Epiphytic Bacteria on the Green Alga Ulva Australis: Biofilm Formation and Ecology

Epiphytic bacteria on the green alga Ulva australis: biofilm formation and ecology

Dhana Rao

A thesis submitted to the University of New South Wales for the degree of Doctor of Philosophy

2005 Abstract

Most surfaces immersed in seawater, including those of many living marine organisms rapidly become colonised by a complex community of fouling organisms. Algae however have evolved a number of antifouling defences and it has been suggested that the green alga Ulva australis exploits antifouling compounds produced by epiphytic bacteria such as Pseudoalteromonas tunicata and Roseobacter gallaeciensis.

The ecology of colonisation and competitive interactions were studied in order to understand the role of these epiphytic bacteria on the surface of the marine alga U. australis. It was found that P. tunicata and R. gallaeciensis exhibited different colonisation strategies. Whilst R. gallaeciensis is capable of colonising the surface of U. australis under a range of conditions, colonisation by P. tunicata is enhanced by high cell densities, presence of cellobiose in the pre-culture, inoculation in the dark and interactions with a natural seawater community in order to attach and persist on the surface of the algae. In competition studies on both artificial and living surfaces, P. tunicata outcompeted other marine strains isolated from U. australis, and was in turn outcompeted by R. gallaciensis. This study was the first to demonstrate that inhibitory compounds produced by marine bacteria can provide a competitive advantage during biofilm formation.

In order to investigate the number of surface attached bacterial cells necessary to prevent fouling, settlement assays were performed with micro- and macrofouling organisms. Assays demonstrated that remarkably low cell densities ( 102-105 cells cm-2) of P. tunicata were effective in preventing settlement of algal spores, marine fungi and invertebrate larvae in Petri dishes. Similarly, low cell densities (103-104cells cm-2) of R. gallaeciensis had anti-larval and anti-bacterial activity, indicating that P. tunicata and R. gallaeciensis can play a role in the defence against fouling on U. australis at cell densities that commonly occur in vivo.

The final part of this thesis aimed to determine whether P. tunicata and R. gallaeciensis are able to invade a complex community or a synergistic mixed species biofilm. It was

11 observed that P. tunicata can colonise a complex seawater community on glass surfaces at in situ densities, but was unable to colonise an epiphytic community. A synergistic mixed species biofilm was able to resist invasion by P. tunicata. However, R. gallaeciensis was an aggressive coloniser and able to dominate an epiphytic community on U. australis and a synergistic mixed species biofilm on both inanimate and animate surfaces.

This study demonstrates that microbial colonisation of the plant surfaces is a dynamic process where differences in attachment, colonisation and competitive biofilm formation can markedly affect the establishment and organisation of epiphytic microbial communities. The results presented here suggest that P. tunicata and R. gallaeciensis may indeed provide an antifouling defence to U. australis.

Ill Acknowledgements

The successful completion of this project was possible because of the help I received from many people. Saying thank you does not do justice to the gratitude I feel for my supervisor Staffon Kjelleberg, for his advice, encouragement and enthusiasm. He has been an excellent supervisor and scientific role model.

Many thanks to Jeremy Webb, my co-supervisor, who has also been brilliant. He has been an inspiration, mentor, and most importantly a good friend. I am grateful to him for providing perceptive, painstaking and generously detailed responses to drafts of this work and for his endless encouragement.

I would like to thank Carola Holmstrom for her advice on assays, proofreading various chapters of this thesis, for her support and kindness and being such a wonderful friend.

I am greatly indebted to all those who collaborated in this project, including Peter Steinberg, Ingela Dahllof, Mette Burm0lle and Torben Skovhus. They offered crucial support and advice at different junctures. Likewise, Carola Holmstrom and Niina Tujula are to be thanked for expert and enjoyable assistance in the field. Su Egan, Charles Svenson and Ashley Franks are also to be thanked for the use of their strains.

Many thanks are also due to Diane, Scott, Su and Torsten for their advice and encouragement. Diane has also been a fantastic office mate and friend.

I would like to thank my wonderful friends Johnny, Anne and Kregor for all their help and encouragement. Evi, Doralyn, Bee, Flavia, Sacha, Niina, Sharon, Kirsty, Nick and Sohail are also to be thanked for their friendship and support which has made the whole PhD experience a lot more enjoyable.

I would also like to thank all the present and past members of the SK lab for their friendship over the last few years, particularly Sally, Maurice, Mary, Lan, Mathew, Ashley, Dacre, Nidhi. Leena, Maria, Kin, Andre, Alicia, Ana Maria, Anni, Greg, Mike, David, Muoi, Caarsten and Marcus.

lV I must also thank members of the PS Lab for their assistance and advice: Mike, Sharon, Megan, Anis, Paul, Jacinta, Bee Swanson, Jennifer, Tim, Menuk, Neda and Lachlan,

I would also like to thank Nick Paul for help with statistics who devoted quite a bit of time teaching me how to analyse data.

Bill O'Sullivan is to be thanked for his proof-reading of this thesis and Paul Halasz for assistance with microscopy.

A heartfelt thanks to Adam and Julie, for their help in all things administrative and also their encouragement and friendship.

I would also like to thank Sharat Babu for his advice and help.

No amount of gratitude can ever be enough for my family whose unquestioning faith, support and understanding were unwavering. I will be eternally grateful to Mum, Dad and Hem for their love and encouragement, without which I would never have accomplished this. To them I dedicate this thesis.

V Table of Contents

Abstract 11

Acknowledgements lV

Table of Contents Vl

List of Figures IX

List of Tables Xl

Chapter One: General Introduction ...... 1 1.1 Background ...... 1 1.2 Establishment of biofilms and their role in biofouling ...... 3 1.2.1 Bacterial biofilms produce cues for macrofouling organisms 4 1.3 Bacterial biofilm development ...... 7 1.3 .1 Attachment ...... 7 1.3.2 Microcolony formation ...... 9 1.3.3 Maturation ofbiofilms ...... 10 1.3.4 Detachment ...... 11 1.4 Microbial interactions in biofilms ...... 13 1.4.1 Cooperation ...... 14 1.4.2 Competition in biofilms ...... 14 1.4.3 Antagonism ...... 15 1.4.4 Inhibitory compounds produced by marine bacteria ...... 15 1.4.5 Succession ...... 17 1.5 The organisms ...... 18 1. 5 .1 Ulva australis ...... 18 1.5.2 Pseudoalteromonas genus ...... 19 1.5.3 Roseobacter clade ...... 22 1.6 Ai1ns of this study ...... 24 1. 7 References ...... 25

Chapter Two: Competitive interactions in mixed-species biofilms containing the marine bacterium Pseudoalteromonas tunicata ...... 47 2.1 Introduction ...... 47 2.2 Materials and methods ...... 49 2.2.1 Isolation of marine strains ...... 49 2.2.2 Fluorescent labelling of marine isolates ...... 50 2.2.3 Biofilm experiments ...... 51 2.3 Results and Discussion ...... 52 2.3.1 Characterisation of marine strains ...... 52 2.3.2 Mono-species biofilm development ...... 55 2.3.3 Mixed-species biofilm development ...... 60 2.3.4 Role of the P. tunicata antibacterial protein AlpP in competitive biofilm development ...... 60 2.3.5 Role of microcolonies in competitive interactions within biofilms 62 2.4 Conclusions ...... 63 2.5 References 65

Vl Chapter Three: Microbial colonisation and competition on the marine alga Viva australis ...... 70 3.1 Introduction ...... 70 3.2 Materials and methods ...... 73 3 .2.1 Collection of plants and generation of axenic U. australis ...... 73 3.2.2 Colonisation experiments ...... 73 3.2.3 Effect of multispecies consortia on colonisation ...... 75 3.2.4 Competition in dual-species biofilms on U. australis ...... 75 3.3 Results ...... 77 3.3.1 Obtaining axenic plant tissue ...... 77 3.3.2 Factors influencing attachment and colonisation ...... 77 3.3.3 Competitive biofilm development on U. australis ...... 84 3.4 Discussion ...... 85 3.4.1 Effect of cell densities on attachment ...... 85 3.4.2 Attachment in the dark ...... 86 3.4.3 Effect of cellobiose on attachment ...... 86 3.4.4 Synergistic biofilm formation ...... 87 3.4.5 Competition in biofilms on U. australis ...... 88 3.5 Conclusions ...... 89 3.6 References 91

Chapter Four: Effects of bacterial cell density on antifouling by epiphytic marine bacteria ...... 96 4.1 Introduction ...... 96 4.2 Materials and Methods ...... 98 4.2.1 Preparation of marine strains ...... 98 4.2.2 Establishment ofbiofilms ...... 99 4.2.3 Anti-algal bioassays ...... 100 4.2.4 Anti-larval assays ...... 101 4.2.5 Antifungal assays ...... 101 4.2.6 Antibacterial assays ...... 101 4.2. 7 Statistical analysis ...... 102 4.3 Results and discussion ...... 102 4.3.1 Effect of bacterial biofilms on an inanimate surface ...... 103 4.3.2 Effects ofbiofilms on the surface of U. australis ...... 118 4.3.3 Density and antifouling activity ...... 120 4.4 Conclusions ...... 121 4.5 References ...... 122

Vll Chapter Five : Effects of introducing inhibitory bacteria into mixed species marine biofilms ...... 127 5.1 Introduction ...... 127 5.2 Materials and Methods ...... 129 5.2.1 Establishment of a natural seawater community on glass slides 129 5.2.2 Biofilm sampling and DNA extraction of the natural seawater community on glass surfaces ...... 130 5.2.3 Diversity of the natural seawater community on glass slides 130 5.2.4 Microscopy of the natural seawater community on glass slides 131 5.2.5 Epiphytic communities on U. australis ...... 131 5.2.6 Synergistic mixed species biofilms established on glass surfaces 132 5.2.7 Synergistic mixed species biofilms on U. australis ...... 133 5.3 Results ...... 134 5.3.1 Diversity of the natural seawater community on glass surfaces 134 5.3.2 Morphology of the natural seawater community on glass surfaces 135 5.3.3 Introducing P. tunicata and R. gallaeciensis into an epiphytic community on U. australis ...... 136 5.3.4 Mixed species biofilms established on glass surfaces ...... 136 5.3.5 Synergistic mixed species biofilms on U. australis ...... 142 5.4 Discussion ...... 142 5.4. l Natural seawater community on glass surfaces ...... 142 5.4.2 Epiphytic community on U. australis ...... 144 5.4.3 Invasion of synergistic mixed species biofilms ...... 145 5.5 Conclusions ...... 146 5.6 References 148

Chapter Six : General Discussion ...... 151 6.1 Host-bacterial interactions ...... 151 6.2 Interactions within biofilm communities ...... 154 6.3 Interactions between bacteria and fouling organisms ...... 157 6.4 Future directions ...... 159 6.5 References ...... 160

Vlll List of Figures

Fig. 2.1 Single-species biofilm development for bacteria isolated from the 56 marine alga Ulva australis

Fig. 2.2 Competitive biofilm development in two-species biofilms 57 containing Pseudoalteromonas tunicata and other epiphytic bacterial isolates

Fig. 2.3 Competitive bioifilm development by Pseudoalteromonas 58 tunicata AlpP mutant that does not produce the antibacterial protein

Fig. 2.4 Competitive biofilm development in pre-established biofilms in 58 glass flow cells

Fig. 3.1 Effect of axenic treatment on the Ulva australis epiphytic 81 community

Fig. 3.2 A comparison of attachment and colonisation by 81 Pseudoalteromonas tunicata grown in cellobiose as compared to glucose

Fig. 3.3 A comparison of colonisation of Ulva australis by 82 Pseudoalteromonas tunicata cells suspended in sterile seawater, natural seawater and in combination with a mixture of epiphytic strains

Fig. 3.4 Competitive biofilm development in pre-established biofilms on 83 the surface of Ulva australis

Fig 4.1 Pseudoalteromonas tunicata biofilms established at different 106 densities on plastic Petri dishes inhibit Polysiphonia sp. spore settlement

Fig. 4.2 Roseobacter gallaeciensis biofilms on plastic Petri dishes do not 107 affect Polysiphonia sp. spore settlement

Fig. 4.3 Pseudoalteromonas tunicata biofilms established at different 108 densities on plastic Petri dishes inhibit Ulva australis spore settlement

Fig. 4.4 Roseobacter gallaeciensis biofilms established at different 109 densities on plastic Petri dishes stimulate Ulva australis spore settlement

Fig. 4.5 Synthetic acyl homoserine lactoness (AHLs) stimulate Ulva 110 australis spore settlement

IX Fig. 4.6 High density Pseudoalteromonas tunicata biofilms on plastic 111 Petri dishes inhibit Bugula neretina larval settlment

Fig. 4.7 Roseobacter gallaeciensis biofilms established at different 112 densities on plastic Petri dishes inhibit Bugula neretina larval settlement

Fig. 4.8 Pseudoalteromonas tunicata biofilms established at different 112 densities on plastic Petri dishes inhibit fungal attachment

Fig. 4.9 Roseobacter gallaeciensis biofilms established at different 113 densities on plastic Petri dishes inhibit bacterial attaachment

Fig. 4.10 Roseobacter gallaeciensis biofilms established on the surface of 114 Ulva australis stimulate Ulva australis spore settlement

Fig.4.11 Roseobacter gallaeciensis biofilms established on the surface of 115 Ulva australis inhibit bacterial attaachment

Fig. 4.12 Pseudoalteromonas tunicata biofilms established on the surface 116 of Ulva australis inhibit fungal attachment

Fig. 5.1 A comparison of DGGE profiles of eubacterial communities in 137 the absence of Pseudoalteromonas tunicata and with the addition of low and high densities of P. tunicata

Fig. 5.2 A comparison of DGGE profiles of Pseudoaltermonas 138 communities in the three biofilm system

Fig. 5.3 A comparison of invasion of the epiphytic community by 139 Pseudoalteromonas tunicata, P. tunicata AlpP mutant and Roseobacter gallaeciensis

Fig. 5.4 Invasion of a synergistic mixed species biofilm by 140 Pseudoalteromonas tunicata

Fig. 5.5 Invasion of a synergistic mixed species biofilm by Roseobacter 141 gallaeciensis

X List of Tables

Table 2.1 16S rRNA gene identification of bacteria isolated from the 53 marine alga Ulva australis

Table 2.2 Drop test activity for the detection of extracellular inhibitory 53 compounds active against Pseudoalteromonas. tunicata and sensitivity of each strain to the P. tunicata antibacterial protein AlpP

Table 3.1 Factors influencing attachment and colonisation of 78 Pseudoalteromonas tunicata on Ulva australis

Chapter One

General Introduction

1.1 BACKGROUND

Surfaces immersed in marine waters become rapidly colonised by orgamsms - a process known as biofouling. Biofouling is an ancient problem and it has plagued mariners since boats were put into water. The problem goes beyond just mere aesthetics - naval battles have been won and lost based on the effects of fouling on ship speed and manoeuvrability. Biofouling of ships hulls and other artificial surfaces remains a major problem to this day and the economic and environmental impact on global maritime and coastal industries is severe.

All submerged surfaces, both natural and artificial, become fouled in the marme environment. Macroalgae are particularly susceptible to fouling because they are sessile and restricted to the photic zone where conditions for fouling are optimal (de Nys et al., 1995). Also, as space is often a limiting resource, their large surface areas present a relatively long-lived and stable habitat for colonisation (Seed and O'Connor, 1981). Besides providing surfaces for attachment, algae can also be a source of nutrients and may provide shelter for epibionts. But biofouling communities on surfaces of marine plants car. have detrimental effects on the host organism. Fouled seaweeds can experience reduced rates of growth, photosynthesis and reproduction due to increased shading and competition for nutrients (Schmitt et al., 1995; Williams, 1996). Further, fouling by calcareous epibionts causes an increase in weight and reduces buoyancy and enhances breakage in a turbulent environment (D'Antonio, 1985).

While some seaweeds are heavily fouled, other species in the same habitat remain remarkably free from fouling, indicating that they possess defence mechanisms against colonising organisms. Marine algae have evolved a number of ways to protect themselves from fouling, primarily by avoidance and physical and chemical defence mechanisms. The brown algae Laminaria digitata and L. saccharina grow rapidly enough to produce tissue at a rate higher than the local fouling rate. In this way, a photosynthetically active zone is maintained at the meristem, while fouling is restricted to the older parts of the thallus (Russell, 1983; Russell and Veltkamp, 1984). Physical defences include the production of mucus which serves to remove newly settled epiphytes as the mucus sloughs away (Barthel and Wolfrath, 1989), continuous shedding of outer layers of cells (Johnson and Mann, 1986; Holmstrom and Kjelleberg, 1994; Keats et al., 1997; Dobretsov and Qian, 2002) and entire blades being abandoned (Littler and Littler, 1999).

The red alga Graci/aria conferta utilises chemical defence by releasing a burst of hydrogen peroxide and other active oxygen species (Weinberger and Friedlander, 2000a; Weinberger et al., 2005). This response is induced by the presence of agar oligosaccharides, which are degradation products from its own cell walls, and leads to a substantial decrease in bacterial epiphytes (Weinberger and Friedlander, 2000b). Another well studied red alga, Delisea pulchra, produces a class of compounds called furanones that mimic and interfere with the quorum-sensing systems of bacteria (Kjelleberg et al., 1997). It was shown that the furanones interfered with communication between bacterial species during colonisation of the algal thalli through competition with cognate signal molecules (Givskov et al., 1996). Furanones also behave as an acyl homoserine lactone (AHL) blocker in the non marine bacteria Pseudomonas aeruginosa and a synthetic furanone was found to affect the architecture of P. aeruginosa biofilms and enhance detachment (Hentzer et al., 2003). AHL mimics have subsequently been reported in the unicellular green alga Chlamydomonas reinhardtii (Peters et al., 2003; Teplitski et al., 2004) and higher plants (Teplitski et al., 2000; Mathesius et al., 2003). However, in contrast to furanones from D. pulchra, many of the AHL mimics from plants stimulate quorum sensing-regulated responses m bacteria. AHL mimics such as the furanones have attracted significant interest as possible alternatives to antibiotics because of their ability to interfere with bacterial biofilm development.

Although effective, algal defences are thought to be costly as they divert limiting resources away from the primary functions of growth, resource capture and reproduction (Bazzaz et al., 1987). It has been proposed that some algae may rely on

2 microbial mediated defence by exploiting compounds produced by the bacterial community on their surface. This would protect the host by interfering with the development of a mature biofouling community. Such interactions have been described for the marine crustaceans Palaemon macrodactylus and Homarus americanus where symbiotic bacteria have been shown to defend embryos from a lethal fungal infection (Gil-Turness et al., 1989; Gil-Turness and Fenical, 1992). Furthermore, bacteria isolated from marine algae have been shown to form biofilms on artificial surfaces that strongly inhibit settlement of macrofouling organisms (Maki et al., 1989; Holmstrom and Kjelleberg, 1994; Neal and Yule, 1994a,b), suggesting that analogous biofilms on eukaryotic surfaces may contribute to the fouling defence of their hosts.

