC 1950
1951 TTACTAGCGCA 1961
Figure 2-5. Nucleotide sequence of the genomic-DNA regions flanking the transposon in the SM4 mutant. The nucleotide sequence with the translated amino acid sequence in one-letter code is shown. The arrow ( ) indicates where the mini-TnlO transposon insertion occurred. Specific open reading frames are highlighted as follows: ORF1 is shown in red, ORF2 is shown in blue and ORF3 is shown in green. The potential promoter region and putative ribosomomal binding site (RBS) are underlined.
2.3.3.1.4. Sequence analysis of the SM5 mutant
Sequence analysis indicated that the transposon in the SM5 mutant had disrupted an ORF with homology to mshJ, a MSFIA biogenesis protein of V cholerae. The detailed genotypic analysis of theSM5 mutant is discussed in Chapter 4.
2.3.3.1.5. Sequence analysis of the SM6 mutant
The DNA sequence analysis of the consensus nucleotide sequence of this mutant showed that the transposon had disrupted a 431 bp open reading frame (ORF) which is 100 % identical to the dppD of P. tunicata (GenBank accession number AAL76240) (Egan, 2001). This gene was identified in a dark purple transposon mutant generated by Egan (2001). The ORF gene product is similar to a putative methyl transferase from Streptomyces coelicolor (GenBank accession number CAA 16186). The dppD gene is located downstream of dppB, the gene disrupted in the SMI mutant. The SM6 mutant strain is purple pigmented, making it identical to the dark purple mutant generated by Egan (2001). The genotypic characteristics of this mutant have been reported by Egan et al (2002a).
70 2.3.3.2. DNA sequence analysis of the P. tunicata AlpP-resistant mutants
The four selected AlpP-resistant mutants were analyzed for their genotypic characteristics. Using the pan-handle PCR method, the initial sequence analysis of the regions flanking the transposon in each of the four mutants revealed that they have been disrupted in different DNA regions. Two of the AlpP-resistant mutants (RM3 and RM4) were disrupted in a different region of the same ORF. Primer walking strategy and additional pan-handle PCR of the known sequenced regions from both sides of the transposon in the mutants, provided substantial DNA sequence for analysis. The DNA sequences were assembled, aligned and analyzed using the Staden package software (Medical Research Council-Laboratory of Molecular Biology, Cambridge, England). The resulting consensus nucleotide sequence was submitted to the ORF-finder and BLASTX and BLASTP programs of NCBI. The ExPASy (Expert Protein Analysis System) program packages of the Swiss Institute of Bioinformatics (SIB) were used for the analysis of the deduced amino acid sequence of the identified open reading frames.
2.3.3.2.1. Sequence analysis of the RM1 mutant
A total of 1774 bp of DNA surrounding the transposon in the RM1 mutant was sequenced and identified using the panhandle PCR and primer walking with oligonucleotides shown in Table 2-4. This nucleotide sequence was submitted to the ORF-finder program of NCBI for analysis. The transposon had disrupted a 1557 bp ORF encoding for 519 aa. The ORF starts at base position 175 and ends at base position 1731. A putative ribosomal binding site (5’GAACA3’) was identified 1 bp upstream of the start codon. The putative region was located 20 bp upstream of the RBS. The transposon was inserted at base position 680 (Figure 2-6).
The deduced amino acid sequence of the ORF showed that the protein had a molecular weight of 58.89 kDa and a predicted pi of 5.04. The gene product has a 46 % similarity to a hybrid sensory kinase of Synechocystis sp. PCC 6803 (GenBank accession number BAA17998). Other protein homologies are the probable sensor/response regulator hybrid of Pseudomonas aeruginosa with a 45 % similarity (GenBank accession number AAG07499) and the sensor histidine kinase/response regulator of Caulobacter
71 crescentus CB15 with a 47 % similarity (GenBank accession number AAK25064). PSORT analysis showed that the predicted location of the gene product is in the bacterial cytoplasm.
A signal transduction histidine kinase domain was identified within the open reading frame. This domain is located between amino acids 212 and 517. The multiple amino acid sequence alignments of the deduced amino acid sequence of the signal transduction histidine kinase of the interrupted gene of RM1 with histidine kinase domains of other bacteria is presented in Figure 2-7.
Table 2-4. Oligonucleotides used in the primer walking strategy to sequence DNA regions flanking the inserted transposon in the RM1 mutant.
Oligonucleotide Sequence (5’-3’) Target Region
RMlTnlOD-D8 TT AACG AA A AAT AC AGGGGGG 886-907
RMlTnlOD-D8A CTCCGATGAATGCAATTATTGG 1151-1172
RMlTnlOD-D8B AATCAACCGCTAGCAATCAAGG 1373-1394
RMlTnlOD-D8C CGT AAATTT GGT GG AAC AGGG 1677-1698
RMlTnlOC- D9 ATCGGTCCTTTCGACAAGC 546-563
RMlTnlOC-D9A GAT ATTT GCGCC ACT G ACT GC 282-303
RMlTnlOC-D9B ACTCTGCACCTAATTGTCCCG 92-112
RMlTnlOKl CAACTACCCGTAAATTTGGTGG- 1670-1698 AACAGGG
RMlTnlOJ GTTCTGTGTGAGGCGCTAACCCG 369-391
72 1 CTGCTTGGTTTTAACAAAGATCAATTAAATGGCGTTAATTTACATAAAAT 50
51 ATTCAATTTTAATCATGAAATTTTATTTTCTCCCACTTCAACGGGACAAT 100
-35 region -10 101 TAGGTGCAGAGTCAGCGCTGGTAACTACAGATAGTTTTCGTTCTTTTTTA 150
region RBS 151 CTTACAAAGATCTATATAGAACAAatggatagcttaaaatccattgttgt 200 "mdslksivv
201 tactgtcgctgagacaacacaactctcaattacaaataatatcagtactc 250 TVAETTQLSITNNI STQ
251 aagtcaaaagtgatttcaacttactgcaaaatgcagtcagtggcgcaaat 300 VKS DFNLLQNAVSGAN
301 atcggtatttggcgctacaacataatgagtaaaaatactgatttctcaca 350 IGIWRYNIMSKNTDFSQ
351 aaaatcaaaagagttactcgggttagcgcctcacacagaactgacatggc 400 KSKELLGLAPHTELTWQ
401 agacatttattgaaacagtccacccgcatgaccaagtgctttttgaagct 450 T FI ETVHPHDQVLFEA"
451 tttttcgataatcatcttgaatttagattattgcttcattttgagtttag 500 FFDNHLEFRLLLHFEFR
501 aatcattgttgaaggcgtttatcaatggtttgaactcagaggtgagcttg 550 I IVEGVYQWFELRGELV
551 tcgaaaggaccgatggtgatagtcatatttacggcacactcataaattgc 600 ERTDGDSHIYGTLINC
601 catcaagaaaaagaaatggtgattgcattaaatgatgcaaacgaaagtaa 650 HQEKEMVIALNDANESK i 651 agctttggcaatggaggcaggaaaaataggcacttggcgtgcatctaaat 700 ALAMEAGKIGTWRASKS
701 ctggaaatgattggacatggaattgggataagctcactaatgatatcttc 750 GNDWTWNWDKLTNDI F
751 caattagatgatgatgatataggttgcttagaaaaatgggctgagcgggt 800 QLDDDDI GCLEKWAERV
801 tcaccctgaggacatagatcaagtacttaagcaacttgaatcctcattaa 850 HPEDIDQVLKQLESSLI
851 taactggacgagagtttaacgaaaaatacaggggggtattgccttcagga 900 tgrefnekyrgvlpsg'
73 901 gaaataatttatgtatcagctcgaggtgtggttggtaaaaatgcattaga 950 El I YVSARGVVGKNALD
951 tgaaaactatcgtattgatggtttttgtgtagatgaaactgaaatttatg 1000 ENYRIDGFCVDETEIYE
1001 aagcgcatgcgcaattaaaaaaactaaacctagaactcgaagctcgggtt 1050 AHAQLKKLNLELEARV
1051 gaagaaagaacggcagagctaactcaagcaattacgcaagctgaactagc 1100 EERTAELTQAITQAELA
1101 aagtaaaattaaatctgaatttctagccatgatgagtcacgagttaagaa 1150 SKIKSEFLAMMSHELRT
1151 ctccgatgaatgcaattattggttcattagagttgctttctttatcaatt 1200 PMNAI IGSLELLSLS I
1201 aacgcacatgaagagagtgaacttattaatactgcatctatgtctgcaca 1250 NAHEESELINTASMSAH
1251 caacctagtcaatattctaaatgacatcctcgacattaataagattgagt 1300 NLVNILNDILDINKIES
1301 caggtaaactagaaatagaaacccatgattttgatcatcaacagcttata 1350 GKLEIETHDFDHQQLI
1351 tataatatagtaaaaacctttgaatcaaccgctagcaatcaaggtgtaac 1400 YNIVKTFESTASNQGVT
1401 actgattattcatgaagatacaaagatgccaccgattgtgtctggtgatg 1450 LI IHEDTKMPPIVSGDE
1451 aaattagagttaggcaaattattatgaatatcataagtaatgcggttaaa 1500 IRVRQI IMNIISNAVK
1501 tttacggctggtaatgataaagaaataaagattgtaaccttctgtattca 1550 FTAGNDKEIKIVTFCIQ
1551 gtggaggcaaataggaggttgtctatttgaaataatttatgaaattattg 1600 WRQIGGCLFEIIYEIID
1601 atactggtattggtattaatcaagaaactcaaaaaagactctttacacct 1650 TGIGINQETQKRLFTP
1651 tttatccaagctgaaaagtcaactacccgtaaatttggtggaacagggtt 1700 FIQAEKSTTRKFGGTGF
1701 tagggcctaccatctcaggaaaattggctgaTATGATTGGGGGCTCAATC 1750 RAYHLRKIG*
1751 CGAGCTAAAACGCCTGCCGGTATA 1774
74 Figure 2-6. Nucleotide sequence of the genomic-DNA region flanking the transposon in the RM1 mutant. The nucleotide sequence with the translated amino acid sequence in one-letter code is shown. The arrow (i) indicates where the mini-TnlO transposon insertion occurred. The potential promoter region and putative ribosomomal binding site (RBS) are underlined.
D2RM1 272 ' GI 16127421 229 1 GI 15598658 371 1 GI 16329398 324 ■
D2RM1 332 -INAH-- EESELINTASMSAHNLVNILNDILDINKIESGKLEIETH GI 16127421 285 : -KLSDD—GRRLLTEALSCGEMLSALLNDIIDFSKIEAG-KLELSAE GI 15598658 427 : -PLDRG—QAAYVETIASSGSALMSVINDILDYARIESG-KLHLERI GI 16329398 384 :
D2RM1 385 GI 16127421 338 GI 15598658 480 GI 16329398 438
D2RM1 440 ------VTFCIQWRQIGG GI 16127421 392 ---- ARLRLRFEIDDTGVG- GI 15598658 535 ------ERLLYSVSDSGIG- GI 16329398 493 -TEPKNKYWLDFTIKDTGKG-
D2RM1 466 GI 16127421 431 -1GEEAGARLFERFRQGDGSTTRRFGGSGLGLAICRRLAELMGGEV GI 15598658 571 -ISAQAQKTLFESFSQADSSTTRRYGGSGLGLAISRELVQVIMGGRI GI 16329398 542 -ISEVELSHLFEAFSQTESGRNAQ-EGTGLGLAITRQFIKLMGGDI
Figure 2-7. Multiple amino acid sequence alignments of the deduced amino acid sequence of the signal transduction histidine kinase of the interrupted gene of P. tunicata RM1 with histidine kinase domain of other bacteria. (*) identical amino acids, (:) conserved substitutions, (.) semi-conserved substitutions, (-) denote gaps. GI 16127421 - Caulobacter crescentus CB15, GI 15598658 - Pseudomonas aeruginosa PAOl, GI 16329398 - Synechocystis sp. PCC 6803
75 2.3.3.2.2. Sequence analysis of the RM3 and RM4 mutants
A total of 3013 bp of DNA were sequenced and identified from the RM3 and RM4 mutant strains (Figure 2-8) using panhandle primers and specific oligonucleotides for primer walking as shown in Table 2-5. Analysis of the consensus sequence indicated that the transposon mutants RM3 and RM4 were disrupted in different regions of ORF2. This ORF consists of a 1925 bp encoding 641 aa. The ATG start site is at base position 989 and the ORF ends at base position 2914. The putative ribosomal binding site (5’GAGCGC3’) was identified 9 bp from the start codon, which is preceded by a putative promoter region 48 bp upstream.
The deduced aa sequence of the ORF2 had a molecular weight of 70.92 kDa and a predicted pi of 5.14. It showed high similarity to peptide synthetase proteins including 46 % identity and 63 % similarity (over 409 aa residues) to the SyrE, syringomycin synthetase of Pseudomonas syringae pv. syringae (GenBank accession number AAC80285.1) and 47 % identity and 63 % similarity (over 409 aa residues) arthrofactin synthetase C of Pseudomonas sp. MIS38 (GenBank accession number BAC67536.)
Two conserved domain sites were identified in the ORF2 (Figure 2-9). An amino acid adenylating domain was identified between aa 1 and 378. This region catalyzes the adenylation of the amino acids to form thioesters. The domain is similar to the amino acid adenylating domain present in the SyrE module of P. syringae (Bender et al., 1999) (Figure 2-10). Each SyrE module contains 5 core sequences. The ORF2 in the RM3 and RM4 mutants contains three SyrE core signature sequences. The core 2 sequence (SGTTGxPKGV) was identified between amino acids 97 and 106. The glycine-rich motif contains a potential phosphate binding loop involved in ATP binding. Core 4 sequence (TGD) was detected in the ORF protein between aa 329 and 331. A motif associated with ATPase activity. The other conserved domain identified was a pp- binding domain, a phosphopantetheine attachment site between aa 469 and 528. This prosthetic group acts as a - 'swinging arm' for the attachment of activated fatty acid and amino-acid groups, specifically serine. This domain forms a fourhelix bundle. In some protein members, the attachment amino acid is an alanine rather than a serine (Bender et al., 1999). In the P. syringae SyrE module, the core 6 sequence (LGGHSL) is present in
76 the pp-binding domain. Such a core 6 sequence was also detected in the ORF protein between aa 494 and 500 (Figure 2-11).
Directly upstream of ORF2 is ORF1 consisting of 393 bp. ORF1 encodes for 130 aa, with a start codon at base position 144, preceded by a putative ribosomal binding site 3bp upstream. The putative promoter region was identified 51 bp upstream of the RBS. The gene product had a predicted pi and a molecular weight of 6.39 and 13.86 kDa, respectively. A condensation domain was identified in this ORF. The condensation domain (CD) catalyses a condensation reaction to form peptide bonds in non-ribosomal peptide biosynthesis (Bender et al., 1999).
Table 2-5. Oligonucleotides used in the primer walking strategy to sequence DNA regions flanking the inserted transposon in the RM3 and RM4 mutants.
Oligonucleotide Sequence (5’-3’) Target Region
RM4TnlOC-D12 GGCT A ATTT CTTT G AGTT GGGT GG 2449-2469
RM4TnlOC-D12A GCTACCGAAGCTGCAATTGAGC 2846-2866
RM4Tnl0D-D10 GTTTTCCTATCGGCACACTGC 1755-1775
RM4Tnl0D-D10A GCTGTGACGTTAACCCTATTTCC 1231-1253
RM4Tnl0D-D10B GGTAGTTGGTTCGCTCTTGGC 539-559
RM4TnlOJ GGGTGGCCACTCTTTGATGAT- 2477-2492 TATGG
RM4TnlOK GCCTTTT GGTT G ACCT GT AG A- 1275-1303 CCCTGAGG 1 CCGCGTTTTTCAGCCAACAACTTGGTGAGGTGAGTGAGCCGACCCTGCCT 50
-35 region -10 region 51 TATGGGTTAAGCGATGTACTGAATGATGGTAAAGCGGTCACGAGTGTCAA 100
RBS 101 AGTCAGATTGGACAAAGCCCAAGCGCAACAGGTACGGCAGTTAatgcgtg 150 M R E
151 agcacaacagcagcccagcagcgttgtttcacttagtatgggccaaagtg 200 HNSSPAALFHLVWAKV
201 ttatcggtgtgcagcggtcagtcacaagtggtgtttggcacggttttatc 250 LSVCSGQSQVVFGTVLS
251 aggacgcatgaatgggctccccggcatcgaacgcatgatggggatgttaa 300 GRMNGLPGIERMMGMLI
301 tcaacaccttgccattaaaagtggacttaggcagtcacagagccgcaacg 350 NTLPLKVDLGSHRAAT
351 ttattaagccaactgaacagcgacttacaagacttagtaccgtatgagca 400 LLSQLNSDLQDLVPYEQ
401 agtttcattagcagaagcacaacgttacagcggcgtcagtggacaactgc 450 VSLAEAQRYSGVSGQLP
451 cgttattcagcgcgatgttaaattaccgtcactcagcaatagcagagcaa 500 LFSAMLNYRHSAIAEQ
501 ggcgcagaaagcacaggcgggatttcgggtccttgaAAGCCAAGAGCGAA 550 GAESTGGISGP*
551 CCAACTACCCGGTTTAACTTATTAGTGGATGATTTTGGTGATGGCTTTGC 600
601 ATTTGAAGTGCAAATAGATACACGGGTCCCAGCGCAGCAGATAGCGGCGT 650
651 ATGTACAGGTTACACTGACAAACGTACTCGATTTATTAGCAAAAGACTCA 7 00
701 ATACAATCCGTGCACAGTGTGTGTGTATTATCAGAGGCCGAGCGCCAGCA 750
751 GCAGCTGGTAGATTGGAACAATACTGCGCTGAGCTACCCTAAAGAATTGT 800
801 GCATTCATGAATTATTTGAAGCGCAAGTGCAACACGCGCCAGAGCGCACC 850
-35 region 851 GCGGTATGGTTTGAAGAGCAGTGTTTAAGTTATGGCGAACTTAATGCCAA 900
-10 region 901 AGCCAACCAGTTGGCACATTATTTACGCGCTGAGCATGGTGTTGGGCCTG 950
RBS 951 ACAGTTTAGTGGGCTTATGCACTGAGCGCTCATTGGAAatggtcatcggg 1000 M V I G
78 1001 atttgggggattttaaaagccggcggcgcttatgtaccattggaccccga 1050 IWGILKAGGAYVPLDPE
1051 gttaccgtctgctcgattgcagtatttggtcagtgatacccaggccaatg 1100 LPSARLQYLVS DTQANV
1101 tggtattaagtacccaagcccttaaagagcacatcacgttaggcgaagca 1150 VLSTQALKEHITLGEA
1151 caagtggtctatcttgatggcttggggagtcaggttacacattcgtttag 1200 QVVYLDGLGSQVTHS FS
1201 cgagtatagcgaagagaatattaatctaaaggaaatagggttaacgtcac 1250 EYSEENINLKEIGLTSQ
1251 agcaccttgcctacatgatttacacctcagggtctacaggtcaaccaaaa 1300 HLAYMIYTSGSTGQPK
1301 ggcgtattgttggcccatcaagccttgcacaaccgtattgattggatgga 1350 GVLLAHQALHNRI DWMD
1351 ccgagaatatggctgtgacagcctagatgttattttacaaaagaccccgt 1400 REYGCDSLDVILQKTPY
1401 acagttttgatgtgtcggtctgggaatttatttggcccatgctaaagggc 1450 S FDVSVWE FIWPMLKG~
1451 gcaaaactagtggtagccaaaccgcaaggacacaaagacccaagttacct 1500 AKLVVAKPQGHKDPSYL >1 1501 caccgaactgatagttgccaccggcgtgaccaaattgcattttgtgcctt 1550 TELIVATGVTKLHFVPS
1551 ccatgttaggcgtgatgttagcccatggtgacttgcatcgatgtcagtca 1600 MLGVMLAHGDLHRCQS
1601 ctgaaacaggttttttgtagtggtgaagcgttacaaatcagccatgttga 1650 LKQVFCSGEALQI SHVE
1651 acaatttagacatcagttaccagaggttggattacataacttatatggcc 1700 QFRHQLPEVGLHNLYGP
1701 caacagaggccgctattgatgtgagctactgggattgctcgcagccactg 1750 TEAAI DVSYWDCSQPL
1751 ggcagcagtgtgccgataggaaaaccaatacaaaacatccagctctatat 1800 GSSVPIGKPIQNIQLYI
1801 attggatgacgaattaaatctattaccacaaggagcctgtggcgagttac 1850 LDDELNLLPQGACGELH
1851 atattggtggcgacggcttagcccgcggctatttaaatcggccagagctg 1900 IGGDGLARGYLNRPEL
1901 acacaagagcggtttatcgccaacccattttatcaagcagaagaaggcaa 1950 TQERFIANPFYQAEEGN
79 1951 cagcagtgagcgcttgtataaaacaggagatttggtgcgctacaaagaag 2000 SSERLYKTGDLVRYKED
2001 atggcaacatagaatacatggggcgtttagaccatcaagtgaagataaga 2050 GNIEYMGRLDHQVKIR
2051 ggctttcgcatagagttgggcgaaatagaatatcaggtcgcacaacataa 2100 GFRIELGEIEYQVAQHK I 2101 gcaaatagactcagcgctggttgttgcccaagcggataaagcaggcaatc 2150 Q I DSALVVAQADKAGNQ
2151 aacggctgatagcctatgccaaaaccgtgcaagcagacgcgtcaaaagag 2200 RLIAYAKTVQADASKE
2201 gacgttattgcctcattaaaagctgagttagcggcggtattaccagaata 2250 DVIASLKAELAAVLPEY
2251 catggtgccagcgaacgttattttggtctcagaatggccattgacgccaa 2300 MVPANVILVSEWPLT PN
2301 atggtaaaatagaccgcagaggactgccctcggtggatgagagtatcgat 2350 GKI DRRGLPSVDES I D
2351 agcacagcgtatgtcgcgccacacggtgagacagaattgatgctggttga 2400 STAYVAPHGETELMLVE
2401 gatttgtgctgaacttttaagaatcaataaattagaaattagtattatgg 2450 ICAELLRINKLEI S IMA
2451 ctaatttctttgagttgggtggccactctttgatgattatggatttagtt 2500 NFFELGGHSLMIMDLV
2501 agccgcctaaaaaaacgcggttttagtacatcagttcagtctttatttgc 2550 SRLKKRGFSTSVQSLFA
2551 ggcaaaagtattaaaagagatgaccctagagctagtgcctaataaagaga 2600 AKVLKEMTLELVPNKE I
2601 tccagagtgacgcattactacctgaaaatttaatccctgcagcgtgtaca 2650 QSDALLPENLIPAACT
2651 tatttaacacctcagatggtcaaccttgcttcagtgtctcaacaagagct 2700 YLTPQMVNLASVSQQEL
2701 tgataagattagtactttagtacctggcggggcacaaaacattcaagata 2750 DKI STLVPGGAQNIQDI
2751 tttatccgttggcgccgttacaagaaagcatctttttgatccattgtgcc 2800 YPLAPLQESIFLIHCA
2801 acagaaggaacagatccttatgtgacaattattacgttagaatttgctac 2850 TEGTDPYVTIITLEFAT
80 2851 cgaagctgcaattgagcaatttgttgtgcgcttaaataaagtgattaaac 2900 EAAIEQFVVRLNKVIKR
2901 gccatgacatctaaGAACAATGATAAGTTGGCCGTGGTTGTCCACAGCCG 2950 H D I *
2951 CTGCAAGAAGCCTTTACGTGATGTTACATTAATACCAAGCTGGTTACCGT 3000
3001 TTTTTAGGCGAGA 3013
Figure 2-8. Nucleotide sequence of the genomic-DNA regions flanking the transposon insert in the RM3 and RM4 mutants. The nucleotide sequence with the translated amino acid sequence in one-letter code is shown. The arrow ( ) indicates where the mini-TnlO transposon insertion occurred. Specific open reading frames are highlighted as follows: ORF1 is shown in red and ORF2 is shown in blue. The potential promoter region and putative ribosomomal binding site (RBS) are underlined.
0 500 1000 1500 2000 2500 30C L till*, . , 1 < , 1 i 1 i i i i 1 ii i i 1 i i J_Ll
mm Amino acid at ig domain ■
Figure 2-9. Conserved domain identified in the RM3 and RM4 DNA regions disrupted by the mini-TnlO transposon. The arrangement of the domains is similar to the SyrE module in Pseudomonas syringae, which is involved in the biosynthesis of syringomycin, a peptide pore-forming cytotoxin. The condensation domain (CD) catalyses a condensation reaction to form peptide bonds in non-ribosomal peptide biosynthesis. The amino acid adenylating domain contains 5 core sequences, which are involved in adenylation and substrate recognition. The pp-binding (PB) domain is a phosphopantetheine attachment site, which is associated with covalent binding of the substrate amino acid. Open reading frames are indicated by the bold coloured lines; gray = ORF1; yellow = ORF2.
81 D2 RM4 1 ------MVIGIWGILKAGGAYVPLDPELPS 24
GI 351062 9 1510 TYAELNQQANQLAHRLIELGVEPDTRVAVSLRRGAEMWALLGILKAGGAYVPIDPDLPD 1569 GI 4481934 497 TYAELDARAERLAGALTARGAGPERFVAVAVERSAELWALLAVLKSGAAYVPVDPGYPA 556 GI 6136084 1527 TYRELNARANQLARLLRSHGTGPDTLIGIMVDRSPGMVVGMLAVLKAGGAYTPIDPSYPP 1586 GI 2522212 1418 TYKELNEQANRVAWELIDRGVKAETTVAIMGRRSPEMLIGIYAILKAGGAYLPIDPDYPE 1477
• ~k • ~k ★
D2 RM4 25 ARLQYLVSDTQANWLSTQALKEHI----- TLGEAQWYLD------GLGSQVTH 68 GI 3510629 1570 ARQAYMLSDSAPRAVLTSHELLADLPDLGVPALVLDGR------DDSA 1611 GI 4 481934 557 DRIAHILRDAGAMLVLTTRDTAERLPGDGTPRLLLDEPAA------AGTTAA 602 GI 6136084 1587 ERIQYMLSDSQAPILLTQRHLQELAAYQGEIIDV------1620 GI 2522212 1478 ERIRFLLKDSDSDILLIQSDNLDIPAFDGEIVHL------S 1512
D2 RM4 69 SFSEYSEENINLKEIGLTSQHLAYMIYTSGSTGQPKGVLLAHQALHNRIDWMDREYGCDS 128 GI 3510629 1612 LLKKQPTGNPDAKALDLQPNHLAYVLYTSGSTGTPKGVMNEHLGWNRLLWARDAYQVNS 1671 GI 4481934 603 GAPAPPGTLPRALPAP-- GHPAYVIYTSGSTGRPKGVVISHRAIVNRLAWMQDTYGLEP 659 GI 6136084 1621 DEEAIYTGADTNLDNVAGKDDLAYVIYTSGSTGNPKGVMISHQAICNHMLWMRETFPLTT 1680 GI 2522212 1513 QLNSGLKRRLSNPNVEVYPDSLAYMIYTSGSTGRPKGVQVEHQSAVNFLNSLQFRYPLNQ 1572
D2 RM4 129 LDLD—--- VILQKTPYSFDVSVWEFIWPMLKGAKLVVAKPQGHK-DPSYLTELIVATGVTKLH 184 GI 3510629 16721672 QDQD—--- RVLQKTPFGFDVSVWEFFLPLLTGAELVMARP-SGHQDPDYLAQVISDAGITLLH 1727 GI 4481934 660 SDSD—--- RVLQKTPSGFDVSVWEFFWPLVQGATLVVARP-GGHTDPAYLAGTVRREGVTTLH 715 GI 6136084 16811681 EDED---— AVLQKTPFSFDASVWEFYLPLITGGQLVLAKP-DGHRDIAYMTRLIRDEKITTLQ 1736 GI 2522212 15731573 SDSD—---VILHKTSYSFDASIWELFWWPYGGASVYLLPQ-GGEKEPDMILKVIEEQQITAMH 1628 *.*■*•. **★.**. . * ...... *..
D2 RM4 185 FVPSMLGVMLAHGDLH—R—CQSLKQVFCSGEALQISHVEQFRHQLPEVGLHNLYGPTE 240 GI 3510629 1728 FVPSMLDVFLEHRST--- RDFPQLRRVLCSGEALPRALQRRFEQHLKGVELHNLYGPTE 1783 GI 4481934 716 FVPSMLDVFLREPAAAA-LGGATPVRRVFCSGEALPAELRARFRAVS-DVPLHNLYGPTE 773 GI 6136084 1737 MVPSLLDLVMTDPGW--- SACTSLQRVFCGGEALTPALVSRFYETQ-QAQLINLYGPTE 1791 GI 2522212 1629 FVPSMLHAFLEYLKNGPVPIKTNRLKRVFSGGEQLGAHLVSRFCELLPDVTLTNSYGPTE 1688
.***.* •'k'k»'k-k’k-k'k
D2 RM4 241 AAIDVSYWDCSQPL-- GSSVPIGKPIQNIQLYILDDEL-NLLPQGAC 284 GI 3510629 1784 AAIDVTAWECRPTDP-- GDSVPIGRPIANIQMHVLDALG-QLQPMGVA 1828 GI 4481934 774 AAVDVTYWPCAEDTG-- DGPVPIGRPVWNT RMYVL DAAL-RPVPAGVP 818 GI 6136084 1792 TTIDATYWPCPRQQE-- YSAIPIGKPIDNVRLYVVNASN-QLQPVGVA 1836 GI 2522212 1689 ATVEAAFFDCPLDEK-- LDRIPIGKPIHHVRLYILNQKQ-KMLPAGCI 1733
• ★ ★ ★ ★ ★ • ★ ■k
82 D2 RM4 285 GELHIGGDGLARGYLNRPELTQERFIANPFYQAEEG------NSSERLYKT 329 GI 3510629 1829 GELHIGGIGVARGYLNQPQLSAERFIADPFSNDP------QARLYKT 1869 GI 4481934 819 GELYIAGVQLARGYLGRPALSAERFTADPHGAPG------SRMYRT 858 GI 6136084 1837 GELCIAGDGLARGYWQREELTKASFVDNPFEPG------GTMYRT 1875 GI 2522212 1734 GELYIAGAGVARGYLNRPELTEERFLDDPFYPG------ERMYKT 1772
•k k k k • k • • k k k k...... k • k • ★ • k k
D2 RM4 330 GDLVRYKEDGNIEYMGRLDHQVKIRGFRIELGEIEYQVAQHKQIDSALV 378 GI 3510629 1870 GDVGRWLANGALEYLGRNDFQVKIRGLRIEIGEIEAALAKHPAVHEAW 1918 GI 4481934 859 gdlarwnhdgsldylgradhqvklrgfrielgeieaalvrqpeiaqaav 907 GI 6136084 187 6 GTMVRYLPDGHIEYLGRIDHQVKIRGHRIELGEIEATLLQHEAVKAVW 1924 GI 2522212 1773 GDLARWLPDGQVEFLGRLDDQVKIRGYRIEPGEIEAALRSIEGVREAAV 1821 * * * . . * *.***.*■* *■**.****._.
Figure 2-10. Multiple amino acid sequence alignments of the deduced amino acid sequence of the amino acid adenylating domains of the interrupted gene of RM3 and RM4 with AMP-binding domain of other bacteria. CLUSTAL W (1.82) (*) identical amino acids, (:) conserved substitutions, (.) semi-conserved substitutions, (-) denote gaps. RM4 mutant - P. tunicata AlpP-resistant mutant, GI 3510629 - SyrE, syringomycin synthetase of Pseudomonas syringae pv. syringae, GI 4481934 - CDA peptide synthetase (Streptomyces coelicolor A(3)2, GI 6136084 - Tyrocidine synthetase III (Brevibacillus parabrevis), GI252212 - fengycin synthetase FenA (.Bacillus subtilis). SyrE core 2 sequence (SGTTGxPKGV) is indicated in red and core 4 sequence (TGD) is indicated in blue.
83 D2RM4 4 67 lmLVEICAELLRINKLEISIMANFFE-LGGHSLMIMDLVSRLKKR-GFSTSVQSLF—AA 522 lAcp 5 ERVKKIIGEQLGVKQEeVTNNASFVEdLGADSLPTVELVMALEEEFDTEIPDEEAE—KI 62 GI 113205 11 TILVACAGEDDGVDLSGDILDITFEE-LGYDSLALMESASRIERELGVALADGDIN—EE 67 GI 728785 13 RILVEAAGADESAGPDDI-LDTTFAL-LGYESLALLETGGCIEREIGISLDDDTLT--DA 68 .. *.**..** * ... •
D2RM4 523 KVLKEMtlel 532
1ACP 63 TTVQAAIDYI 72 GI 113205 68 LTPRVLLDLV 77 GI 728785 69 LTPRELIDHV 78
Figure 2-11. Multiple amino acid sequence alignments of the deduced amino acid sequence of the pp-binding domains of the interrupted gene of RM3 and RM4 with pp- binding domain of other bacteria. CLUSTAL W (1.82) (*) identical amino acids, (:) conserved substitutions, (.) semi-conserved substitutions, (-) denote gaps. RM4 mutant - P. tunicata AlpP-resistant mutant, 1ACP - Escherichia coli B, GI 113205- Streptomyces violaceoruber and GI 728785 - Streptomyces cinnamonensis. SyrE core 6 sequence (LGGHSL) is indicated in red.
2.3.3.2.3. Sequence analysis of the RM6 mutant
A total of 1027 bp DNA was sequenced around the transposon inserted in the RM6 mutant using pan-handle PCR method and primer walking with specific primers; RM6TnlODl 1 -(5’GGC CAT CCA AGA TAC CCT CTG G’3), RM6TnlOD DI 1A (5’GGC ATG TAT TTT TGC TGA CCG’3), RM6TnlOD-D14 (5’CGTTAATTGATTTAGACTCCACTCC’3) and RM6TnlO-D14A (5’CGCTGGTATCAGTTGTATCACGG’3). The transposon had disrupted a non coding frame in the DNA region specifically at base position 279 (Figure 2-12). Upstream of this mutation is a 116 bp ORF (ORF1). The ATG start site is at base position 111 bp and the stop codon at base position 227. The deduced amino acid sequence has a theoretical pi and molecular weight of 9.75 and 4.5 kDa, respectively. This protein has an unknown function and does not have any similarity with any
84 proteins presently found in GenBank. The predicted location of the protein is in the bacterial cytoplasm.
Downstream of the transposon insertion is another ORF (ORF2). This is a 149 bp ORF with an ATG start site at base position 304 and the stop codon is at base position 453. The putative RBS is 6 bp upstream of the start codon. There was no putative promoter region identified from this ORF indicating that ORF2 and ORF1 may be co-transcribed. The deduced amino acid sequence of ORF2 has a theoretical pi and molecular weight of 4.56 and 5.78 kDa, respectively. The protein is similar to the putative acetyl transferase of Bacteriodes thetaiotaomicron showing 64 % similarity (GenBank accession number NP_813417.1) and acetyl transferase, from the GNAT family of Pseudomonas putida showing 63 % similarity (GenBank accession number NP 743352.1). No transmembrane protein was detected, thus the predicted location of the protein is the bacterial cytoplasm.
An additional ORF was identified immediately down stream of ORF2. This ORF (ORF3) consists of 146 bp with the ATG codon start site at base position 774 and the stop codon at base position 920. The putative RBS is 1 bp upstream of ATG, which is preceded by a putative promoter region 7 bp upstream. This ORF has no similarity with any proteins submitted in GenBank. The deduced amino acid sequence has a theoretical pi and molecular weight of 9.22 and 5.97 kDa, respectively. This protein is also predicted to be located in the bacterial cytoplasm.
85 -35 region 1 CAGCATCGATTTTGTATTGATTTTTAATACAAATTATTTAAAATCAACGA 50
-10 region RBS 51 GATATTTATGAGTGGCATGTATTTTTGCTGACCGGAGTATGTATTCATGA 100
101 ACTAATTTTAatgaagcactatctggccatccaagataccctctggaaat 150 MKHYLAIQDTLWKL
151 taatttttatgaagacccagattggtctgatttgtctaaattactcgcgg 200 I FMKTQIGLICLNYSR
201 atttcaacaatgccngctaacaagtagCTGCACCGGAAAAAATACTCGCT 250 I STMPANK* i RBS 251 GCGCTCATATTTTTCCGGTGAGCTAAGCGTTAAATTTTCAATCGGGTTTA 300
301 GTCatgaatattgagatcgtaaaaaaagcagatcaccttaagctaataga 350 MNIEIVKKADHLKLIE
351 aatctgggaatcatcagtccgagcaactcatgattttctggctgaagaag 400 IWESSVRATHDFLAEED
401 atttacaagagttaaagccgttaatcttggagcagtattttgacgccgtt 450 LQELKPLILEQYFDAV
4 51 taaAAAAAATACTAGACACCCATAAATATCTTGTTGAAAAATCACAATAG 500 *
501 TGGGTCTAATTTTAATTAATCACTTCCTGTTTTGGTGGCTACTTTTTGCA 550
551 AAAAAAAGGGGCACAAGCAAGATGGTCGTTAATTGATTTAGACTCCACTC 600
601 CTTCTTTCCTTTAATAACATAAGCACTGCATCAGTCGATGTGTACGCAGG 650
651 GCTTTTTTAGCGGGTTTTGATAAGTATTCCGGTCAAAATTTTGAACACAG 700 -35 region 701 ACGGGCTTGGTTAGTTGAGCGATTTAAACGGCTATCGCAAGTGTTTGCCA 750
-10 region RBS 751 TTGATATTGCGGCCTATGCGGTGatgtcgaatcattaccatttggtgctt 800 MSNHYHLVL
801 agggtcgatagaagtagggcattaaactggtcaaaggatgaagtgattga 850 RVDRSRALNWSKDEVIE
851 gcgctggtatcagttgtatcacggcacaattttggttgaccgctatcgta 900 RWYQLYHGTILVDRYRK
901 aaggtgaaaagctcgaataaGCCTATATGTACAGCGTTGATAAAACGGTA 950 G E K L E *
951 GAAGTTTGGCGAAATCGTTTATATGACATCAGTTGGTATATGCGACTTTC 1000
1001 GTTTTAAAATAAGTAAACGAGTTTATT 1027
86 Figure 2-12. Nucleotide sequence of the genomic-DNA regions flanking the transposon inserted in the RM6 mutant. The nucleotide sequence with the translated amino acid sequence in one-letter code is shown. The arrow (1) indicates where the mini-TnlO transposon insertion occurred. Specific open reading frames are highlighted as follows: ORF1 is shown in red, ORF2 is shown in blue and ORF3 is shown in green. The potential promoter region and putative ribosomal binding sites (RBS) are underlined.
87 2.4. Discussion
It has previously been shown that the wild-type P. tunicata is sensitive to the autolytic protein, AlpP, during logarithmic phase of growth and develops resistance to AlpP during stationary phase of growth (James et al., 1996). To better understand the response of P. tunicata to its autolytic protein, AlpP, transposon mutagenesis was performed. This chapter describes in detail the phenotypic and genotypic characteristics of P. tunicata transposon mutants with altered sensitivity to the autolytic protein, AlpP. The P. tunicata AlpP-sensitive and -resistant mutant strains were selected from the transposon library based on their response to AlpP, relative to the response of P. tunicata wild type. Some of the P. tunicata mutants selected from the transposon library were phenotypically unstable after a series of subculturing. Only five AlpP- sensitive and four AlpP-resistant mutant strains retained their phenotypic characteristics. The AlpP-sensitive mutants were shown to have different degrees of sensitivity to AlpP. Three of the AlpP-sensitive mutants displayed an approximately two-fold increase in sensitivity to AlpP as compared to the P. tunicata wild type strain, while two AlpP- sensitive mutant strains had an increased sensitivity of approximately four-fold. Given that the variable increase was not the same for any of the mutant strains suggests that the genes disrupted by the transposon in each of these AlpP-sensitive mutants may have different functions in protecting the cell against AlpP.
Only one P. tunicata transposon mutant strain (RM1) displayed a complete resistance to AlpP when compared to the P. tunicata wild type strain, suggesting that the gene disrupted in RM1 AlpP-resistant mutant plays a major role in the sensitivity of P. tunicata to AlpP. The three resistant mutants (RM3, RM4 and RM5) demonstrated the same response to AlpP when tested in both logarithmic and mid-stationary phase of growth. The finding suggests that the genes mutated in these mutants may be involved in the same mechanism of rendering sensitivity response to AlpP.
The genotypic identity of the AlpP-sensitive and -resistant mutants was determined using the pan-handle PCR method which is described in section 2.2.6.4. This genetic tool was employed to identify the genes disrupted by the insertion of the mini-TnlO transposon. In addition, primer walking was also employed to extend the identification
88 of the DNA regions flanking the transposon insertion sites. The genes disrupted in the AlpP-sensitive mutants may be directly linked to the development of resistance by the bacterium to AlpP. Similarly, the genes disrupted in AlpP-resistant mutants may be involved in the sensitivity response of P. tunicata to AlpP.
The DNA region flanking the transposon in the P. tunicata SMI AlpP-sensitive mutant is involved in protein secretion across the cell membrane of bacteria. The SMI AlpP-sensitive mutant was disrupted in an open reading frame with 100 % identity to dppB gene, a putative ABC transporter of P. tunicata. This gene was identified in one of the dark purple mutants of P. tunicata generated by Egan (2001) for analysis of the production of antifouling products in P. tunicata. The dppB gene is also similar to other genes encoding ABC transporters in Gram-positive bacteria such as yvrO of B. subtilis and tptC of S. cristatus. The ATP-binding cassette (ABC) protein transporters in Gram negative bacteria are considered as general export pathways of proteins across the cell membrane. This export pathway completely by-passes the periplasm and directly secretes proteins from the cytoplasm across the outer membrane. The ABC protein transport apparatus consists of an ABC protein, which is an ATP-driven protein translocator, found in the inner membrane, and two accessory proteins which mediate the activity of the ABC protein exporter (consisting of a membrane fusion protein (MFP) and an outer membrane protein (OMP)) (Thanassi and Hultgren, 2000). The ABC protein transporters mediate the transport of several proteins such as toxins, proteases and lipases (Binet et al., 1997). These exoproteins contain a secretion signal in the C-terminal, specifically in the last 60 amino acids. Moreover, these exoproteins have a domain consisting of a Gly-rich sequence that is repeated 4-36 times (Welch, 1991). The ABC protein transporters are also employed in the uptake of substrates such as sugars (e. g. maltose, glucose, cellobiose), which are needed for cell growth (Seo et al., 2002). It has been proposed that DppB, a putative ABC transporter in P. tunicata, acts as an exporter of the yellow pigment and is involved in the uptake of a precursor to the synthesis of the yellow pigment (Egan et al., 2002a). Given that the ABC transporter is involved in both the export and import of molecules into and out of the bacterial cell, it is possible that the putative ABC transporter protein in P. tunicata is involved in the secretion of protective or immunity proteins against the autolytic protein, AlpP (see Chapter 3).
89 The DNA region flanking the transposon in the P. tunicata SM2 AlpP-sensitive mutant is similar to chiA, a chitinase gene of several bacteria such as Vibrio harveyi, V. cholerae and Aeromonas hydrophila. There was only one ORF identified in this sequence, followed by a termination sequence downstream of the ORF. Chitinases are found in various bacteria such as Serratia marcescens (Brurberg et al., 1994), V. cholerae (Folster and Connell, 2002), V. harveyi (Soto-Gill and Zyskind, 1984), A. caviae (Sitrit et al., 1995) and Pseudoalteromonas sp. strain S91 (Techkamjanaruk et al., 1997). These enzymes hydrolyze chitin polymers into small oligosaccharides, especially chitobiose. The hydrolysis of the p-(l-4)-glycosidic linkage of chitin can be carried out by exo- or endochitinase activity. The degradation of chitin is due to modules present in the chitinase protein complex. This complex consists of a catalytic domain, which is usually made up of proteins belonging to the hydrolyse family and a chitin-binding domain, which is involved in the binding of the enzyme to the substrate. Neither a catalytic domain, nor a chitin-binding domain was identified in the putative chiA gene of P. tunicata. However, it is possible that other chitinase genes are present upstream of the chiA in P. tunicata, comprising a chitinase operon. Although no chitin catalytic domain was identified, it is possible that the gene product has an active site that carried out the hydrolysis of the AlpP, making it inactive and hence provides a link between chiA and the resistance behavior of P. tunicata to AlpP. Another possible explanation is that steric hindrance prevents AlpP from coming into contact with the cell. It has been shown that chitinase is one of the catalytic enzymes in the cellulosome formed by the cell (Zverlov et al., 2002b). Cellulosomes are extracellular protuberant structures known to anchor enzymes such as cellulases and chitinases to the cell surface so that they act together to break down complex substrates (Shoham et al., 1999), see also Chapter 5. If cellulosome-associated enzymes are involved in the degradation of AlpP it is possible that the disruption of one of these enzymes could affect the function of the cellulosome.
The region disrupted by the transposon in the P. tunicata SM4 AlpP-sensitive mutant encodes for a protein similar to a hypothetical protein of Magnetococcus sp. MC-1. The ORF contains a tetratricopeptide repeat (TPR) domain. Proteins containing a TPR domain are involved in a wide spectrum of cellular activities, which include cell cycle control, protein transport, regulation of phosphate turnover and protein folding. In
90 bacteria and fungi, the TPR containing proteins are involved in protein transport (Blatch and Lassie, 1999). It was reported that a TPR domain is present in the magnetosomes of M. magnetotacticum (Okuda et al., 1996). However, the specific function of the TPR domain in the magnetosomes of this bacterium has not been established. As observed by phenotypic analysis, the SM4 mutant showed the highest MID value in a series of drop test assays which might suggest that this gene may have a major function in the response of P. tunicata to AlpP. It is also suggested that the gene product might be involved in the transport of protective or immunity proteins against the autolytic activity of the AlpP protein. There are two ORFs identified flanking the disrupted gene in P. tunicata SM4 AlpP-sensitive mutant. Upstream is an open reading frame, which has similarity to a hypothetical protein of Shewanella oneidensis. Downstream of the disrupted gene is an ORF, which has no similarity to any proteins submitted in the GenBank. There is no indication that the three ORFs identified in the SM4 AlpP-sensitive mutant belong to a single operon, as a putative promoter region precedes each of the ORF. However, the products of these genes might be involved in the transport of proteins, as they are putatively membrane-bound proteins. In this context, the phenotype of the SM4 AlpP- sensitive mutant may be explained by its nonfunctional membrane-bound proteins, which are possibly involved in the transport of proteins that protect P. tunicata cells against AlpP.
The DNA region disrupted by the transposon in the P. tunicata SM5 AlpP-sensitive mutant has homology to mshJ, a MSHA biogenesis protein of Vibrio cholerae. This gene is a component of the secretory operon of MSHA biogenesis protein of V. cholerae that is involved in the expression of type 4 pili (Marsh and Taylor, 1999). It is suggested that this gene encodes for proteins leading to the production of the extracellular pili. These extracellular structures may provide steric hindrance and thus preventing contact of AlpP with the bacterial cell. A detailed discussion of the SM5 mutant is presented in Chapter 4.
The region flanking the P. tunicata SM6 AlpP-sensitive mutant is 100 % identical to dppD, a methyl transferase gene of P. tunicata. It has previously been suggested that this gene encodes a product involved in the synthesis of the yellow pigment in P. tunicata (Egan, 2001), in which the putative methyl-transferase transfers methyl groups from a donor molecule to a recipient molecule. Furthermore, it is hypothesized
91 that dppB (the gene disrupted in the SMI mutant) and the dppD belong to a single putative operon which is involved in the synthesis and transport of the yellow pigment in P. tunicata (Egan, 2001). It is possible that this putative operon may also be involved in the synthesis and transport of protective or immunity proteins in P. tunicata.
The sequence information obtained from AlpP-resistant mutants gives insight into the sensitivity response of P. tunicata to AlpP. In RM1, the most AlpP-resistant mutant, the region flanking the transposon is similar to genes encoding the sensor histidine kinase/response regulator of several bacteria, including Synechocystis, P. aeruginosa and Caulobacter crescentus. The two-component sensor-regulator system is involved in processing intracellular information that links external stimuli to specific adaptive responses (West and Stock, 2001). The signaling pathway involves the phosphorylation of two conserved proteins: a histidine protein kinase and a response regulator protein. The two-component proteins are involved in the regulation of various cellular processes such as chemotaxis, metabolism, transport and osmoregulation. Moreover, it is reported that this sensor-regulator system controls the expression of toxins and other proteins essential for pathogenesis (Parkinson and Kofoid, 1992). For example, the GacS/GacA two-component system found in the majority of fluorescent Pseudomonas species is involved in multiple cellular functions including the production of virulence factors and secondary metabolites (Heeb and Haas, 2001). In P. syringae, the GacS/GacA regulates toxigenesis through the production of syringomycin, a peptide phytotoxin with the ability to cause necrosis in plant tissues (Rich et al., 1994). The gene disrupted in the RM1 AlpP-resistant mutant may have an important role in establishing sensitivity response of P. tunicata to AlpP. As shown by the drop test assays, this mutant showed total resistance to AlpP, thus the disrupted gene has an important link with the sensitivity of P. tunicata to AlpP. The presence of a putative two-component system in P. tunicata would suggest that the sensitivity of P. tunicata to AlpP is an adaptive response which is stimulated by the presence of AlpP. It is possible that AlpP is a signaling molecule, which is sensed by the bacterium through the two-component system and elicits a specific response such as autolysis. This would explain the phenotypic behavior of the RM1 AlpP-resistant mutant, i.e. the lack of non-functional two-component sensor-regulator system makes the mutant resistant to AlpP because the signal molecule (AlpP) is not perceived by the bacterium.