Such defence strategies have led to considerable interest in exploring whether "natural" mechanisms could be adapted to protect artificial surfaces. Current control strategies for biofouling in marine and coastal industries are sometimes only partially effective, and in many cases introduce toxic chemicals into the environment. Therefore, environmental concerns and increasing regulatory requirements mean that there is a need to develop alternative antifouling technologies to prevent or minimise biofouling. An understanding of the dynamic processes involved in biofouling, may lead to the development of novel environmentally benign technologies, for the control of fouling in marine environments. One approach to controlling attachment of macrofouling organisms would be to manipulate bacterial biofilms so that they are inhibitory to invertebrate larvae and algal spores. In order to do this, a better understanding of biofilm formation and the processes that determine the composition of bacterial communities is required. Also, understanding colonisation strategies by epiphytic bacteria and their interactions with the host, as well as their efficacy in deterring the settlement of macrofouling organisms, would be an important step in determining the role of epiphytic microbial communities on undefended algal surfaces.

1.2 ESTABLISHMENT OF BIOFILMS AND THEIR ROLE IN BIOFOULING

Biofouling is a dynamic process and the sequence of events leading to the formation of a biofouling community has been well studied (Wahl, 1989; Clare et al., 1992). Within minutes of immersing a clean surface in seawater, it adsorbs a conditioning film, which

3 is made up of organic molecules (Marshall, 1985). Once the conditioning film is in place, bacteria and single-celled diatoms may encounter the surface and attach. Attached cells divide rapidly and form colonies, which eventually coalesce into a compact biofilm, representing the second stage of fouling. The biofilm community is a mixture of adsorbed macromolecules, attached bacteria and diatoms enmeshed in a matrix of extracellular polymers (Mihm et al., 1981 ). Biofilm formation on a marine substratum modifies its chemistry (Characklis and Cooksey, 1983; Anil and Khandeparkar, 1998), and mediates the settlement of a large number of secondary fouling organisms (Steinberg et al., 2002). The different phases of bacterial biofilm formation are described in more detail in Section 1.3.

1.2.1 Bacterial biofilms produce cues for macrofouling organisms

The vast majority of marine invertebrates have complex life cycles that alternate between a sessile, benthic adult phase and a planktonic larval phase. The larval phase of marine invertebrates can last anywhere from hours (Thiyagarajan et al., 2003) to years (Chiswell et al., 2003). The process whereby larvae are able to locate a suitable habitat for the benthic adult is a key stage in the life cycle.

Settlement of invertebrate larvae is affected by many physical factors such as light, hydrophobicity, texture and orientation of the surface. However, the most important determinants of metamorphosis in competent larvae (referred to as settlement in this study) are specific chemical cues associated with the substratum (Kirchman et al., 1982b; Maki and Mitchell, 1985; Szewzyk et al., 1991b; Pawlik, 1992; Pechenik and Qian, 1998; Qian et al., 2000). Cues include the presence of other invertebrates of the same species (conspecifics) (Pawlik, 1992), plant compounds (Williamson et al., 2000), and bacterial biofilms (Tamburri et al., 1992). Selective settlement on biofilms has been shown for larvae of marine invertebrates from a number of phyla, including the polychaetes Janua brasilliensis (Kirchman et al., 1982b ), and Hydroides elegans (Huang and Hadfield, 2003), the bryozoan Bugula neritina (Maki et al., 1989), the barnacle Ba/anus amphitrite (Wieczorek et al., 1995) and the ascidian Ciona intestinalis (Szewzyk et al., 1991 b ). Biofilm cues utilised by invertebrate larvae include water borne products (Neumann, 1979), and substances associated with the

4 bacterial cell surface (Kirchman et al., 1982a; 1982b; Maki and Mitchell, 1985a; Szewzyk et al., 1991b).

Most larvae prefer to settle on substrata that possess a well developed biofilm (Clare et al., 1992), possibly because biofilms reflect aspects of prevailing environmental conditions (Neal and Yule, 1994a,b) and may indicate a relatively stable underlying substratum with a good water flow regime above the surface (Wieczorek and Todd, 1998). The presence of a biofilm may be a general signal that a surface is suitable and larvae may use more specific chemical signatures from characteristic microbial assemblages to indicate preferred ecological conditions at a site {Lau and Qian, 1997; Unabia and Hadfield, 1999; Lau et al., 2002). Many larvae are able to distinguish between biofilms of varying composition, physiological condition and growth phase (Neumann, 1979; Maki et al., 1988; Pearce and Scheibling, 1991; Szewzyk et al., 1991a; Holmstrom et al., 1992; Wieczorek et al., 1995) indicating that bacteria serve as important signposts for larvae seeking a settlement substratum. The inhibitory or stimulatory effects of bacterial biofilms are amplified with age of the biofilm (Maki et al., 1989; Pearce and Scheibling, 1990; Holmstrom et al., 1992; Wieczorek et al., 1995; Wieczorek and Todd, 1998; Hamer et al., 2001) and this seems to be correlated with the density of cells in biofilms, with older biofilms being more attractive for settlement (Hadfield and Paul, 2001 ). Research in some laboratories suggests that bacterial biofilms have to be alive to induce larval settlement (Qian, 1999; Unabia and Hadfield, 1999) but other studies have found that larvae will readily settle on biofilms killed with formalin or treated with antibiotics (Kirchman et al., 1982a ; Hamer et al., 2001 ).

While there is evidence that larvae of specialist marine herbivores settle in response to compounds secreted by their host alga (Williamson et al., 2000; Swanson et al., 2004), it has also been suggested that generalist marine herbivores respond to broadly distributed habitat cues such as bacterial biofilms (Steinberg et al., 2002). A recent study demonstrated that larvae of the sea urchin Heliocidaris erythrogramma, which is a generalist herbivore, responds to biofilm cues distributed across many algal species and inanimate substrata such as rocks and shells (Huggett, 2005).

Similar to invertebrate larvae, zoospores of algae settle through a process that involves exploration and sensing of a surface (Callow et al., 1997). The process has been well 5 documented for Ulva spores and many factors influence zoospore surface selection, including negative phototaxis, thigmotaxis, surface chemistry and topography (Callow and Callow, 2000; Callow et al., 2000a,b; Callow et al., 2002). However, one of the most significant factors is the presence of bacterial biofilms and several studies have focused on the influence of biofilms on algal spore settlement (Dillon et al., 1989; Holmstrom et al., 1996; Joint et al., 2000; Egan et al., 2001b; Patel et al., 2003; Tait et al., 2005).

Microbial films have been shown to enhance settlement of algal spores (Dillon et al., 1989; Holmstrom et al., 1996; Joint et al., 2000; Egan et al., 2001b; Patel et al., 2003; Tait et al., 2005). Monospecies biofilms of strains ascribed to Pseudomonasl Alteromonas established on glass surfaces enhanced spore germination in Enteromorpha, whilst other strains were inhibitory (Thomas and Allsopp, 1983). Mixed species biofilms also enhanced settlement of Enteromorpha spores (Dillon et al., 1989). A quantitative study conducted by Joint and co-workers, showed that there was a correlation between the number of bacteria that formed mixed species biofilms and the number of Enteromorpha spores that attached. Promotion of attachment was attributed to the production of diffusible signal molecules (Joint et al., 2002). It was also shown that living biofilms were more stimulatory than dead biofilms (Tait et al., 2005). In an extensive study utilising 99 isolates, Patel and co-workers (2003) showed that spore settlement stimulation by monospecies biofilms was strain specific and not taxon specific. Pseudoalteromonas species showed a range of effects ranging from stimulatory to neutral to inhibitory and the activity of spore settlement was dependent on the age of the biofilm (Patel et al., 2003).

Similarly, Pseudoalteromonas species have also been found to inhibit algal spore settlement in other studies (Holmstrom et al., 1996; James et al., 1996; Egan et al., 2001b). Testing a wide range of isolates, Holmstrom et al., (1996) found that 66% of strains were inhibitory to U lactuca spore settlement with Pseudoalteromonas tunicata displaying the strongest activity (Holmstrom et al., 1996). The biologically active component was isolated and it was found to prevent the germination of not only Ulva spores but also spores from the red alga Polysiphonia spp. (James et al., 1996; Egan et al., 2001b).

6 Biofilm communities function as pioneer communities developed during the first stages of succession on a newly cleared substratum and they act as both an attractant and the actual substratum upon which algae and larvae subsequently settle. The attachment of macrofoulers results in a complex fouling community (Kirchman et al., 1982a) and the transition from a microbial biofilm to the more complex mature fouling community is dependent on properties of the biofilm. Thus biofilms may be either stimulatory, or inhibitory to the development of fouling communities.

1.3 BACTERIAL BIOFILM DEVELOPMENT

Biofilms are the predominant mode of growth for bacteria in the environment and provide important advantages to organisms. Bacteria growing in biofilms manifest marked resistance to antimicrobial agents (Stewart, 2002), increased synthesis of protective matrix materials (Costerton et al., 1999) and may exhibit enhanced metabolic cooperation (Shapiro, 1998). Furthermore, biofilm growth may facilitate horizontal gene transfer and intercellular communication (Hausner and Wuertz, 1999; Parsek and Greenberg, 2000) and increase genetic diversity of bacterial populations (Boles et al., 2004). All of these characteristics would enhance the survival of bacterial communities in harsh environmental conditions. Overall biofilm development is a well-regulated process that results from the integration of both external and internal signals. Although mixed species biofilms predominate in most environments, single species biofilms have been the focus of most current research. Thus, the published literature on biofilm development is based almost exclusively on single species biofilms.

Biofilm development has been depicted as a linear process that proceeds through a series of discrete events that can be categorised into 4 phases: i) initial attachment of bacteria to the substratum ii) microcolony formation iii) biofilm maturation iv) dispersal

1.3.1 Attachment The first step in biofilm formation involves the adhesion of bacterial cells to a conditioning film on a surface. The film can change the properties of the surface and 7 can affect the affinity between the bacterium and the substratum. Adhesion to a conditioned surface can be described to occur in two stages : reversible and irreversible adhesion (Busscher et al., 1992). In reversible adhesion the bacterium interacts weakly with the surface and is not committed to adhesion and can detach. The change from reversible to irreversible attachment has been characterised as the transition from a transient interaction of the cell with the substratum to a permanent bonding (Characklis, 1990). This commitment to irreversible attachment is a crucial step in biofilm formation because initial surface colonisers are likely to be the foundation upon which the mature biofilm will be built. Microbial adhesion is a complex process and is also influenced by many factors including the physico-chemical properties of the cell surface (Fletcher, 1991) and the substratum (Harkes et al., 1992; Mueller et al., 1992; Doss et al., 1993). More recently, it has been shown that the transition from reversible to irreversible attachment has a genetic determinant in both P. fluorescens (Hinsa et al., 2003) and P. aeruginosa (Caiazza and O'Toole, 2004), but the mechanism by which they make the transition appears to differ between the two organisms.

Once at the surface, stable attachment must occur for successful colonisation. There is evidence to indicate that bacteria can sense contact with a surface and alter gene expression to promote stable cell-surface interactions (Kuchma and O'Toole, 2000; O'Toole et al., 2000; Otto and Silhavy, 2002) but the genes that are required for stable cell-surface interactions remain unknown. Stable cell-surface interactions can also be regulated by environmental conditions, specifically increased osmolarity (Prigent Combaret et al., 2001 ). It has been proposed that starvation is the trigger for surface colonisation (Costerton et al., 1995), while other research suggests that bacteria prefer to form biofilms when nutrient availability is high (Wimpenny and Colasanti, 1997). Depending on the system, both starvation and nutrient availability may be important for biofilm formation. Other environmental signals that can also influence initial attachment are pH, iron availability, oxygen tension and temperature (Costerton et al., 1995). Although environmental triggers for biofilm formation can vary from organism to organism, it is evident that environmental parameters have a huge impact on the transition from planktonic to biofilm growth.

8 1.3.2 Microcolony formation

Microcolonies are basic structural units of biofilms and they develop following the adhesion of bacterial cells to a surface (Davey and O'Toole, 2000). Structurally, microcolony formation can occur by at least three mechanisms. One means is by surface translocation in which surface associated cells utilise Type IV pili, or some other means of swarming or twitching motility, to join existing microcolonies. In P. aeruginosa, flagella and Type IV pili mediated twitching motility both play a role in surface aggregation (O'Toole and Kolter, 1998a), and in E. co/i, flagella, Type 1 pili and curli fimbriae have been implicated in biofilm formation (Jackson et al., 2002b). Although motility appears to assist colonisation by Gram-negative organisms, it does not seem to be a prerequisite for biofilm formation as several nonmotile bacteria such as streptococci, staphylococci and mycobacteria readily form biofilms (Rupp et al., 1999; Cucarella et al., 2001).

A second method of microcolony formation is by clonal growth through cell division of existing resident bacteria. As cells divide, daughter cells spread outward and upward from the attachment point to form microcolonies, in a manner similar to microcolony formation on agar plates. This type of growth has been monitored in Mycobacterium fortuitum (Hall-Stoodley and Stoodley, 2005) and P. aeruginosa (Heydom et al., 2000; Tolker-Nielsen et al., 2000). The clonal growth model showed that although Type IV pili are not necessary for initial attachment, twitching motility was required later for the expansive migration of colonies along the surface to form a mature biofilm (Klausen et al., 2003a; Klausen et al., 2003b ).

The third method is when cells are recruited into the microcolony, so that planktonic cells (Tolker-Nielsen et al., 2000), or cell floes (Stoodley et al., 2001), directly attach to cell clusters from the bulk fluid. However, in both of these studies, recruitment appeared to play a minor role in colonisation. The relative contribution of each of these mechanisms will vary depending on the organisms involved, the surface being colonised and the physical and chemical conditions of the environment (Stoodley et al., 2002).

9 1.3.3 Maturation of biofilms

With time, microcolonies develop into a mature biofilm that is often associated with the increased production of extracellular polymeric substances (EPS). The chemistry of EPS is complex and includes polysaccharides, nucleic acids and proteins (Sutherland, 2001; Flemming and Wingender, 2002). Recently genes proposed to be involved in the synthesis of the polysaccharide component of the EPS have been reported (Friedman and Kolter, 2004; Jackson et al., 2004; Matsukawa and Greenberg, 2004). The polysaccharide polymer alginate produced by P. aeruginosa is the best studied component of biofilm EPS and appears to play an important role in determining biofilm structure (Nivens et al., 2001). Studies with P. aeruginosa show that overexpression of alginate gives rise to highly structured biofilms (Hentzer et al., 2001) and upregulation of alginate synthesis has also been linked with the downregulation of flagellum synthesis (Garrett et al., 1999). One of the earliest and most thoroughly documented changes of a sessile lifestyle is the repression of flagellar gene expression (Sauer and Camper, 2001 ). It seems that as cells adjust to an immobile life on a surface, they lose their flagella and increase production of EPS.

Although polysaccharides are the best studied components of EPS, data suggest a large diversity in EPS produced by different species under different growth conditions (Sutherland, 2001 ). The role of proteins and nucleic acids in biofilm structure remains largely unknown, but studies suggest that extracellular DNA may play a structural role in the early events of biofilm formation (Whitchurch et al., 2002).

Another important step is the formation of the characteristic biofilm architecture with "mushroom" and "pillar" microcolonies separated by water channels. Recent reports have showed that the mushroom-like structures consist of stalks on which caps form. Apparently cells in the stalk do not move and a subpopulation of cells use twitching motility to migrate up the stalk and form a cap (Klausen et al., 2003a, b). Rhamnolipid surfactants produced by cells in the biofilm have been shown to be important in development of mushroom-like structures (Davey et al., 2003). Rhamnolipid genes were shown to be expressed primarily in the stalks of the mushroom-like structures (Lequette and Greenberg, 2005). Rhamnolipids also play a role in maintaining open spaces or channels surrounding microcolonies by inhibiting colonisation of invading

10 planktonic cells (Davey et al., 2003) and in swarming motility by allowing flagellum based propulsion of P. aeruginosa over solid surfaces (Caiazza et al., 2005).

It has been shown that for some bacteria, cell-cell communication is essential to establish a well-ordered surface community. There is a growing body of evidence that signalling is important for biofilm development (Davies et al., 1998; Loo et al., 2000; De Kievit et al., 2001; Huber et al., 2001; Kjelleberg and Molin, 2002; Li et al., 2002; Lynch et al., 2002; Prouty et al., 2002; McNab and Lamont, 2003; Kuchma et al., 2005; Lequette and Greenberg, 2005; Merritt et al., 2005). However, it has been suggested that the dependence on signalling is not absolute and nutrient conditions may affect whether quorum sensing is required (Purevdorj et al., 2002). For example, in E. coli K- 12, biofilm formation was observed in relatively rich media (Pratt and Kolter, 1998) but in contrast, E. coli 0517 :H7 only forms biofilms under conditions of low nutrients (Dewanti and Wong, 1995). In addition, a change in carbon source has also been shown to alter the biofilm structures formed. Biofilm formation was restored in P. fluorescens mutants by the addition of supplements such as iron, citrate, and glutamate to minimal media (O'Toole and Kolter, 1998b) and P. aeruginosa failed to form the mushroom type biofilm architecture when citrate was used as the sole carbon source instead of glucose (Davies et al., 1998; Klausen et al., 2003a). More recently it was demonstrated that growth of cells in complex medium can prevent many of the AHL regulated genes in P. aeruginosa (Yarwood et al., 2004). In Serratia marcescens it was observed that cell chain type biofilm could be converted to a microcolony type biofilm when the amount of glucose and casamino acids in the medium is reduced (Rice et al., 2005). It seems that biofilm structures can be manipulated by changing the medium. Thus the control of biofilm architecture is a complex process involving a combination of nutrient components and master regulators, such as those of quorum sensing systems.

1.3.4 Detachment

While there are a number of benefits associated with the biofilm mode of growth, these advantages come at some cost (Boles et al., 2005). For example when local conditions deteriorate, biofilm bacteria have a reduced ability to evade stresses, as they are physically confined by the matrix and their motility functions are repressed (Whiteley et al., 2001; Sauer et al., 2002). Thus it is vital that bacteria possess mechanisms to

11 separate from biofilms and resume planktonic life. This process is termed detachment (some researchers use the term dispersal). Detachment can occur by both passive and active mechanisms. Active mechanisms are triggered by environmental conditions, to bring about the release of cells (either individually or in groups) from a biofilm. Examples of passive mechanisms include sloughing, which is the random removal of large portions of biofilm, due to shear forces. The process of detachment is perhaps the least understood of the biofilm life cycle. Detachment has been extensively studied because of industrial and medical implications in biofilm removal, but this area of biofilm development has been difficult to define biochemically and genetically.

There are a number of environmental factors that trigger detachment. In particular, starvation for several hours of nutrients such as nitrogen and carbon has been shown to induce detachment in Pseudomonas spp., Escherichia coli, and Acinetobacter spp. (Delaquis et al., 1989; Sawyer and Hermanowicz, 2000; Jackson et al., 2002b). In contrast, Sauer and coworkers showed that increasing nutrient levels could induce detachment and that the degree of detachment was carbon specific (Sauer et al., 2004). Dispersal in P. aeruginosa could also be induced by a rapid decrease in pH and was accompanied by induction of genes involved in flagellar swimming motility (Purevdorj Gage et al., 2005). In E. coli, lack of oxygen has been demonstrated to result in increased flagellar and LPS synthesis, which correlates with a decrease in biofilm formation (Landini and Zehnder, 2002). Recently it was shown that a sudden downshift in molecular oxygen concentration could induce cells to rapidly disperse from Shewanella oneidensis MR-I biofilms (Thormann et al., 2005). It is possible that flagella synthesis is switched on as oxygen becomes limiting. Thus, dispersal seems to be triggered by several different cues that allow bacteria to revert to the planktonic state as environmental conditions change.

The process of detachment is complex, and there appear to be a variety of dispersion/dissolution mechanisms utilised by different species. Even within a single bacterial species, multiple detachment patterns have been observed (Stoodley et al., 2001; Boles et al., 2005). One form of detachment is believed to be the process of programmed cell dispersal (Allison et al., 1998). The death of a sub population of cells has been reported as a normal feature in the development of a number of biofilms including P. aeruginosa (Webb et al., 2003), Vibrio cholerae (Sodhi, N, Rice, S, 12 McDougald, D. unpublished data), Pseudoalteromonas tunicata (Mai-Prochnow et al., 2004) and mixed species biofilms in the oral cavity (Auschill et al., 2001). AHL mediated quorum sensing has also been shown to be involved in dispersal (Schooling et al., 2004; Rice et al., 2005), and may be more general than initially thought. An overexpression of alginate lyase in P. aeruginosa resulted in increased detachment (Boyd and Chakrabarty, 1994) and detachment was attributed to a loss of EPS in P. fluorescens (Allison et al., 1998). A general mechanism by which biofilms may detach is through the production of surfactants such as rhamnolipids (Boles et al., 2004). The latter authors demonstrated that addition of exogenous rhamnolipid could induce premature dispersion in wild type P. aeruginosa biofilms (Boles et al., 2005). However, it remains to be seen whether rhamnolipids can induce detachment of natural biofilms in a medical setting or a mixed species biofilm community in the marine environment.

1.4 MICROBIAL INTERACTIONS IN BIOFILMS

In nature, colonisation of habitats by mixtures of bacterial populations is the rule rather than the exception (Marsh and Bowden, 2000). Excluding certain types of infections or symbiosis, most biofilms consist of multiple species of both eukaryotic and prokaryotic organisms. Microbial communities are said to display emergent properties (Odum, 1969), that is, the properties of the community are more than the sum of its component populations. Existing in close proximity within a microbial community can have profound consequences for component populations. The potential for interspecies communication, competition, and cooperation is high (Watnick and Kolter, 2000), and the physiology and metabolism of multi-species biofilm communities are immensely complex.

Alexander (Alexander, 1971), has described a range of potential interactions among populations in microbial communities. These include mutualism, commensalism, protocooperation, competition, antagonism and parasitism. Here, I will focus on cooperation, competition, antagonism, inhibitory compounds produced by marine bacteria and describe succession in microbial communities.

13 1.4.1 Cooperation

When different organisms interact in biofilms, such collaborations can be beneficial to one or more of the participating populations. Potential benefits include a broader habitat range for growth through the modification of their local environment (Bradshaw et al., 1994; Caldwell et al., 1997; Palmer et al., 2001; Gilbert et al., 2002; Velicer, 2003). Studies of cooperative bacterial interactions have also shown that they result in enhanced resistance to environmental stress and inhibitors (Erb et al., 1997; Cowan et al., 2000; Leriche et al., 2003) and play a role in the persistence of species under hostile conditions (Bradshaw et al., 1997; Karthikeyan et al., 1999). Other examples of cooperation include biodegradation through co-metabolism (Cowan et al., 2000; Nielsen et al., 2000; Christensen et al., 2002) and synergistic biofilm formation (Palmer et al., 2001; Filoche et al., 2004; Sharma et al., 2005; Yamada et al., 2005). Thus interacting populations in biofilms are able to defy constraints imposed by the external microenvironment and survive conditions that are inhibitory to the same cells growing in pure culture. It has been further suggested recently, that metabolic interactions in biofilms may be strong driving forces in microbial evolution (Hansen et al., 2005).

1.4.2 Competition in biofilms

Microorganisms can compete for resources within biofilms. Strategies of competition within biofilms may be different from those in planktonic cells. In competition studies, where Pseudomonas putida was allowed to invade a Hyphomicrobium sp. biofilm, P. putida was found to dominate, even though the biofilm-associated Hyphomicrobium numbers remained relatively constant (Banks and Bryers, 1991 ). In dual species biofilms, Klebsiella pneumoniae and P. aeruginosa were able to coexist in a stable community even though P. aeruginosa growth rates were much slower in the mixed culture biofilm than when grown as a pure culture biofilm (Stewart et al., 1997). Competition for substrate in freshwater biofilms resulted in a non-uniform spatial distribution of populations (Zhang et al., 1994). Thus competing strains, which are normally outcompeted in the planktonic state, are able to coexist in a biofilm.

Experimental evidence based on mathematical modelling also shows that the biofilm lifestyle with growth of bacteria in microcolonies with its attendant substrate gradients

14 can result in slow growing bacteria with higher yield (YS) dominating. This outcome is very different to the situation in continuous systems such as chemostats, where bacteria which have a high growth rate at a low yield strategy (RS) will always outcompete bacteria which have a high yield at a low growth strategy (YS) (Kreft, 2004).

1.4.3 Antagonism

Antagonism among microorganisms is also a major contributing factor in determining the composition of microbial communities. The production of antagonistic compounds such as bacteriocins can give an organism a competitive advantage when interacting with other microorganisms. For example, studies with human volunteers showed that the degree of colonisation of teeth by oral streptococci was proportional to their level of in vitro activity (Hillman et al., 1987). The production of antagonistic factors may also be a mechanism whereby exogenous species are prevented from colonising pre established microbial communities. This would contribute to the phenomenon of "colonisation resistance" (van der Waaij et al., 1971 ).

In dual species biofilms, it has been found that bacteriocin-producing strains do have a competitive advantage over bacteriocin-sensitive strains within a biofilm (Tait and Sutherland, 2002), both in gaining a foothold in a new environment and in preventing the colonisation of a potential competitor into a pre-established biofilm. Similarly, inhibitory compounds were shown to give bacteria a competitive edge when established biofilms of Burkholderia cepacia were dispersed by P. aeruginosa known to produce a B. cepacia growth-inhibitory substance (Al-Bakri et al., 2004). However, despite the production of inhibitory factors, the existence of discrete microhabitats within a biofilm can result in organisms being spatially organised and therefore bacteria can coexist in a community with species that would be incompatible with one another in a homogeneous environment (Tait and Sutherland, 2002).

1.4.4 Inhibitory compounds produced by marine bacteria

It is now recognised that many groups of marine bacteria produce antibiotics that inhibit the growth of other marine bacteria (Engel and Pawlik, 2000; Engel et al., 2002). The growing interest in marine microbial products is demonstrated by the number of novel metabolites reported from marine bacteria (Wratten et al., 1977;

15 Kalinovskaya et al., 1995; lsnansetyo and Kamei, 2003; Spyere et al., 2003; Kalinovskaya et al., 2004; Longeon et al., 2004; Jensen et al., 2005; Williams et al., 2005). A number of reviews have covered this area (Jensen and Fenical, 1994; Beman et al., 1997; Paul and Puglisi, 2004), and only a brief description of the distribution of inhibitory bacteria will be given. Similar to their soil counterparts, marine bacteria appear to produce antibiotics in nutrient rich hotspots such as organically rich marine snow particles or aggregates or surfaces of algae. A remarkably large fraction (53.5%) of marine bacterial isolates was found to exhibit antagonistic properties against other pelagic bacteria (Long and Azam, 2001). Investigations suggest that particle-attached bacteria are more likely to produce inhibitory compounds than their free-living counterparts (Nair and Simidu, 1987; Long et al., 2003). Furthermore, particle-attached isolates produced broad range inhibitors suggesting that antibiotic production may be a mechanism used by particle specialists to dominate the particle phase by deterring other potential colonizers (Long and Azam, 2001 ). While antibiotic production is distributed throughout all major cultivable heterotrophic phylogenetic groups, the most potent producers belong to the y-proteobacteria with the Alteromonadales and Vibrionales groups being the dominant producers, and also the least sensitive to inhibition (Long et al., 2003). Thus, they may dominate the particulate matter and biofilms in the ocean. Giovannoni and Rappe (Giovannoni and Rappe, 2000), found that y-proteobacteria were the dominant cultivable marine bacteria and the bias towards these groups observed in their findings has been suggested to be due to their ability to produce antibiotics and their resilience to antibiotics (Long and Azam, 2001).

Similarly, it has been found that compared to planktonic strains, a large percentage of bacteria growing on algal surfaces produce antibacterial compounds (Lemos et al., 1985; Boyd et al., 1998; Boyd et al., 1999). Lemos and co-workers (1985) showed that the highest number of active strains were isolated from Enteromorpha intestinalis and were Pseudomonas-Alteromonas strains. Some seaweed extracts were found to enhance the growth of bacteria found on their surfaces (Sakami, 1996), and Boyd and co workers ( 1999) demonstrated that strains would produce inhibitors if they were grown attached to a surface, but not in planktonic cultures. Furthermore, surface attached bacteria had broader-spectrum antagonistic interactions than free-living bacteria. Generally, it has been observed that bacteria attached to surfaces are found to produce

16 more activity, suggesting that surfaces offer microenvironments where antibiosis is most intense.

1.4.5 Succession

Studies on ecological succession in microbial communities lag behind those of plant or animal communities and there is little experimental evidence from which to construct models. The sequence of events in microbial succession is well understood for human dental surfaces, and the primary colonists and the participation of these organisms in subsequent colonisation stages have been elucidated (Xie et al., 2000; Li et al., 2001; Filoche et al., 2004). However, this detailed level of understanding has not been attained for other types of surfaces.

The following successional model has been proposed for microbial communities in aquatic systems (Jackson et al., 2001). The process begins with the primary attachment of a number of species recruited from the bulk water population. Since bacteria are initially spatially separated, the community is likely to have a high level of diversity. Metabolism of primary colonisers can modify the habitat to make conditions more suitable for growth of secondary colonisers. In the secondary community, competition for resources or space may exclude less competitive organisms causing a reduction in diversity. As the biofilm community matures, more niches are created due to the formation of gradients and the internal recycling of resources. At this stage, species richness increases again, reflecting a complex spatial structure with many functional groups of bacteria (Jackson et al., 2001; Jackson, 2003). This model fits data from investigations conducted by Santegoeds co-workers (Santegoeds et al., 1998) and Jackson (2003). These researchers conducted studies over similar periods of time (approximately 3 months) and used comparable techniques. Data from a longer term study, conducted over a three year period on drinking water biofilms, also agreed with the above conceptual model for succession (Martiny et al., 2003).

Studies on succession m the marme environment are limited to early colonisation events. Dang and Lovell (2000) studied early succession in marine waters primarily to assess the diversity of primary colonisers and to identify the most common early colonists. Their investigations suggest that bacteria within the Alteromonas group were

17 early colonists but were displaced by the Roseobacter group which were dominant in all phases of colonisation.

While multispecies biofilms abound in most environments, there is relatively little known about these communities in the marine environment. The strategies by which marine bacteria form multispecies biofilms are poorly understood, and cooperative and competitive interactions within these communities, remain largely unstudied. Although, molecular techniques such as DGGE have revealed the complexity of microbial communities on the surfaces of algae (Fisher et al., 1998), little is known about their ecological role.

1.5 THE ORGANISMS

1.5.1 Ulva australis

Ulva australis (previously known as Ulva lactuca and commonly known as the sea lettuce), is a cosmopolitan green alga, which is responsible for green tides and marine fouling (Callow et al., 1997; Blomster et al., 2002). It is highly tolerant of variable salinity, temperature and water quality and grows quite rapidly in nutrient-rich waters (Tan et al., 1999). Its life history consists of morphologically similar haploid and diploid phases, both of which reproduce prolifically by haploid and diploid asexual zoospores (Blomster et al., 2002). Sexual reproduction involves fusion of opposite mating types of haploid gametes, which can also develop parthenogenetically into adult thalli.

Ulva sp have been used as model organisms in studies of algal morphology (Provasoli and Pintner, 1977; Nakanishi et al., 1996). There is evidence from a number of studies that bacteria have important consequences for macroalgal growth and morphogenesis in culture. Axenic cultures of Ulva do not grow normally, a condition that can be alleviated by the addition of uncharacterised bacterial epiphytes (Provasoli and Pintner, 1980; Tatewaki et al., 1983). Nakanishi and co-workers (1996) found that 78% of the bacteria capable of inducing morphogenesis in Ulva pertusa belonged to the Flavobacterium group. Matsuo and coworkers (Matsuo et al., 2003) also showed that differentiation in the green algae Monostroma, Ulva and Enteromorpha depends on

18 strains from the Cytophaga-Flavobacterium-Bacteroides (CFB) complex. More recently, these workers reported the isolation of a highly potent differentiation inducer, termed thallusin, from an epiphytic marine strain belonging to the CFB group (Matsuo et al., 2005). Thus certain epiphytic bacterial strains can play an important role in the growth and development of Ulva communities.

Ulva spp. have also played a prominent role in studies conducted on spore bioadhesion (Stanley et al., 1999; Callow et al., 2000a). An unexpected finding from Ulva spore settlement assays was that spores are attracted to AHLs produced by bacterial biofilms (Joint et al, 2002) and the nature of the response appears to be chemokinetic rather than chemotactic (Wheeler et al., 2006). While it is clear that AHLs and related molecules affect some eukaryotes, the ecological relevance of these interactions remains unknown. However, seeing that Ulva is dependant on some members of its epiphytic community for morphological development, it has been proposed that spores may be attracted to certain biofilms to facilitate a close relationship between the developing germling and certain essential bacteria (Tait et al., 2005). However, AHL dependent signalling mechansims have not been identified in the CFB group (Gram et al., 2002; Manefield and Turner, 2002), so it remains to be established whether the morphogenesis inducing strains produce AHLs.

The extensive highly diverse microbial community associated with U. australis, makes it an interesting study organism for addressing questions of surface colonisation and host association. The dynamics of surface colonisation by marine bacteria, particularly of algal surfaces is poorly understood. Compared to higher plants, (see review by (Brencic and Winans, 2005), relatively little is known about colonisations strategies, host-microbe and microbe-microbe interactions on the surface of algae. Surprisingly, the ecology of U. australis remains largely unknown, despite being common and widespread in distribution in Australia. The research undertaken in this study, focuses on two organisms that are frequently isolated from U. australis, Pseudoalteromonas tunicata and Roseobacter gallaeciensis.

19 1.5.2 Pseudoalteromonas genus

The genus Pseudoalteromonas belonging to the y-proteobacteria, was established in 1995, after a phylogenetic reorganisation based on new available 16S rRNA gene sequences (Gauthier et al., 1995). Members of this genus are ubiquitous and successful colonisers of living marine surfaces (Holmstrom and Kjelleberg, 1999), commonly expressing either mutualistic or saprophytic relationships with the host (Egan et al., 2000; Mitsutani et al., 2001; Negri et al., 2001; Ivanova et al., 2002b; Lau et al., 2005). Some Pseudoalteromonas species have also been isolated from non-living marine surfaces such as sediments and rocks and the genus is widespread worldwide (Acinas et al., 1999; DeLong et al., 1999). Pseudoalteromonas spp. only constitute a minor fraction of pelagic seawater samples, despite being dominant in bacterioneuston where it has been suggested that they form biofilms at the sea surface interface (Franklin et al., 2005).

Pseudoalteromonas species produce a range of bioactive compounds, which include antibiotics, agarases, proteases, tetradotoxin and EPS (Holmstrom and Kjelleberg, 1999). Several species within this genus demonstrate antibacterial activities, including P. aurantia, P. luteoviolacea, P. rubra, P. citrea, P. ulvae, P. tunicata, P. maricaloris and P. phenolica (Gauthier and Flatau, 1976; Ivanova et al., 1998; Egan et al., 2001a; lvanova et al., 2002a; Isnansetyo and Kamei, 2003; Sobolevskaya et al., 2005). On the other hand, some Pseudoaltermonas species produce compounds which have a stimulatory effect on larval settlement (Szewzyk et al., 1991b; Leitz, 1997; Lau and Qian, 2001; Negri et al., 2001).

In an analysis of the antifouling activity of ten different species of Pseudoalteromonas, Holmstrom and co-workers (Holmstrom et al., 2002) showed that eight species contained at least one of the four investigated antifouling properties : growth inhibition against bacteria and fungi and inhibition of algal spore and larval settlement. The most effective of the ten species were P. tunicata and P. ulvae. These constitute the antifouling subgroup of the genus Pseudoalteromonas.

20 1.5.2.1 Pseudoalteromonas tunicata

The Pseudoalteromonas species that I have investigated in this thesis was 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 Ulva australis (Egan et al., 2001a). P. tunicata has also been isolated from an algal bloom in Tasmania, Australia and its 16S ribosomal DNA (rDNA) sequences have been found in clone libraries sampled from biofilms in saline water caves in Australia and Antarctic sea ice (Brown and Bowman, 2001; Holmes et al., 2001). More recently, the species was also detected on the surfaces of U lactuca, C. intestinalis and Ulvaria fusca in Aarhus Bay, Denmark (Skovhus et al., 2004). P. tunicata produces at least five target specific compounds that inhibit bacteria, fungi, algal spores, invertebrate larvae, diatoms and protozoa and the production of antifouling compounds has been linked to the production of pigments.

1.5.2.2 Antibacterial protein inP. tunicata

The antibacterial protein (AlpP) is a 190 kDa multi-subunit protein that is produced during the stationary phase of growth and has a broad spectrum of activity (James et al, 1996). P. tunicata cells are also sensitive to their own antibacterial protein when the cells are in the exponential phase of growth (James et al., 1996). A database search of sequence similarities between the alpP sequence in P. tunicata compared to other organisms suggests that AlpP is a conserved protein among several other bacterial species. These include Marinomonas mediterranea, Chromobacterium violaceum, Caulobacter crescentus, Magnetococcus sp. MC- I, Rhodopseudomonas palustris, Rhodopirellula baltica, Microbulbifer degradans and Nitrobacter hamburgensis (Mai Prochnow et al, submitted). The mode of action of AlpP is that ofH2O2 generated stress and lysis (Mai-Prochnow, A., Lucas-Elio, P., Webb, J.S. and Kjelleberg, S., unpublished data) similar to the mode of action of marinocine the AlpP homologous protein produced in M Mediterranea (Lucas-Elio, P., Gomez, D., Solano, F. and Sanchez-Amat, A., unpublished data).