92 The transposon inserted in the P. tunicata RM3 and RM4 AlpP-resistant mutants occurred in different regions of the same gene. The sequence information of this region suggests strong homology to SyrE, a syringomycin peptide synthetase in P. syringae pv syringae. The ORE has core sequences and domain regions similar to those of SyrE. In the plant pathogen P. syringae pv syringae, SyrE is one of the modules involved in the production of a syringomycin, a pore-forming peptide, which causes necrosis of plant tissues (Rich et al., 1994). P. tunicata may also have other modules similar to SyrE, because a domain similar to SyrE has been identified upstream of the putative syrE gene. Based on these results it may be proposed that P. tunicata produces syringomycin- like autolysins which cause lysis of the bacterial cell.
The transposon inserted in the AlpP-resistant P. tunicata mutant, RM6, occurred in the non-coding frame of the DNA region. Further sequencing of this region, identified three ORFs, of which only one, when translated, has similarities with the proteins in the GenBank database. The ORF2 is situated downstream of the transposon insert. The gene product of this ORF is similar to the acetyl transferases of P. putida and Bacteroides thetaiotaomicron. It is suggested that this gene may be involved in the biosynthesis of autolysins, in which an acetyl group is transferred from the donor molecule to the recipient molecule.
To conclude, this chapter has initiated phenotypic and genotypic studies of the resistance and sensitivity responses of P. tunicata to AlpP. The genes identified in the AlpP-sensitive mutants appear to encode traits involved in the protein transport pathway and biosynthesis of proteins which suggest that the gene products are linked with the synthesis and transport of protective or immunity proteins against AlpP. Conversely, the genes disrupted in the AlpP-resistant mutants may encode proteins involved in sensory and regulatory' mechanisms in the cell. The genetic information obtained from the transposon mutagenesis outlined in this chapter forms the basis of a proposed model of the mode of action of AlpP in P. tunicata. This hypothetical model is discussed in detail and presented in the next chapter.
93 3. Elucidation of the mode of action of AlpP and the response of Pseudoalteromonas tunicata to its autolytic protein product
3.1. Introduction
Many marine microorganisms have been reported to produce antibacterial agents. Examples include loloatins, cyclic decapeptide antibiotics from a marine Bacillus (Gerard et al., 1999) and 5-indomycinone, a new member of the pluramycin family of antibiotics from marine Streptomyces species (Biabani et al., 1997). Specifically, many species of the genus Pseudoalteromonas have been reported to produce antibacterial agents and several bioactive molecules (Holmstrom and Kjelleberg, 1999). It has been suggested that the production of bioactive compounds by Pseudoalteromonas species assist the bacterial cells in their competition for nutrients and space and aids in the colonization of marine surfaces (Holmstrom and Kjelleberg, 1999). Pseudoalteromonas species which produce antibacterial compounds include P. rubra (Gauthier, 1979), P. luteoviolacea (Gauthier and Flatau, 1976), P. aurantia (Gauthier and Breittmayer, 1979) and P. citrea (Gauthier, 1979). The antibacterial agents produced by these species have a broad range of activity. Auto-inhibition has also been observed in Pseudoalteromonas species such as P. luteoviolacea (Gauthier and Flatau, 1976) and P. denitrifricans (Enger et al., 1987). It has been suggested that the mode of action of auto-inhibition contributes to the maintenance of microbial diversity in the marine ecosystem (Holmstrom and Kjelleberg, 1999). A powerful competitor is P. tunicata. This marine bacterium has been shown to produce several different bioactive compounds, including an antibacterial agent. The antibacterial agent produced by P. tunicata has been demonstrated to be a protein, which is produced during the stationary phase of growth (James et al., 1996). The antibacterial protein is known to have a broad spectrum of activity against many marine, soil and medical isolates (James et al., 1996; Stelzer, 1999). P. tunicata ceils were also found to be sensitive to its own antibacterial protein, however, it was observed that cells developed resistance during the stationary phase of growth (James et al., 1996).
94 It has been reported that phage-tail like structures mediate autolysis in producing bacteria. Recently, autolysis in P. aeruginosa biofilm has been linked to the expression of a Pfl -like filamentous prophage of P. aeruginosa (Webb et al., 2003b). This autocidal system led to the hypothesis that the antibacterial protein AlpP may be related to phage or phage tail-like molecules. To test this hypothesis, the ultrastructure of AlpP was investigated in this chapter using transmission electron microscopy (TEM) and immunolabelling studies.
Another well-studied autolytic mechanism in bacteria is the toxin-antidote systems, which elicits bacterial cell death in E. coli. This "self-destructive" behavior is controlled by a plasmid, which encodes for a stable protein toxin and an unstable protein antitoxin. This controlled autolytic mechanism may also explain the sensitivity and resistance of P. tunicata to AlpP. The production and release of an antidote that counteracts the autolytic protein may explain the resistance developed by P. tunicata against its own autolytic protein. This chapter describes studies to identify the protective or “antidote” molecule in P. tunicata. Furthermore, the hypothesis in chapter 2 that SMI (dppB) and SM6 (dppD) mutants are involved in synthesis and export of an antidote was investigated.
Finally, the transposon mutational analysis carried out in Chapter 2 is discussed in detail in this chapter, leading to several hypothetical models to explain the mechanism of action of the antibacterial protein AlpP.
95 3.2. Materials and Methods
3.2.1. Bacterial strains and culture conditions
The Escherichia coli strains BL21 (DE3) (F" ompT r"e rn'e) (Mardones and Venegas, 2000) and ML-35p (Tsuji et al., 2001) were grown in Luria-Bertani broth (LB) with shaking at 37°C. The P. tunicata transposon mutants SMI and SM6 were grown at room temperature in VNSS medium containing 85 pg/ml kanamycin (Km) and 100 pg/ml of streptomycin (Sm) and the wild type P. tunicata was grown at room temperature in VNSS medium.
3.2.2. Preparation of cell-free concentrated supernatant from P. tunicata wild type, SMI and SM6 Alp-sensitive mutants
The hypothesis that SMI (dppB) and SM6 (dppD) mutants are involved in synthesis and export of an antidote was investigated. The P. tunicata wild type, SMI and SM6 mutants cell-free concentrated supernatant were prepared as described in section 2.2.2. An aliquot of each cell-free concentrated supernatant was tested for autolytic activity using the drop test assay, and stored at -20°C for further analysis.
3.2.3. Fractionation of the AlpP protein from the cell-free concentrated supernatant
The fractionation of AlpP protein from the cell-free concentrated supernatants of P. tunicata wild type and SMI and SM6 mutants was performed using the method described in section 2.2.3. Three different fractions were collected and analyzed. The first fraction (FI) was eluted with 0.1 M NaCl. The second fraction (F2), which contained the AlpP protein, was eluted with 0.3 M NaCl and the third fraction (F3) was eluted with 0.7 M NaCl. The different fractions were assayed for autolytic activity using the drop test assay, and stored at -20°C.
96 3.2.4. Determination of bacteriolytic or bacteriostatic activity of the AlpP protein
Investigation of the hypothetical lytic effect of the AlpP protein was performed using a chromogenic plate test (Mardones and Venegas, 2000), in which the activity of AlpP was compared with other antibiotics with known bacteriolytic and bacteriostatic activities.
The chromogenic plate assay is based on the use of B-galactosidase as a marker of cellular lysis, by following the hydrolysis of the chromogenic compound 5-bromo-4- chloro-3-indoyl-B-D-galactoside (X-Gal) to form a blue product. The target cells used in this assay express B-galactosidase, which is released into the medium when cell lysis occurs. The released B-galactosidase hydrolyzes X-Gal, which is included in the agar medium, to form a blue stain at the edge of the antibiotic inhibition zone.
The E. coli strain BL21 (DE3) (F' ompT r"B m'B), which expresses cytoplasmic p-galactosidase, was used in this assay. Antibiotics known to employ bacteriolytic and bacteriostatic effects such as ampicillin and chloramphenicol, respectively, were used as positive controls. The target bacterial strain was grown, shaking, in 2 ml of LB medium at 37°C overnight. Fifty microlitres of bacterial inoculum was mixed with 10 ml of 0.8 % agar (previously melted and stored at 45°C), 10 pi of 1 mM IPTG and a final addition of 50 pi of 50 mg/ml X-Gal. This soft-agar incubation mixture was carefully overlayed onto LB plates containing solidified 1.5 % agar. Ten microlitres of each of the antibiotics and AlpP protein sample were then applied to the solidified soft agar surface using fine disposable tips. The antibacterial agents used were ampicillin at a concentration of 6 pg/ml (bacteriolytic) and chloramphenicol at a concentration of 37.5 pg/ml (bacteriostatic) and the AlpP protein at a concentration of 410 pg/ml. The plates were incubated at 37°C for 9-16 h. The chromogenic assay was carried out in triplicate.
97 3.2.5. Assay of inner-membrane permeability in the presence of AlpP protein
In order to determine the permeability of bacterial inner-membrane in the presence of AlpP protein, the leakage of p-galactosidase from the cytoplasm of the target cell was assessed. The E. coli strain ML-35p was used as the target strain as it constitutively expresses periplasmic P-lactamase and cytoplasmic p-galactosidase. In this experiment, the expression of the p-lactamase was not relevant because the integrity of the inner membrane was assessed by the release of cytoplasmic p-galactosidase into the liquid substrate (Tsuji et al., 2001). Cells of E. coli ML-35p were grown in LB 10 and incubated for 16-18 h at 37°C with shaking. After incubation, the cells were centrifuged at 10, 000 x g for 5 min and resuspended in 50 ml of fresh LB 10 to a final concentration
o of approximately 1x10 cfu/ml (OD620nm = 0.35). One hundred pi of active cell-free concentrated supernatant (1.64 mg/ml, total protein concentration) from P. tunicata was added to the cell suspension. The mixture was incubated at 37°C with shaking. One ml of sample was collected every 20 min from 0 to 120 min of incubation. The sample was centrifuged at 10, 000 x g for 5 min and the supernatant was assayed for p- galactosidase. The P-galactosidase assay mixture was made up of 75 pi of supernatant, 125 pi Z-buffer (Miller, 1972) (Appendix I) and 40 pi onitrophenyl-/?-D- galactopyranoside (ONPG, 4 mg/ml dissolved in phosphate buffer, pH 7.0). Incubation of the assay mixtures was done at 37°C for 5-10 min and the optical density measured at 420 nm. The AOD values presented here were calculated by subtracting OD 420 nm at 0 h from OD 420 nm at each time point. The assay was carried out in triplicate.
3.2.6. Transmission electron microscopy studies of P. tunicata AlpP protein
In Gram-positive bacteria, the phage tail-like antibiotics, bacteriocins, mediate autolysis of the producing cells (Guder et al., 2000). Recently, autolysis in P. aeruginosa biofilm has been linked to the expression of a Pfl-like filamentous prophage of P. aeruginosa (Webb et al., 2003b). This autocidal system led to the hypothesis that the antibacterial protein AlpP may be related to phage or phage tail-like molecules. To investigate this hypothesis, transmission electron microscopy (TEM) and immunolabelling studies were
98 carried out. The P. tunicata active cell-free concentrated supernatant containing the AlpP protein was examined using electron microscopy. The sample was placed on a carbon-coated formvar copper grid and negatively stained with 2 % phosphotungstic acid for 30 sec. The grid was examined using a Hitachi H700 TEM at an accelerating voltage of 75 kV.
The partially purified AlpP-containing fractions obtained from ion-exchange chromatography were visualized on native-PAGE using the protocol described in James (1996). The band corresponding to the AlpP protein was excised, placed in a 3 ml syringe and pushed through the opening without a needle into a second syringe to disintegrate the gel. The procedure was repeated five times. The gel material was collected in a 2 ml Eppendorf tube and 1 ml of 20 mM Tris-HCl, pH 7.5 was added. The mixture was vortexed for 30 sec and incubated overnight at room temperature. The gel material was centrifuged at 12, 000 x g for 1 min and the supernatant collected and resolved on a native-PAGE using the protocol described in James (1996) to detect the presence of the AlpP protein band.
The fractions from ion-exchange chromatography and the AlpP protein eluted from native-PAGE were placed on the carbon-coated formvar copper grid and fixed using PBS with 0.1 % glutaraldehyde for 5 min. The grid was rinsed with 50 mM phosphate buffer solution, pH 7.0 containing 1 % bovine serum albumin (PBS + BSA) for 5 min. The grid was transferred onto drops of P. tunicata AlpP protein polyclonal antibody (1:100) (Mai-Prochnow et al., 2004) and incubated for 2 h. After incubation the grid was rinsed three times with 1 % BSA in PBS for 3 min and then the grid was rinsed twice with PBS (pH 7.0). The goat anti-rabbit IgG conjugated with 12 nm gold, EM Grade (Jackson Immuno Research Laboratories, Inc., PA, USA) was used as the secondary antibody at a 1:20 dilution, and incubated for 1 h. The grid was rinsed five times with PBS, pH 7.0. The grid was fixed for 5 min on a drop of 2 % glutaraldehyde, and then rinsed twice with milli-Q water for 5 min prior to staining for 30 sec with 2 % phosphotungstic acid. The stained grid was examined using a Hitachi H700 TEM at an accelerating voltage of 75 kV.
99 3.2.7. Transposon mutagenesis and genetic analysis
To investigate how P. tunicata responds to the AlpP protein, transposon mutagenesis and genetic analysis were carried out as described in Chapter 2 (section 2.2.4.) The genetic information obtained from the transposon mutants led to a hypothetical model addressing the responses of P. tunicata to its autolytic protein, AlpP.
3.2.8. Investigation of the toxin-antidote system in P. tunicata
The development of resistance towards its autolytic protein is an interesting characteristic of P. tunicata. One means by which resistance could be achieved is through the production and release of an antidote that counteracts the autolytic protein. Two different studies were carried out to identify a putative antidote. The first study examined the effect of dilution upon the equilibrium between AlpP activity and hypothetical antidote. Secondly, the concentrated supernatant was fractionated using ion exchange chromatography and the fractions were examined for protective effects against the activity of AlpP.
3.2.8.1. Identification of a putative antidote activity in active cell-free concentrated P. tunicata supernatant
The concentrated supernatant was prepared as described previously (section 2.2.2). One hundred pi of fresh VNSS containing 5 pi of P. tunicata cells from an overnight culture (OD = 0.35) was inoculated in each well of a 96-well microtitre plate. One hundred pi of the diluted supernatant was then added into each of the wells, making a final volume of 200 pi. The dilutions used in the experiment were Ox (undiluted supernatant), 2x, 4x, 8x, 16x, 32x, 64x, 128x, 256xx, 512x and 1024x. Each treatment was done in triplicate. Wells containing P. tunicata cells without the supernatant were used as controls. The optical density was measured at 600 nm from 0 h until 15 h of incubation. The plate was incubated at room temperature with shaking during the reading interval.
100 3.2.8.2. Fractionation of the cell-free concentrated supernatant to identify a putative antidote factor
The fractionation of the concentrated supernatant was described in section 2.2.3. Three different fractions were collected, namely: FI- (non-autolytic) eluted with 0.1 M NaCl, F2 - (autolytic) eluted with 0.3 M NaCl and F3 - (non-autolytic) eluted with 0.7 M NaCl. The non-autolytic and autolytic fractions were mixed in 50:50 volume ratios. The pooled fractions were diluted in different serial dilutions and assayed using the drop test assay. The active fraction and buffer in a 50:50 volume ratio was used as control. A reduction in the activity of the AlpP-containing fraction when mixed with the non-AlpP fraction was scored as an indication of an antidote factor.
Another approach to validate the presence of antidote in the inactive fractions was by heat treatment, which is based upon its chemical stability, as bacterial antidotes are usually unstable molecules (Engelberg-Kulka and Glaser, 1999). Non-active fractions were exposed to 90°C prior to the addition of active fractions containing the AlpP protein for the drop test assay. Mixtures with clearing zone on the surface of the bacterial lawn indicate that the antidote was inactivated by heat.
3.3. Results
3.3.1. Chromogenic plate assay to evaluate bacteriolytic activity of P. tunicata AlpP protein
A chromogenic plate assay was carried out to distinguish whether the mode of action of P. tunicata AlpP protein is bacteriolytic or bacteriostatic. It was demonstrated that AlpP killed E. coli BL21 effectively by lysing the bacterial cells. P. tunicata AlpP displayed similar pattern of clearing zones as the control bacteriolytic antibiotic, ampicillin (Figure 3-1). However, P. tunicata AlpP protein displayed a clearing zone of only 5 mm in diameter, which is 2 mm diameter less than that of ampicillin. The P. tunicata AlpP showed rapid killing, where a blue stain was observed as early as 9 h after incubation. The blue stain continuously intensified up to 16 h of incubation. Interestingly, the
101 intensity of the blue color was more pronounced for the P. tunicata AlpP protein than for ampicillin. The bacteriolytic activity of P. tunicata AlpP protein may result in the leakage of cytoplasmic (3-galactosidase from the target cell. The release of P-galactosidase suggests that the P. tunicata AlpP protein disrupts the inner membrane of target cells leading to a rapid efflux of ions, solutes and other cellular metabolites and thus causing cell death.
Figure 3-1. Chromogenic plate assay showing the bacteriolytic activity of P. tunicata AlpP protein. Note the presence of a blue-colored edge at the clearing zone produced by A) P. tunicata AlpP protein, 410 pg/ml and B) ampicillin, 6 pg/ml. No blue color was detected at the clearing zone of the bacteriostatic agent, C) chloramphenicol, 37.5 pg/ml).
102 3.3.2. Inner-membrane permeability assay
To further evaluate the bacteriolytic activity of the antibacterial protein, an additional assay was performed. This assay measured the p-galactosidase released from the cell into a liquid substrate. To measure the release of cytoplasmic p-galactosidase, the supernatant from the cell suspension treated with P. tunicata AlpP protein was added to a buffer containing ONPG, and the absorbance of ONP was measured at 420 nm over time. ONP is a break down product of ONPG through the action of p-galactosidase. It was clearly demonstrated that the cytoplasmic p-galactosidase was released from the target cell, as shown by the increased ONP absorbance in treatment samples. The difference between the treatment and control is presented in Figure 3-2. The P. tunicata AlpP protein caused immediate killing of the E. coli ML-35p strain with about 93 % dead cells after 60 min of incubation. The absorbance reading of the cell suspension with P. tunicata AlpP protein and the control containing only the cell suspension determined the killing percentage. The lysis of the cells was detected by the release of cytoplasmic P-galactosidase. p-galactosidase was detected in the treatment samples after 20 min of incubation and levels continuously increased for 60 min. This continuous release of P-galactosidase suggested that the activity of the P. tunicata AlpP protein is bacteriolytic. It supports and confirms the results of the previous chromogenic plate assay.
103 Time (minutes)
Figure 3-2. Effect of P. tunicata active cell-free concentrated supernatant on bacterial inner-membrane permeability. A culture supernatant of E. coli ML-35p, expressing cytoplasmic p-galactosidase, exposed to P. tunicata concentrated supernatant, was added to ONPG, a substrate for that enzyme. An increase in OD 420 nm represents enhanced release of P-galactosidase, suggesting disruption of bacterial cell-membrane.
3.3.3. Transmission electron microscopy studies of P. tunicata AlpP protein
As phage or phage tail-like molecules have been shown to mediate autolysis in some bacteria, such as P. aeruginosa (Webb et al., 2003b), it was considered that P. tunicata AlpP may be associated with phage or phage tail-like molecules. To determine if this hypothesis is true, TEM and immunolabeling studies were performed. The negatively stained samples from concentrated cell-free supernatant and the ion-exchange fraction were shown to contain different types of phage-like structures. The TEM image of the phage-like structure from concentrated cell-free supernatant showed that it has a head of -75 nm in diameter, which is joined to a contractile tail of -150 nm (Figure 3-3A). Some appeared to have no heads, just a long hollow tail of -90 nm (Figure 3-3B). The ultrastructure of the phage-like structures from ion exchange chromatography samples
104 revealed another type of phage. In this case, the phage head had a diameter of ~110 nm and a -120 nm long tail (Figure 3-3C and Figure 3-3D). Several hollow tails with no heads were also observed in the sample. To verify if these phage and phage tail-like structures are associated with P. tunicata AlpP, an immunolabeling experiment was carried out on samples containing the protein using the AlpP protein antibody, as the primary antibody (Mai-Prochnow et al., 2004). A 1:100 dilution of the P. tunicata AlpP antibody was shown to be the optimal concentration for labeling, under which conditions there was no background of unspecific binding seen in the samples. To label the binding of primary antibody to the AlpP protein, a secondary antibody, goat anti rabbit IgG conjugated with 12 nm of gold, was used. The dilution of the secondary antibody that demonstrated the optimal concentration for labeling the AlpP protein was 1:20. The TEM image of the ion-exchange fractionation sample revealed that the phage and phage tail-like structures were not labeled (Figure 3-3E and Figure 3-3F) however a labeled aggregate complex structure was observed. Similar aggregate structures were observed in all immunolabeled samples which could be explained by the precipitation of the AlpP protein during the first stage of fixation of sample using glutaradehyde. To further validate this observation, the protein band from the native-PAGE gel, which corresponds to the AlpP protein was eluted into buffer solution and used for immunolabeling. It was clearly shown that there were no phage or phage tail-like structures in the sample (Figure 3-3G and Figure 3-3H). Images from immunolabeling revealed the presence of a labeled aggregate mass of complex structure, similar to that observed in immunolabeled samples from ion-exchange chromatography. These additional immunolabeled images confirm that the P. tunicata AlpP protein is not associated with the phage-like structures and is structurally distinct from phage or phage tail-like autolysins.
105 Figure 3-3. Transmission electron micrographs and immunolabeling studies of P. tunicata AlpP protein. Phage-like structures were observed in P. tunicata cell-free concentrated supernatant (A and B) and the antibacterial fraction from ion exchange chromatography (C and D). Immunolabeling of the AlpP fraction purified using ion- exchange chromatography (E and F) and the AlpP protein band cut from native-PAGE (G and H). Phage-like particles were not labeled (arrow head) and these structures were not seen on the pure AlpP protein. Bar = 200 nm
106 3.3.4. P. tunicata genes associated with resistance and sensitivity to AlpP
Although the above assays demonstrate that the P. tunicata AlpP protein kills target cells by lysis, additional studies are needed to clarify how P. tunicata responds to its own autolytic protein and develop resistance. Transposon mutagenesis was employed to further study the mechanism of the autolytic activity. In this chapter, a hypothetical model of the mechanism of action of AlpP is proposed based on the information obtained from the transposon mutagenesis described in Chapter 2. Three genes (dppB, dppD and mshJ), identified by using transposon mutagenesis and found to be associated with AlpP resistance, are involved in the protein transport and secretion (Table 3-1). The gene disrupted in another AlpP-sensitive mutant, SM4, showed no homology when translated with any of the proteins in GenBank. However, the predicted location of all proteins encoded by the disrupted genes is in the cytoplasmic membrane and periplasmic space, suggesting that these proteins may be involved in the secretion and assembly of proteins in the cell. The genes identified in AlpP-resistant mutants are involved in bacterial sensing and regulatory mechanisms and biosynthesis of bioactive metabolites. The gene disrupted in the RM5 AlpP-resistant mutant encodes a protein which showed no homology with any of the proteins submitted to the GenBank.
107 Table 3-1. Phenotypic characteristics of P. tunicata transposon mutants.
MID value (drop test) Strain Logarithmic Stationary Sequence similarity - protein phase cells phase cells and organism
WT 4x Ox
SMI 8x 8x Putative ABC transporter, DppB, P. tunicata ABC-transporter proteins, YvrO, Bacillus subtilis ATP-binding protein, TptC, Streptococcus cristatus SM2 8x 4x Chitinase, ChiA, Vibrio harveyi, Vibrio cholerae, Alteromonas hydrophila SM4 16x 4x Hypothetical protein, Magnetococcus sp. MC-1 SM5 8x 8x MSHA biogenesis protein, MshJ, Vibrio cholerae SM6 16x 8x Methyl transferase, DppD P. tunicata, Streptomyces coelicolor RM1 Ox Ox Two component hybrid sensor and regulator, Synechocystis sp. PCC 6803, Pseudomonas aeruginosa and Caulobacter crescentus CB15 RM3 2x 2x Syringomycin synthetase, SyrE, Pseudomonas syringae RM4 2x 2x Syringomycin synthetase, SyrE, Pseudomonas syringae RM5 2x 2x No significant sequence homology Sequence similarities determined by NCBI GenBank searches using the BlastX program MID value = maximum inhibitory dilution (using drop test assay) WT = wild type Pseudo alter omonas tunicata SM = AlpP-sensitive mutant RM = AlpP-resistant mutant
108 3.3.5. Investigation of the toxin-antidote system in P. tunicata
It was previously found that P. tunicata is sensitive to its own autolytic protein, AlpP, during the logarithmic phase of growth and becomes resistant during the stationary phase of growth (James et al., 1996). This suggests that P. tunicata has mechanisms to protect itself from autolysis. Because stationary phase cells were already known to have increased resistance to AlpP exposure (James et al., 1996), the concentrated supernatant from a high-density stationary phase cell suspension was tested for protective effects. The P. tunicata wild type cells and serial dilutions of the concentrated supernatant were added to microtitre wells, along with some fresh nutrients and their growth monitored over time. The growth pattern of the cells exposed to serial dilutions of concentrated supernatant occurred into two phases (Figure 3-4). There was an unexpected resistance observed between 0 and 2-fold dilution as shown by the increase in final cell density of the P. tunicata indicating that cells were less inhibited. One possibility to explain the unexpected resistance is the presence of an antidote factor that counteracts the toxic effect of AlpP. In E. coli, antidotes are expressed simultaneously with the toxin, and subsequently counteract the lethal effect of the toxin (Engelberg-Kulka and Glaser, 1999). In the second phase (between 4-fold and higher), the P. tunicata cells were strongly inhibited at the 4-fold dilution and progressively less inhibited at increasing dilutions with a plateau obtained at 32-fold dilution. This result suggests that an antidote factor may be diluted out of its active concentration range earlier than the AlpP protein, as antidotes are usually small and unstable molecules (Engelberg-Kulka and Glaser, 1999), thus the toxic effect of AlpP is triggered at low-fold dilution (e.g. 4-fold dilution),
109 0.50
Dilutions
Figure 3-4. Growth of P. tunicata cells in the presence of serial dilutions of the concentrated supernatant after 16 h. Error bars represent the standard error of triplicate cultures.