21 1.5.3 Roseobacter clade

The Roseobacter clade falls within the a-3 subclass of the class Proteobacteria. Roseobacter species are important members of the aquatic microbial community (Shiba, 1992), and except for strains of the genus Ketogulonicigenium (Urbance et al., 2001 ), have all been isolated from marine or high-salt environments such as seawater, marine sediments, and surfaces of marine organisms (Moran et al., 2003). They are very abundant and widely distributed (Fuhrman et al., 1993; Mullins et al., 1995; Giuliano et al., 1999; Gonzalez et al., 1999; Eilers et al., 2000; Giovannoni and Rappe, 2000). Further, they are dominant in coastal bacterial communities (Dang and Lovell, 2000) and it is estimated that, in some areas, Roseobacter clade-affiliated organisms account fer 7 to 30% of the microbial population (Gonzalez et al., 1997; Gonzalez et al., 2000; Selje et al., 2004), though they may also be present in lower proportions (i.e., 1 % )(Eilers et al., 2001 ). Significant growth of Roseobacter species has been recorded in unenriched seawater cultures or nutrient depleted medium in several marine areas (Pinhassi and Berman, 2003). The Roseobacter group is found in diverse marine habitats which include free living, particle associated or in commensal relationships (Alavi et al., 2001; Alavi, 2004; Selje et al., 2004; Alonso and Pemthaler, 2005) Furthermore, members of this group are readily cultured (Giovannoni and Rappe, 2000).

The exact ecological role of the Roseobacter group is unclear, but there is an increasing body of evidence to suggest that they play a major role in the transformation of sulfur compounds in nature. Although members of this group possess diverse metabolic capabilities, the ability to utilise dimethylsulfoniopropionate (DMSP), as both a carbon and sulfur source, appears to be a characteristic of a large number of strains (Moran et al., 2003). DMSP is an abundant organic sulfur compound produced as an osmolyte, by marine algae and coastal vascular plants (Ledyard et al., 1993). DMSP is released due to algal senescence, predation or stress and is degraded by both algal and bacterial enzymes -as reviewed in (Y och, 2002). Field studies have provided a link between Roseobacters and sulfur cycling in situ (Gonzales et al., 2000). Roseobacter phylotypes were found to be abundant in a DMSP-rich algal bloom, accounting for almost 50% of the 16S rDNA gene sequence pool and abundance was positively correlated with DMSP degradation (Gonzales et al., 2000). Direct evidence of DMSP uptake was recently

22 demonstrated by microautoradiography (Vila et al., 2004). Furthermore, some Roseobacter spp. exhibit close physical and physiological relationships with toxic DMSP-producing dinoflagellates, including Procentrum spp. (Lafay et al., 1995), Alexandrium spp. (Gallacher et al., 1996), and Pfiesteria spp. (Alavi et al., 2001; Miller and Belas, 2004; Miller et al., 2004). Thus, DMSP-producing algae appear to be a niche area for Roseobacter in the marine environment, but the basis for some of these interactions remains unclear.

Several Roseobacter strains also produce AHLs (Gram et al, 2002). The possible involvement of AHL regulation in Roseobacter metabolism can have major implications for turnover of organic material in the ocean, as well as their colonisation of, and growth on, organic aggregates and phytoplankton (Gram et al, 2002). While AHLs are clearly involved in bacterium-bacterium interactions, these regulatory systems may also be a key to understanding specific bacterium-eukaryote interactions (Fuqua and Winans, 1994; Steidle et al., 2001; Hoffmann et al., 2005). Roseobacter strains form symbiotic relationships with diverse eukaryotic marine organisms, including algae (see review by Buchan et al., 2005), and Roseobacter species are required for dinoflagellate growth (Alavi et al., 2001; Miller and Belas, 2004). Colonisation and growth on algae may be regulated by AHLs similar to that of epiphytic bacteria in higher plants (Quinones et al., 2004).

1.5.3.1 Roseobacter gallaeciensis

This species was first found in seawater from larval cultures of the scallop Pecten maximus in Galicia, Spain (Ruiz-Ponte et al., 1998), and then later isolated from the German Wadden sea (Brinkhoff et al., 2004) and in Australian waters from the surface of the green alga Viva australis. Organisms belonging to the R. gallaeciensis cluster are widespread in the marine environment (Ruiz-Ponte et al., 1998; Long and Azam, 2001; Sekiguchi et al., 2002; Pinhassi and Berman, 2003; Brinkhoff et al., 2004). It is likely that R. gallaeciensis has a dual-niche existence alternating between free-living and attached growth forms. Cells are gram-negative; ovoid-rod shaped and are motile by means of polar flagella (Ruiz-Ponte et al., 1998). Unlike several members of the Roseobacter clade, R. gallaeciensis cells do not contain bacteriochlorophyll a. When grown on marine agar, colonies appear smooth, convex and brownish and produce a

23 diffusible pigment (Ruiz-Ponte et al., 1998). R. gal/aeciensis has strong antagonistic activity against a number of bacterial strains, mostly marine (Ruiz-Ponte et al., 1999; Brinkhoff et al., 2004). The inhibitory compounds are proposed to be a peptide (Ruiz Ponte et al., 1999) and tropodithietic acid, respectively (Brinkhoff et al., 2004 ).

1.6 AIMS OF THIS STUDY

The major hypothesis addressed in this study is that the biofilm community on Ulva australis is able to defend the host against fouling organisms. It is likely that inhibitory bacteria that live on the algal surface are able to influence the colonisation of other organisms and hence influence the population composition of the microbial community.

The specific aims were:

1. To investigate if P. tunicata is an effective competitor against other marine bacterial isolates during biofilm formation (Chapter 2).

2. To investigate if P. tunicata and R. gallaeciensis are effective colonisers of U. australis and are able to compete and dominate over other marine bacterial isolates during biofilm formation on the plant surface (Chapter 3).

3. To assess if P. tunicata and R. gallaeciensis biofilms, at ecologically relevant cell densities, are effective at preventing the attachment of fouling organisms (Chapter 4).

4. To determine if P. tunicata and R. gallaeciensis are able to colonise and invade a complex seawater community or a synergistic mixed species biofilm (Chapter 5).

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Chapter Two

Competitive interactions in mixed-species biofilms containing the marine bacterium Pseudoalteromonas tunicata1

2.1 INTRODUCTION

Biofouling is ubiquitous in the manne environment and bacteria are among the first organisms to foul surfaces. They form biofilms which serve as a focus for the attachment and growth of other organisms such as invertebrates, sessile plants and animals (Davis, 1989). Mature marine biofouling communities are complex, highly dynamic ecosystems and once established are extremely difficult to eradicate (Holmstrom et al., 2002).

Many marine organisms have evolved efficient strategies to combat epibiosis. Seaweeds employ a number of physical and chemical defence systems to prevent fouling such as the shedding of outer layers of cells (Keats et al., 1997) or production of inhibitory compounds (Dworjanyn et al., 1999). However, antifouling defence is energetically costly (Wahl, 1989), and it has been suggested that the seaweed Ulva australis (formerly known as Ulva lactuca) which has neither physical nor chemical defences, relies on microbial defence (Holmstrom et al., 1992; Egan et al., 2001). It has been demonstrated that bacterial biofilms are present on the surface of U. lactuca (Sieburth, 1975), and it has been shown that they can be beneficial for their hosts by enhancing their antifouling strategies (Armstrong et al., 2001; Gil-Tumess et al., 1992; Nakasano et al., 1993; Holmstrom and Kjelleberg, 1994; Boyd et al., 1999; Dobretsov and Qian, 2002).

1 This chapter is a modified version of: Rao, D., Webb, J.S., and Kjelleberg, S. (2005) Competitive interactions in mixed-species biofilms containing the marine bacterium Pseudoalteromonas tunicata. Appl. Environ. Microbial. 71: 1729-1736.

47 One important group of marine bacteria that is found in association with living surfaces is Pseudoalteromonas sp. This genus produces a diverse range of biologically active compounds that specifically target marine fouling organisms. Perhaps the most extensively studied species within the genus is Pseudoalteromonas tunicata. Pseudoalteromonas tunicata is a green-pigmented gram-negative gamma proteobacterium. It colonises living marine surfaces, including U. australis (Holmstrom et al., 1998), and produces at least five novel inhibitory compounds. One of these compounds is a 190-kDa multi-subunit antibacterial protein designated AlpP, which is effective against both gram-negative and gram-positive bacteria from a range of environments (James et al., 1996). However, this protein was also found to be active against P. tunicata itself, which raises the question as to its ecological role. It is speculated that AlpP might provide a competitive advantage to P. tunicata during biofilm growth in the marine environment.

P. tunicata has been shown to form complex differentiated biofilms (Mai-Prochnow et al., 2004). Biofilm formation appears to typically follow a sequence. Single cells attach to a surface and differentiate into mature matrix-enclosed microcolonies separated by a network of open water channels. Dispersal of bacteria from the interior regions of the microcolonies has been observed, resulting in formation of hollow voids inside the microcolonies (Mai-Prochnow et al., 2004). Microcolony development is thought to be a coordinated adaptive response that facilitates continued biofilm development and dispersal (Webb et al., 2003). Very little is known about the role of microcolonies within biofilms. However, recent findings suggest that their formation by Pseudomonas aeruginosa allows it to escape predation by protozoans and flagellates (Matz et al., 2003). Generally, microcolonies may be an adaptive strategy for competing under stressful conditions.

The microcolonies that constitute the biofilm can be composed of single-species populations or mixed populations with varying degrees of interaction, depending on the environmental conditions under which they were formed. Two species formed mixed microcolonies when they were grown under commensal conditions and formed separate microcolonies under non commensal conditions (Nielsen et al., 2000). Thus, commensal relationships can play a role in determining the spatial distribution of the organisms in microbial communities. Meanwhile, in competitive interactions between 48 bacteriocin-producing and bacteriocin-sensitive strains, the bacteria formed mixed biofilms in which each bacterial strain formed its own microcolonies (Tait and Sutherland, 2002). Generally, surprisingly little is known about the factors that govern the establishment and distribution of bacteria within multispecies biofilms in marine environments. A few studies on epiphytic microbial communities present on macroalgae have highlighted the complex spatial distribution of bacterial populations (Corre and Prieur, 1996), but the strategies that marine bacteria use to colonise surfaces and to compete with other bacteria are poorly understood.

I explore here the hypothesis that P. tunicata competes effectively with other marine bacterial isolates during biofilm formation, based on the isolation of this organism on marine plants in repeated sampling and based on the production of the potent antibacterial compound AlpP by this species. In pure culture all the marine isolates formed biofilms within 72 hours. P. tunicata was found to be the dominant isolate and totally removed a competing strain unless the competitor was not sensitive to the P. tunicata antibacterial protein or exhibited strong inhibitory activity against P. tunicata. In competition studies in which microcolonies were allowed to form before the introduction of P. tunicata, biofilms coexisted for greatly extended periods. Studies with a P. tunicata AlpP mutant suggest that AlpP provides an advantage during colonisation of biofilms formed by other marine bacteria. The data also suggests that microcolonies may be protective structures during bofilm development that enhance persistence of an organism during competitive interactions.

2.2 MATERIALS AND METHODS

2.2.1 Isolation of marine strains

Pseudoalteromonas tunicata and other isolates were originally isolated from the surface of the common marine alga Ulva australis, which was collected from the rocky intertidal zone near Sydney, Australia. The algal thallus was suspended in sterile nine salt solution (NSS) and surface bacteria removed by vortexing. Aliquots of the samples were then spread on the complex marine medium Vaatanen nine salt solution (VNSS) (Marden et al., 1985) containing 1.5% agar and incubated at 23°C for 48 h. Morphologically distinct bacterial colonies were selected. Bacteria were stored at -

49 70°C in 30% glycerol. While over 50 different colony phenotypes were observed, only 20 isolates survived repeated subculture on VNSS medium. These bacterial strains were routinely grown and maintained on VNSS agar at 25°C. In addition to P. tunicata, four of the 20 culturable strains were chosen to be sequenced as these strains are commonly found growing on algae. The strains were: Pseudoalteromonas gracilis, Alteromonas sp., Cellulophaga fucicola, and Roseobacter gallaeciensis. Sequencing of the remaining isolates formed part of another project and was carried out by Niina Tujula (Tujula, N., Webb, J. S., Dalhoff, I., Holmstrom, C. and S. Kjelleberg unpublished data).

2.2.2 Fluorescent labelling of marine isolates

Marine strains were labeled with a green fluorescent protein (GFP) color tag by transconjugation using the constitutive GFP expression plasmid pCJS 10. This plasmid contains the gfpmut3 gene (Cormack et al., 1996) on an RSFlOIO backbone from the broad-host-range vector pHRP304 (Bagdasarain et al., 2003). In addition, strains were labeled with a red fluorescent protein (RFP) color tag by using the pCJS 10-derived plasmid pCJSlOR. This plasmid contains the RFP gene dsred (Clontech) in place of gfpmut3 on pCJS 10. Plasmids pCJS 10 and pCJS 1OR were gifts from Charles Svenson, University of New South Wales, Sydney, Australia. GFP triparental conjugations were carried out as described previously (Egan et al., 2002), and labeled transconjugants were grown on VNSS agar plates containing 15 µg of chloramphenicol per ml and 100 µg of streptomycin per ml. GFP- and RFP-labeled strains showed bright fluorescence after overnight culture and no differences were observed in the growth rates or surface attachment properties of the labeled strains compared with the unlabeled parent strain.

2.2.2.1 Detection of inhibitory compounds produced by marine bacterial isolates

In order to detect inhibitory compounds produced by marine bacterial isolates, concentrated supernatant from P. tunicata and other marine isolates was prepared by the method described previously (James et al., 1996), with some modifications. The strains were grown in marine minimal medium (3M) with 0.2%w/v trehalose and 0.2%w/v glucose as the carbon source for 48 h, harvested by centrifugation (12,000 x g for 20 min) and resuspended in 3M trehalose at a density of 0.7 g/ml (weight/volume). Each concentrated cell suspension was incubated, without shaking, at 28°C for 24 h.

50 Cells were removed by centrifugation (14,000 x g for 1.5 h) and the concentrated supernatant sterilised by using a 0.2-µm-pore-size sterile filter (Millipore).

Supernatant samples were assayed for inhibitory activity using the drop test assay (James et al., 1996). Briefly, 100 µl of an overnight broth culture of the target strain was spread on a VNSS agar plate and the plate dried at 30°C for 30 min. Drops containing 20 µl of the concentrated supernatant, as well as a control (nine-salt solution), were placed on the agar surface and incubated overnight at room temperature to allow formation of inhibition zones.

2.2.3 Biofilm experiments

Bio films were grown in continuous-culture flow cells ( channel dimensions 1x4 x 40 mm) at room temperature as previously described (Moller et al., 1999). Channels were inoculated with overnight cultures and incubated with no flow for 1 h at room temperature. Cultures were adjusted so that biofilms were established with a flow rate of 150 µl min- 1• R. gallaeciensis, which could not be labelled with a fluorescent colour tag, was stained with Syto 59 diluted to a concentration of 3 µl mr1 in biofilm media. (Bio films were grown in 3M medium containing 0.01 % trehalose, 0.01 % glucose and 0.01 % fructose). Biofilms were visualised with a confocal laser scanning microscope (CLSM) (Olympus) using fluorescein isothiocyanate and tetramethyl rhodamine isocyanate optical filters.

To cultivate mixed biofilms, flow chambers were inoculated with 500 µl of a mixture of stationary-phase cultures of P. tunicata and a competitive marine strain. In order to ensure that the initial ratio of attached cells of the two competing strains was 1: 1, initial attachment of the mixed culture was monitored after a 1 h adhesion period. In most cases, initial levels of attachment to the glass surface was approximately equal, as determined by counting the numbers of red- and green- labelled cells by epifluorescence microscopy using an eyepiece grid. However, for C. fucicola and Alteromonas sp. it was necessary to increase the ratio of cells to 4: 1 and 2: 1, respectively, in order to achieve equal levels of attachment for the two competing strains.

51 To investigate in more detail whether microcolonies improved the competitiveness of test strains, marine bacterial strains were allowed to pre-establish for 24 h (P. gracilis) and 48 h (C. fucicola and Alteromonas sp.). Each pre-formed biofilm was inoculated with - 10 7 cells of the wild type P, tunicata and the flow stopped for 1 h. After resumption of the flow, the biofilm was examined for red and green fluorescence. The experiments were run in three separate rounds with three independent flow cells running in parallel.

2.3 RESULTS AND DISCUSSION

2.3.1 Characterisation of marine strains

2.3.1.1 16S rDNA sequencing

I first carried out partial 16S rDNA sequencing of bacteria isolated from Ulva australis (Table 2.1) and found that the four isolates studied all exhibited 16S rDNA homology with organisms that are commonly isolated from marine eukaryotic surfaces. Cellulophaga fucicola belongs to the Flavobacterium group of the Bacteroides, a diverse group with members commonly found in coastal marine regions (Pinhassi et al., 1997). C. fucicola is frequently found on the surface of marine benthic macroalgae (Bolinche5 et al., 1988) and has been found to decompose highly polymeric material from the brown alga Fucus serratus (Johansen et al., 1999). The sequenced isolates in my collection also contained Pseudoalteromonas and Alteromonas spp. The newly derived genus, Pseudoalteromonas resulted from the division of the genus Alteromonas into two genera, Alteromonas and Pseudoalteromonas based on phylogenetic comparison by Gauthier et al., ( 1995). Numerous bacteria of these genera are frequently isolated from marine waters and are found in association with marine invertebrates, algae, plants and animals (Holmstrom and Kjelleberg, 1999). They are readily culturable and are also capable of surviving in nutrient poor environments. P. gracilis is frequently found on algae such as Graci/aria and produces disease symptoms due to its agarolytic activity (Schroeder et al., 2003). Members of the genus Roseobacter are often found on the surfaces of algae (Brinkmeyer et al., 2000; Meusnier et al., 2001 ).

52 Table 2.1 16S rRNA gene identification of bacteria isolated from the marine alga Ulva australis.

Highest Percentage identity to Strain Closest relative sequence in database 2.06 97 Cel/ulophaga facicola 2.1 99 Roseobacter gal/aeciensis 2.14 95 Pseudoalteromonas gracilis 2.19 99 Alteromonas sp.

Table 2.2 Drop test activity for the detection of extracellular inhibitory compounds active against Pseudoaltermonas tunicata and sensitivity of each strain to the P. tunicata antibacterial protein AlpP, as determined by the drop test assay. a A score of 3 indicates a high level of sensitivity (complete inhibition of growth in the region of the droplet), and a score of 1 indicates slight sensitivity (partial inhibition of growth in the region of the droplet, which appears turbid with bacterial growth).

Scores with the following targetsa Alteromonas Producer P. tunicata P. gracilis C.facicola R. gallaeciensis s . P. tunicata 3 0-1 1 3 1 P. gracilis 1 1 0 2 1 Alteromonas sp. 1 0 0 2 1 C.facicola 0 0 0 0 0 R. gallaeciensis 3 3 1 0 1

53 2.3.1.2 Production of extracellular antibacterial compounds

I examined the ability of the bacterial isolates from U australis to inhibit the growth of each of the other isolates from the plant. Studies of the antibacterial activity in concentrated supematants indicated that P. tunicata and R. gallaeciensis were the most inhibitory (Table 2.2). Each of these strains could inhibit the growth of several of the bacterial strains used in this study. P. tunicata exhibited strong inhibitory activity against itself and C. fucicola, and weak inhibition of P. gracilis. The 190 kDa protein (AlpP) responsible for the antibacterial and autotoxic activity of P. tunicata has now been well characterized (James et al., 1996). This protein contains at least two subunits of 60 and 80 kDa, which are joined by noncovalent bonds. It was shown to be released during stationary phase (James et al., 1996). P. tunicata possesses a ToxR-like regulon, which appears to control determinants for the expression of iron uptake, and also regulates expression of AlpP (Stelzer, S., Egan, S., and Kjelleberg, S. unpublished data).