The next study was aimed at separating the antidote factor from AlpP by fractionating the concentrated supernatant using anion exchange chromatography. Three different fractions were collected: a) FI - non-autolytic fraction, b) F2 - autolytic fraction and c) F3-non-autolytic fraction. The non-autolytic fractions (FI and F3) from ion- exchange chromatography were mixed separately with the active fraction (F2) containing the AlpP protein. In Table 3-2, it was shown that fraction containing autolytic activity (F2) became less toxic (i.e. lost its activity more quickly upon dilution) when mixed with the non-toxic fraction F3, as compared to the control sample (F2 mixed with buffer). A two-fold reduction in its activity was observed (i.e. from 8x MID to 4x MID), possibly suggesting the presence of an antidote factor in fraction F3. The molecules eluted in this fraction (0.7 M NaCl) are likely to be strongly positively charged molecules, as they remained adhered to the anion exchange column until a
110 relatively high salt concentration was used. When F1 was added to the active fraction (F2), the mixture displayed a MID value of 16 which is higher than the MID value of F2 with buffer. This suggests that there are undefined factors present in FI that support the effect to AlpP.
An additional experiment to identify the presence of a putative antidote in heat-treated supernatant was performed. This test is based on the fact that antidotes are often unstable proteins (Engelberg-Kulka and Glaser, 1999). In this study the inactive fractions, FI and F3, were heat treated prior to the drop test assay. The result showed that there was no change in the auto lytic activity of F2 either when mixed with FI or F3 (Table 3-3). The unchanged MID value of F2 when mixed with heat-treated F3 suggests that the reduction in activity caused by untreated F3 in the previous experiment must be due to a factor which is inactivated by heat-treatment. This is consistent with the presence of a heat-labile antidote factor. Similarly, the unchanged MID value of F2 after mixing with the heat-treated FI fraction suggests that the supported antibacterial effect observed in FI (non heat treatment experiment) is lost after heat treatment.
Table 3-2. Relative activity of cell-free concentrated P. tunicata culture supernatant fraction mixes.
Fraction mix F2 + buffer F2 + F1 F2 + F3
MID (drop test) 8x 16x 4x
MID = Maximum inhibitory dilution (using the drop test assay) FI = Fraction 1, non-autolytic, eluted with 0.1 M NaCl F2 = Fraction 2, autolytic fraction, eluted with 0.3 M NaCl F3 = Fraction 3, non-autolytic eluted with 0.7 M NaCl All fractions were mixed to a 50:50 volume ratio (F2 only mixed with buffer)
111 Table 3-3. Drop test assay of the heat-treated (90°C) inactive and active fractions of the P. tunicata cell-free concentrated supernatant after ioni exchange chromatography.
The active fraction, F2 was not heat-treated prior to drop test assay.
Fraction mix F2 + buffer F2 + F1 F2 + F3
MID (drop test) 8x 8x 8x
MID = Maximum inhibitory dilution (using the drop test assay) FI = Fraction 1, non- autolytic, eluted with 0.1 M NaCl F2 = Fraction 2, autolytic fraction, eluted with 0.3 M NaCl F3 = Fraction 3, non- autolytic eluted with 0.7 M NaCl All fractions were mixed to a 50:50 volume ratio (F2 only mixed with buffer)
Finally, an experiment using the concentrated supernatant of the SMI and SM6 mutants was performed to explore the presence of an antidote in these mutant strains. It was hypothesized that the genes (dppB and dppD) disrupted in these mutants may be involved in the synthesis and transport of the hypothetical antidote. The autolytic fraction F2 became less toxic (i.e. lost its activity more quickly upon dilution) when mixed with fraction F3 from both the SMI and SM6 mutants, as compared to the control sample (F2 mixed with buffer), as shown in Table 3-4. These results demonstrated that the mutants had similar antidotal properties to the wild type, indicating that the disrupted genes are not involved in the synthesis and transport of the hypothetical antidote.
112 Table 3-4. Relative activity of cell-free concentrated P. tunicata culture supernatant fraction mixes.
F3 as taken from concentrated supernatant of the wild type, the SMI and SM6 mutants.
Fraction Mix MID (drop test assay)
F2 (D2 wt) + buffer 4x
F2 (D2 wt) + F3 (D2 wt) 2x
F2 (D2 wt) + F3 (SMI) 2x
F2 (D2 wt) + F3 (SM6) 2x
MID = Maximum inhibitory dilution (using the drop test assay) F2 = Fraction 2, autolytic fraction, eluted with 0.3 M NaCl F3 = Fraction 3, non- autolytic eluted with 0.7 M NaCl All fractions were mixed to a 50:50 volume ratio (F2 only mixed with buffer)
113 3.4. Discussion
3.4.1. The AlpP is bacteriolytic to target cells
Microorganisms produce inhibitory agents to compete against bacteria with the same nutritional requirements or colonizers of the same ecological niche (Lemos et al., 1991). The surface colonizing marine bacterium P. tunicata produces an antibacterial agent, AlpP, which is suggested to give P. tunicata a competitive advantage against other surface colonizers (James et al., 1996). This bacterium was previously found to have a broad-spectrum of inhibitory activity against other bacteria, including important medical and marine isolates (James et al., 1996). As described in this chapter, the mode of action of the antibacterial activity of AlpP was studied and found to be bacteriolytic. This was demonstrated using a chromogenic plate assay and an inner-membrane permeability assay, in which the leakage of cytoplasmic p-galactosidase, a marker of cellular lysis, was observed from the target cell. The release of P-galactosidase suggests that the P. tunicata AlpP protein may disrupt the inner membrane of target cells which leads to rapid leakage of ions, solutes and other cellular metabolites and thus cause cell death. A group of antibacterial agents with similar mechanism of activity to P. tunicata AlpP are the bacteriocins. Bacteriocins are produced by Gram-positive bacteria and usually occur as small peptides of 2-5 kDa in size (Guder et al., 2000). The killing mechanism is initiated by the formation of pores in the cytoplasmic membrane leading to rapid efflux of ions, solutes, amino acids, ATP and other nucleotides, followed by immediate arrest of all biosynthetic processes and cellular death (Ruhr and Sahl, 1985; Sahl and Bandis, 1982). The bacteriolytic effect of AlpP is suggested to be of great advantage for P. tunicata in its competition with other bacteria for nutrients and space.
114 3.4.2. P. tunicata AlpP is not associated with phage or phage tail like structures
AlpP protein and bacteriocins are both auto-toxic. Bacteriocins kill cells of the same or closely related species and they appear as phage tail-like structures and can be induced by mitomycin C (Daw and Falkiner, 1996). Recent studies in our laboratory have shown that auto-toxicity in P. aeruginosa biofilms is linked to activity of a P. aeruginosa bacteriophage (Webb et al., 2003b). The similar role of AlpP in killing cells in biofilm of P. tunicata to that of bacteriophage in P. aeruginosa led to the hypothesis that AlpP may be a phage or phage tail-like structure. In order to investigate this hypothesis, TEM and immunolabeling experiments were performed. It was shown that phage and phage tail-like structures were present in the cell-free concentrated supernatant and the ion exchange chromatography active fraction. However, the immunolabeled TEM image of the ion-exchange fractionation sample revealed the presence of unlabeled phage-like structures suggesting that AlpP protein is not the phage-like structures. This observation was confirmed by immunolabeling the protein band from the native-PAGE gel corresponding to AlpP protein. The images revealed no phage tail-like structures present in the samples and thus AlpP protein is very distinct from bacteriocins. Moreover, it is unlikely that AlpP originates from a phage because there are no phage flanking regions identified around the alpP gene (S. James, unpublished data).
3.4.3. P. tunicata genes encoding traits which mediate altered sensitivity to autolytic protein, AlpP
To further explore the role of AlpP production in P. tunicata and to understand the mechanism of the autolytic activity, transposon mutagenesis analysis was employed. The information obtained from AlpP-sensitive mutants facilitates the identification of genes involved in resistance to the AlpP protein and possibly the genes involved in the production or regulation of an “antidote” molecule. The information obtained from AlpP-resistant mutants identifies genes linked to bacterial sensing and regulatory mechanisms and the biosynthesis of bioactive metabolites. The summary of the phenotypic and genotypic information obtained from transposon mutational analysis is
115 presented in Table 3-1. Several genes were identified from the selected mutants, including those involved in bacterial transport and signal mechanisms, the production of extracellular structures, a two-component hybrid sensor-regulator system and the synthesis of metabolites. A hypothetical model of the mode of action of AlpP against P. tunicata is presented in Figure 3-5.
3.4.3.1. Transport and signal mechanisms are linked with AlpP activity
The mutated genes of the two AlpP-sensitive mutants SMI and SM6 are related to each other. The mutants were disrupted in the genes dppB and dppD, respectively, which have previously been identified in P. tunicata (Egan et al., 2002a). The dppB gene encodes a putative ABC transporter similar to the yvrO, which encodes an ABC transporter in B. subtilis. The dppD gene encodes a putative methyl transferase proposed to be involved in the biosynthesis of antifouling agents in P. tunicata (Egan et al., 2002a). The dppB and dppD genes were predicted to belong to the same operon (Egan et al., 2002a). Several hypotheses are suggested to explain the involvement of these genes in the production of the autolytic activity of AlpP. A possible mechanism is based on the hypersensitivity response observed in B. licheniformis, which uses bcrABC genes encoding for ABC transporter proteins for transporting the antibiotic (bacitracin) out of the cell. It was shown that a mutation on the ABC transporter resulted in hypersensitivity to bacitracin (Ohki et al., 2003). As a result of this observation it was proposed that the ABC transporter proteins protect B. licheniformis against its own product by facilitating the export of bacitracin out of the cell (Ohki et al., 2003). A similar mechanism may occur in P. tunicata, where an ABC transporter may transport out the AlpP protein out of the cell. However, as shown previously (Egan et al., 2002a), the dppB mutant maintained the antibacterial activity and was able to export the AlpP protein. Moreover, the amino acid sequence of AlpP does not contain the carboxy-terminal secretion signal common to substrates of this transport pathway (Binet et al., 1997). These results suggest that a dp/?i?-associated transporter does not excrete the AlpP.
The second possible mechanism of AlpP resistance involves the synthesis and export of an antidote, which protects P. tunicata from AlpP autolytic activity. It is suggested that
116 the products of the dppD and dppB genes are jointly involved in the synthesis and export of antidote. In such a case, the dppD gene would be involved in antidote synthesis and the dppB gene would be involved in export of the antidote. This is consistent with the results so far obtained in screening the transposon mutant library, as the dppD mutant would no longer encode for synthesis of the antidote and the dppB mutant would consequently no longer encode for products involved in export of the antidote, hence explaining their hypersensitivity to AlpP. Investigation of the presence of a hypothetical antidote was therefore carried out. The supernatants of both the dppD (SM6) and dppB (SMI) mutants were fractionated and screened for antidote activity, and both were found to be equally able to neutralise the activity of AlpP, indicating that this hypothesis is incorrect.
A third possible resistance mechanism involving the dppD and dppB genes, is the synthesis and export of autolysins, if AlpP were to signal an autolytic response that in turn was mediated by a peptide autolysin (like bacitracin). This proposed mechanism would be consistent with the sensitivity of the dppD (biosynthesis) mutant and dppB (ABC-transporter) mutant to AlpP exposure, due to altered autolysin biosynthesis and export. It is possible that AlpP may function as a signal molecule for P. tunicata to produce autolysins. Very small amounts of AlpP protein can result in autoinhibition - a feature that is consistent with the action of a signal molecule. It was found that the Minimum Inhibitory Concentration (MIC) of AlpP against P. tunicata is only 4 pg/ml (James et al., 1996), which considering that the molecular weight of AlpP is 190kDa, corresponds to approximately 13 molecules per pm .
3.4.3.2. Autolysis upon AlpP exposure may be mediated by a two-component hybrid sensor regulator
If AlpP act as a signal molecule for autolysin production, another possible explanation arises from the data obtained from the AlpP-resistant mutants. The RM1 AlpP-resistant mutant had an insertion in a gene similar to a putative two-component hybrid sensor and regulator in Caulobacter crescentus. The RM3 and RM4 AlpP-resistant mutants were mutated in the same ORF, which had high similarity to the syringomycin synthetase gene, syrE, in the plant pathogen Pseudomonas syringae. Syringomycin is a cyclic lipodepsinonapeptide phytotoxin, which forms pores in the plasma membrane of the
117 target cells (Bender et al., 1999). In P. syringae, production of syringomycin toxin is believed to be mediated by phenolic glycosides (commonly produced by plants), which act via a two-component sensor regulator, SyrP, resulting in its transport out of the cell through an ABC transporter, SyrD. The toxin then causes plant necrosis via its pore forming activity (Bender et al., 1999). The P. syringae sensor is associated with the syringomycin synthetase operon, indicating that a similar association may exist between the hybrid sensor and syringomycin synthetase-like genes in P. tunicata, which results in increased resistance to AlpP. This raises the possibility that AlpP may function as a signal for an autolysin-mediated response, where AlpP is perceived by a two-component hybrid sensor that induces a syringomycin-like gene to express an autolysin. The dppD (biosynthesis) gene may also contribute to the biosynthesis of autolysis as suggested in section 3.4.3.1. In this case, autolysis would occur after the syringomycin-like autolysin leaves the cell, via the ABC transporter as suggested above (Figure 3-5). Such a mechanism would explain the increased resistance of cells mutated in either the sensor or autolysin-synthetase gene, as the mutants would be unable to sense AlpP or synthesize autolysin respectively.
3.4.3.3. Expression of extracellular structures may be linked with resistance to AlpP activity
Other possible resistance mechanisms to AlpP are proposed based on the analysis of the SM5 and SM2 mutants. It is suggested that the resistance to AlpP is linked to the production of extracellular structures. The SM5 AlpP-sensitive mutant had an insertion in a gene with high similarity to the mshJ gene of Vibrio cholerae. In V. cholerae, mshJ is a member of the MSHA secretory operon in the biogenesis of mannose-sensitive hemagglutinin (MSHA) type 4 pilus (Marsh et al., 1996). The MshJ of P. tunicata is an outer membrane protein similar to the type II secretory proteins. The type II secretory proteins are known to export toxins, proteases and cell metabolites (Thangassi and Hultgren, 2000). In this case, it is possible that MshJ exports AlpP out of the cell in order to initiate resistance. However, substrates for the type II secretory pathway require a cleavable N-terminal signal sequence for Sec-dependent translocation across the inner membrane (Thangassi and Hultgren, 2000), which the P. tunicata alpP gene does not encode. Thus, a similar transporter in P. tunicata would not be involved in exporting AlpP. It was shown that the mshJ gene is required for the biogenesis of pili in
118 P. tunicata (Chapter 4). Considering that MshJ is a pili biogenesis protein in P. tunicata, one possible hypothesis is that the expression of pili could potentially provide steric hindrance to incoming AlpP protein, and in this way provide some protection to P. tunicata cells. A detailed discussion of this mutant is presented in Chapter 4.
The sensitive P. tunicata mutant (SM2) was disrupted in a gene similar to the chitinase gene chiA, in Vibrio cholerae. Although it is difficult to make a connection between the function of this enzyme and resistance to the AlpP protein, it is possible that this enzyme is associated with putative cellulosome structures that have been observed on the P. tunicata cell surface (Chapter 5). Cellulosomes are large extracellular structures known to anchor enzymes such as cellulases and chitinases to the cell surface so that they function together to break down complex substrates (Shoham et al., 1999). If cellulosome-associated enzymes are involved in the degradation of AlpP then it is possible that the disruption of one of these enzymes could affect the function of the cellulosome as a whole.
3.4.4. P. tunicata demonstrates mechanisms to prevent self killing
It was observed that P. tunicata colonies of different ages exhibit different levels of tolerance to AlpP exposure (James et al., 1996), suggesting that control mechanisms are present in the bacterium. Although the selected AlpP-sensitive mutants did not reveal any gene encoding an antidote in P. tunicata, the identification of the hypothetical antidote was based on the hypothesis that it is expressed simultaneously with AlpP, as the E. coli toxin-antidote system involves the production of a stable protein toxin and an unstable protein antitoxin (Engelberg-Kulka and Glaser, 1999).
The cell-free concentrated supernatant was shown to have inhibitory and protective activities. This was shown by the growth pattern of P. tunicata cells when added with serially diluted concentrated supernatant. The protective activity was demonstrated when there was an unexpected resistance observed on P. tunicata cells when exposed to concentrated supernatant at 0 to 2-fold dilution. This finding suggests that an antidote may be present in the concentrated supernatant that counteracts the toxic effect of AlpP.
119 The toxicity appeared to increase as the dilution factor was increased as shown between 4 and 8-fold dilution suggesting that a protective or “antidote” factor was being diluted out of its active concentration range earlier than the AlpP protein. Although the ratio of AlpP to any protective substance must be the same at all dilutions, it is possible that each has an effective threshold concentration that is reached at different stages - especially if the “antidote” is less stable than AlpP which is typical of bacterial toxin- antidote systems (Gerdes, 2000). To investigate this h>pothesis further, a fractionation of the cell-free supernatant was carried out using ion-exchange chromatography to separate the hypothetical antidote factor from the autolytic protein, AlpP. It was demonstrated that anion exchange chromatography can be used to separate the toxic (AlpP-containing) fraction of cell-free supernatant from another fraction that appears to have antidote-like characteristics. This “antidote” fraction reduced the toxicity of the AlpP fraction when the two were mixed. As a control, equal volumes of the AlpP fraction and a buffer did not reduce the toxicity of the AlpP protein to the same extent.
3.4.5. Ecological roles of P. tunicata autolytic protein, AlpP
Lewis (2000) suggested that “altruistic” or programmed cell death (PCD) occurs in a fraction of the stationary phase population of many bacterial species, to release nutrients for cells in the vicinity as a response to nutrient limitation. Autolysis in response to starvation has been linked with the toxin-antidote system in bacteria. It was shown that the mazEF genes are the best-characterized toxin-antidote system in E. coli (Aizenman et al., 1996), forms an operon upstream of relA gene, which encodes an ATP:GTP 3’- pyrophosphotransferase, for the synthesis of ppGpp (Aizenman et al., 1996), which is activated during amino acid starvation. It was also demonstrated that mazEF mediated cell death is induced by overproducing ppGpp, indicating that mazEF elicits PCD during nutrient deprivation (Aizenman et al., 1996). The expression of AlpP during stationary phase is proposed to represent an example of bacterial PCD, as the bacterial cells employed this mechanism in response to poor nutrient conditions. Moreover, as suggested above, P. tunicata appears to have “programmed death signals” through the activity of its putative sensor-regulator system, which mediates autolysis in response to AlpP production in stationary phase of growth. The expression of AlpP also denotes an important ecological role of the product. Firstly, the expression of AlpP during
120 stationary phase of growth may be a nutrient deprivation response, suggesting that P. tunicata possess starvation regulatory mechanisms. Secondly, expression of AlpP displays control mechanisms for population density and is an altruistic process. This offers an explanation for how P. tunicata can produce am autoinhibitory product without killing itself, and possibly reflects a balance between the levels of alpP and “antidote” expression within a starved P. tunicata population. A control mechanism for population density is significant in the biofilm formation of P. tunicata as this would prevent bacterial overgrowth on the surfaces of host organisms. Equally, the altruistic suicide of a portion of the bacterial population would result in the release of nutrients from the lysed cells, hence maintaining the bacterial community and aiding dispersal of the species. Thirdly, AlpP expression may allow P. tunicata colonies to behave as “individuals” able to defend themselves from invading bacteria and from other P. tunicata cells.
In summary, this chapter discusses the mode of action of AlpP and enhances our understanding of its ecological role. The antibacterial properties of AlpP coupled with the known predatory lifestyle of P. tunicata indicate that this protein could clearly aid in competitive interactions with other bacteria, either by directly attacking their membranes or by inducing autolytic pathways. With regards to auto-toxicity, there are clearly control mechanisms in place, possibly mediated via an “antidote” molecule, which are able to prevent AlpP activity during growth and colony-formation. Finally, transposon mutant analysis has begun to address possible mechanisms for the regulation and transport of AlpP, along with raising the possibility that its activity is mediated via a sensor and regulatory systems controlling the expression of other autolytic products. Many of these findings will provide a starting point for future research on P. tunicata and its ecology. In particular, the identification of the mshJ gene disrupted in the SM5 mutant reveals mechanisms of colonization and offers an explanation for the ecological distribution of P. tunicata in the marine environment. Suggested colonization mechanisms are presented in the next chapter.
121 AutoJytic protein, AIpP A) Syringomycin-like peptide autolysin lysis
External environment * Two-component f ABC 'N hybrid sensor y transporter y Cytoplasm A To syringomydn-like peptide autolysin Two-component hybrid regulator 15
syrE ? hypothetical ? dppD dppC dppB
ExtraceHular structures (pili and cellulosome) vtfiich may provide steric hindrance to AIpP chiA mshi V
B)
Autoiytic protein, AIpP
Figure 3-5. Hypothetical model for the mode of action of AIpP in P. tunicata. A) The proposed model for sensitivity, where in the AIpP is a signal molecule, which mediates the production of autolysin peptide, causing autolysis. B) The proposed model for resistance, where in AIpP is prevented from coming in contact with the cell by steric hindrance brought about by the expression of extracellular structures. ? = Unknown gene or protein T = Putative transferases. See section 3.4 for a detailed discussion.
122 4. Identification and characterization of a putative mannose-sensitive hemagglutinin (MSHA) pilus biogenesis gene cluster and the role of MSHA pilus in the colonization of living marine surfaces
4.1. Introduction
Marine surfaces are colonized by a diversity of microorganisms and sessile marine organisms, collectively known as biofouling communities. The biofouling process is initiated by the attachment of bacteria to a surface followed by the settlement and adherence of diatoms, free swimming algal spores and invertebrate larvae (Bryers and Characklis, 1982). Some sessile higher organisms employ chemical defenses against biofouling through the production of secondary metabolites which inhibit the development and formation of a biofouling community (Harrison, 1992; Mary et al., 1993; Maximilien et al., 1998). For example, furanones produced by the red alga Delisea pulchra have been reported to inhibit the settlement of common fouling organisms (de Nys et al., 1994). For marine organisms without intrinsic defense mechanisms, it has been proposed that protection against fouling is maintained by the secondary metabolites produced by surface associated bacteria (Egan et al., 2001a; Holmstrom et al., 1992; 1996; 1998; Holmstrom and Kjelleberg, 1999; James et al., 1996).
Surfaces of higher marine organisms have been reported to harbor Pseudoalteromonas species and many of these species have been found to produce bioactive compounds against different classes of fouling organisms (Holmstrom and Kjelleberg, 1999). A well-studied surface associated bacterium is P. tunicata. This green pigmented bacterium was first isolated from the surface of a tunicate, Ciona intestinalis, in Sweden (Holmstrom et al., 1996) and later from a green alga, Ulva lactuca, in Australian waters (Egan et al., 2001a). Recently, it has been reported that the same species was isolated from Aarhus Bay, Denmark on the surfaces of U. lactuca, C. intestinalis and Ulvaria fusca (Shovhus et al., 2004), indicating that P. tunicata is widely distributed in different
123 marine waters. The antifouling activity characteristic of thiis bacterium is due to the production of a number of different extracellular bioactive compounds, each of which has a specific inhibitory activity against target organisms siuch as algal spores, fungi, invertebrate larvae or bacteria (Egan et al., 2001a; 2001b;, Holmstrom et al., 1992; James et al., 1996). While several studies have addressed th The successful colonization of bacteria on surfaces is often mediated by cell surface appendages such as pili and flagella. For example, the attachment of E. coli to abiotic surfaces is promoted by the presence of both type I pili and flagella (Pratt and Kolter, 1998). In Pseudomonas aeruginosa, flagella are believed to be important for initial attachment to a surface, while type 4 pili promote the formation of microcolonies on the surface (O’Toole and Kolter, 1998). It has also been reported that type 4 pili mediate attachment of pathogenic bacteria such as Neisseria gonorrhoeae (Morand et al., 2001) and P. aeruginosa (Zolfaghar et al., 2003) to host epithelial cells. The adherence of Vibrio cholerae to environmental surfaces is directly associated with the presence of the mannose-sensitive hemagglutinin (MSHA) pilus, which belongs to a family of type 4 pili (Marsh and Taylor, 1999). The MSHA pilus has been demonstrated to play a role in the colonization and subsequent biofilm formation of V. cholerae on abiotic as well as biotic surfaces (Watnick et al., 1999). The attachment of bacteria to abiotic surfaces is accelerated by the presence of MSHA pili (Watnick and Kolter, 1999). Additionally, the MSHA pilus was recently found to promote attachment of V. cholerae to zooplankton with a chitinous exoskeleton as well as mucilage-sheathed phytoplankton (Chiavelli et al., 2001). This chapter describes the non-piliated mutant of P. tunicata that carries a transposon insertion in an open reading frame termed mshJ which shows homology to the V. cholerae mshJ, an MSHA pilus biogenesis protein encoding gene. DNA sequencing of the region flanking the transposon revealed the presence of a gene locus with six potential open reading frames that are homologous to the secretory operon for the MSHA pilus biogenesis gene locus. This chapter also reports the detailed genetic 124 characterization of these ORFs and suggests that this gene locus is involved in the assembly and transport of MSHA pilus in P. tunicata. The identification and characterization of the MSHA pilus produced by P. tunicata is also described, and its role for attachment of the bacterium to abiotic as well as biotic surfaces, specifically to the surface of green alga U. lactuca. Finally, it is proposed that P. tunicata demonstrates surface sensing mechanisms as revealed by increasing pilus production in the presence of cellulose, one of the major surface polymers of U. lactuca. 4.2. Materials and Methods 4.2.1. Bacterial strains, plasmids and culture conditions P. tunicata wild type strain was grown on VNSS agar plates as described in section 2.2.1. The P. tunicata generated transposon mutant, SM5, was grown on VNSS agar plates containing 100 pg/ml streptomycin (Sm) and 85 pg/ml kanamycin (Km) (section 2.2.4) The P. tunicata wild type strain and SM5 mutant cells were tagged with green fluorescent protein (GFP) by trans-conjugation using the GFP expression plasmid, pCJSlOG. The SM5 mutant was also tagged with red fluorescent protein (RFP) using the RFP expression plasmid, pCJSlOR. The labeled transconjugants were grown on VNSS agar plates containing 15 pg/ml of chloramphenicol (Ch) and 100 pg/ml streptomycin (Sm). Prior to the attachment assay, bacterial isolates were grown in Marine Minimal Media (MMM), each medium containing 0.05 % (w/v) glucose, 0.25 % microcrystalline cellulose (Avicel, Fluka) or 0.1 % cellobiose (Sigma) as the sole carbon source. 4.2.2. Panhandle PCR, DNA sequencing and sequence analysis To obtain sequence information from the genes disrupted by the mini-TnlO transposon in the SM5 mutant, panhandle polymerase chain reaction (panhandle-PCR) was 125 employed using adaptor specific primer, API, 5’-GGA TCC TAA TAC GAC TCA CTA TAG GGC-3’ and transposon specific primers: TnlOC, 5’-GCT GAC TTG ACG GGA CGG CG3’ and TnlOD, 5’-CCT CGA GCA AGA CGT TTC CCG-3’) (section 2.2.6.2). Primer walking strategy using the oligonucleotides presented in Table 4-1 was employed to continue sequencing further along the DNA regions flanking the transposon in P. tunicata SM5 mutant. 4.2.3. Transmission and scanning electron microscopy studies The ultrastructures of P. tunicata wild type and the SM5 mutant were examined using transmission electron microscopy (TEM). The bacterial cells were grown on VNSS plates for 24 h, harvested and resuspended in phosphate buffer solution (PBS), pH 7.4 (8.00 g NaCl, 0.20 g KC1, 1.44 g Na2HP04,0.24 g KH2P04 per 1 liter) to a concentration of about 106 cells/ml. Alternatively, bacterial cells were grown in static conditions in MMM medium containing glucose, cellobiose or cellulose as the sole carbon source. The bacterial cells grown in liquid medium were centrifuged (12, 000 x g for 5 min), washed twice and resuspended in PBS. Carbon-coated formvar copper grid was placed on a drop of the cell suspension for 5 min and negatively stained with 2 % phosphotungstic acid for 30 sec. The cells were examined using a Hitachi H7000 transmission electron microscope. Scanning electron microscopy (SEM) studies were performed on the bacterial samples grown in liquid medium prepared as described above. The bacterial cells were placed on poly-L-lysine coated cover slips and fixed in 2.5 % glutaraldehyde for 30 min. The cells were dehydrated for 10 min in different ethanol concentrations. The samples were critical point dried in liquid C02 and coated with chromium. Samples were examined using a Hitachi H900 scanning electron microscope. 126 Table 4-1. Oligonucleotides used in the primer walking strategy to sequence DN; regions flanking the inserted transposon in the SM5 mutant Oligonucleotide Sequence (5’-3’) Target region SM5F AGT C A AGGCTTT GT GC CGC 1730 -1748 SM5TnlOC-D6 CACAGCAGCCATTTTGTAAG 1749-1768 SM5TnlOC-D6A GGCTTTT ATTTGCTT AATCCGC 1523-1544 SM5TnlOC-D6B TTTGCAATTCAGTTAACGTCG 1027-1048 SM5TnlOC-D6C AAATAGCTCGTAATGGCGGC 652-672 SM5TnlOD-D7 AT GGCT ATT ACGCTCG AAGGC 1889-1910 SM5TnlOD-D7A GTCAGTGATCCGAAAGCAGGG 2195-2216 SM5TnlOK GGCT GT G AA ATT ACGC A A AAA 2419-2445 -GACCC SM5TnlOJ CCTTCGAGCGTAATAGCCATG- 1885-1909 CCG SM5TnlOJl GGCAAAAAAATGCTCAATCCT 1557-1586 -TAATCGCG SM5TnlOKl CCCGCATGTATCCGCTAGATG- 2756-2794 TATTTCG SM5TnlOK2 GGTTCACAAATGAATGGTGCC 3019-3040 SM5TnlOK3 CATGCCTTACCAAGTGAAATT- 3163-3189 GCGGC SM5TnlOKC CCTT ACC AAGT G A AATT GCG 3167-3187 SM5 TnlOK4 CAGGTAATGCGACAACCACGA 3575-3600 CACC SM5 TnlOK5 AGC A ACGC AAAGT GATT ACC 3713-3733 SM5 TnlOK7 GGGC ATTT ATTT AAAAGT GT G 3901-3930 CGAAAGCG SM5 TnlOK8 T AGCGGGT GAT GT GGT GGT GG 4246-4267 127 4.2.4. Hemagglutination assay The hemagglutination assay was performed as described previously (Gardel and Mekalanos, 1996). The bacterial strains were grown in VNSS medium at 28°C in static conditions and assayed during both the exponential and stationary phase of growth. Briefly, the cells were washed twice and resuspended in Krebs-Ringer solution (KRT) (Appendix I). An initial concentration of 1010 cells/ml was serially diluted in a 96-well microtitre plate with each well containing 100 pi of 3 % (vol/vol) washed horse erythrocytes suspension. The mixture was incubated at room temperature for 30 min and scored for hemagglutination. The titer was recorded as the reciprocal of the dilution factor of the sample still showing hemagglutination. The V. cholerae Ml 615 strain was used as a positive control. To determine if the pili were sensitive to the presence of mannose and fucose during the hemagglutination assay, 50 pi of 1 % a-methyl D- mannoside, a non-metabolized derivative of mannose or L-fucose solution was added to the hemagglutination mixture. The hemagglutination assay for each strain was carried out in triplicate. 4.2.5. Attachment assays The attachment of the P. tunicata wild type and the SM5 mutant strains to abiotic surfaces was tested by modifying an attachment assay described previously (Taylor et al., 2002). The bacterial isolates were grown in static conditions at 28°C on VNSS medium and harvested after 24 h. The cell suspension was centrifuged (6, 000 x g for 5 min), washed twice and resuspended in 10 ml of PBS (pH 7.4) to an optical density (600nm) of 0.6-0.7. One ml of cell suspension was added to each well of a 24-well polystyrene microtitre plate. The plate was shaken slowly for 1 h at room temperature. The wells were washed twice using sterile distilled water and air-dried for 45 min. The attached cells were fixed at 80°C for 30 min and stained with 0.1 % crystal violet for 45 min. The cells were destained with 95 % ethanol and quantified by measuring the optical density at 590 nm. The attachment assay for each of the strain was carried out in triplicate. 128 Attachment of GFP tagged P. tunicata wild type and RJFP tagged SM5 mutant cells to microcrystalline cellulose were performed by modifying the method of Bayer et al (1983). The glucose and cellobiose grown cells were harvested after 24 h and the cells grown in cellulose were harvested after 48 h of growing in static conditions. The cells were then washed twice with PBS and resuspended in 10 ml of PBS solution. The assay mixture consisted of 1 ml of cells (~107 cells/ml) (pregrown in glucose, cellobiose or microcrystalline cellulose), 1 ml of 20 % microcrystalline cellulose in PBS, and 1 ml of PBS. The mixture was shaken slowly for 1 h and incubated statically for 1 h at room temperature. The optical density of the suspension was measured at 400 nm and compared to a control (identical cell suspension in PBS). To test the effect of mannose on the attachment of the cells to cellulose, a-methyl D-mannoside at a final concentration of 100 mM was added to the attachment assay mixture. The attachment assay for each strain was carried out in triplicate. The samples were mounted on a glass microscopy slide and transmission fluorescent images were captured using an Olympus LSMGB200 confocal laser scanning microscope (CLSM). 4.2.6. Preparation of axenic thallus of the green alga Ulva lactuca To investigate the attachment of P. tunicata wild type and SM5 mutant to living marine surfaces, an attachment assay with U. lactuca was carried out, in collaboration with Andre Scheffel, Centre for Marine Biofouling and Bio-Innovation at the University of New South Wales. Plants of the green alga U. lactuca were collected from rocks at Clovelly Bay, Sydney, Australia. The collected plants of U. lactuca were rinsed with 50 ml autoclaved seawater and thallus discs of around 0.6 cm diameter were excised from the lower part of U. lactuca and used in the experiments. To obtain axenic plant material of U. lactuca, a sterilization protocol was developed (Scheffel, 2003). Briefly, the surfaces of the collected plants were swabbed with sterile cotton tips and exposed to 0.3 % (v/v) NaOCl for 5 min. The recovered plant pieces were incubated in an antibiotic mixture (ampicillin 300 mg/1, polymyxin 30 mg/1 and gentamycin 60 mg/ml) for 24 h followed by 1 h recovery in 20 ml sterile seawater to dilute the remaining antibiotic and NaOCl residues. 129 4.2.7. Attachment assay with axenic Ulva lactuca GFP-tagged P. tunicata wild type and SM5 mutant were grown in VNSS medium containing chloramphenicol (15 mg/L) for 24 h. After 24 h the cell suspensions were centrifuged at 6, 000 x g for 7 min. The bacterial cells were rinsed twice with 10 ml PBS and resuspended in PBS to an optical density of 0.35 - 0.45 at 610 nm. The assay mixture consisted of 1 ml of washed cells, 1 axenic thallus disc of U. lactuca and 1 ml of PBS. The mixture was incubated for a period of 2 h with slow shaking and for 1 h without shaking at room temperature. The U. lactuca pieces were rinsed twice with 10 ml PBS to remove unattached cells. For each sample, ten images were manually analysed by counting the bacteria attached to the surface using CLSM. The attachment assay was repeated four times with triplicate for each bacterial strain. 4.3. Results 4.3.1. DNA sequence analysis of the regions flanking the TnlO insert in the P. tunicata SM5 mutant DNA sequencing of the panhandle PCR products (Figure 4-1) yielded a 4,291 bp nucleotide sequence with six potential open reading frames (ORF) (Figure 4-2). No putative promoter region or terminator sequences were identified indicating that the six ORFs belong to a single operon. The deduced amino acid sequence of the six ORFs showed homology to the secretory operon for the MSHA pilus biogenesis gene locus of various bacteria. A summary of the genetic characteristics of the MSHA gene cluster of P. tunicata is presented in Table 4-2. The DNA sequence generated has been entered in the GenBank databases under accession number AY695819. 130 MW 1 2 3 4 5 6 >3130 bp 9461 bp 6557 bp 4361 bp 2322 bp 2027 bp 564 bp Figure 4-1. Panhandle PCR products of the digested and ligated genomic DNA of P. tunicata SM5 mutant. MW = molecular weight marker, lane 1 = PvuW digest and SM5Tnl0J, lane 2 = Rsal digest and SM5Tnl0Jl, lane 3 PvuW and SM5TnlOK2, lane 4 = PvuW digest and SM5Tnl0K3, lane 5 = PvuW and SM5TnlOK4, lane 6 = Dra\ and SM5Tnl0K5. 131 1 cacttctgtttgtatggagaaaaagcaaaaaaactggttgatagtggagc 50 MEKKQKNWLI VEH 51 actattcagcgcaaactcacatcagatttagtgtaacatcggcaatagtg 100 YSAQTHIRFSVTSAIV 101 acttgtctaaaaaaaatagcagccccttctgctttagtatccttaatatt 150 TCLKKIAAPSALVSLIL 151 acctagccaaagttaccagttggtacaaattgataagcctaatttatctc 200 PSQSYQLVQIDKPNLSP 201 cagaagaaattcagcagtcattaccatggacagtaaaagacttagtaaca 250 EEIQQSLPWTVKDLVT 251 attaattcagcagatattattgccgattatatcgatcaccctgttaagca 300 INSADI IADYIDHPVKQ 301 agggaatcaagataaaataagtgtttttgtgaccaatcgctcgttcatta 350 GNQDKISVFVTNRSFIT 351 cccctattattgacgccataaaaaaagcttcagctaaattagagctatta 400 PI IDAIKKASAKLELL 401 agttgcgaagaaataatgctcactacattagttggcaaacacagcgctgc 450 SCEEIMLTTLVGKHSAA 451 aaatttgattatttcacaagagctcggccaagagcctagcttattaattg 500 NLIISQELGQEPSLLIV 501 tgcgcgacggtgcagtactgtttgcaaggcgtttacgtggttttagtcga 550 RDGAVLFARRLRGFSR 551 atttcttctatgactttagaagatctccaacatggtttattagactcgtt 600 I SSMTLEDLQHGLLDSL 601 aagcttagaaattcaacgttcaatcgatttttttgaaagccaactaaaac 650 SLEIQRSIDFFESQLKQ 651 agccgccattacgagctattttattgagtttaccaagcccttatttggtg 700 PPLRAILLSLPSPYLV 701 gagctcgccacagagctagggcaacattttcctgtaaaagttgaaaaatt 750 ELATELGQHFPVKVEKF 751 cactactgcactagctgcttgtgaagggcaggatcctaattatcatttag 800 TTALAACEGQDPNYHLA 801 cgattgctgccgcgatggagttaattggagttgacgatgaaaactaggat 850 IAAAMELIGVDDEN* M K T R I 851 taacttatatttagctgaacttcgtccaaaaaaagatccgttgagtttaa 900 NLYLAELRPKKDPLSLN 901 atcgagtaacggtatttttacttactcttttgttggttatggttttgctt 950 RVTVFLLTLLLVMVLL 132 951 tcaagtgtgcttaaaactaaagtggtggcgcagcaaaaaatgctacaaaa 1000 S SVLKTKVVAQQKMLQN 1001 ccagcagcaattagtagcacagcaagcgacgttaactgaattgcaaacag 1050 QQQLVAQQATLTELQTA 1051 ccttagcgagaaaacaagataagaatgtactgtcgcaacaactcgctcag 1100 LARKQDKNVLSQQLAQ 1101 ataaaagctgaaattcagcacaagcaattagttatggagtttatttcaac 1150 IKAEIQHKQLVM.EFIST 1151 ccatgagcaaactattttatatgctgaggtaatggaagatttagcgcggc 1200 HEQTILYAEVMEDLARL 1201 ttcacgatcctcttatttggttaactggatttaaatttaataatcagcat 1250 HDPLIWLTGFKFNNQH 1251 attattttagaagggcaaacagacgagccagcacaaattccccattggtt 1300 I ILEGQTDEPAQI PHWL 1301 agatgggttaaaagcatcaagttatttttcaggtaagcttctatccgaaa 1350 DGLKASSYFSGKLLSEM 1351 tgaagtttgaacaacgtgatggcgttacctattttcqagttgctagcgag 1400 KFEQRDGVTYFRVASE 1401 ccagttatggaggcaaaatgaagcagcaatggcaagtttatagagagaag 1450 P V M E A K * MKQQWQVYREK 1451 tttgcagccttacaaatgagagaaaaatacctcatttttggtgttggttt 1500 FAALQMREKYLI F G V G L 1501 gtttttaatttgttatctttttggcttttatttgcttaatccgctttatt 1550 FLICYLFGFYLLNPLYL 1551 taaaatggcaaaaaaatgctcaatccttaatcgcgatagaaaaaaaactt 1600 KWQKNAQSLIAI EKKL 1601 gttgcgaatacagctcaaataaccttatttagtgacgcattaactcgaga 1650 VANTAQITLFSDALTRD 1651 ttatacccaagaacttagaagtgaaattgcgacagctgagctagctttaa 1700 YTQELRSEIATAELALK 1701 agcaagttgatgagaaattaaatcagtttagtcaaggctttgtgccgcct 1750 QVDEKLNQFSQGFVPP I 1751 tacaaaatggctgctgtgcttaaaaaagtattgcttgataatcgcaagtt 1800 YKMAAVLKKVLLDNRKL 1801 gactttaaaagcgttcaaattagtcggcgtagaaccaatcattatcggtt 1850 TLKAFKLVGVEPI I IGL 1851 tagaacaacagcagaaagttgcattttatgagcacggcatggctattacg 1900 EQQQKVAFYEHGMAI T 133 1901 ctcgaaggcgagtattttgatttattaaagtatttaaatgcggtgcaaaa 1950 LEGEYFDLLKYLNAVQN 1951 tttagaagaaaaactgtttattaaagagtatagctatcaggtgattcaat 2000 LEEKLFIKEYSYQVIQY 2001 atccgatagcgcaattaagtttggttataacaacggtgagtgcagatgaa 2050 PIAQLSLVITTVSADE M K 2051 gagtttttatctatttagtatttggctgttgttgattttacctgctttat 2100 E F L S I * SFYLFSIWLLLILPALS 2101 cgtttgggaaaagctatcaagatccgaccaaacctaatattaagcatcag 2150 FGKSYQDPTKPNIKHQ 2151 aaaaatgtatccgttgccgggtctttaggtacattgcgtttagagtcagt 2200 KNVSVAGSLGTLRLESV 2201 gatccgaaagcagggaaaagtcaaagtggtcatctcaggcaaaatatatg 2250 I RKQGKVKVVISGKI YA 2251 ctaaaggtgagcaagtgggcgaatacgttcttagtaaaattaacgcagat 2300 KGEQVGEYVLSKINAD 2301 actgtttatttaactcgtggtagcgaacaactaaaattagaattatatca 2350 TVYLTRGSEQLKLELYH 2351 tcatgaaattaaacgttaaaaaagcatttaaattagctatatttccttat 2400 H E I K R * MKLNVKKAFKLAI FPY 2401 atcgcattgggtttagtgggctgtgaaattacgcaaaaagacccagatat 2450 IALGLVGCEITQKDPDI 2451 ccgccctcatattgcagatgagctgacatcttctgtggcgcaaaatgccc 2500 RPHIADELTSSVAQNAQ 2501 aacaaaaaaactcagctaagatgccagctgaattaaccgaagaattactc 2550 QKNSAKMPAELTEELL 2551 tcatctgtaaagtctaatttattcgcgcctaatgaacttgatatgaagcg 2600 SSVKSNLFAPNELDMKR 2601 ttttgaaattgctgccaatgaagtcgacgttagaagcttttttgctagct 2650 FEIAANEVDVRSFFASL 2651 tagttgatggcacaccttacagtgttgcgcttcatcctgaagtagaagga 2700 VDGTPYSVALHPEVEG 2701 aaaataagcctcaatctgaaagatgttactcttgatgaagtgattaaaat 2750 KISLNLKDVTLDEVIKI 2751 cgttacccgcatgtatccgctagatgtatttcgtgatggtaaagtggtgc 2800 VTRMYPLDVFRDGKVVQ 2801 aagtattgcccgcgagaatgcgaacagagtctatcgcagttaattattta 2850 VLPARMRTESIAVNYL 2851 atgatgaagcgcaccggaatttctaccgtcagtgtggttgctggtggcgt 2900 MMKRTGI STVSVVAGGV 134 2901 gagtcagtttggtcaaagtggcagcggaggctcagcaaattctggcgcac 2950 SQFGQSGSGGSANSGAR 2951 gaggtaataaccaaaattctggtgcaaacggtaatcagagttcaggtaat 3000 GNNQNSGANGNQSSGN 3001 ggaaatggcatgagtaacggttcacaaatgaatggtgccagtattcagac 3050 GNGMSNGSQMNGASIQT 3051 atcatcagaatttgatttttggactgatttaaaaaaagcgcttgaatcat 3100 SSEFDFWTDLKKALESL 3101 tagttggcgtagataaaggccgctacatcattgttagcccacaagcaagc 3150 VGVDKGRYI IVS PQAS 3151 ttggttactgtacatgccttaccaagtgaaattgcggcaatgaaagagtt 3200 LVTVHALPSEIAAMKEF 3201 tttacgtttaggtgaagaaagtctgcagcgccaagttattttagaagcca 3250 LRLGEESLQRQVILEAK 3251 aaattattgaagtaaccttgaaggacgattatcagcaaggtgttaattgg 3300 I I EVT LKDDY QQGVNw" 3301 aaagaggtgcttggtcatattggtagtacagatttaacgtttgccaccac 3350 KEVLGHIGSTDLTFATT 3351 agcttcggggcaaatcggcaatactattaccgcaggtattggtggtgtct 3400 ASGQIGNTITAGIGGVS 3401 ctagtttggtctttaaaaatgctgatttttctggagttgtcagcttgctg 3450 SLVFKNADFSGVVSLL 3451 tcgactcaaggtgatgtgcaaatgctgtcaaatcctcgagttaccgcaac 3500 STQGDVQMLSNPRVTAT 3501 aaacaaccaaaaagcggtcataaaagtaggacaagatgagtactttgtga 3550 NNQKAVIKVGQDEYFVT 3551 ctgaggtttctagcaccacagtgacaggtaatgcgacaaccacgacacct 3600 EVSSTTVTGNATTTTP 3601 gaaatatcactaacgccttttttctcaggtattgcactggatgtcacacc 3650 EISLTPFFSGIALDVTP 3651 acaaattgataaatatggctcggtgattttacatgtgcacccatctgtca 3700 Q I DKYGSVILHVHPSVT 3701 ctgaaacagcagagcaacgcaaagtgattacccttaataatgaagagttt 3750 ETAEQRKVITLNNEEF 3751 gtactgcctttggcgcaaagtaacattcgagagtctgatactgttattcg 3800 VLPLAQSNIRESDTVIR 3801 agctggcagcggtgaaatcgtagtgataggtggtttaatgcaaaccgtga 3850 AGSGEIVVIGGLMQTVT 3851 ctaccgatgaggaatcaaaaaccccactcttaggtgatatgccggtactt 3900 TDEESKTPLLGDMPVL 135 3901 gggcatttatttaaaagtgtgcgaaagcgtcaggataaaaaagagctcat 3950 GHLFKSVRKRQDKKELI 3951 tattttaatcaaaccaacagtcgtgatggctgatacttggcaacaacagc 4000 I LIKPTVVMADTWQQQQ L A T T A 4001 aacagcgctcaatgcagctgttaaaaagttggtattcaaactaattatgt 4050 QRSMQLLKSWYSN* TALNAAVKKLVFKL I MY 4051 atttaagctttttttcactacaggaaatgcctttttcgctgacgccgaat 4100 LSFFSLQEMPFSLTPN 4101 acgcaatttttttgcgctctacagccgcataatgaagcgatgcaagtgct 4150 TQFFCALQPHNEAMQVL 4151 gttaactgcattaaaaatgggggaaggttttattaaggttacaggtgagg 4200 LTALKMGEGFIKVTGEV 4201 ttggtacaggcaaaaccttgttatgccgcaagctgcttaatagtttagcg 4250 GTGKTLLCRKLLN SLA 4251 ggtgatgtggtggtggcgtatttacctaatccttatttacc 4291 G DVVVAYLPNPYL Figure 4-2. Nucleotide sequence of the genomic-DNA regions flanking the transposon in the SM5 mutant. The nucleotide sequence with the translated amino acid sequence in one-letter code is shown. The arrow (v) indicates where the mini-TnlO transposon insertion occurred. The six open reading frames arc highlighted as follows: ORF1 (mshl\) = dark red, ORF2 (mshI2) = blue, ORF3 (mskJ) = pink, ORF4 (mshK) = bright green, ORF5 (mshL) = orange and ORF6 (mshM) = violet. 136 Table 4-2. Characteristics, location, size and predicted location of the gene products of the ORFs. _G o o O PQ '55 .§ ’ 00 P 4 53 G CD a) G < CO ^ w 22 _o 4 ? 'o * •4 & CD P CO ‘ ,G go tW x> ^3 O o ^ a. O o 00 P, G -s: -a s •a ^ w S O'* C pH ro so U ’ ^ G- CO o o r- 'j * H. 5 co O 5 ? s w G o g "5; ^ & PS U « o as o r- r- -$2 5 b S' PP O SO O Os Os SO PQ < so s 5 o a a a co -a co rv oj o mshK, V. vulnificus (BAC95712.1) cr> (N rnshK, V. parahaemolyticus (BAC60968.1) The 0RF1 termed mshl 1 is an 833 bp open reading frame encoding a protein with 277 amino acids residues. The ATG start site is at base position 14, which is preceded by a putative ribosomal binding site (RBS) 5’CTGTT3\ When compared with the GenBank databases, the gene product of ORF1 is similar to Mshl, a biogenesis protein for MSHA found in the Vibrio species (Table 4-2). The similarity of the gene products reflects the regions between 51 to 278 amino acid positions of Mshl of V. cholerae 01 biovar El Tor strain; between 53 to 282 amino acid positions of Mshl of V. parahaemolyticus\ and between 69 to 265 amino acid positions of Mshl of V vulnificus. The protein product contains a conserved domain (over 250 amino acid residues) of a type IV pilus (Tfp) pilus assembly protein with ATPase activity. This domain is found in PilM, a type 4 pili biogenesis protein which is linked to the production of polar type 4 fimbriae in P. aeruginosa (Martin et al., 1995). Downstream of mshll is a contiguous open reading frame, ORF2 termed mshll. Its initiation codon overlaps the TAG stop codon of mshl\ which is preceded by a potential RBS 5’GACGA3’ 8 bp upstream of the mshll stop codon. The 584 bp mshI2 gene encodes a protein with 194 amino acid residues similar to Mshl, which encodes the MSHA biogenesis protein found in Vibrio species (Table 4-2). The similarity of the gene product corresponds to the regions between 287 to 463 amino acid positions of Mshl of V. cholerae 01 biovar El Tor strain; between 288 to 471 amino acid positions of Mshl of V parahaemolyticus\ and between 290 to 480 amino acid positions of Mshl of V. vulnificus. Downstream of mshll is another contiguous open reading frame termed mshJ, within which the transposon insertion had occurred. The ATG start codon and putative RBS 5’GGAGGC3’ of mshJ are located within the coding region of mshll. The mshJ gene is a 650 bp long and encodes a protein with a predicted length of 216 amino acids residues. This gene product is similar to MshJ of the Vibrio species (Table 4-2). 139 A contiguous open reading frame (ORF4) was identified downstream of mshJ. The ORF4 termed mshK, consists of 324 bp with an ATG start codon and predicted RBS 5’GGTGAG3’ located within the coding region of mshJ. The gene product shares similarities with MshK of the Vibrio species. No transmembrane region was identified and the predicted location of the gene product is in the bacterial periplasmic space as deduced using PSORT analysis. The ORF4 is designated as mshK. ORF5 termed mshL, is located downstream of mshK. The ATG codon overlaps the stop codon of mshK, which is preceded by a putative RBS 5’GCGAAC3’ located in the coding region of mshK. The mshL gene consists of 1691 bp encoding a protein with 563 amino acids residues. The gene product shares homology with MshL of Vibrio species (Table 4-2). The mshL gene has a conserved secretion protein domain similar to several proteins of type II secretory pathway and pilus assembly proteins. The secretion protein domain (GG(X12)VP(L/F)LXXIPXIGXL(F/L)) (Huang et al., 1992) starts at amino acid residue 493. Its amino acid sequence is similar to that of PulD, a protein involved in the secretion of pullulanase in Klebsiella oxytoca (Pugsley, 1993) and CpaC, a protein involved in the fimbrial assembly in Caulobacter crescentus (Viollier et al., 2002). The similarities with these domains are localized in the C-terminal region. Downstream of mshL is an incomplete contiguous open reading frame, ORF6 termed mshM. No obvious stop codon was identified, suggesting that this gene continues downstream. The start codon of mshM overlaps the stop codon of mshL, which is preceded with RBS 5’GATGGC3’ located within the coding region of mshL. The incomplete ORF showed homology with mshM of V. cholerae 01 biovar El Tor and contains a domain similar to ExeA, a type II secretory protein. 