R. gallaeciensis exhibited strong activity against both P. tunicata and P. gracilis. Production of secondary metabolites by members of this group has been reported previously (Ruiz-Ponte et al., 1999; Boettcher et al., 2000; Gram et al., 2002). One of the inhibitory compounds (proposed to be a peptide) has been reported to be produced only in the presence of other bacteria (Ruiz-Ponte et al., 1999), although in this study I observed strong inhibitory activity with pure culture supematants of this strain. More recent work indicated that R. gallaeciensis also produces a new antibiotic called tropodithietic acid (Brinkhoff et al., 2004). This compound showed strong inhibitory activity against marine bacteria of various taxa and marine algae.

C. fucicola did not demonstrate antibacterial activity against the strains used in this study. However, a Cytophaga strain (RB 1057) has previously been shown to produce an extracellular inhibitor of colony expansion of closely related Cytophaga strains (Burchard and Sorongon, 1998). The inhibitor (a glycoprotein), inhibited the competing strain's ability to adhere to, and glide on, a substrate. The inhibitor had no measurable impact on several other related strains of gliding bacteria. Thus inhibitors produced by this genus may only be effective against closely related bacteria, which may explain

54 why no activity was detected for the Cellulophaga strain tested in this study. In their studies on antagonistic interactions among marine pelagic bacteria, Long and Azam found that the members of the Bacteroidetes group were the most sensitive to inhibition by other marine bacteria and were also the least inhibitory (2001). Similarly, Grossart and coworkers found that the Flavobacterium-Sphingobacterium group had the lowest percentage of inhibitory strains among a diverse group of 51 marine bacterial isolates (Grossart et al., 2004).

Alteromonas sp. showed activity against P. tunicata, C. fucicola and R. gallaeciensis. Alteromonas spp have been shown to produce a wide range of inhibitory compounds. Some species, such as A. citrea (now known as Pseudoalteromonas citrea) and A. rubra, produce only a macromolecular polyanionic substance, whereas other species such as A. luteoviolaceus, produce both a diffusible macromolecule and two intracellular low molecular weight brominated compounds (Gauthier and Flatau, 1976). Barja et al., (1989) found that Alteromonas species isolated from seaweeds produced thermolabile low molecular weight inhibitors (molecular sizes less than 2000), whereas strains isolated from seawater produced a high molecular weight glycoprotein (90,000), which displayed a broad inhibitory spectrum against clinical and environmental isolates. An Alteromonas strain (SW A T5) derived from particulate organic matter was found to produce 2-alkyl-quinolinols. The antibiotics were produced only on polysaccharide matrices and were found to be hydrophobic (Long et al., 2003).

2.3.2 Mono-species biofilm development

Biofilm development of each of the marine species in monoculture was monitored in glass flow cells (Fig. 2.1 ). For all strains, single cells were observed attached to the substratum after inoculation. Each test strain established a stable biofilm, with a characteristic architecture, after 72 h. All of the strains exhibited microcolony formation to various extents during the course of biofilm development. P. tunicata and P. gracilis each formed well-defined spherical microcolonies of up to 50 µm in diameter and 100 µm high within 48 h after inoculation. Alteromonas sp. also formed large, distinct microcolony structures, which were frequently over 100 µm in diameter and 50 µm high after 72 h of biofilm development. In contrast, biofilms formed by R. gallaeciensis

55 P. gracilis

Alteromonas sp.

P. tunicata

C.fucicola

R. gallaeciensis

24 h 48 h 72 h

Fig. 2.1 Single-species biofilm development for bacteri a isolated from the marine alga Viva australis expressing GFP. The test strains are Pseudoalteromonas gracilis, Alteromonas sp., Pseudoalteromonas tunicata, Cellulophaga fucicola and Roseobacter gallaeciensis. Magnification, X600

56 P. tunicata/ P. gracilis

P. tunicata/ Alteromonas sp.

P. tunicata/ C.fucicola

P. tunicata/ R. gallaeciensis

2h 72 h

Fig. 2.2 Competitive biofilm development in two-species biofilms containing Pseudoalteromonas tunicata (expressing GFP) and other marine bacterial isolates (expressing RFP). Magnification, approximately X600.

57 Fig. 2.3 Competitive biofilm development by the P. tunicata AlpP mutant strain that does not produce the antibacterial protein. P. tunicata AlpP mutant (green) had reduced ability to compete against P. gracilis (red) strains. Magnification, X 600.

P. tunicata/ P. tunicata/ P. tunicata/ P. gracilis Alteromonas sp. C.fucicola

Fig. 2.4 Competing marine strains (red) were allowed to form microcolonies before introduction of the superior competitor P. tunicata (green) into the flow cell. Microcolonies enhanced the coexistence of strains in competition with P. tunicata. Microcolonies of test strains (red) were able to persist for more than 72 h in competition with P. tunicata (green). Images were taken at 72 h after inoculation of P. tunicata into a pre-established biofilm. Magnification, X 600.

58 and C. fucicola were less structured. C. fucicola biofilms consisted of both flat unstructured regions of biofilm as well as many small microcolonies up to l O µm in diameter and up to l O µm high. R. gallaeciencis initially formed cell chains which later developed into small microcolonies (10-20 µm in size) during the early stages of biofilm development. These clusters then formed a relatively flat, mat-like biofilm whose thickness approached l O µm. For each test strain, the characteristic architecture formed after 72 h persisted for at least a further 2 days within flow cells. However, after 8 days the majority of the biomass had detached from the flow cells for all of the strains studied, which left only single cells attached to the flow cell reactors.

The marine strains studied here compnse both orgamsms that form microcolonies rapidly and organisms that form less defined microcolonies. Previously it has been proposed that microcolonies can result from gradients of carbon-flux during growth within biofilms (van Loosdrecht et al., 2002). In addition, many genetically encoded regulatory and structural determinants of microcolony formation have been revealed over the past few years. Recent examples include the role of conjugative plasmids (Reisner et al., 2003; Ghigo, 2001) and antigen 43 (Reisner et al., 2003; Danese et al., 2000) in enhancing microcolony formation in Escherichia coli biofilms, suggesting that these cell surface structures might act as cellular adhesins in stabilizing microcolony structures. Generally, the formation of multicellular structures such as microcolonies is thought to be an adaptive response that mediates survival under unfavourable conditions.

Of the marine species studied here, only P. tunicata has previously been characterized for biofilm development under continuous culture conditions. In agreement with the present study, P. tunicata was observed to form prominent microcolony structures during biofilm development (Mai-Prochnow et al., 2004). Biofilm development has been well characterised in the marine bacterium Vibrio cholerae, which also can form microcolonies (Watnick and Kolter, 1999; Watnick et al., 1999; Moorthy and Watnick, 2004). However, there have been no reports that have considered the ecological role of biological structures in mediating interactions between bacteria in mixed-community marine biofilms. Interactions in marine biofilm communities have been investigated by a number of groups (Dang and Lovell, 2000; Grossart et al., 2003; Kiorboe et al., 59 2003). However, these workers focused mainly on succession events or addressed the issue of attachment and detachment of defined pure and mixed cultures on agar beads. In this study, the role of microcolonies during mixed-species biofilm development was also addressed.

2.3.3 Mixed-species biofilm development

Because of its strong inhibitory activity against a broad range of surface-fouling organisms, I explored the possibility that P. tunicata would exhibit an aggressive biofilm-forming lifestyle and would dominate the other marine strains during competitive biofilm development. I therefore prepared mixed 1: 1 inoculae containing P. tunicata (GFP-tagged) and a second organism (dsRed-tagged) and allowed them to attach in a flow cell. In this experiment, P. tunicata totally outcompeted C. fucicola and Alteromonas sp. (Fig 2.2). No cells of the latter competing test strains were visible after 24 h. This is consistent with my observation that P. tunicata produces inhibitory compounds. The results of the drop test assay (Table 2.2) showed that both C. fucicola and Alteromonas sp. are susceptible to inhibitors from P. tunicata. P. gracilis, which has a lower susceptibility to AlpP, was able to coexist with P. tunicata for up to 72 h before it was removed. The outcome of the competition may be correlated with the results of the drop test assay, which showed that P. gracilis is relatively insensitive to the antibacterial protein.

In contrast to the dominance of P. tunicata over C. fucicola and Alteromonas sp., P. tunicata was outcompeted by R. gallaeciensis. While some cells of P. tunicata persisted on the surface throughout the experimental period, these cells did not grow, and R. gallaeciensis rapidly grew and formed a biofilm. This is again consistent with the results of the drop test assay, which showed that P. tunicata is highly susceptible to the inhibitory compounds produced by R. gallaeciensis.

2.3.4 Role of the P. tunicata antibacterial protein AlpP in competitive biofilm development

To evaluate whether the antibacterial protein AlpP can play a role in competitive biofilm development, a specific ~alpP insertional knockout mutant (Mai-Prochnow et al., 2004) was used in competition studies. I first repeated the mixed-inoculum biofilm

60 experiments above, usmg the 11alpP strain instead of the wild type. For these experiments I observed no difference in the outcome of the biofilm competition experiments when using either wild-type or 11alpP strains. When inoculated together with C. fucicola, Alteromonas sp. or P. gracilis, the 11alpP mutant became dominant after 72 h, similar to the data presented for the wild-type strain. Also, R. gallaciences was dominant over the P. tunicata 11alpP strain, in a manner similar to that observed with wild-type P. tunicata.

However, I found that AlpP can play an important role in the colonisation of pre established biofilms of competing marine bacterial strains. As shown in Fig. 2.3, the 11alpP strain had a greatly reduced ability to colonize 24 h-old biofilms of P. gracilis. Under these conditions, wild-type P. tunicata was able to establish within the biofilm, form microcolonies, and completely remove P. gracilis after 120 h. In contrast, introduction of the 11alpP strain led to the establishment of a mixed species biofilm containing separate microcolonies of each strain. However, after 120 h the 11alpP mutant strain was completely overtaken by P. gracilis (Fig. 2.3). When the P. tunicata alpP strain was introduced into preestablished biofilms of C. fucicola and Alteromonas sp., mixed-species biofilms were formed that persisted for over 9 days. In contrast, the wild-type strain was able to remove these competitors after 120 h.

These results suggest that AlpP can provide a competitive advantage in certain ecological situations such as the colonisation of established biofilms. The production of inhibitory compounds by marine organisms appears to be a response to various ecological pressures in the environment. Epiphytic bacteria live in a highly competitive environment where space and access to nutrients are limited. It is possible that P. tunicata up-regulates its production of the antibacterial protein in the presence of competitors; future studies will examine AlpP expression in mixed species biofilms by using a Gfp reporter protein fused to the alpP promoter. Some bacteria demonstrate an inducible chemical antagonism when they are grown in the presence of competing marine bacteria (Mearns-Spragg et et al., 1998; Slattery et al., 2001 ). It is also possible that the antibacterial protein is a key factor when there is competition between microcolonies. Evans and co-workers (Evans et al., 1994) compared protease production in planktonic and biofilm cells and found that it was higher in the latter cells, suggesting that antimicrobial agents were important in biofilms. Furthermore, it 61 has been shown that strains that are normally considered organisms to not produce inhibitors, can express inhibitory activity when growing as a biofilm (Yan et al., 2003). Moreover, there is compelling evidence that mature microcolonies may be the site of inhibitor production in P. tunicata. Production of inhibitors has been linked to pigmentation in P. tunicata (Egan et al., 2001) and mature microcolonies have been observed to become pigmented in biofilms (Webb J.S. and Kjelleberg, S., unpublished data).

2.3.5 Role of microcolonies in competitive interactions within biofilms

To evaluate the role of microcolonies in competition, microcolonies of Alteromonas sp., P. gracilis and C. fucicola were allowed to pre-form before being exposed to P. tunicata. My data suggest that microcolony formation may enhance the ability of the organism to compete against P. tunicata and persist within the flow cell reactor. It was found that P. gracilis biofilms containing microcolonies could co-exist with P, tunicata for more than 5 days, in contrast to biofilms without microcolonies (Fig 2.4). Moreover, pre-established biofilms of C. fucicola and Alteromonas sp. (which normally persist for no more than 24 h in the presence of P. tunicata) were able to persist up to 72 h before being dispersed. Thus, high densities of cells within microcolonies may allow for enhanced persistence during co-culture with the superior competitor, P. tunicata.

The finding that microcolonies enhance the competitiveness of an otherwise poor competitor was confirmed when doing co-culture experiments with the AlpP mutant strain, where pre-established biofilms of C. fucicola and Alteromonas sp. persisted for up to 8 and 9 days respectively (Fig 2.4). It suggests that enhanced microcolony formation would be an advantage in a highly competitive environment where space is limiting and such a conclusion is supported by the outcome of competition studies conducted by other researchers. In dual species biofilms, bacteriocin producing enteric bacteria prevented the colonisation of a potential competitor into a pre-established biofilm (Tait and Sutherland, 2002). Al-Bakri et al., (2004) also found that pre established biofilms of P. aeruginosa conferred colonisation resistance to Burkholderia cepacia.

62 2.4 CONCLUSIONS

By labelling marine bacteria isolated from the green alga Viva australis with genetic colour-tags, I was able to examine the colonisation, biofilm forming behaviour and competitive interactions of these strains during mixed-species biofilm development in real time. This study is the first to demonstrate that inhibitory compounds produced by marine bacteria can provide a competitive advantage during competitive growth within biofilms. It is further suggested that microcolony formation may represent one adaptive strategy that increases the ability of bacteria to persist in mixed-species biofilms under conditions of competitive biofilm formation.

My studies show that the AlpP protein, a potent antibacterial protein produced by P. tunicata, can enhance the ability of P. tunicata to colonize biofilms formed by other bacteria. The wild-type P. tunicata strain aggressively colonized and dominated strains that were sensitive to the AlpP protein. In contrast, a P. tunicata alpP mutant was defective in the ability to colonize and dominate biofilms under the same conditions. Moreover, marine strains that were tolerant to the AlpP protein in drop-plate assays were recalcitrant to invasion and displacement by P. tunicata during biofilm development.

Previous studies from this laboratory have suggested that bacteria present on the surface of some higher marine eukaryotes may play an important role in the chemical defence against biofouling in the marine environment (Holmstrom and Kjelleberg, 1999). Many members of the genus Pseudoalteromonas, for example, produce multiple extracellular inhibitory compounds that target different classes of marine fouling organisms (Holmstrom et al., 1998). Competitive interactions between bacteria such as those demonstrated in this study may play an important role in determining the composition of such "antifouling" communities, because they likely lead to a predominance of inhibitory bacteria on the host surface.

It is also suggested that specific interactions between inhibitor-producing bacteria and eukaryotic host surfaces may enhance the antifouling defense of the host organism. For example, recent studies in our laboratory have revealed that P. tunicata possesses a mannose-sensitive type IV pilus that promotes attachment to the cellulose-containing

63 surfaces of U. australis and Ciona intestinalis, the principal hosts of P. tunicata in the marine environment (Saludes, D., Scheffel, A., James, S., Webb, J. S. Holmstrom, C., and S. Kjelleberg, unpublished data). Both U. australis and C. intestinalis are thought to rely on inhibitor-producing bacteria for defense against biofouling; thus, it may be an advantage to these organisms to become associated with antifouling bacteria such as P. tunicata. The present study provides a platform for further studies of interactions between marine bacteria in surface-associated communities and between biofilms and their eukaryotic host surfaces.

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Chapter Three

Microbial colonisation and competition on the marine alga Ulva australis

3.1 INTRODUCTION

Biofouling is caused initially by bacterial growth and biofilm formation on natural and artificial surfaces. Bacterial biofilms serve as a focus for the attachment and growth of a range of other fouling organisms such as diatoms, invertebrate larvae and algal spores (Davis et al., 1989). Strategies to prevent bacterial biofilms therefore represent a powerful approach to the prevention of biofouling.

Biofouling communities on surfaces of marine plants and animals can have detrimental effects on the host organism. Physical damage to the host can result from the production of toxins, digestive enzymes and waste products by the microbial community. However, the extent of biofouling on marine organisms is markedly less than that on inanimate structures. Often the later stages of biofouling, such as the attachment of algae and barnacles, do not occur (Armstrong et al., 2000a). Contrary to inanimate surfaces, which are colonised in a rapid and predictable manner by a diverse assemblage of marine microbes (Wahl, 1989), biotic surfaces frequently harbour species-specific microbial communities (Wahl, 1995; Taylor et al., 2004; Grossart et al., 2005). These communities can be variable and distinct from those found in the surrounding environment. Thus it is evident that many algal and invertebrate species are able to regulate the bacterial colonisation of their surfaces (Wahl et al., 1994; Jensen et al., 1996; Steinberg et al., 1997).

This chapter is a modified version of: Rao, D., Webb, J.S., and Kjelleberg, S. (Submitted) Microbial colonization and competition on the marine alga Ulva australis. Appl. Environ. Microbiol.

70 Microalgal surfaces are typically covered in bacteria with abundances of approximately 107 bacteria cm-2 (Armstrong et al., 2000b). Studies on the epiphytic microbial communities present on macroalgae have emphasised the spatial distribution of bacteria with specific parts of the thallus playing host to specific bacterial populations (Corre and Prieur, 1990). In some cases the bacterial populations change with the season or the age of the host (Sieburth and Conover, 1965; Sieburth and Tootle, 1981). Associations between algae and bacteria are common and studies have generally focused on the benefits afforded to the bacteria, such as support of bacterial growth by dissolved organic carbon released by algal cells (Lange, 1967; Jones and Cannon, 1986; Rier and Stevenson, 2002).

Although some of the bacterial-algal interactions have been characterised, the ecological significance of most naturally occurring epiphytic bacterial communities is unclear and in many cases the bacterial species involved have not been identified (Fisher et al., 1998). It has been demonstrated that bacterial biofilms are present on the surface of U. australis (previously known as U. lactuca) (Sieburth, 1975), and it is speculated that the seaweed uses microbial defence to protect against fouling (Holmstrom et al., 1992; Egan et al., 2002). U. australis has no known physical or chemical defence systems against fouling organisms and it has been suggested that the host in some way manipulates the bacterial community on its surface, which in tum protects the host by interfering with the development of a mature biofouling community. Such interactions are not uncommon in the marine environment (Gil Tumess and Fenical, 1992; Nakasono et al., 1993; Holmstrom and Kjelleberg, 1994; Boyd et al., 1999; Armstrong et al., 2001; Dobretsov and Qian, 2002).

One important species of marine bacteria that is found in association with U. australis is Pseudoalteromonas tunicata. This bacterium produces a diverse range of biologically active compounds that specifically target marine fouling organisms (Holmstrom et al., 1998). One of these compounds is a 190 kDa multi-subunit antibacterial protein designated AlpP which is effective against both Gram-negative and Gram-positive bacteria from a range of environments (James et al., 1996; Mai-Prochnow et al., 2004). Recent work has demonstrated that AlpP can provide a competitive advantage to P. tunicata during biofilm growth in laboratory biofilm experiments (Rao et al., 2005).

71 Roseobacter gallaeciensis is also frequently isolated from the surface of U. australis. Roseobacter spp. are cosmopolitan and have been isolated, for example, from green seaweeds (Shiba, 1992), marine snow particles (Gram et al., 2002) and dinoflagellates (Lafay et al., 1995; Alavi et al., 2001; Miller and Belas, 2004). Roseobacter spp. are able to metabolise dimethylsulfoniopropionate (DMSP), and their presence and activity is significantly correlated with (DMSP)-producing algae, including dinoflagellates and prymnesiophytes (Gonzalez and Moran, 1997).

The extensive, highly diverse microbial community associated with Ulva australis makes it an interesting study organism for addressing questions of surface colonisation and host association. The dynamics of surface colonisation in natural systems, particularly during the early stages of biofilm establishment are poorly understood in marine bacteria. For example, an assessment of the ability of distinct bacteria to colonise algal surfaces would be useful in identifying bacterial traits that contribute to epiphytic fitness. Although competitive biofilm interactions of P. tunicata and other marine strains have been studied in a laboratory glass flow cell system (Rao et al., 2005), little is known about the ecology of colonisation and competition on a living surface. Further, it is not known whether P. tunicata is a dominant competitor in ecologically relevant settings.

This study aimed to investigate, for the first time, the colonisation biology of marine bacteria on the surface of a marine plant, in this case U. australis. Here, I investigated the hypothesis that P. tunicata and R. gallaeciensis are effective colonisers of U. australis and are able to compete and dominate over other marine bacterial isolates during biofilm formation on the plant surface. It was found that P. tunicata requires the presence of a natural seawater community to colonise effectively, whereas R. gallaeciensis is an aggressive coloniser under all conditions tested. Results presented in this chapter highlight the differences in colonization strategies exhibited by P. tunicata and R. gallaeciensis and demonstrate a role for the antibacterial compounds in the colonization of U. australis. Thus microbial colonisation of plant surfaces is a dynamic process where differences in attachment, colonisation and competitive biofilm formation can markedly affect the establishment and organisation of epiphytic microbial communities.

72 3.2 MATERIALS AND METHODS

3.2.1 Collection of plants and generation of axenic U. australis

The common marine alga Ulva australis, was collected from the rocky intertidal zone at Shark Point, Sydney, Australia (33°91 '21" S, 151 °25'72"E). The method used to make U australis axenic was modified from that described by Scheffel (Scheffel, 2003). Briefly, plants were rinsed in 50 ml autoclaved seawater and cut into 5 cm long fragments. These were swabbed with cotton buds and 0.6 cm discs were punched out of the thallus using a cork borer. The discs were rinsed in autoclaved seawater and incubated in 0.012% sodium hypochlorite (NaOCl) for 5 minutes. Discs were allowed to recover in sterile seawater for 1 hour before being incubated in an antibiotic cocktail

(Ampicillin 300 mgr1, Polymyxin B 30mgr1, Gentamycin 60 mgr1 dissolved in sterile seawater) for 18 hours in 24 well tissue culture plates (Falcon). Discs were then incubated in 0.008% NaOCl for 5 minutes. These discs were recovered in sterile seawater for at least 3 hours to remove traces of oxidants and suspended in 2 ml of sterile seawater and agitated at room temperature at 60 rpm.

Examination of plant tissue viability was carried out by staining treated U australis discs with 0.25% (w/v) Evans blue (Swain and De, 1994). The blue coloured dye penetrates into dead and damaged plant cells while intact cells are able to exclude the dye. All colonisation and competition experiments were done in triplicate, and three to five individual discs were randomly sampled at each sampling time.

3.2.2 Colonisation experiments

Bacterial strains were isolated from the surface of U australis as described previously (Rao et al., 2005). Cultures were stored at -80°C in 50% (vol/vol) glycerol in VNSS media (Marden et al., 1985) and maintained on VNSS plates. P. tunicata and R. gallaeciensis were labelled with a green fluorescent protein (GFP) colour tag as described previously (Rao et al., 2005). Bacteria were cultured for 24 h at 25°C in VNSS broth for preparation of inoculae. Cells were harvested by spinning down the culture and resuspending the pellet in seawater. Cell concentration was estimated by counting under epifluorescence microscopy using a haemocytometer and adjusted by dilution with seawater to the desired concentration. Experiments were conducted in

73 Falcon 24 well plates with axenic discs and the bacteria were applied by immersing the discs in a suspension of P. tunicata and R. gallaeciensis for 3 hours. The discs were then rinsed twice in filtered seawater and transferred to fresh Falcon 24 well plates containing 2 ml of filtered seawater. The plates were incubated on a shaker at 60 rpm at

25°C (16 h photoperiod at 20 µEm· 2s- 1).

3.2.2.1 Factors affecting colonisation

In order to investigate some of the factors that were essential for colonisation of P. tunicata, a range of conditions were tested:

3.2.2.1.1 Densities of P. tunicata required for attachment P. tunicata was inoculated on axenic U australis discs in 24 well tissue culture plates at

a range of concentrations (104 - 108 cells mr1). Discs were incubated for 3 h, rinsed twice in sterile seawater and then incubated at 25°C at 60 rpm for the duration of the experiment.

3.2.2.1.2 Axenic vs non-axenic plant surfaces Attachment of P. tunicata cells, which had been inoculated at 107cells mr', was compared on the two surfaces of U australis discs.

3.2.2.1.3 U. australis discs vs whole plants Attachment was tested on axenic discs in 24 well plates as well as on axenic whole plants (which had been treated in the same way as the discs). The plants were oriented upright in beakers and immersed in filtered seawater.

3.2.2. 1.4 Dark vs Light Inoculation was conducted in the dark at 107cells mr1 with incubation in the dark for 3 hours. This was compared to inoculation and incubation for 3 h in the light at the same densities.

3.2.2.1.5 Carbon source Standard culture conditions for marine strains were on VNSS media containing glucose. However, cells were also grown for 48 h in minimal media (Neidhart et al., 1974) with cellobiose as the sole carbon source in a still culture. The cells were inoculated onto U.

74 australis discs and attachment was compared to that of cells grown in minimal medium with trehalose and glucose as carbon sources.

3.2.2.1.6 Incubation time Discs were inoculated with 107cells mr1 and incubated for 1, 3, 12 and 24 h respectively.

3.2.3 Effect of multispecies consortia on colonisation

Axenic algal discs were inoculated with overnight cultures of GFP-labelled P. tunicata and 17 unlabelled strains isolated from the surface of U. australis (Rao et al., 2005) in filtered seawater. P. tunicata was also inoculated in a natural seawater community. The resulting mixed species biofilm was visualised by staining with acridine orange. Attachment and colonisation of GFP-labelled P. tunicata within the mixed biofilm was visualised by observing unstained biofilm under a confocal microscope. Attachment, biofilm formation and microcolony development were monitored by epifluorescence microscopy every 24 h for a period of up to three weeks. At each sampling time, three discs were randomly selected and a total of 12 random fields of view counted for each disc. Samples were viewed by epifluorescence microscopy with an Axiophot microscope equipped with a 1Ox, 1.30 numerical aperture objective (Leica).

As R. gallaeciensis was an aggressive coloniser of U. australis, attachment of this bacterium was not tested under a range of conditions. I did however examine the effects of different densities on colonisation and tested a range of concentrations of cells in the innoculae (104 - 108 cells mr1).

3.2.4 Competition in dual-species biofilms on U. australis

The other marine strains used for competition experiments were Pseudoalteromonas gracilis, Alteromonas sp. and Cellulophaga fucicola, and were isolated from U. australis as described previously (Rao et al ., 2005). The strains were labelled with red fluorescent protein (DsRed) and green fluorescent protein (GFP) as described previously (Rao et al., 2005). Wild type marine isolates and their labelled derivatives showed no significant differences in their growth rate on the surface of axenically treated U. australis.

75 Overnight cultures of bacteria were inoculated onto U. australis at densities of 108cells mr1 in filtered and natural seawater and incubated for 3 hours without shaking at room temperature. Discs were rinsed three times with filtered seawater and transferred to sterile seawater in Falcon 24 well plates and incubated at 25°C at 60 rpm for the duration of the experiment. Unlabelled biofilms were visualised on the surface of the plant by staining with acridine orange. Biofilms were grown in filtered seawater and visualised with a confocal laser scanning microscope (CLSM) (Olympus) using fluorescein isothiocyanate and tetramethyl rhodamine isocyanate optical filters for green and red fluorescent proteins, respectively. GFP-labelled P. tunicata was inoculated onto the surface with competing marine strains, including R. gallaeciensis. To investigate whether the antibacterial protein produced by P. tunicata improved the competitiveness of this organism on the surface of U. australis, competition studies were also conducted with the AlpP mutant of P. tunicata (Mai-Prochnow et al., 2004).

To cultivate mixed biofilms, discs were inoculated with 500 µI of a mixture of stationary phase cultures of P. tunicata and the competitive marine strain in a suspension of filtered seawater. In order to ensure that the initial ratio of attached cells of both competing strains was 1: 1, initial attachment of the mixed culture was monitored after a 3 h adhesion period. In most cases, initial attachment to the plant surface was deemed to be approximately equal as determined by counting the numbers of both red and green labelled cells under epifluorescence microscopy.

To investigate whether P. tunicata and R. gallaeciensis were able to invade and colonise an established biofilm, marine strains (P. tunicata, C. fucicola and Alteromonas sp.) were allowed to pre-establish for 48 hours on U. australis. The pre formed biofilm was inoculated with - 10 8 cells of the wild type P. tunicata or R. gallaeciensis and incubated for 3 h without shaking and then incubated at 60 rpm for the duration of the experiment. After the resumption of shaking, the biofilm was examined for red and green fluorescence. Experiments were repeated in three separate rounds with three independent 24-well plates running in parallel. Viability of bacterial cells was determined using a Live-Dead staining kit (Molecular Probes Inc, Eugene, Oreg).

76 3.3 RESULTS

3.3.1 Obtaining axenic plant tissue

In order to conduct the attachment and colonisation experiments it was necessary to remove the majority of bacteria on the U. australis thallus surface. Observation of untreated algal tissue demonstrated a complex community of epiphytic bacteria (Fig.

3. lA). However, treated axenic tissue had negligible numbers (< 50 cells cm-2) of attached solitary cells on the plant surface (Fig 3.1B). Antibiotic treatment resulted in approximately 90% of epiphytic bacteria being removed while causing minimal damage to the U. australis tissue (as determined by the Evans blue test for plant viability). Axenic discs that were not inoculated with bacteria in the course of the experiments bleached rapidly and died within a week, indicating that a surface population of bacteria is necessary for plant survival.

3.3.2 Factors influencing attachment and colonisation.

Preliminary experiments showed that P. tunicata colonised the surface of U. australis poorly, which led me to test attachment under different conditions. The data provides quantitative information on the attachment of P. tunicata on the surface of U. australis under a range of different environmental conditions (Table 3.1). P. tunicata harbouring a GFP reporter gene was visualised by epifluorescence microscopy to differentiate P. tunicata cells from indigenous bacteria on the algal surface. Cells were observed after a 3 h attachment period, and it was seen that they were not randomly scattered over the U. australis thallus surface. Rather, cells were distributed in patches over the plant surface. Observations after a 24 h growth period showed that some of the attached cells had divided and formed microcolonies, which occurred in a wide range of sizes (5 -50 µm). Observations after 3 days of incubation revealed that vast areas of U. australis thallus tissue remained uncolonised with microcolonies restricted to certain regions. On the other hand, R. gallaeciensis formed microcolonies which were dispersed evenly over the surface of the thallus tissue. At 24 h, R. gallaeciensis had formed cell chains, which later developed into small microcolonies (10-20 µm in size), by day 3. By day 7, these microcolonies had merged to form a relatively flat, mat~like biofilm which covered a large proportion of the algal surface.

77 Table 3.1 Factors influencing attachment and colonisation of Pseudoalteromonas tunicata on Viva australis. Note: For testing all the other factors apart from density, a density of 107 cell mr1 was used. Counts were average values of 12 fields of view for each disc, which were then converted to number cm-2. * indicates significant values.

Factors No of cells attached ( Cells cm-2) Mean ± SE 24 h 3 days 7 days Density 104 cell mr1 0 0 0 Density 105cell mr1 0 0 0 Density 106 cell mr1 52 ± 3.0 13 ± 2.5 0 Density 107 cell mr1 224 ±4.9 92 ± 4.9 0 Density 108 cell mr1 1880±58 * 848 ± 43 0 Axenic discs 290 ± 11.4 125 ± 3.9 0 Non-Axenic discs 270 ± 10 131 ±9.7 0 Carbon source - Trehalose 250 ± 6.9 112 ± 3.6 0 Carbon source - Cellobiose 460 ± 7.7 * 276 ± 9.9 0 Incubation time - 1 h 184 ± 6.5 72 ±4.7 0 Incubation time - 3 h 302 ± 9.3 * 118 ± 3.9 0 Incubation time - 12 h 289 ± 11.4 125 ± 3.9 0 Incubation time - 24 h 263 ± 9.2 99 ± 4.4 0 Inoculation in light 250 ± 6.8 79 ± 7.2 0 Inoculation in the dark 374 ± 12.6 * 171±7.2 0 P tunicata in autoclaved seawater 250 ± 6.6 92 ± 5.2 0 P tunicata in filtered seawater 263 ± 5.8 92 ± 5.8 0 P tunicata in natural seawater 539± 15.1 * 934 ± 26.2 1880 ± 74 *

78 3.3.2.1 Effects of inoculum cell density on colonisation of U. australis

Direct in situ observation of epiphytic bacteria revealed that the density of cells within the inoculum affected microcolony formation. After an initial attachment period (3 h), I incubated discs for 24 h and then examined the plant surface for bacterial cells and microcolony formation. Surprisingly, no attachment took place when P. tunicata was inoculated at low densities (104 or 105 cells mr1) (see Table 3.1). At densities of 106 cells mr1 I observed that few cells attached (52 cells cm-2). These were mostly solitary, with no microcolonies forming and cells did not usually persist beyond 3 days. At densities of 107 cells mr1 some of the attached cells were clustered together and formed small microcolonies consisting of about 15-20 cells after 24h. With 108 cells mr1, microcolony sizes were considerably larger (up to l00µm across) after the same growth period (24 h). In spite of the large size attained by microcolonies derived from high cell densities, P. tunicata did not persist beyond 5 days. Although R. gal/aeciensis was able to colonise at lower densities ( 104 cells mr I and 105cells mr 1), persistence was much improved at higher densities (106 - 108cells mr1). At higher densities R. gallaeciensis rapidly formed microcolonies that survived and persisted indefinitely on the plant compared to those at low density, which did not persist beyond 5-6 days. Thus, both organisms exhibited density dependent colonisation with microcolony formation enhanced at high cell densities.

3.3.2.2 Colonisation in the dark

On the basis of my observation of surprisingly low levels of attachment of P. tunicata on the plant surface, I investigated the possiblity that P. tunicata attachment may be affected by the presence of DMSP on the plant surface. It is known that DMSP is released in senescent algae or when algae undergo undergo oxidative stress particularly under high light intensities (Karsten et al., 1990). I therefore examined colonisation of U. australis by P. tunicata in the dark and observed that attachment of P. tunicata improved when cells were inoculated in the dark, with the number of cells attaching increasing from 250 cells cm·2 to 374 cells cm·2 (Table 3.1). However, the cells did not persist and form biofilms. Most of the microcolonies formed were small, although in

79 greater numbers (about 16-20 microcolonies per cm-2 with 12-25 cells in each microcolony compared to light incubated plant tissue which had about 3-6 microcolonies per cm-2 with 50-90 cells in each microcolony). I repeated these sets of experiments using the Live-Dead stain, to establish whether cells in smaller microcolonies were dying or whether they were simply failing to attach. However, the results were inconclusive as it was difficult to visualise the dead cells against the red autofluorescence of the algae. In contrast, R. gallaeciencis was able to attach and colonise at 106 cells mr1 regardless of whether inoculation was conducted in the light or dark.

3.3.2.3 Carbon source

It was found that P. tunicata cells pre-grown in media containing cellobiose as a sole source of carbon were able to attach better on the surface of U australis. As illustrated in Table 3.1 and Fig. 3.2A, P. tunicata cells grown in cellobiose had a much higher number of cells that were able to attach to the surface of the algae, as compared to those grown in glucose or trehalose as a carbon source (Fig 3.2B). However, similar to glucose/trehalose grown cells, these cells did not persist beyond 5 days on the plant surface.

3.3.2.4 Inoculation in natural seawater and mixed species biofilm development

I compared colonisation of U. australis by P. tunicata cells suspended in each of sterile-filtered, autoclaved and natural seawater. I observed that in sterile-filtered water, cells did not persist beyond 5 days (Fig. 3.3A) (Table 3.1) and colonisation was most effective in natural seawater. Not only was there a higher level of attachment in this system, but cells also became established and persisted indefinitely on the surface of the algal thallus. Together the bacteria formed a mixed biofilm, which was stable and attained a thickness of up to 20 µm (Fig. 3.3B). When P. tunicata was inoculated in combination with a mixture of 17 uncharacterised marine strains isolated from U australis, it was also found to form mixed biofilms and persist on the surface indefinitely (Fig. 3.3C). These were more stable than single species or dual species biofilms. In contrast to P. tunicata, R. gallaeciensis was able to colonise the surface of the alga regardless of whether the cells were inoculated in sterile or natural seawater.

80 A R

Fig. 3.1 Effect of axenic treatment on the U. australis epiphytic community. (A) Non-axenic U. australis with the natural microbial community present. (B) Axenic U. australis with very few bacteria remaining. Only plant cells are visible. Specimens were stained with 0.01 % acridine orange. Images were taken at 600X magnification.

24h 48h 72h

Fig. 3.2 A comparison of attachment and colonisation by GFP-labelled P. tunicata grown in cellobiose (A), as compared to glucose (B). Images were taken at 600X magnification.

81 A

Day 2 Day 7 Day 11 Day 18

Fig. 3.3 A comparison of attachment and biofilm formation by P. tunicata cells suspended in sterile seawater, natural seawater and in a mixture of 17 epiphytic strains. (A) P. tunicata inoculated in filtered seawater does not persist beyond 3 days. (B) P. tunicata inoculated in natural seawater is able to attach to V. australis and form biofilms that persist for up to 3 weeks. (C) P. tunicata inoculated with a defined mixture of 17 strains isolated from V. australis, formed a complex biofilm that is able to persist indefinitely. Specimens were stained with 0.01 % acridine orange. Images were taken at 600X magnification.

82 A

Day 3 Day 5 Day7 Day9

Fig. 3.4 Competitive biofilm development in pre-established biofilms on U. australis. (A) In competitive biofilm development, P. tunicata (Green) is able to dominate in certain situations, but more commonly it is able to coexist with P.gracilis (Red). (B) P. tunicata (Green) is however outcompeted by R. gallaeciensis (Red). (C) The P. tunicata AlpP mutant strain (Green) is less competitive compared to the wild type P. tunicata as seen in competitive biofilm development with P. gracilis (Red). Images were taken at 600X magnification.