140 4.3.2. Detection and analysis of cell surface pili Following the detection of six contiguous genes (mshll, mshI2, mshJ, mshK, mshL and mshM) in P. tunicata chromosomal DNA, which are presumed to encode secondary proteins for MSHA pilus biogenesis, the presence of pili on the cell surface was explored. Transmission electron micrographs of P. tunicata wild type and SM5 mutant cells using negative staining are shown in Figure 4-3. It was observed that P. tunicata wild type cells were surrounded by 8 to 10 flexible pili on their surface. Each pilus had a diameter of 7 to 8 nm and a length of 90 to 150 nm. These pili are arranged over the whole surface of the bacterial cell. The SM5 mutant cells were found to be nonpiliated, suggesting that the region of mutation is important to pilus expression. Further electron microscopy studies revealed an increase in peritrichously arranged flexible pili on the surface of P. tunicata wild type cells pregrown on cellulose or cellobiose as the sole carbon source. There was a considerable increase in the number of pili on the cell surface of cells grown in these carbon sources compared to cells grown in glucose (Figure 4-3). 141 Figure 4-3. Transmission electron micrographs of P. tunicata cells. A) P. tunicata wild type cell expressing pili on its surface. Bar, 0.2 pm B) P. tunicata SM5 mutant cell displaying nonpiliated phenotype. Bar, 0.5 pm. Both strains were grown on VNSS agar plates for 24 h. C) A hyperpiliated P. tunicata wild type cell grown in cellulose as the sole carbon source for 24 h in static condition. Bar, 2.0 pm D) P. tunicata wmpR mutant cell displaying nonpiliated phenotype. Bar, 0.5 pm. All samples were negatively stained with 2 % phosphotungstic acid. 142 4.3.3. Assay for agglutination of red blood cells In order to further characterize the identity of the P. tunicata pilus, hemagglutination assays were performed. It was found that P. tunicata wild type cells cause agglutination of the horse red blood cells. Hemagglutination was observed with both exponential and stationary phase cells, and P. tunicata was shown to cause agglutination of red blood cells at a 20 time dilution (Table 4-3). No hemagglutination was observed in the presence of the SM5 mutant when the cells were assayed during exponential phase, while in stationary phase a slight hemagglutination effect was observed. When mannose and fucose were added to the assay mixture, no hemagglutination was observed for the P. tunicata wild type. Table 4-3. Hemagglutination of horse erythrocytes by bacteria at different growth stages3 Bacterial Stationary Log Mannoseb Fueosec strain phase phase Stationary Log Stationary Log Vibrio cholerae >50 >50 - - >50 >50 M1615 (+ control) P. tunicata wild type 20 20 - - - P. tunicata mshJ <2 ND ND ND ND ND (SM5) mutant JValues represent the reciprocal of the dilution in which hemagglutination can be observed. bAssay performed in the presence of 1 % a-methyl D-mannoside. c Assay performed in the presence of 1 % L-fucose dND, not detectable 143 4.3.4. Pili promote the attachment of P. tunicata to abiotic and biotic surfaces The polystyrene surface in a microtitre plate, a widely used attachment assay system (O’Toole and Kolter, 1998; Taylor et al., 2002), was used as an abiotic test surface. After 1 h of attachment, the P. tunicata wild type cells were more densely attached to the polystyrene surface, than the SM5 mutant cells as detected macroscopically by crystal violet staining. The crystal violet stained P. tunicata wild type cells displayed an optical density of 0.972 ± 0.093, which was 65 % higher than the crystal violet stained SM5 mutant cells (Figure 4-4). The attachment of P. tunicata wild type and SM5 mutant cells to cellulose was also tested. Cellulose is a major surface polymer of C. intesrinalis (de Leo et al., 1977) and U. lactuca (Chapman, 1979). As demonstrated in Figure 4-5, P. tunicata wild type cells have the ability to attach efficiently to microcrystalline cellulose as shown by the 22 % ± 0.498 reduction of the final OD of the assay mixture. The SM5 mutant cells however, attached less effectively to cellulose showing only 4 % ± 2.004 reduction. It was further demonstrated that attachment to cellulose is greatly increased for the wild type cells when grown on the substrates cellulose and cellobiose. The P. tunicata wild type cells displayed 30 % ± 0.419 and 38 % ± 4.731 reduction of the final OD of the assay mixture when the cells were pregrown in cellobiose and cellulose, respectively. In contrast, the SM5 mutant cells pregrown in cellobiose and cellulose showed a 26 % ± 0.990 and 14 % ± 9.022 reduction in the final OD of the assay mixture, respectively. It was also demonstrated that mannose significantly inhibits attachment of P. tunicata wild type cells to cellulose displaying only 4 % ± 1.687 reduction of the final OD of the attachment assay mixture. 144 wild type mshJ mutant Figure 4-4. Attachment of P. tunicata wild type and mshJ (SM5) mutant cells to polystyrene microtitre plate surfaces. Bacterial strains attached to polystyrene surface were stained with 1 % crystal violet and destained with 95 % ethanol for quantification. Attached crystal violet stained cells are shown above. Quantification of the crystal violet associated with attached cells is shown below. The bar represents the measure of OD 590 nm. Error bars represent the standard error of triplicate cultures. 145 □ P. tunicata wild type □ P. tunicata mshJ (SMS) mutant Growth Media Figure 4-5. Attachment of P. tunicata wild type and mshJ (SM5) mutant to microcrystalline cellulose (Avicel). The microcrystalline cellulose was mixed with bacterial cells (~10 cfu/ml) for 1 h and the final OD of cellulose and bacterial cells suspension was measured after 1 h of incubation. The attachment of bacteria to cellulose was scored by the reduction of the final OD of the suspension. Error bars represent the standard error of triplicate cultures. aBacteria were grown on this specific medium prior to the attachment assay. bBacteria were grown on VNSS medium prior to the attachment assay. Attachment mixture contained 100 mM a-methyl D-mannoside. 146 A qualitative assessment of the attachment of the two strains to cellulose using fluorescently tagged cells and confocal laser scanning microscopy was made. The resulting images revealed that a substantial number of P. tunicata wild type cells were attached to microcrystalline cellulose (Figure 4-6B). In contrast, the SM5 mutant cells attached less to cellulose when assayed alone or together with wild type cells (Figure 4- 6A and 6C). Examination by SEM clearly demonstrated the role of pili in the attachment of the wild type cells to cellulose (Figure 4-7). It was shown that these cell surface appendages promote attachment of bacteria to cellulose surfaces. 4.3.5. Attachment of P. tunicata to the surface of axenic green alga U. lactuca Axenic U. lactuca thallus discs were incubated with GFP tagged P. tunicata wild type and GFP tagged SM5 mutant and attachment assays were carried out in PBS (pH 7.4). The numbers of surface attached cells of P. tunicata wild type and the SM5 mutant are presented in Figure 4-8. P. tunicata wild type cells demonstrated an enhanced attachment to the surface of U. lactuca compared to the SM5 mutant cells. It was found that the attached P. tunicata wild type cells formed cell aggregates or microcolonies on the surface of U. lactuca. In contrast, the attached SM5 mutant cells were distributed as solitary cells. The attachment of the SM5 mutant cells was only 42.2 ±8.1 % of the total number of attached wild type cells. 147 Figure 4-6. Transmission fluorescent micrographs using confocal laser scanning microscopy showing attachment of bacterial strains to microcrystalline cellulose (Avicel) after completion of the attachment assay. A) GFP tagged P. tunicata wild type and RFP tagged mshJ (SM5) mutant mixed in 1:1 mixture, B) GFP tagged P. tunicata wild type and C) RFP tagged mshJ (SM5) mutant. Bar, 20 pm. 148 Figure 4-7. Scanning electron micrographs of P. tunicata cells grown in cellulose. A) P. tunicata wild type grown in cellulose as the sole carbon source. B) P. tunicata wild type adhering to microcrystalline cellulose. 149 Figure 4-8. Confocal laser scanning microscopy images of (A-l) GFP tagged P. tunicata wild type and (A-2) GFP tagged mshJ (SM5) mutant attached on the surface of treated U. lactuca. Green fluorescing cells were manually counted on images taken with a LSMG200 confocal laser scanning microscope. B) The bar graph represents the number of adhered cells to treated U. lactuca pieces. 150 4.4. Discussion This chapter describes a gene locus proposed to be involved in the export and assembly of an MSHA pilus in P. tunicata. The examination of the DNA regions flanking the transposon insert of the P. tunicata SM5 mutant revealed the presence of an ORF with high homology to mshJ, a member of the secretory operon for the MSHA pilus biogenesis gene locus of V. cholerae (Marsh and Taylor, 1999). Further sequencing of the DNA flanking the transposon revealed six contiguous open reading frames: ms hi l, 12, J, K, L and M, with similar genetic organization and high homology to the MSHA pili biogenesis secretory proteins of V. cholerae, V. parahaemolyticus and V. vulnificus. The best characterized MSHA pilus gene locus is that of V. cholerae El Tor (Marsh and Taylor, 1999). The six P. tunicata ORFs identified in this study share major similarities with the MSHA pili secretory genes of V. cholerae, including: (1) homology of the predicted amino acid sequences, (2) similarity in the predicted location of the gene products, (3) similarity in their organization, orientation and arrangement, and (4) the presence of polycistronic genes with overlapping ORFs. Electron microscopy studies revealed the presence of flexible pili on the cell surface of P. tunicata, with ultrastructural characteristics similar to the pili of Gram-negative bacteria. In contrast, the P. tunicata SM5 mutant disrupted in the mshJ gene did not express pili on its surface. The biogenesis of pili requires numerous gene products, including structural prepilin subunits, ancillary proteins with prepilin-like leader sequences, inner and outer membrane proteins and nucleotide binding proteins (Aim and Mattick, 1997). It has been reported that a mutation in any of these genes is sufficient to prevent the assembly of functional pili (Aim and Mattick, 1997; Strom and Lory, 1993). For example, mutation in the P. aeruginosa pilus biogenesis secretory genes pilO, pilP, or pilQ resulted in phenotypes that were lacking pili, confirming the importance of these genes in pilus biogenesis (Martin et al., 1995). These secretory genes belong to an operon for secretion and export which includes two other genes (pilM and pi IN) required in the biogenesis of fimbriae in P. aeruginosa (Martin et al., 1995). In V. cholerae, it was reported that the expression of MSHA to form functional pili on the bacterial surface is completely dependent on the transcription and expression of two operons tenned secretory and structural, respectively (Marsh and Taylor, 1999). Deletion in any of the putative promoter regions upstream of mshl, a secretory gene, or mshB, a 151 structural gene, abolished MshA pilus assembly, secretion and expression (Marsh and Taylor, 1999). Additionally, mutation in mshE, a secretory gene in MSHA biogenesis locus was shown to abolish hemagglutination (Hase et al., 1994). The fact that P. tunicata SM5 mutant displayed a nonpiliated phenotype and was unable to mediate hemagglutination suggests that the mshj gene is involved in pilus biogenesis in P. tunicata. Although the mutated gene in SM5 is not in a structural gene of MSHA pilus biogenesis, it is likely that the mutation of a secretory gene would also result in a nonpiliated phenotype. Gram-negative bacteria bind to surfaces via the tip adhesins of the pili (Strom and Lory, 1993). These adhesins may have different specific receptors. For example, V. cholerae El Tor strains produce a hemagglutinin pilus with preference for mannose receptors (Jonson et al., 1991). Additionally, V. cholerae classical strains express hemagglutinin pilus with a preference for fucose (Hanne and Finkelstein, 1982). A hemagglutinin pilus with preference for both mannose and fucose is expressed by the V. cholerae 01 strain Bgdl7 (Ehara et al., 1991). We have demonstrated that the P. tunicata pilus causes hemagglutination of horse red blood cells. As for V. cholerae 01 strain Bgdl7, the clumping of red blood cells was abolished in the presence of mannose and fucose, suggesting the recognition of more than one receptor domain or the presence of accessory adhesin subunits in the P. tunicata pilus. The MSHA pili expressed by V. cholerae are known to be involved in the attachment to and colonization of surfaces. The MSHA pilus is an important attachment factor in the aquatic environment and has been shown to mediate attachment to solid substrates (Chiavelli et al., 2001; Watnick et al., 1999a). It was reported that the V. cholerae El Tor mshA mutant strain is unable to form biofilms and shows decreased adherence to abiotic and biotic surfaces (Watnick et al., 1999a). In this study, we first tested attachment of P. tunicata wild type and SM5 mutant cells to polystyrene and to cellulose, the major surface polymer of U. lactuca (Chapman, 1979) and C. intestinalis (de Leo et al., 1977) from which P. tunicata had been isolated. The SM5 mutant cells showed less attachment to both the polystyrene and cellulose surfaces, in contrast to the piliated wild type strain, which displayed significant attachment to both surfaces. This suggests that the P. tunicata MSHA pilus plays a key role in the attachment of P. tunicata to both surfaces. The attachment of the SM5 mutant, while being lower than 152 that of the wild type, was sufficient to suggest that other undefined attachment factors are also involved in the attachment process of P. tunicata to these surfaces. Surface associated bacteria on marine sessile organisms may provide benefits to their hosts. For example, P. tunicata produces a range of inhibitory compounds and may offer protection against the colonization by biofouling and pathogenic organisms (Egan et al., 2001a; 2001b; Holmstrom et al., 1992; James et al., 1996). Such interactions are initiated by the attachment and initial colonization of bacteria on the host surface, and may be facilitated by the expression of surface adhesins. In V. cholerae El Tor, the attachment to phytoplankton and zooplankton surfaces is mediated by the MSHA pilus (Chiavelli et al., 2001). Recently, it was reported that the MSHA pilus promotes interaction between V. cholerae El Tor and hemolymph of the mussel Mytilus galloprovincialis (Zampini et al., 2003). In this study, the role of the MSHA pilus in the attachment of P. tunicata to the surface of U. lactuca was investigated. The attachment assays revealed significantly higher numbers of attached P. tunicata wild type cells compared to the number of SM5 mutant cells at the surface on the U. lactuca surface. The findings of the investigation also indicate that the MSHA pilus facilitates the initial stage of colonization. The wild type, but not the MSHA mutant cells, formed microcolonies, a behaviour typical of bacterial biofilm formation, on the surface of axenic plant tissue of U. lactuca (Costerton et al., 1995; Stoodley et al., 2002). The ability of P. tunicata SM5 cells to attach to the surface of U. lactuca suggests that, although this attachment is reduced compared to the wild type, other hitherto undefined surface structures and mechanisms may be involved in the adhesion of P. tunicata to the algal surface. This study also showed that specific growth conditions affect the expression of P. tunicata MSHA pilus. P. tunicata cells grown in cellulose or cellobiose were found to be hyperpiliated when examined under TEM.-These specific substrates also promoted the attachment of P. tunicata to cellulose. We observed that wild-type cells pregrown in cellulose or cellobiose displayed an increased attachment to cellulose compared to cells grown on other carbon sources. The minor increase in adhesion to cellulose displayed by the SM5 mutant grown in cellulose or cellobiose rather than complex media was unclear, however the results of these studies suggest that cellulose or cellobiose may serve as an environmental signal, that induces the expression of MSHA pilus and thus 153 promotes the attachment of P. tunicata to cellulose and specific marine living surfaces. The findings of this study concur with previous reports that chemosensory mechanisms control pilus expression. In P. aeruginosa, expression of pilA is controlled by a two- component sensor-regulator gene pair pilS and pilR (Hobbs et al., 1993). The PilS protein is a sensor protein located upstream of the regulator protein, PilR, thought to be responsible for sensing unknown environmental signals (Boyd, 2000). In V. cholerae, the response of tcp (toxin-coregulated pili) genes to environmental stimuli involves the ToxR regulon (Strom and Lory, 1993). The ToxR protein is a major sensor and regulator protein that transmits signals from the periplasmic side of the inner cell membrane, and regulates transcription of virulence factors such as TCP. Interestingly, a putative transcription regulatory gene, wmpR has been identified in P. tunicata (Egan et al., 2002b). This gene is homologous to toxR from V. cholerae and cadC from E. coli and is involved in the regulation of the expression of the antifouling compounds in P. tunicata (Egan et al., 2002b). The P. tunicata wmpR mutant is devoid of pili on its surface when examined in TEM (Figure 4-3D). It is possible that the regulatory activity of wmpR may explain the surface sensing mechanisms demonstrated by P. tunicata in response to environmental stimuli. In the marine ecosystem, the P. tunicata wmpR gene product may sense an environmental signal (e.g., cellulose or surface polymers of C. intestinalis and U. lactuca) and respond by increasing the expression of MSHA pili on the cell surface, hence promoting the attachment of P. tunicata to the surfaces of these marine organisms. In summary, the results described in this chapter have shown that the expression of the MSHA pilus on the cell surface of P. tunicata requires the activity of a putative MSHA gene locus for pilus biogenesis. MshJ may be directly involved in the secretion and assembly of P. tunicata pilin subunits. The findings of this study demonstrate that the production of MSHA pilus mediates the attachment of P. tunicata to surfaces, and that this pilus is a key attachment determinant for the colonization of P. tunicata on both abiotic and living surfaces. In addition, it has also been demonstrated that the expression of the MSHA pilus is induced by environmental conditions, indicating that P. tunicata possesses a surface sensing mechanism. It is proposed that the MSHA pilus as an attachment factor and the surface sensing mechanisms of P. tunicata have a profound effect on the distribution of this bacterium in marine systems. In addition, the ability of P. tunicata to grow on cellulose as a carbon source, as well as the possibility that it has a 154 surface sensing mechanism for this polymer, suggest that it may possess mechanisms to degrade the cellulose into soluble substrates and utilize them as an energy source. The next chapter of this thesis addresses these questions. 155 5. The role of a putative cellulosome and its ecological importance in association of Pseudoalteromonas tunicata with higher marine organisms 5.1. Introduction Cellulose is the most abundant polymer in nature and is a key source of carbon and energy in many ecosystems (Sheehan and Himmel, 1999). It is a very stable polymer composed of long unbranched homopolymers of D-glucose, linked with p-1,4- glucosidic bonds. Efficient and complete hydrolysis of this polymer requires the synergistic activity of numerous cellulolytic enzymes known as cellulases. To date, hundreds of different cellulases have been identified in a wide variety of cellulolytic microorganisms, occurring either as free enzymes or within enzyme complexes known as cellulosomes (Doi and Tamaru, 2000). One well-studied example of a free cellulase enzyme producer is the filamentous fungus Trichodemia reesei (Xu et al., 1998). Several Gram-positive cellulolytic Clostridia have been observed to produce cellulase enzymes within cellulosome complexes, including Clostridium thermocellum (Bayer and Lamed, 1986), C. cellulovorans (Doi and Tamaru, 2000), C. josiu (Kakiuchi et al., 1998), C. cellulolyticum (Belaich et al., 1997) and C. acetobutylicum (Sabathe et al., 2002) as well as other anaerobic bacteria including Ruminococcus flavefaciens, Ruminococcus albus, Acetivibrio cellulolyticus and Bacteriodes cellulosolvens (Ohara et al., 2000; Ding et al., 1999; 2000; 2001). The cellulosome enzyme complex is made up of a scaffolding protein, which has a cellulose binding domain and numerous catalytic enzymes, which act synergistically to degrade cellulose (Bayer et al., 1998). Transmission and scanning electron microscopy have revealed that this complex enzyme system occurs as protuberant structures tightly attached to the cell surface (Bayer and Lamed, 1986). Cellulose and other polymeric substances such as chitin, agar and lignin are abundant in the marine environment, where they serve as a nutrient source for microorganisms (Gooday, 1994; Gonzales et al., 1997). Marine bacteria convert these polymers into 156 soluble monomers, which can be further broken down and assimilated by the cell through the activities of ectoenzymes (extracellular hydrolases) such as proteinases, tyrosinases, cellulases, chitinases, agarases and glucosidase (Martinez et al., 1996). Members of the genus Pseudo alt eromonas have been reported to produce extracellular enzymes, which degrade polymers in marine environments. Some species namely P. agarolyticus, P. antarctica, P. carrageenovora and P. atlantica have been shown to produce agarase that hydrolyze agar, a surface component of red algae (Akagawa- Matsushita et al., 1992; Vera et al., 1998). Enzymes such as alginases and glycosidases have been reported in P. issachenkonii, a bacterium isolated from the degraded thallus of the brown alga Fucus evanescens (Ivanova et al., 2002d). Chitinases have also been detected in Pseudo alt eromonas sp strain S91. The cluster gene of this enzyme has been identified and well investigated (Techkamjanaruk and Goodman, 1999). A cold-active cellulase with endocellulase activity has recently been isolated and identified in the antarctic psychrophile P. haloplanhtis (Violot et al., 2003). Additionally, P. haloplanktis produces a cold-active (3-galactosidase demonstrating high catalytic efficiency on natural and synthetic substrates (Hoyoux et al., 2001). It has been suggested that hydrolytic ectoenzyme activities in Pseudoalteromonas species are linked to not only the taxonomic classification of the strain but also the environment from which it is isolated (Ivanova et al., 2003). It was proposed in Chapter 4 that P. tunicata is capable of growing on cellulose and cellobiose as the sole carbon source, and that growth on these substrates triggers the expression of the MSHA pilus - a key colonization factor that mediates attachment to U. lactuca. Cellulose is one of the major components of the U. lactuca surface (Chapman, 1979) and this polymer may be utilized as a nutrient source for attached P. tunicata cells. This chapter describes additional means by which P. tunicata uses cellulose for attachment and as a source of carbon and energy. Electron microscopy studies revealed that P. tunicata grown on cellulose as the sole carbon source produces protuberant-like surface structures similar to the cellulosomes of cellulolytic clostridia. To date, cellulosomes have only been observed in a restricted number of anaerobic bacteria, and the finding of a possibly similar surface structures in P. tunicata is therefore novel. This chapter identifies and describes this structure and its putative role in the utilization of cellulose by P. tunicata. It is proposed that the production of this 157 protuberant-like surface structure represents an ecopbysiological response of the bacterium, allowing it to adapt to life in association with higher marine organisms that have cellulose polymers on their surfaces. 5.2. Materials and Methods 5.2.1. Bacterial isolates and culture conditions P. tunicata wild type was grown in Marine Minimal Medium (MMM) (Appendix I), containing 0.05 % (w/v) glucose, 0.1 % (w/v) cellobiose (Sigma) or 0.25 % (w/v) microcrystalline cellulose (Avicel, Fluka) as the sole carbon source. The bacterial culture was incubated for 24 h with shaking (150 rpm) at room temperature. 5.2.2. Transmission and scanning electron microscopy studies The ultrastructures of the wild type P. tunicata cells grown in cellobiose and cellulose were examined using transmission electron microscopy (TEM). The bacterial cells were grown overnight on VNSS agar plates and transferred in liquid medium of MMM with glucose, cellobiose or cellulose as the sole carbon source and incubated for 24 h with shaking (section 5.2.1). The bacterial cells grown in liquid medium were centrifuged at 12, 000 x g for 5 min, washed twice and resuspended in PBS. A carbon-coated formvar copper grid was placed on a drop of the cell suspension for 5 min and negatively stained with 2 % phosphotungstic acid for 30 sec. The cells were examined using a Hitachi H7000 transmission electron microscope. Scanning electron microscopy (SEM) studies were performed on the bacterial samples grown in liquid medium prepared as described above. The bacterial cells were placed on poly-L-lysine coated cover slips and air-dried for 10 minutes and coated with chromium using EMITECH type K575x sputter coater. The final metallic film had a thickness of 15 nm. Samples were examined using a Hitachi H900 scanning electron microscope. 