83 3.3.3 Competitive biofilm development on U. australis.

I examined competitive interactions in mixed species biofilms during co-colonisation of the surface of U australis in viva. After initial attachment to the plant surface, P. tunicata totally out-competed C. fucicola, Alteromonas sp. and P. gracilis, and no cells of the latter strains were visible after a 24 h incubation period. In contrast, P. tunicata was out-competed by R. gallaeciensis in these experiments. When C. fucicola, Alteromonas sp., or P. gracilis were each allowed to pre-establish a biofilm on U lactuca for 48 hours before being challenged with P. tunicata, they were able to co exist with P. tunicata for the duration of the experiment. Under these conditions, each of the competing strains persisted on the plant surface in discrete monospecies microcolonies, without mixing of the competing organisms. P. tunicata microcolonies were found to coexist with P. gracilis microcolonies on the plant surface (Fig. 3.4A). Generally, microcolonies appeared as protective structures because, once established, removal of microcolonies by competing strains was rarely observed under the conditions used in the experiments. An exception to this finding occurred in competition experiments involving R. gallaeciensis; this organism was able to colonise and out-compete a pre-established biofilm as effectively as it colonised the axenic surface of U australis (3.4B).

To investigate the role of the antibacterial protein AlpP of P. tunicata in competitive biofilm development on U australis, I repeated the mixed-inoculum biofilm experiments above, using the 11alpP strain instead of the wild type. When competing strains were inoculated simultaneously, no difference in the outcome of the biofilm competition experiments when using either wild-type or 11alpP strains was observed (data not shown). However, I found that AlpP can play an important role in the colonization of pre-established biofilms of other marine strains on U australis. The 11alpP strain had a greatly reduced ability to colonize and establish in 48 hour-old biofilms of P. gracilis, C. fucicola and Alteromonas sp. For example, the 11alpP strain is unable to colonise and invade a pre-established biofilm of P. gracilis (Fig. 3 .4C). The mutant was unable to form microcolonies and the few attached cells had little or no impact on the proliferation of competing strains.

84 3.4 DISCUSSION

Little is known about the ecological role of microbial communities on the surfaces of marine algae. However, the abundance of bacteria that produce extracellular inhibitory compounds on the surface of the marine alga U. australis has prompted speculation that they may protect the algae against fouling (Holmstrom et al., 1992; Egan et al., 2002). In this study I explored some of the factors that influence attachment and colonisation of U. australis by epiphytic bacteria, as well as the competitive interactions that occur between bacterial strains on the plant surface.

3.4.1 Effect of cell densities on attachment

One of the most important factors that influenced the attachment of P. tunicata, was the density of cells in the inoculae. Not surprisingly, I found that the number of cells attached to the surface was higher when inoculae with higher concentrations was used. However, I also observed that surprisingly high densities of cells were necessary for P. tunicata to establish and grow in microcolonies. Although R. gallaeciensis was able to attach at lower densities, biofilm formation and colonisation was much improved at higher densities in a similar manner to P. tunicata, perhaps also because high cell density inoculae allowed for the formation of larger numbers of microcolonies by this organism. The dependence on high cell densities for microcolony formation has been previously reported for biofilms on leaf surfaces (Wilson and Lindow, 1994b; Monier and Lindow, 2003, 2004). In these studies, there was evidence that cells in aggregates exhibited a much greater ability to survive periodic desiccation stress, and increased tolerance was correlated with the size of aggregates (Monier and Lindow, 2003). Presumably cells in larger microcolonies are more tolerant of oxidative stress than those located in smaller microcolonies (Costerton et al., 1999) and may also be more capable of concentrating nutrients from dilute sources which can have important consequences in a nutrient poor environment.

The mechanism by which high cell densities allow for enhanced microcolony formation is unclear, but may point to a role for quorum sensing in the colonization of U. australis. Quorum sensing is known to play a role in the formation of multicellular structures within biofilms (Davies et al., 1998; Hentzer et al., 2002). To date there is no evidence that P. tunicata produces signalling molecules (Franks, 2005), but recent

85 studies from our laboratory have shown that R. gallaeciensis produces N-acyl homoserine lactones (AHLs) suggesting that it is capable of quorum sensing (Case, R., Low, A., Kjelleberg, S., unpublished data). In some epiphytic bacteria, quorum sensing has been shown to play a role in colonisation of the plant and studies with Pseudomonas syringae suggest that AHL production is important in contributing to epiphytic fitness in the early stages of colonisation of bean leaves (Quinones et al., 2004).

3.4.2 Attachment in the dark

Inoculation in the dark clearly enhanced attachment of P. tunicata, indicating that this organism may be sensitive to DMSP or reactive oxygen species present on the surface of the algae. DMSP occurs in many diverse species of algae including U australis (Dickson and Kirst, 1986) and is released in senescent algae or under conditions of oxidative stress, particularly under high light intensities (Karsten et al., 1990; Sunda et al., 2002). There is evidence that DMSP provides protection against photooxidation for algae (Sunda et al., 2002) but its effects on epiphytic bacterial communities on Ulva spp. are not fully understood (Nishiguchi, 1994). I observed that attachment of P. tunicata in the light resulted in fewer but larger microcolonies, whereas microcolonies established from cells inoculated in the dark were smaller and more numerous. One possibility is that cells in larger more resistant microcolonies survived exposure to oxidative stress, which increases with higher light intensities. R. gallaeciensis was able to attach and colonise effectively regardless of light conditions suggesting that it may be able to tolerate and/or metabolise DMSP (Sunda et al., 2002). In studies of the diversity of OMS-producing bacteria from oceanic and estuarine waters it was found that all the isolates tested from the Roseobacter group were DMS producers and therefore could utilise DMSP (Gonzalez et al., 1999). Another proposed ecological role for DMSP is that it acts as a precursor of cues for chemosensory attraction between algae and certain specific bacteria (Simo, 2001). Thus, DMSP may be one of the chemical signals released by U australis which is recognised by R. gallaeciensis.

3.4.3 Effect of cellobiose on attachment

It is evident that P. tunicata attachment improved considerably when cells were grown in cellobiose, but the cells did not persist on the surface of U australis. Both U lactuca 86 and C. intestinalis, from which P. tunicata is most commonly isolated, contain accessible cellulose polymers in their cell walls ( de Leo et al., 1977; Baldan et al., 2001). A cellulose binding protein with a high binding affinity for microcrystalline cellulose was discovered in P. tunicata (Dalisay-Saludes, 2004). Clearly, cellobiose plays a role in attachment, but its effects may be restricted to the early stages of colonisation where it has been suggested to function as an anchorage or cue for P. tunicata attachment (Skovhus, 2004). Recent studies have found that cellulose stimulated the production of pigment in this organism which is known to be coregulated with the expression of the AlpP (Skovhus, 2005) and also induced the expression of MSHA (Type IV-like) pili in P. tunicata (Dalisay-Saludes, 2004). Thus, cellulose appears to play an important role in the colonisation of U australis by providing a cue for the initial attachment of P. tunicata to the plant surface.

3.4.4 Synergistic biofilm formation

P. tunicata persisted on the surface of the algae only if it was inoculated in seawater that contained a natural seawater community, or if it was inoculated in filtered seawater that contained a mixture of epiphytic strains isolated from U australis. It suggests that P. tunicata requires the presence of diverse bacteria within the inoculum for effective colonisation, which may allow for succession and or cooperative colonisation of the surface. Co-colonising bacteria could modify the habitat and create a 'microenvironment' that encourages the attachment and growth of other colonising microorganisms. I tested whether a pre-existing biofilm enhances the attachment and persistence of P. tunicata by allowing it to colonise non-axenic U australis discs with the intact natural community on the surface. However, P. tunicata was unable to colonise under these conditions, indicating that an established biofilm of other bacteria does not facilitate its colonisation. Another possibility is that co-operative interactions during the process of colonisation (for example co-metabolism or co-aggregation) can occur between P. tunicata and other co-colonising marine strains. The phenomenon of cooperative biofilm formation in mixed consortia has recently been described for other bacteria (Filoche et al., 2004; Sharma et al., 2005). Of the 17 strains present, it is not clear which individual strains or mixtures of strains is responsible for allowing P. tunicata to colonise. The underlying mechanisms of cooperative biofilm formation are the basis of ongoing experiments at the Centre for Marine Biofouling and Bio-

87 the basis of ongoing experiments at the Centre for Marine Biofouling and Bio innovation.

3.4.5 Competition in biofilrns on U. australis

In co-inoculation competition studies, I observed that P. tunicata out-competed all of the other marine isolates tested, except R. gallaeciensis. The results are similar to my previous findings of competition on glass surfaces within a laboratory flow cell model (Chapter 2). In that study, it was seen that competition was largely controlled by the production of inhibitory compounds and the relative sensitivity of competing strains towards the inhibitors (Rao et al., 2005).

However, I did observe some important differences between bacterial competition on the surface of the marine plant when compared with laboratory systems in the pre established biofilm studies. Unlike laboratory flow cells, which allow for a continuous flow of nutrients, the surface of algae likely presents nutrient limited conditions. My studies demonstrated that exogenous application of nutrients resulted in a rapid increase in microcolony size of P. tunicata (data not shown), suggesting that microcolonies on the plant surface are indeed nutrient limited. Furthermore, my observation of the patchy distribution of microcolonies might reflect the spatial heterogeneity of nutrients available on algal surfaces. Several studies conducted on higher terrestrial plants suggest that most areas of a leaf harbour only small amounts of nutrients (Weller and Saettler, 1980; Leben, 1988) and bean plants inoculated with Pseudoamonas syringae strain B728A, had aggregates of bacteria distributed nonrandomly in a wide range of cluster sizes, which roughly corresponded to nutrient availability (Leveau and Lindow, 2001 ). My observations that P. tunicata colonised a pre-established biofilm poorly, suggest that the initial colonisers could deplete a large percentage of carbon sources and make it more difficult for the invading bacteria to sequester themselves in the existing biofilm community. This has been described for epiphytic bacteria on bean leaves where a pre-established biofilm can deplete resources leading to pre-emptive exclusion (Wilson and Lindow, 1994a).

In competition studies where P. tunicata was allowed to invade a pre-established biofilm, it did not out-compete the other strains and coexisted with the competing strain

88 for the duration of the experiment. When invading a pre-established biofilm, P. tunicata tends to colonise and establish microcolonies in areas which remain free from colonisation by a pre-established biofilm. This resulted in limited interactions between spatially segregated microcolonies of competing strains and led to coexistence of strains. My observations of competitive interactions on the surface of U. australis are supported by recent reports on spatial segregation of epiphytic bacteria on leaf surfaces (Monier and Lindow, 2005a; Monier and Lindow, 2005b). The observations differ from flow cell experiments where P. tunicata is able to dominate and eventually remove certain competing stains (Rao et al., 2005). Evidence of reduced ability to colonise a pre-established biofilm was even more pronounced for the AlpP mutant. The mutant was impaired in its ability to form microcolonies in a pre-established biofilm and had minimal impact on the competing strain, which remained dominant. Thus my results indicate that AlpP enhances colonisation by P. tunicata of a pre-established biofilm.

In contrast, R. gallaeciensis does not seem to be constrained by nutrient limitations and formed a biofilm which eventually covered most of the surface of the algae. An ability to utilise a wide range of carbon sources (Ruiz-Ponte et al., 1998) and metabolise DMSP, may enable this organism to colonise regions of the plant that are inhospitable to other bacteria. Even when colonising a pre-established biofilm, R. gallaeciensis might be able to access nutrients inaccessible to other epiphytic bacteria and this ability might contribute to its success and epiphytic fitness (Beattie and Lindow, 1999).

3.5 CONCLUSIONS

P. tunicata and R. gallaeciensis were found to exhibit different colonisation and competition behaviours during the colonisation of U. australis. Whilst R. gallaeciensis is capable of colonising U. australis under a range of conditions, colonisation by P. tunicata is enhanced by high cell densities, presence of cellobiose in the pre-culture, inoculation in the dark and interactions with a natural seawater community to attach and persist on the surface of the algae. The epiphytic fitness of R. gallaeciensis may be attributed to several factors, including its versatility in utilising a number of carbon sources (particularly those not available to competing strains) and the production of antibacterial compounds and signalling molecules. 89 Colonisation of algal surfaces appears to be inducd by cues produced at those surfaces. In the case of R. gallaeciensis, sulphur compounds such as DMSP appear to be the key for initial selection, and it is likely that cellulose may function as a cue for the attachment of P. tunicata.

Competition studies in which a pre-established biofilm is challenged with P. tunicata results in the coexistence of competitors. This may be partly due to the protective nature of microcolonies which may resist invasion. A diffusion gradient exists within microcolonies so that metabolically active cells at the outer edge of the microcolony may perish whilst cells in the deeper regions are likely to be protected from the antibacterial protein. Limited nutrients on the surface of U australis leads to strains inhabiting distinct niches on the plant and this may also result in a coexistence of competing strains. Microcolonies of competing strains are spatially separated which may limit microbial interactions and provide refuges for competitors. R. gallaeciensis does not seem to be constrained by low nutrient conditions and is able to invade and disperse competing strains suggesting that its antibacterial protein is able to diffuse through microcolonies.

The environmental factors that influence nutrient accumulation on the surface of the seaweed, and in turn microbial colonisation and competitive interactions are complex. Environmental factors may have a profound impact on the composition of the epiphytic community and consequently the antifouling activity of host surfaces. It suggests a possible mechanism by which host modulation of the nutritional status of the algal surface could modulate the antifouling activity of epiphytic bacteria.

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Chapter Four

Effects of bacterial cell density on antifouling by epiphytic marine bacteria

4.1 INTRODUCTION

Surfaces never remain pristine in the marine environment, but become quickly colonised by a film of marine bacteria. Such biofilms serve as a focus for the attachment and growth of other organisms such as invertebrates, sessile plants and animals that leads to the establishment of a mature biofouling community (Davis et al., 1989). It has long been known that bacterial biofilms produce chemical cues that may modify the behaviour of settling invert~brate larvae and algal cells (Wieczorek and Todd, 1998) and significantly influence the final composition of the biofouling community. These cues can be either positive (Maki and Mitchell, 1985; Szewzyk et al., 1991; Lau and Qian, 1997) or negative (Maki et al., 1988; Holmstrom et al., 1992), depending on the species of fouling organism and bacteria concerned. Previous studies suggested that algal and larval settlement is a function of bacterial species composition (Keough and Raimondi, 1996; Lau and Qian, 1997; Huang and Hadfield, 2003) and biofilms have been shown to inhibit and induce settlement of macrofoulers in a species-specific manner (Egan et al., 2001). Thus biofilms play an important role in the development of fouling communities on marine surfaces.

As well as being a common fouling organism, the intertidal green alga Ulva australis (previously known as Ulva lactuca) is itself susceptible to fouling, because it is sessile

This chapter is a modified version of: Rao, D., Webb, J.S., Holmstrom, C. Steinberg, P. D. and Kjelleberg, S. (Submitted) Effects of bacterial cell density on antifouling by epiphytic marine bacteria. Appl. Environ. Microbiol.

96 and restricted to the photic zone where conditions for fouling organisms are optimal ( de Nys et al., 1995). Many seaweeds have evolved efficient strategies to combat epibiosis and employ a number of physical and chemical defence systems to prevent fouling such as the shedding of outer layers of cells (Keats et al., 1997) or production of inhibitory compounds (Dworjanyn et al., 1999). However, antifouling defences are energetically costly (Wahl, 1989) and it has been proposed that Ulva australis relies on microbial defence (Holmstrom et al., 1992; Egan et al., 2001). Such interactions have been described for the marine crustaceans Palaemon macrodactylus and Homarus americanus where symbiotic bacteria were shown to defend embryos from fungal infection (Gil-Tumess et al., 1989; Gil-Tumess and Fenical, 1992).

Inhibitory bacteria have been isolated from U. australis and one of the best characterised of these is Pseudoalteromonas tunicata (Egan et al., 2000; 2001; 2002a, 2002b ). This bacterium produces a diverse range of biologically active compounds that specifically target marine fouling organisms (Holmstrom et al., 1998), including a polar, heat-stable anti-larval molecule of less than 500 Da (Holmstrom et al., 1992); a heat-sensitive anti-algal peptide between 3-10 kDa (Egan et al., 2001 ); an uncharacterised anti-diatom compound (Egan, S and Kjelleberg, S. unpublished data); a yellow antifungal compound (Franks, 2005) and a large autolytic and antibacterial protein (James et al., 1996). The antibacterial protein is a 190 kDa multi-subunit protein, designated AlpP, which is effective against both Gram-negative and Gram positive bacteria from a range of environments (Mai-Prochnow et al., 2004). Roseobacter gallaeciensis is also frequently isolated from the surface of U. australis and has known antibacterial activity (Ruiz-Ponte et al., 1998; Brinkhoff et al., 2004; Rao et al., 2005). Roseobacter spp. are cosmopolitan and have been isolated from green seaweeds (Shiba, 1992), marine snow particles (Gram et al., 2002) and dinoflagellates {Lafay et al., 1995; Alavi et al., 2001; Miller and Belas, 2004).

Molecular investigations based on Real-Time Quantitative PCR have shown that the genus Pseudoalteromonas is present throughout the marine environment (Skovhus et al., 2004), whilst P. tunicata mostly inhabits living surfaces that are relatively free from fouling such as green algae (Ulva lactuca and Vivaria fusca) and tunicates (Ciona intestinalis) (Skovhus, 2004). The same study found that the in situ density of the genus Pseudoalteromonas accounted for only 0.57% of the total eubacterial abundance and P. 97 tunicata constituted only a minor part of the total Pseudoalteromonas community. Studies based on a newly developed method combining catalysed reporter deposition with of fluorescence in situ hybridisation (CARD-FISH) suggested that R. gallaeciensis may be present in higher numbers as the genus Roseobacter comprised 12% of the epiphytic bacterial community on U. australis (Tujula, N., Crochetti, G., Holmstrom, C., Dahllof, I., and Kjelleberg, S. unpublished data).

Although the antifouling effects of P. tunicata are well established (Holmstrom et al., 1996; Egan et al., 2002), antifouling effects have not been investigated at ecologically relevant bacterial cell densities. The observation that P. tunicata is present at such low densities, raises the question as to whether it has any significant impact on antifouling activity at these densities. Moreover, the antifouling effects of biofilms on marine eukaryotic surfaces remain largely unstudied and little is known about the efficacy of P. tunicata and R. gallaeciensis against fouling on the living surface of U. australis.

In the work described here, it is shown that monospecies biofilms of P. tunicata and R. gallaeciensis on Petri dishes can inhibit the settlement of algal spores and invertebrate larvae as well as attachment of fungi and bacteria at ecologically relevant numbers of bacteria. Inhibition by P. tunicata biofilms is attributed to the production of antifouling compounds, as mutants defective in the production of the extracellular inhibitors do not display inhibitory activity. The data support the hypothesis that P. tunicata and R. gallaeciensis play a role in the antifouling defence of U. australis in vivo.