158 5.2.3. Attachment assay Attachment of P. tunicata cells to microcrystalline cellulose was measured by modifying the method of Bayer et al (1983). This assay is the same as that described in section 4.2.5, the only difference being that the bacterial cultures were incubated at room temperature with shaking (150 rpm) prior to the attachment assay. TEM studies revealed that high speed shaking shears off pili from the cell surface. The attachment experiment was carried out in triplicate. 5.2.4. Growth of P. tunicata in cellobiose or cellulose as the sole carbon source The growth curve assay was performed by growing P. tunicata in 50 ml side arm flasks containing 10 ml of Marine Minimal Media (MMM) with 0.25 % microcrystalline cellulose (Avicel, Fluka) or 0.1 % cellobiose (Sigma) as the sole carbon source. Ten microlitres of an overnight culture was inoculated into each flask and incubated with shaking at room temperature. Growth was monitored by measuring the optical density at 610 nm over 3 days. Experiments here were carried out in triplicate. 5.2.5. Isolation of bacterial fractions and preparation of cellulose-bound proteins The method of protein preparation used by Mosoni and Gaillard-Martinie (2001) was adopted with some modifications. P. tunicata cells were grown as described in section 5.2.1. The glucose- and cellobiose-grown cell cultures were incubated for 24 h with shaking at room temperature, except for cellulose-grown cells which were incubated for 48 h. The glucose and cellobiose-grown cultures were centrifuged at 12, 000 x g for 10 min. The cellulose-grown culture was centrifuged at 500 x g for 10 min to separate cellulose from the cells and the cellulose-free cell culture was centrifuged at 10, 000 x g for 10 min at 4°C. The cellulose separated from the cell culture was washed five times with PBS, pH 6.5 to remove adhering cells. The washed cellulose was boiled for 5 min 159 with SDS sample buffer according to the method described by Miron and Forsberg (1999) to elute out the cellulose-bound proteins. To prepare the membrane-bound proteins, the pelleted cells grown from different substrates were resuspended in 5 ml of PBS (pH 6.5), vortexed vigorously for 15 min and centrifuged at 12, 000 x g for 10 min at 4°C. The protein profiles of the different samples (cell-free supernatant, membrane proteins and cellulose-bound proteins) were analyzed using SDS-PAGE. 5.2.6. Identification of cellulose-binding proteins by affinity based assay In order to identify the cellulose-binding proteins from the different protein fractions prepared above, an affinity based assay was carried out as described previously (Pegden et al., 1998). Briefly, an aliquot of about 250 pi of the proteins eluted from the residual cellulose was mixed with 300 mg of microcrystalline cellulose (Avicel) suspended in PBS (pH 6.5) with 4 mM calcium chloride and 2 mM dithiothreitol, in a total volume of 1 ml. The suspension was mixed slowly for 1 h at room temperature and the cellulose particles were harvested by slow centrifugation (500 x g for 5 min). The supernatant containing unbound proteins was removed and analyzed. The cellulose particles were washed with 1 ml of the following: a) PBS b) Triton X-100 (0.1 % w/v) and c) SDS- PAGE sample buffer. The mixture was centrifuged at 500 x g for 5 min and the washed fractions were examined using SDS-PAGE. 5.2.7. Peptide sequencing In order to know the protein identity of the cellulose bound protein, peptide sequencing was carried out. Briefly, the protein was excised from the SDS-PAGE gel using a sterile razor blade and dried in a vacuum. The protein was trypsin digested and the peptide fragments were subjected to liquid chromatography and mass spectroscopy (LC-MS), made available through the Bioanalytical Mass Spectroscopy Facility of the University of New South Wales. The Mascot Program and NCBI program BlastP (search for short 160 nearly exact matches) were used to find the homologies and matches of the sample protein with the proteins in the GenBank. 5.2.8. Cellulase activity assay Cellulase activity was detected qualitatively using the congo-red staining method in a gel diffusion assay (Carder, 1986). Twenty microlitres of the membrane bound proteins from cellulose-grown culture was added to a petri dish containing solidified agarose gel (1 %) with carboxymethylcellulose (CMC) (1.5 % w/v) in 50 mM sodium acetate, pH 5.0 and incubated for 48 h at room temperature. The positive control was 20 pi of endo- cellulase (EGII) (1000 U/ml) (Megazyme) and the negative control 20 pi of endo- cellulase (EGII) boiled with SDS buffer. After 48 h the gel was stained with congo-red (0.1 % w/v) for 30 minutes and destained with 1 M NaCl until a zone of clearing was observed. 5.2.9. SDS-PAGE analysis and the zymogram assay Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was performed as developed by Laemmli (1970). The samples used for the SDS-PAGE were boiled with the sample buffer and analyzed using a 10 % w/v gel. Samples intended for zymogram analysis were incubated in sample buffer for 1 h at 45°C (mild denaturation) before SDS-PAGE (Pohlschroder et al., 1994). CMCase activity was detected using SDS-PAGE zymograms which contained 1 % of CMC as substrate. The denatured samples were loaded in the prepared gels and run at 20 mA for 2 h at 4°C. The gels were renatured in 20 % isopropanol in 50 mM NaAc, pH 5.0, for 30 min. The renaturation process was repeated three times. The gels were then washed twice in 50 mM NaAc (pH 5.0) for 30 min. The washed gels were stained with congo-red (0.25 % w/v) for 30 min and destained with 1 M NaCl until clearing zones were observed. 161 5.3. Results 5.3.1. P. tunicata produces cellulosome-like surface structures The negatively stained P. tunicata cells grown in cellulose as the sole carbon source were observed to have protuberant-like structures on their surfaces when examined by TEM (Figure 5-1 A). Each of the spherical structures was closely bound to the surface of the cell, clearly showing the point of its origin on the cell surface (Figure 5-1 A, arrow). Some structures were large (~80 to 100 nm) possibly indicative of maturity, while others appear as small (~30 to 50 nm) newly developed spherical structures. These structures were also observed on the surface of P. tunicata cells grown in cellobiose, however in this case the spherical particles appeared to be smaller and less developed than those of the cellulose-grown cells (Figure 5-IB). P. tunicata cells grown in glucose as the sole carbon source appeared to have no protuberant structures on their surfaces (Figure 5-1C). The spherical particles were examined in further detail using SEM. This analysis confirmed that cells grown in cellulose had numerous spherical particles, apparently covering the entire cell surface (Figure 5-ID) and being closely attached to the surface of the microcrystalline cellulose (Figure 5-IE). Some non-cell associated spherical particles closely bound on the surface of the cellulose were also observed (Figure 5-1F). 162 3 3 c o 5 t/d o C attaching O X 2Q ocS O •*-*»K jd 3 o tunicata P. O E) Vd vd cd 3 the u 3 0 U T3 .£ c cd 0) covering Vd 1 o 3 o jjn 3 3 o o structures a oa ss; 4-i o cellulosome-like xcx cd•— 00 C/d with Oh 3 cell Vi 3 c c/d o _£h 0 3 x-i—> 3 tunicata 3 3 00 o P. .3 c/d 3 D) 3 1 o c/d X source: 3 sole § 3 o the X as -t—> ir< 00 3 a> ose s- 3 ’? 3 .Sf o jo u. X 3o 3 5.3.2. P. tunicata attaches to cellulose The attachment assay used in this chapter is different from that described in Chapter 3, where cells were grown without shaking in order to promote formation of pili. In this chapter, the bacterial cultures were shaken from the start of incubation until the time of harvest with the intention of shearing off the pili (Figure 5-2). The attachment assay was carried out in order to determine any other possible mechanisms by which P. tunicata attaches to cellulose. It was demonstrated that P. tunicata attached to cellulose as measured via the reduction of the optical density of the culture suspension (Figure 5-3). The reduction in optical density reflects increased attachment of the cells to cellulose which sediments out of suspension during incubation. Cells grown in cellobiose or cellulose as the sole carbon source displayed an OD reduction of 33 % and 37 %, respectively, whereas the glucose-grown cells attached less, with only a 16 % reduction in optical density. Figure 5-2. Transmission electron micrographs of P. tunicata grown in cellulose as carbon source in different incubation conditions. A) Cells incubated without shaking showing the presence of pili. Bar, 1.0 pm. B) Cells incubated with shaking (150 rpm) showing no pili, Bar, 0.5 pm. 164 100 80 c 60 o 5 3 ■o 0> DC a 40 o a? 20 0 ------ Glucose Cellobiose Cellulose Growth media Figure 5-3. Attachment of P. tunicata wild type to microcrystalline cellulose (Avicel). The microcrystalline cellulose was mixed with bacterial cells (~10 cfu/ml) for 1 h and the final OD of cellulose and bacterial cells suspension was measured after 1 h of incubation. The attachment of bacteria to cellulose was scored by the reduction of the final OD of the suspension. Error bars represent the standard error of triplicate cultures. 5.3.3. Growth of P. tunicata in different substrates There was a different growth pattern observed in P. tunicata grown in cellobiose compared to cellulose (Figure 5-4). The P. tunicata cultures in each of the substrates commenced growth on the first day of inoculation and reached their stationary phase on the second day, however the growth of the cellobiose-grown P. tunicata culture started to decline after the third day of incubation. In contrast, the cellulose-grown cells underwent another logarithmic phase after 30 h of incubation, reaching a plateau on the third day. 165 1.5 E c o Q o Tirre(hcxrs) Figure 5-4. Growth curves of P. tunicata in different substrates. The growth curve assay was performed by growing P. tunicata in 50 ml side arm flasks containing 10 ml of Minimal Marine Medium (MMM) with 0.25 % microcrystalline cellulose (Avicel, Fluka) or 0.1 % cellobiose (Sigma) as the sole carbon source. The optical density was measured at 610 nm. 166 5.3.4. Cellulose-binding protein facilitates the attachment of P. tunicata to cellulose Aliquots of the cell-free supernatant and membrane bound proteins sampled from the cellulose-grown P. tunicata cells, as well as the proteins eluted from cellulose particles, were analyzed using SDS-PAGE. The sample containing proteins eluted from the cellulose particles gave a single band of molecular weight approximately 46-48 kDa band upon SDS-PAGE analysis, indicating that this band corresponds to the cellulose binding protein (Figure 5-5, lane C). Based on SDS-PAGE analysis, the cellulose binding protein was present in lower amounts in the supernatant of the cell culture grown in cellulose (Figure 5-5, lane A) as shown by a lighter stained 46-48 kDa protein band. A heavy stained 46-48 kDa protein band was present in membrane bound proteins, suggesting a higher abundance of the protein in this fraction (Figure 5-5, lane B). The 46-48 kDa protein band was also found in the cell free supernatant and membrane bound protein fractions taken from cellobiose-grown cells (Figure 5-6, lane B). The proteins harvested from the glucose-grown cells showed, relatively, the least amount of the 46-48 kDa protein (Figure 5-6, lane C), as shown by a faint protein band in SDS-PAGE. The 46-48 kDa protein appeared to be tightly bound to the cellulose as deduced from the result of the affinity based assay (Figure 5-7). Protein samples eluted from residual cellulose was used in the affinity-based assay. It was found that the 46- 48 kDa protein was bound to the microcrystalline cellulose. Triton X and SDS largely removed the bound protein from the cellulose (Figure 5-7, lane C and E). Taken together, the results of these experiments indicate that the 46-48 kDa protein is a main cellulose-binding protein. 167 MW A B C kDa Figure 5-5. SDS-PAGE-Silver nitrate analysis of proteins from cellulose grown P. tunicata. Lane A, cell-free supernatant; lane B, membrane-bound proteins; and lane C, proteins eluted from cellulose residues using SDS sample buffer. MW, Molecular weight markers expressed in kilodalton (kDa). 168 kDa MW ABC 203 90 **3 >a£ 51.7 34.1 Figure 5-6. SDS-PAGE-Coomassie blue analysis of membrane-bound proteins of P. tunicata grown in different substrates. Lane A, cellulose; lane B, cellobiose; and lane C, glucose. MW, molecular weight markers expressed in kilodalton (kDa). 169 kDa MW ABC D E ■ «v 203 120 * 90 51.7 34.1 28 Figure 5-7. SDS-PAGE-Silver nitrate analysis of the affinity-based assay of P. tunicata proteins eluted from the cellulose residues. Lane A, proteins prior to the affinity based assay; lane B, unbound proteins; lane C, proteins removed from the cellulose by Triton X-100; lane D, proteins removed from the cellulose by PBS; and lane E, proteins removed from the cellulose by SDS sample buffer. Proteins eluted from cellulose were mixed with 300 mg of microcrystalline cellulose (Avicel) suspended in PBS (pH 6.5) with 4 mM calcium chloride and 2 mM dithiothreitol, making a total volume of 1 ml. The suspension was mixed slowly for 1 h at room temperature and washed with 1 ml either Triton X-100, PBS or SDS-PAGE sample buffer. MW, Molecular weight markers expressed in kilodalton (kDa). 5.3.5. Peptide sequencing of 46-48 kDa protein The peptide sequence of the 46-48 kDa protein was identified using LC-MS. Four peptide sequences were obtained, 1) EDTLTFQFSQK 2) FANNLTWVLAVK 3) PMLWQDFTVK and 4) LQQTALG. These peptide sequences were compared with proteins deposited in the GenBank using the Mascot Program. The results did not reveal any high scoring matches. Analysis using the NCBI BlastP program (search for short nearly exact matches) also did not allow the identification of the protein with full confidence. However, it is interesting to note that two peptide sequences showed similarities with protein segments of cellulases. The FANNLTWVLAVK peptide sequence, specifically the NNLTWV segment, is similar to the amino acid 352 to 357 segment of the endoglucanase family 5 of C. acetobulyticum (GenBank accession no. AAK78802.1). The PMLWQDFTVK, particularly the PMLWQD segment, is similar to the amino acid 510 to 515 segment of the glucoamylase TGA of Thermoactinomyces vulgaris (GenBank accession no. BAB40639.1). The segment similarities apply to the C-terminal region of the subject proteins. 5.3.6. Cellulase activity and zymogram assay The membrane proteins from P. tunicata grown with cellulose as the sole carbon source, were shown to have cellulase activity when assayed using the CMCase assay (Figure 5- 8) as shown by the clearing zone on the drop of the sample protein. This activity was lost when the sample was boiled with SDS sample buffer prior to the assay, indicating that the activity becomes unstable under conditions that cause protein denaturation. This result was further verified by the SDS-PAGE zymogram assay results. These results indicate the existence of two proteins (with molecular weight approximately 120 and 90 kDa) with CMCase activity. A clearing zone was observed corresponding to the position of these two proteins on the SDS-PAGE gel. No activity was detected in the protein corresponding to putative CBP (approximately 46-48 kDa) (Figure 5-9). 171 Figure 5-8. Congo-red plate diffusion cellulase activity assay of membrane bound proteins of P. tunicata grown in cellulose as sole carbon source. Sample 1 - membrane bound proteins boiled with SDS; sample 2 - membrane bound proteins. The positive control was endo-cellulase (EGII) (1, 000 U/ml) and the negative control endo-cellulase (EGII) (1, 000 U/ml) boiled with SDS sample buffer. Membrane-bound proteins were added in solidified agarose gel (1 %) with carboxymethylcellulose (CMC) (1.5 % w/v) in 50 mM sodium acetate, pH 5.0 and incubated for 48 h at room temperature. The gel was stained with congo-red (0.1 % w/v) for 30 min and destained with 1 M NaCl. 172 kDa MW A B Figure 5-9. SDS-PAGE and zymogram analysis of membrane-bound proteins of P. tunicata grown in cellulose as the sole carbon source. A) SDS-PAGE-Coomassie blue analysis of membrane bound proteins. B) Zymogram gel of membrane bound proteins after running SDS-PAGE. Samples for zymogram analysis were incubated in sample buffer for 1 h at 45°C (mild denaturation) before SDS-PAGE. The SDS-PAGE zymograms contained 1 % of CMC as substrate. The denatured samples were loaded in the prepared gels and run at 20 mA for 2 h at 4°C. The gels were renatured in 20 % isopropanol in 50 mM NaAc, pH 5.0, for 30 min and washed twice in 50 mM NaAc (pH 5.0) for 30 min. The washed gels were stained with congo-red (0.25 % w/v) for 30 min and destained with 1 M NaCl. 173 5.4. Discussion In Chapter 4 it was shown that P. tunicata is able to attach to cellulose particles and use cellulose or cellobiose as its sole carbon source. This chapter adds further information on the means by which P. tunicata is able to attach to and utilize cellulose. It was found that P. tunicata produces spherical-shaped, protuberant structures on its surface when grown in cellulose or cellobiose, but not when grown on glucose as the sole carbon source. The ultrastructure of these protuberant-like structures appeared to be similar to the cellulosome complex produced by the cellulolytic Clostridium species (Bayer and Lamed, 1986; Doi and Tamaru, 2001; Kakiuchi et al., 1998; Belaich et al., 1997), which contains numerous catalytic enzymes known to interact in a synergistic manner for the efficient degradation of cellulosic substrates (Bayer et al., 1998). Cellulosomes mediate the attachment of cellulolytic cells to cellulose and the degradation of this polymer into soluble substrates (Bayer et al., 1998). Cellulosomes were first observed in C. thermocellum using TEM, SEM and immunolabeling (Bayer and Lamed, 1986; Mayer et al., 1987). The cellulosomes of C. thermocellum were observed in cells grown in cellulose and cellobiose (Mayer et al., 1987). These structures are closely associated with the cell surface during the early stage of growth (Bayer and Lamed, 1986) and subsequently detach from the cell surface during stationary phase. Loosely bound cellulosomes were found attached to the residual substrate (Lamed and Bayer, 1986). There are many examples of cellulolytic bacteria able to attach to cellulose through their cellulose-binding proteins (CBP). For example, R. albus attaches to crystalline cellulose through its cellulose-binding protein, CbpC, which belongs to the Pil protein family (Pegden et al., 1998). It was demonstrated that CbpC is present in cellulose and cellobiose-grown cultures of R. albus. It has been reported that a ruminal bacterium Fibrobacter intestinalis DR7 also attaches to cellulose through its cellulose-binding proteins (CBP). The efficiency by which this bacterium binds to cellulose is due to the presence of 16 CBPs present in its outer membrane, six of which exhibit endoglucanase activity (Miron and Forsberg, 1999). In this thesis, it was demonstrated that P. tunicata produces a cellulose-binding protein when grown in cellulose or cellobiose as the sole carbon source. It was clearly shown that P. tunicata produces a cellulose-binding 174 protein as demonstrated by the sole appearance of this protein from the residual cellulose. The cellulose-binding protein of P. tunicata adhered strongly to the cellulose and could only be removed by treatment with detergents such as SDS and Triton X-100. Based on SDS-PAGE analysis, the approximately 46-48 kDa CBP is proposed to be a membrane bound protein, as deduced from its relative abundance in the membrane bound protein samples. The cell-free supernatant also showed the presence of a cellulose-binding protein, however it is relatively less abundant in the supernatant, suggesting that the protein is released into the supernatant or may have detached from the cell. Peptide sequence analysis of the approximately 46-48 kDa protein did not match with full confidence with any proteins in the database. However, segments from the identified peptide sequences showed similarities with amino acid segments of endoglucanase and glycoamylase. Although, these peptides have low confidence hits, it is interesting that they match with segments of an endoglucanase in C. acetobutylicum which is present in the cellulosome complex of this organism (Sabathe et al., 2002). It was demonstrated that the cellulose and cellobiose-grown P. tunicata cells attached to cellulose to a greater extent, than did the glucose-grown cells. While the SDS-PAGE analysis showed that the cellulose-binding protein is present in all of the membrane protein samples prepared from bacterial cultures grown in different substrates, the 46- 48 kDa CBP is more abundant in the cellulose- or cellobiose-grown cells compared to glucose-grown cells. This finding suggests that increased expression of CBP in P. tunicata is dependent on the presence of cellulose or cellobiose. It was previously reported that the expression of CBP in R. albus increased in response to the presence of cellulose (Pegden et al., 1998). The presence of cellulose also resulted in an increase in the expression of the 180 kDa CBP of Fibrobacter succinogenes (Gong et al., 1996). In addition, the formation of cellulosome is induced by growth on cellulose as well as cellobiose (Bhat et al., 1993). However, cellobiose-grown C. thermocellum forms an undeveloped protuberant-like structure, similar to the structure observed in cellobiose- grown P. tunicata. It has been reported that the attachment of C. thermocellum to cellulose via its cellulosome complex is mediated by the cellulose-binding domain (CBD) of the noncatalytic scaffolding protein, CipA (Gemgross et al., 1993) and of the CBD of its catalytic enzymes (Bayer et al., 1998). This domain facilitates the binding of cellulosomes to cellulose and contributes to its efficiency in degrading cellulose- containing substrates. Although there is no direct evidence that a scaffolding protein is 175 present in the putative cellulosome complex in P. tunicata, it is tempting to suggest that the approximately 46-48 kDa CBP might be a component of a cellulosome complex facilitating its attachment to cellulose. Cellulases facilitate the hydrolysis of cellulose and this has been reported to be brought about by the synergistic action of three types of enzymatic activities: a) endoglucanase, which acts at random in the polymeric chain b) exoglucanase, which liberates D-glucose and D-cellobiose and c) (3-glucosidase, which liberates D-glucose from cellodextrins (Schwarz, 2001). In the present study, the cellulolytic activity of P. tunicata was investigated. It was shown that the membrane proteins of cellulose-grown P. tunicata cells demonstrate cellulase activity when tested in gel diffusion and zymogram assays. This finding explains the ability of P. tunicata to utilize cellulose as an energy source for its growth. It also suggests that the enzymes are not freely released in the medium to hydrolyze the substrate. Instead, P. tunicata appears to have a bound cellulolytic enzyme complex similar to the cellulosomes found on the surface of cellulolytic Clostridium. There also appears to be a putative chitinase gene in P. tunicata (Chapter 2), which corresponds to one of the enzymes recently found to be present in cellulosome complexes of other bacterial species (Zverlov et al., 2002b). It suggests a correlation between cellulolytic activity and the appearance of cellulosome-like structures in cellulose-grown P. tunicata. Species of Pseudoalteromonas may have the means to utilize polymers in the marine environment as a nutrient source, as these bacteria produce the corresponding enzymes necessary to degrade them. For example, P. antartica, P. agarolyticus, P. carrageenovora and P. atlantica have been reported to produce extracellular agarases that hydrolyze agar, a polymer present in the cell wall of red algae (Akagawa- Matsushita et al., 1992; Vera et al., 1998). It has been suggested that the agarase activity expressed by Pseudoalteromonas species living in association with red algae helps them to secure nutrients from the algal surface (Holmstrom and Kjelleberg, 1999). Another polymer hydrolyzing enzyme previously found to be present in a Pseudoalteromonas species is chitinase produced by Pseudoalteromonas sp strain S91 (Techkamjanaruk and Goodman, 1999). Also, P. haloplanktis was recently found to produce a cold-active cellulase with endoglucanase activity (Violot et al., 2003). All of these species utilize free enzymes to degrade the target polymers. The species 176 P. tunicata appears to have a mechanism of polymer degradation, which is different from the aforementioned Pseudoalteromonas species. The bacterium produces cellulases which are bound on its cell surface possibly via a cellulosome complex. Aside from having cellulolytic activity, the bacterium attaches to the cellulose-based substrate via the cellulose-binding protein. This mechanism provides an advantage with respect to acquiring the soluble sugars as end products of polymer hydrolysis. The proximity of the cell to the substrate, allows efficient accumulation and assimilation of the hydrolyzed product. Additionally, the presence of two or more enzymes (e.g., cellulase and chitinase) not only generates more soluble products as a source of energy but would also offer the producing bacterium a competitive advantage over its competitors. The production of cellulolytic enzymes by P. tunicata may reflect a particular physiological response of the bacterium in the marine environment. It has recently been suggested that the production of extracellular enzymes (e.g., agarase, chitinase, cellulase, amylase, lipase, alginase), which degrade polymers and organic particulate matter in the marine environment, is a consequence of the specific habitat of some Pseudoalteromonas (Ivano et al., 2003). This suggests that the cellulose on the surface of U. lactuca and C. intestinalis from which P. tunicata has been isolated (Holmstrom et al., 1992; Egan et al., 2001a), may trigger cellulase production. While the utilization of the cellulose could lead to bacterial degradation of the host marine organisms, P. tunicata appears to have mechanisms to regulate its own population density (Chapter 3), and thereby prevent overgrowth of the bacterium on the surface of the host. Moreover, the cellulolytic enzyme activity of P. tunicata may be low enough to cause minimal damage to the algal host. In conclusion, P. tunicata expresses protuberant-like structures on its surface which are similar to the cellulosomes of cellulolytic clostridia, when grown in cellulose or cellobiose. This is the first report of a cellulosome like complex being produced by a marine bacterium. Apart from the MSHA pili mediated binding (Chapter 4), it was also demonstrated that a cellulose-binding protein mediates attachment to cellulose. The presence of cellulolytic enzymes aids P. tunicata in the conversion of polymers into substrates that are easily assimilated by the bacterium. In addition, this chapter supports the hypothesis that a mutual interaction may exist between P. tunicata and the higher marine organisms on which it is found, such as U. lactuca and C. intestinalis. This interaction would be mutually beneficial if P. tunicata protects its host against 177 biofoulers through the production of its reported bioactive compounds. The benefit to P. tunicata may be the domination of an ecological niche, as well as the provision of nutrients in the form of degraded algal surface polymers. The production of putative cellulosome complexes on the surface of P. tunicata may be a means by which it is able to survive in the nutrient-poor marine environment. Finally, the findings in this chapter add to our understanding of the ecological role, distribution and physiology of P. tunicata in the marine environment. Despite the evidence shown in this chapter to support the presence of a putative cellulosome-like entity in P. tunicata, additional studies are needed to provide further information and confirm the presence of cellulosomes in this bacterium. Such studies were not carried out due to time constraints of the thesis program. The additional studies should be aimed at identifying the cellulosome signature amino acid sequences (such as cohesin and dockerin domains). This could be achieved by using antibodies specific for the scaffolding protein from cellulolytic Clostridia, sequence candidate proteins and by generating primers based on the protein sequence information for PCR cloning and sequencing. There is also a need to investigate other enzyme activities such as xylanase, chitinase, glucosidase and exoglucanase which are part of cellulosome activities. Finally, TEM immunolabeling using the antibodies for cellulosome or saccharide- specific probes such as cationized ferritin is also recommended, as this will provide further evidence for the presence of a cellulosome-like entity on the cell surface of P. tunicata. 178 6. Summary and General Discussion The genus Pseudoalteromonas has been extensively studied during the last decade. Member species play an important role in the marine environment because of their frequent association with higher marine organisms and their ability to produce bioactive compounds that target a wide range of organisms (Holmstrom and Kjelleberg, 1999). P. tunicata has been used as a model organism in our laboratory, due to its ability to produce several inhibitory products that target a wide variety of surface colonizing organisms. Several studies, which were aimed at exploring the isolation, identification, characterization and regulation of these antifouling agents, have been performed. Also the association of P. tunicata with several higher marine organisms has led to investigations of the ecology and physiology of this bacterium in the marine environment. This thesis in particular has focused on the investigation of the mode of action of an antibacterial protein produced by P. tunicata, known as AlpP, including the colonization mechanisms and general adaptive responses of this bacterium to life in the surface environment. Novel information on the physiology of P. tunicata and its association with higher marine organisms is presented in this thesis. The studies of the mode of action of AlpP against P. tunicata provide a better understanding of the responses by the bacterium to its own product. A possible mechanism by which P. tunicata is able to protect itself against the toxic effects of AlpP has been suggested to be via the production of a putative antidote. The expression of AlpP and a putative antidote suggest that the bacterium is able to regulate its own population density. The self-regulatory mechanism is important in the biofilm formation of P. tunicata and could prevent bacterial overgrowth on the surfaces of host organisms. The discovery of an MSHA pilus in P. tunicata, a key colonization factor, may explain the mechanisms by which this organism is able to adhere to and colonize specific surfaces such as that of the green alga Ulva lactuca. Reports on cellulosomes are limited to a few bacterial genera and the finding of a putative cellulosome in P. tunicata, which appears to have a role in the conversion of cellulose into soluble products, is novel. It is particularly significant as it would be expected to influence the distribution of this bacterium on cellulose-based surfaces in the marine environment. As an important application, these mechanisms, as well as the ability of P. tunicata to produce several different inhibitory compounds, 179 provides a basis for developing strategies to control biofilm-formation and biofouling in the marine environment. The mode of action of AlpP was described in Chapters 2 and 3, through the generation and analysis of AlpP-resistant and -sensitive transposon mutants. Genes involved in the resistance and sensitivity mechanisms of P. tunicata to its own antibacterial product were identified. To further understand how P. tunicata colonizes the surfaces of higher marine organisms, Chapter 4 reported on the investigation of mechanisms involved in the attachment and colonization of P. tunicata to surfaces (biotic and abiotic). It was also shown that P. tunicata is able to grow in cellulose or cellobiose as the sole carbon source. This finding led to the experiments described in Chapter 5, which investigated how P. tunicata adapts to cellulose and utilizes this polymer for the production of cellulosome. This final chapter presents a summary of Chapters 2 to 5. Also, the main findings of this thesis are discussed and suggestions for further studies are provided. 6.1. The antibacterial activity of P. tunicata and the role of the autolytic protein, AlpP P. tunicata has been reported to display a broad range of antibacterial activity against several clinically important pathogens as well as marine isolates, including P. tunicata itself (James et al., 1996). To understand how P. tunicata competes with other surface colonizing bacteria, it is important to understand the mechanism of the antibacterial activity of P. tunicata and the role of its autolytic protein, AlpP. In addition, studies on the mode of action of AlpP against P. tunicata itself would provide information on the physiology of this bacterium, specifically with respect to the possible regulation of its own bacterial density. It was shown that AlpP is bacteriolytic both against itself and against other bacterial species in chapters 2 and 3. This may facilitate P. tunicata in the competition with other bacteria, possibly by directly attacking their membranes and inducing autolysis. Transposon mutagenesis studies were used to further clarify the autolytic action of AlpP. Several hypotheses were proposed to explain the responses of P. tunicata towards its own antibacterial protein based on the genes mutated in the Alp-sensitive and 180 -resistant mutants (see chapters 2 and 3). The proposed model that best explains the autolytic response of P. tunicata upon AlpP exposure is that AlpP may function as a signal for an autolysin-mediated response, in which AlpP is sensed by a two-component sensor regulator system, which induces the production of syringomycin-like peptide autolysins that are released from the cell via an ABC transporter protein (Figure 3-5). The suggestion that AlpP may function as a signal molecule for P. tunicata to produce autolysins is possible, as very small amounts of AlpP protein can result in autoinhibition (MIC 4pg/ml) - a feature that is consistent with the action of a signal molecule (James et al., 1996). An explanation for the resistance of P. tunicata to its own antibacterial product is linked to the production of extracellular structures. Two of the sensitive mutants were observed to have genes involved in the production of extracellular structures: mshJ, encoding for a MSHA pilus biogenesis protein and chi A, encoding for a chitinase, an enzyme present in the enzyme complex called the cellulosome. It is proposed that pili and cellulosomes (structures observed on the surface of P. tunicata, Chapters 4 and 5) prevent contact of AlpP with the bacterial cell, as these structures are closely bound and attached on the cell surface. Although the selected P. tunicata AlpP-sensitive mutants did not identify any genes encoding a putative antidote, P. tunicata appears to prevent self-killing through the expression of a protein antidote to counteract the toxic effect of AlpP. Evidence was obtained for a putative antidote produced simultaneously with AlpP, as is the case for the toxin-antidote system in E. coli involving the production of a stable protein toxin and an unstable protein antitoxin (Engelberg-Kulka and Glaser, 1999). The cell-free concentrated supernatant of P. tunicata was shown to display both inhibitory and protective activities. Confirmation was achieved by fractionation of the cell-free concentrated supernatant was carried out to separate the toxin and an antidote. It was found that the active fraction (AlpP alone) showed reduced activity when combined with fractions containing the putative antidote (non-toxic fraction). The results of this thesis and other reported studies on the activity and function of AlpP, suggest important ecological roles for AlpP in the physiology and survival of P. tunicata. For example, AlpP is involved in the mechanism by which P. tunicata is 181 able to regulate its own cell density. P. tunicata also produces a putative protein antidote to counteract the toxic effect of AlpP. The expression of AlpP most likely also facilitates P. tunicata in the competition with other bacteria in the marine environment. In addition, the expression of AlpP may represent programmed cell-death (PCD), where AlpP is produced during starvation as a “death signal” to induce autolysis, which results in killing of a portion of the bacterial population. This altruistic death could benefit the surviving cells given that nutrients are released. Cell death in biofilms of P. tunicata, as mediated by the autolytic protein AlpP (Mai-Prochnow et al., 2004), may serve a key role in the development of microcolonies and mature biofilms. It has been proposed that autolysis in bacterial biofilms allow for cellular differentiation and for individual cells to interact in a cooperative manner for biofilm development, a trait similar to that of development in higher multicellular organisms (Webb et al., 2003a). In fact, PCD in bacteria and higher eukaryotes display similar features including the involvement of toxin-antidote systems, reactive oxygen species (ROS) and oxidative stress (Webb et al., 2003a). The findings reported in chapters 2 and 3 could lead to other areas of research on the antibacterial activity of P. tunicata. The identification of a syringomycin synthetase-like gene in P. tunicata, which in Pseudomonas syringae is responsible for the biosynthesis of its peptide phytotoxins (Bender et al., 1999), suggests that a similar product may be synthesized by P. tunicata. Should such a peptide be identified in P. tunicata, the peptide gene cluster may be used to engineer new antibacterial compounds to combat bacterial pathogens. The putative two-component sensor and regulator system identified in P. tunicata needs to be explored to reveal whether it is involved in other specific adaptive responses of the bacterium. The proposal that the expression of AlpP is a PCD mechanism should also be further investigated, as P. tunicata would serve as a model of PCD for surface associated and biofilm forming marine bacteria and would potentially provide information for developing means of controlling biofilms on surfaces in waters. Several studies investigating the antibacterial activity of P. tunicata are currently being carried out in our laboratory. For example, the role of AlpP in mixed biofilm formation, in P. tunicata biofilm differentiation and dispersal and the mechanisms of secretion and regulation of AlpP is under investigation. 182 6.2. The role of the MSHA pilus in the colonization of living surfaces by P. tunicata P. tunicata has been found to be associated with the higher marine organisms, Ulva lactuca and Ciona intestinalis (Holmstrom et al., 1992; Egan et al., 2001a; Shovhus et al., 2004), but no data have so far been reported on the means by which P. tunicata is able to specifically colonize these surfaces. However, characterization of one of the P. tunicata transposon mutants, SM5 (mshJ mutant) disclosed one colonization mechanism used by the bacterium (Chapter 4). Phenotypic, genotypic and ultrastructural studies showed that P. tunicata expresses a mannose-sensitive hemagglutinin (MSHA) pilus used as an attachment factor. The gene cluster proposed to be involved in the biogenesis of the MSHA pilus is homologous to the gene locus of the MSHA biogenesis protein located in some Vibrio species including V. cholerae, V. parahaemolyticus and V vulnificus. In V cholerae, this pilus has been demonstrated to facilitate the attachment and colonization of the bacterium to different surfaces in marine waters. The gene locus encoding this structure has been termed “environmental persistent island” in V. cholerae (Marsh and Taylor, 1999). The MSHA pilus of P. tunicata may function similarly as a key colonization factor. This structure facilitates the attachment of P. tunicata and may mediate the formation of microcolonies. P. tunicata also appears to have a sensing mechanism associated with pilus expression, since it was shown that the production of pili increased when cells were grown in cellulose, a major surface polymer of U. lactuca, possibly mediated through the activity of wmpR, a putative transcription regulatory gene (Egan et al., 2002b). In the marine environment, cellulose or surface polymers of C. intestinalis and U. lactuca could facilitate the attachment of P. tunicata to the surfaces of these marine organisms by triggering the production pili. A gene cluster for the biogenesis of a putative MSHA pilus in P. tunicata has been identified in this thesis. However, further studies are needed to identify other additional genes in the operon for example the structural genes for the MSHA pilus, as this would classify the pilus type. The surface sensing mechanism as demonstrated by P. tunicata is valuable for future research pertaining to the ecology of the bacterium. Moreover, investigations of the putative role of the MSHA pilus in P. tunicata biofilm formation 183 are recommended because such studies would provide information about the architecture and mechanisms of biofilm formation at surfaces. 6.3. The role of a putative cellulosome in the hydrolysis of cellulose by P. tunicata In Chapter 5, it was proposed that P. tunicata converts cellulose into soluble products through the activity of a cellulase enzyme contained within a cellulosome complex. It was shown in Chapter 4 that P. tunicata is able to grow in cellobiose or cellulose as the sole carbon source, suggesting that it has a means of converting these substrates into solutes that can be assimilated by the cell. Ultrastructural studies showed that P. tunicata produces protuberant-like structures which are similar to the cellulosomes of cellulolytic Clostridium species (Bayer and Lamed, 1986; Doi and Tamaru, 2000; Kakiuchi et al., 1998; Belaich et al., 1997). This is an enzyme complex consisting of a scaffolding protein, which facilitates binding of the cell to the substrates through its cellulose binding domain, and catalytic enzymes which act synergistically to degrade the polymeric substrate (Bayer et al., 1998). P. tunicata was shown to demonstrate cellulolytic activity, which converts cellulose into soluble products. Additionally, this marine bacterium was also shown to produce a cellulose-binding protein, which would facilitate in the attachment of the cells to cellulose. The peptide sequence of this protein showed similarities with amino acid segments of endoglucanase of C. acetobutylicum. Although, this peptide sequence has low confidence hits, the similarities observed may support that the 46-48 kDa protein is a cellulose-binding protein (CBP). Most CBPs from cellulolytic bacteria have cellulase activities (Miron and Forsberg, 1998). No genetic evidence presented in this thesis proved that the protuberant-like surface structures in P. tunicata are cellulosomes, however their ultrastructural appearance and the presence of catalytic (cellulase) activity and binding components (cellulose-binding protein), suggest that this may be the case. Additionally, the result of the peptide sequencing analysis of the 46-48 kDa protein, while also not conclusive, supports the hypothesis that cellulosomes are present in P. tunicata. One of the peptide sequence matches with the amino acid segments of C. acetobutylicum endoglucanase, an enzyme present in the cellulosome complex of this bacterium (Sabathe et al., 2002). Additional support for the presence of cellulosome derives from the observation that the production 184 of cellulosome-like structures is induced in the presence of cellulose. The possible presence of cellulosomes on the cell surface of P. tunicata is novel, as they have so far only been observed in a restricted number of anaerobic Gram-positive bacteria. Isolation and characterization of the cellulosome-like structures are required to identify the structural subunits and to compare these with the cellulosome subunits of cellulolytic clostridial cells. This would also provide the basis for identifying the scaffolding protein, which contains the cellulosome signature amino acid sequences (such as cohesin and dockerin domains). Such a comparison could be achieved by using antibodies specific for the scaffolding protein from cellulolytic clostridia, sequence candidate proteins and by the generation of primers based on the protein sequence information for PCR cloning and sequencing. TEM immunolabeling studies using the antibodies for cellulosome or saccharide-specific probes such as cationized ferritin would also provide further evidence of the nature of cellulosomes in P. tunicata. Additional studies aiming at investigating the presence of other enzyme activities in P. tunicata, such as xylanase, chitinase, glucosidase and exoglucanase are also needed. Finally, the development of specific P. tunicata probes based on the 16S rRNA gene (T. Skovhus, personal communication) would facilitate studies of P. tunicata on marine organisms. 6.4. A hypothetical model for the occurrence and persistence of P. tunicata on the surfaces of U. lactuca and C. intestinalis Based on the findings presented in this thesis a hypothetical model of the colonization of P. tunicata on surfaces of higher marine organisms such as U. lactuca and C. intestinalis is proposed (Figure 6-1). This model suggests that planktonic P. tunicata cells in search for surfaces and nutrients attach to the surfaces of U. lactuca and C. intestinalis using the MSHA pili. The choice of host organisms may be mediated by P. tunicata cells sensing the cellulose, a major surface polymer of these marine organisms. Cellulose may subsequently act as an inducer for the attachment of P. tunicata. This is supported by the finding that cellulose was found to induce an increased expression of pili. Once 185 attached, the MSHA pili appear to facilitate the formation of microcolonies and subsequent biofilm formation. Nutrients are needed to maintain the bacterial population and it is possible that carbon is obtained from the hydrolysis of the surface polymer (cellulose) of the host organisms. P. tunicata cells appear to produce a putative enzyme complex that converts the cellulose to soluble products and subsequently utilize them as a source of carbon and energy. The benefit for the host organisms might be that P. tiiniccitci protects the host against biofouling by producing an array of antifouling compounds. Importantly, P. tunicata can also control its population to prevent overgrowth and detrimental effects to the host organisms. The bacterial population is controlled and regulated by a mechanism involving the expression of AlpP, a protein that may function as an autolytic signal, and a putative antidotal protein that counteracts the autolytic activity of AlpP. According to this hypothesis, a portion of the bacterial population undergoes lysis, leaving behind live and resistant cells, thus maintaining a bacterial community on the surface of the host organisms. In conclusion, the thesis has identified several distinctive mechanisms to explain the occurrence, survival and persistence of P. tunicata in marine surface ecosystems. It includes mechanisms involved in colonization, surface sensing and the regulation of population density. The findings of this work may also contribute to a better understanding of the ecology, physiology, and distribution of P. tunicata in marine environments and address aspects of its symbiotic relationship with higher marine organisms. 186 Surface of Viva lacluca or Ciona intestinal is A) Planktonic B) Surface sensing and C) Microcolony D) Adaptation E) Control and regulation of cell cells in search colonization via its formation which response to density through the expression of for space and MSHA pilus. is facilitated by surface; AIpP and hypothetical antidote, as nutrients MSHA pilus. formation of shown by the dead cells (red) and cellulosome-like live and persistant cells (green). entity (see magnified figure) and production of bioactive compounds Figure 6-1. A hypothetical model of the occurrence and persistence of P. tunicata on surfaces of U. lactuca and C. intestinalis. See text in section 6.4 for discussion 187 Appendix I Solutions and Buffers LI Nine Salts Solution (NSS) (per litre) 17.6 gNaCl, 1.47 g Na2S04, 0.08 g NaHC03( 0.25 g KC1, 0.04 g KJBr, 1.87 g MgCl2.6H20, 0.41 g CaCl2.2H20, 0.008 g SrCl2.6H20, 0.008 g H3BO3, adjust to pH 7 I.II VNSS (per litre NSS) (Marden et aL 1985) 1.0 g peptone, 0.5 g yeast extract, 0.5 g glucose, 0.01 g FeS04.7H20, 0.02 0.01 g Na2HP04, . for agar plates add 15 g agar before autoclaving I.III Marine Minimal Medium (MMM) 920 ml 1.1 x NSS (i.e. salts for one litre in 920 ml H20) autoclaved 40 ml 1 M MOPS (pH 8.2) sterile filtered 10 ml 0.4 M Tricine + FeS04.7H20 (pH 7.8) sterile filtered 10 ml 132 mM K2HP04 autoclaved (add slowly while stirring) 10 ml 952 mM NH4C1 (pH 7.8) autoclaved 10 ml 20% Trehalose, sterile filtered 188 I.IV Luria Broth (LB) medium (per litre) LB 10 lOgNaCl, 10 g tryptone, 5 g yeast extract -adjust to pH 7.5 -for agar plates add 15 g agar before autoclaving LB 20 20 g NaCl, 10 g tryptone, 5 g yeast extract -adjust to pH 7.5 -for agar plates add 15 g agar before autoclaving I.V. Krebs-Ringer solution (KRT) (per 1 of 10 mM Tris-HCl„ pH 7.4) 7.5 g NaCl 0.383 gKCl 0.318 MgSO4.7H20 0. 305 g CaCl2 1. VI Phosphate Buffer Solution (PBS), pH 7.4 (per liter) 8.00 g NaCl 0.20 g KC1 1.44 g Na2HP04 0.24 g KH2P04 I. VIII 5X TBE buffer (per litre) 54 g Tris base 27.5 g boric acid 20 ml 0.5M EDTA solution (Ph 8.0) 189 I.IX XS Buffer 0.5 g potassium ethyl xanthogenate 10 ml 4M ammonium acetate 5 ml 1M Tris-HCl, pH 7.4 2 ml 0.45M EDTA 2.5 ml 20% SDS add dist water to make 50 ml I.X Z buffer (Miller, 1972) (per liter) 16.1 g Na2HP04-7H20(0.06 M) 5.5 g NaH2P04-H20(0.04 M) 0.75 g KC1 (0.01 M) 0.246 g MgS04-7H20(0.05 M) 2.7 ml P-mercaptoethanol (0.05 M) 190 Appendix II II.I RNase treatment of DNA 1. Prepare a stock solution of RNase at a concentration of 10 mg/ml in 10 mM Tris- HC1, pH 7.5 and 15 mM NaCl. Boil for 15 minutes and cool slowly to room temperature before storing at -20°C. 2. For every 100 pi of DNA solution to be treated, add 1 pi of the previously boiled RNase stock solution. 3. Incubate the mixture at room temperature for 30-60 min. 4. Remove the contaminant RNA from DNA solution by following the phenol:chlorofom:isoamylalcohol extraction and ethanol precipitation. II.II Phenol:chlorofom:isoamylalcohol extraction 1. Place the DNA solution in an Eppendorf tube and add a minimum of 300 pi of milli-Q water. 2. Add an equal amount of phenol:chloroform:isoamylalcohol (25:24:1 (v/v/v)) and mix thoroughly for 1-3 min. 3. Centrifuge the mixture at 14 000 x g for 5 minutes at 15°C to separate the phenol. 4. Transfer the upper aqueous phase (DNA solution) into a fresh Eppendorf tube. 5. Repeat the extraction three times and only use chloroform in the last extraction to remove traces of phenol. II.Ill Ethanol precipitation 1. Add 1/10 volume of ice-cold 3 M sodium acetate (pH 5.2) solution to the cleaned DNA solution. 2. 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