4.2 MATERIALS AND METHODS

4.2.1 Preparation of marine strains

Bacterial strains were isolated from the surface of U. australis as described previously (Chapter 2). The strains selected for this study were: P. tunicata, Pseudoalteromonas gracilis, Alteromonas sp., Cellulophaga fucicola, and Roseobacter gallaeciensis. As controls, P. tunicata mutants defective in the production of the antibacterial protein (AlpP) (Mai-Prochnow et al., 2004), all inhibitory compounds (WmpR) (Egan et al., 2002), and the antifungal compound (FM3) (Franks, 2005) were used. Cultures were stored at -80°C in 50% (vol/vol) glycerol in VNSS media (Marden et al., 1985) and

98 maintained on VNSS plates. For visualisation of bacterial cells under epifluorescence microscopy, P. tunicata and R. gallaeciensis were labelled with a green fluorescent protein (GFP) colour tag as described previously (Rao et al., 2005).

4.2.2 Establishment of biofilms

Bacteria were cultured for 24 h at 25°C in VNSS broth for preparation of inoculae. Cells were harvested by spinning down the culture and resuspending the pellet in seawater. Cell concentration was estimated by counting under epifluorescence microscopy using a haemocytometer and adjusted by dilution with seawater to the desired end concentration. All assays were conducted on both plastic surfaces and on living surfaces of U. australis discs. Biofilms on plastic surfaces were established by inoculating from overnight pre-cultures into 36 mm Petri dishes containing 3 ml of 10% VNSS medium (diluted in seawater) and incubated at 23°C for 24 h. Different densities of cells were inoculated so that the final density of attached cells on the surface ranged from 102 -108cells cm-2• After 24 h incubation, growth media was discarded and biofilms washed three times with sterile filtered seawater and incubated in fresh sterile seawater before conducting the bioassays. Numbers of attached cells were determined by counting the number of GFP-labelled cells under epifluorescence microscopy using an eyepiece grid. Twenty-four hour old biofilms were used for antifouling assays as older biofilms undergo a certain amount of detachment and sloughing, which affected cell densities.

To establish biofilms on the surface of U. australis, axenic discs of the plant were prepared as described by Scheffel (2003). Treated axenic tissue had negligible numbers

( < 50 cells cm-2) of attached solitary cells on the plant surface. Antibiotic treatment resulted in most of the epiphytic bacteria being removed while causing minimal damage to the U. australis tissue (as determined by the Evans blue test for plant viability) (Chapter 3). Axenic discs were immersed in a suspension of either P tunicata or R. gallaeciensis, placed within the wells of Falcon 24 well plates. After an incubation period of 3 h, discs were rinsed twice in filtered seawater and transferred to fresh Falcon 24 well plates containing 2 ml of filtered seawater. The plates were incubated for 48 h on a shaker at 60 rpm at 25°C (16 h photoperiod at 20 µEm- 2s- 1). Because it took longer to establish biofilms on the surface of the algae, 48 h biofilms were utilised

99 in these assays. For the preparation of non-axenic discs containing a natural epiphytic community, discs were obtained as previously described (Scheffel, 2003), but were not treated to make them axenic and were incubated for 24 h in sterile seawater before conducting assays. For all bioassays, at least three experiments were carried out with five replicate Petri-dishes or algal thallus discs in each treatment.

4.2.3 Anti-algal bioassays

The effect of different cell densities of bacteria on attachment and survival of algal spores was assessed on discs and plastic surfaces by exposing spores directly to monoculture biofilms of P. tunicata and R. gallaeciensis at densities ranging from 102 -

108 cells cm-2. Sterile seawater and P. tunicata WmpR mutant biofilms established at 106cells cm-2 served as controls for P. tunicata biofilms. Sterile seawater was the control for R. gallaeciensis. Ulva australis and Polysiphonia spore bioassays were set up as described by (Egan et al., 2001). U. australis spore settlement was assessed after 5 days using an inverted light microscope (Zeiss). Counts of germinated spores were conducted in 10 fields of view under a 40 x magnification and settlement was compared to controls. For assays on Ulva australis, spores were stained with crystal violet to distinguish green spores form the green background of plant tissue. Polysiphonia spore settlement was assessed after 24 h, the number of settled (ie. attached) and unsettled spores were counted under a dissecting microscope and the percentage settlement determined.

In order to determine if Acyl Homoserine Lactones (AHLs) were involved in enhancing settlement of U. australis spores, an assay was conducted with different concentrations of synthetic AHLs. N-oxooctanoyl-L-homoserine lactone (OOHLs) were suspended in a 1% agarose/distilled water support matrix as described in (Tait et al., 2005) at concentrations of 5, 10, 20 and 50 µM, respectively. The control was 50 µm of methanol suspended in 1% agarose. Spore attachment was assessed after 1 h, using an inverted light microscope (Zeiss). Spores were stained with crystal violet and counts were conducted in 10 fields of view under a 40 x magnification and settlement was compared to controls.

100 4.2.4 Anti-larval assays

A range of densities of P. tunicata and R. gallaeciencis were tested on algal discs and plastic surfaces using standard settlement assays of larvae of the bryozoan Bugula neritina (Bryan et al., 1998). Adult broodstocks of Bugula neretina were collected from pilings at Rose Bay, Sydney (33°87'52" S, 151°25'56"E) and larvae were obtained as described in (Bryan et al., 1998). Only newly released larvae were included in the bioassay (ie within 15 minutes of release). Bio films were established as described above, and Petri dishes were washed carefully three times with 2 ml of sterile seawater and then 3 ml of filtered seawater, containing around 20 larvae, were added to each Petri dish and incubated at 25°C for 2 days. Larvae were counted under a dissecting microscope and the percentage settlement determined. Control Petri dishes contained either sterile seawater alone (for both P. tunicata and R. gallaeciensis) or P. tunicata WmpR mutant biofilm with cell densities of 106cells mr1 (control for P. tunicata).

4.2.5 Antifungal assays

Both yeast and filamentous fungi were used to assess the antifungal activity of P. tunicata and R. gallaeciensis on plastic surfaces and algal discs. The yeast test strains (Yl, Y2 and Y3) and the filamentous fungus (Y4) were unidentified marine strains isolated from U. australis by Ashley Franks (Franks, 2005). Fungal strains were maintained on VNSS plates and inoculated into VNSS broth. Biofilms were established as described and Petri dishes were washed three times with 2 ml of sterile seawater and then 2 ml of medium (10% VNSS: 90% seawater), containing 105cells mr1 of fungus was added and incubated for 48 h. The fungi were stained with Syto 59, the numbers of attached cells counted under epifluorescence microscopy and the percentage of surface cover compared to the original inoculum determined. Biofilms of P. tunicata FM3 and WmpR established at 106 cells cm-2 served as controls. Counts were done of 10 fields of view under 40 x magnifications.

4.2.6 Antibacterial assays

Marine strains isolated from U. australis were used as target strains to test for antibacterial activity of P. tunicata and R. gal/aeciensis on plastic and U. australis surfaces. The bacterial challenge strains comprised marine strains isolated from U.

101 australis and consisted of Cellulophaga fucicola, Alteromonas sp., Roseobacter gallaeciensis, Pseudoalteromonas tunicata and Pseudomonas gracilis. Biofilms of P. tunicata, and R. gallaeciensis were established and challenged with test bacteria added to the biofilms at 106 cells mr1 and incubated for 48 h. P. tunicata Alp and WmpR mutant biofilms and culture media were used as controls for P. tunicata whilst culture media was the control for R. gallaeciensis. Test strains (RFP-labelled) could be readily distinguished from biofilm bacteria under epifluorescence microscopy. Counts were done of 10 fields of view under x 10 magnification.

4.2. 7 Statistical analysis

A one-way analysis of variance (ANOVA) and Tukey's pairwise comparisons were performed to determine significant differences between bacterial density counts of biofilms of sequential densities for some of the experiments. ANOVA assumptions of normality and heterogeneity of variance of the data were met. Tukey's pairwise comparison was employed to determine significant differences between individual treatments. Tests were conducted using the statistical program Systat 10 (SPSS).

4.3 RESULTS AND DISCUSSION

The ubiquity of fouling orgamsms m the marme environment and the negative consequences of fouling exert strong evolutionary pressures for marine organisms to develop defences to protect their surfaces from unwanted colonisation (Steinberg et al., 1997). One type of defence is thought to be the exploitation of antifouling compounds produced by epiphytic bacteria such as P. tunicata on Ulva australis. However, there is little evidence for the efficacy of these compounds produced by bacteria at in situ cell densities against naturally occurring micro- and macrofoulers. In this study the cell densities of P. tunicata and R. gallaeciensis that are required to deter settlement and attachment of biofouling organisms were examined. It was found that surprisingly low densities of cells within biofilms of these bacteria are capable of anti fouling effects.

102 4.3.1 Effect of bacterial biofilms on an inanimate surface

4.3.1.1 Settlement of algal spores

On plastic surfaces P. tunicata was inhibitory to Polysiphonia spore settlement at remarkably low cell densities. Spore settlement was inhibited at cell numbers as low as 102 cells cm-2 and at cell densities of 103 cells cm·2, inhibition was more than 90% (Fig. 4.1, ANOVA: Fs,36 =862.70, p < 0.001). At higher densities (106-108cells cm-2), P. tunicata biofilms lysed algal spores suggesting that the anti-algal compound is very potent against certain algal spores. It was found that U. australis spores were less sensitive to inhibition by P. tunicata and relatively higher densities of cells were required to inhibit spore settlement. However, although cell densities of 105 cells cm-2 were required for inhibiting Ulva spore settlement to more than 90%, cell densities as low as 103 cells cm-2 were still capable of some inhibition (Fig. 4.3, ANOVA: F8,36 = 49.27, p < 0.001).

In contrast, R. gallaeciensis biofilms had no effect on Polysiphonia spores at any of the densities tested (Fig.4.2., ANOVA: F1,32 = 6.61, p < 0.001). An inhibitory compound isolated from R. gallaeciensis (tropodithietic acid) has been reported to be toxic towards microalgae (Brinkhoff et al.. 2004) but no evidence of R. gallaeciensis causing inhibition of macroalgal spore settlement was found.

R. gallaeciensis was not inhibitory against Ulva spore settlement, but in fact stimulated spore settlement, an effect that was more pronounced at high cell densities (Fig. 4.4,

ANOVA: F7,32 = 4.00, p < 0.001). Bacterial quorum sensing molecules have been demonstrated to be involved in Ulva zoospore settlement (Joint et al., 2000) and R. gallaeciensis was found to produce AHL signalling molecules N-oxooctanoyl-L homoserine lactone (OOHLs) (Case R., Low, A., Kjelleberg, S. unpublished data). In order to examine in more detail whether OOHLs play a role in Ulva spore settlement, a settlement assay was conducted with different concentrations of OOHL. The settlement of Ulva spores generally increased with increasing concentrations of AHLs (Fig. 4.5). A range of concentrations of synthetic OOHLs were used in this study (5-50 µM), and although these concentrations were higher than those utilised by Joint and co-workers (5 µM) (Joint et al., 2002), spores appeared to be able to discriminate between different

103 concentrations. However, concentrations of OOHL produced by R. gallaeciensis in biofilms remains to be determined. The reasons for enhanced spore settlement and germination in the presence of signalling molecules are unclear but it has been suggested that such responses may facilitate a close relationship between germinating spores and certain essential bacteria (Joint et al., 2002).

4.3.1.2 Invertebrate larval settlement

Biofilms of P. tunicata had an inhibitory effect on Bugula neretina larval settlement. Inhibition of bryozoan larval settlement by P. tunicata biofilms took place at relatively high densities (105-106 cells per cm-2) (Fig. 4.6, ANOVA: F8,36 = 166.09, p < 0.001). P. tunicata has previously been shown to exhibit anti-larval effects at high densities on polystyrene plates (Holmstrom et al., 1992). Studies have demonstrated that several members of this genus can inhibit larval settlement (Holmstrom et al., 1996; Dobretsov and Qian, 2002; Lee and Qian, 2003) whilst other Pseudoalteromonas spp have been identified as being inductive for larval settlement (Leitz, 1997; Negri et al., 2001; Huang and Hadfield, 2003).

R. gallaeciensis also inhibited byrozoan larval settlement and inhibition took place at densities of 103 - 104 cells cm-2 (Fig. 4.7, ANOVA: F7,32 = 126.10, p < 0.001). There have been no previous reports of inhibitory effects of Roseobacter species on larval settlement. In contrast, Harder et al., (Harder et al., 2002) found that Roseobacter sp. at cell densities in the range of 105 -106 cells cm-2 resulted in induction of larval settlement. However, in a study exploring the use of R. gallaeciensis as a probiotic, it was seen that cell extracts from cells at 106 cells cm-2 enhanced scallop larval survival but that extracts from cell densities at 107 cells cm-2 resulted in mortality of Pecten maximus larval cultures suggesting possible toxicity effects on larvae (Ruiz-Ponte et al., 1999).

The findings showed that inhibition increased with increasing bacterial cell densities of P. tunicata and that the minimum number of cells required for inhibition was 1 x 104

cm-2, which is comparable to the results of (Dahms et al., 2004). The latter researchers showed that densities of 2 x 105 cells cm-2 of Pseudoalteromonas sp. were necessary to inhibit settlement of Bugula neretina. Although the possibility of repellents was not discounted, Dahms and co-workers ascribed the higher settlement of larvae at lower

104 densities of bacterial cells to bacteria free space and lower wettability on biofilm coated dishes. Similarly, (Olivier et al., 2000), showed that cyprid settlement in Ba/anus amphitrite was negatively correlated with total density in biofilms and attributed this to free-space availability. The results described here suggest, however, that antifouling compounds produced by P. tunicata result in inhibition of settlement, as the biofilm established by the WmpR mutant was not inhibitory to larval settlement.

Many studies have emphasised the importance of age of biofilms in influencing the settlement of larvae. The rate of settlement seems to be correlated to density of cells within the biofilm. Although changes in cell densities were not monitored as biofilms grew older, generally the older the biofilms, the higher the level of induction (Hamer et al., 2001; Huang and Hadfield, 2003; Lau et al., 2005) or inhibition (Maki et al., 1990; Dahms et al., 2004). Thus larvae of some invertebrates are able to discriminate among biofilms of different ages with settlement levels correlating positively or negatively with bacterial cell densities in the biofilm.

A recent study demonstrated that highly inducing strains for the settlement of the sea urchin Heliocidaris erythrogramma, were dominated by the genera Pseudoalteromonas, Shewanella, and Vibrio. These genera were highly effective at induction, despite being present at low densities ( < 1x 105 cells cm-2) and representing only a small percentage of the whole bacterial community (Huggett, 2005). Field recruitment was correlated with laboratory settlement assays suggesting that cues were present in quantities that larvae are able to detect and respond to.

105 90 d d

80 1

70 1 .... C C QI 60 E QI ;:;.... QI 50 Ill QI C, ,l9 40 • C ..uQI QI II. 30 1 20 1 b I 10 ab ab a a a _o_~ O +-- 0 ~ 0 n 1E+08 1E+07 1E+06 1E+05 1E+04 1E+03 1E+02 Seawater WmpR control Density (cells cm-2 )

Fig 4.1 Percentage settlement (mean ± SE, n = 4) of spores of Polysiphonia sp. after 24 h incubation with P. tunicata biofilms established at different densities on plastic Petri dishes. Sterile seawater served as negative controls and a P. tunicata WmpR mutant biofilm established at 106 cells cm -2 was the positive control. Densities sharing the same letter do not differ at p = 0.05 (Tukey's multiple comparison).

106 90 a

ab b b 80 b b b 70 ... ~ 60 , E (II 50 j t(II Ill (II ...g' 40 j C (II u... QI 30 , Q.

20 1

0 lE+0B 1E+07 1E+06 1E+05 1E+04 1E+03 1E+02 Seawater control

Density (cells cm-2 )

Fig. 4.2 Percentage settlement (mean ± SE, n = 4) of spores of Polysiphonia sp. after 24 h incubation with R. gallaeciensis biofilms established at different densities on plastic Petri dishes. Sterile seawater served as negative controls. Densities sharing the same letter do not differ at p = 0.05 (Tukey's multiple comparison).

107 70

C C C 60 i C

.... 50 ° b C QI E QI b ~ 40 i u, QI

30 EC ~ QI ..u QI Q. 20 j

a a a 0 1E+08 1E+07 1E+06 lE+0S 1E+04 1E+03 1E+02 Seawater WmpR

Density (cells cm-2 )

Fig. 4.3 Percentage settlement (mean± SE, n = 5) of spores of Ulva australis after 5 days of incubation with P. tunicata biofilms established at different densities on plastic Petri dishes. Sterile seawater served as negative controls and a P. tunicata WmpR mutant biofilm established at I 06 cells cm - 2 was the positive control. Densities sharing the same letter do not differ at p = 0.05 (Tukey's multiple comparison).

108 90 a

80 ab ab

70 ab ab b ... ab ~ 60 E QI

EQI so II) QI Cl i 40 1 ...u ~ 30

10 ,

o - lE+0B 1E+07 1E+06 lE+0S 1E+04 1E+03 1E+02 Seawater

Density (cells cm-2 )

Fig. 4.4 Percentage settlement (mean ± SE, n = 5) of spores of Ulva australis after 5 days of incubation with R. gallaeciensis biofilms established at different densities on plastic Petri di shes. Sterile seawater served as negative controls. Densities sharing the same letter do not differ at p = 0.05 (Tukey's multiple comparison).

109 90

80 1

70 I .... ~ 60 E Ill E 5o Ill Ill Ill Cl ~ 40 C Illu... ~ 30

10 j

o L 5 10 20 50 Control Concentration of OOH Ls (µM)

Fig. 4.5 Percentage settlement (mean ± SE, n = 5) of spores of Ulva australis after I h of incubation in the presence of a synthetic AHL (OOHL) suspended at different concentrations in agarose. Methanol (50~tM) suspended in agarose served as a control.

11 0 120

C C C C 100 •

b .., C QI 80 ° E QI ;;.., QI Cl! QI 60 a Cl ..,Ill C QI ..u QI ll. 40 '

a al _ a ~-a []~ 1E+08 1E+07 1E+06 lE+0S 1E+04 1E+03 1E+02 Seawater WmpR

Density (cells cm-2 )

Fig. 4.6 Percentage settlement (mean± SE, n = 5) of Bugula neretina larvae after 48 h incubation with P. tunicata biofilms established at different densities on plastic Petri dishes. Sterile seawater served as ne~ative controls and a P. tunicata WmpR mutant biofilm established at 106 cells cm - was the positive control. Densities sharing the same letter do not differ at p = 0.05 (Tukey' s multiple comparison).

l l l 120

b C 100 b

... C 80 QI E QI ;:;... .,,QI QI 60 1 a C, ...IV C ..uQI QI 40 J 0..

20 , a

a a a 0 t- D lE+0B 1E+07 1E+06 lE+0S 1E+04 1E+03 1E+02 Seawater

Density (cells cm-2 )

Fig. 4.7 Percentage settlement (mean ± SE, n = 5) of Bugula neretina larvae after 48 h incubation with R gallaeciensis biofilms establi shed at different densities on plastic Petri dishes. Sterile seawater served as controls. Densities sharing the same letter do not differ at p = 0.05 (Tukey's multiple comparison).

100 l Ill Yeast strain 1 90 a Yeast st rain 2

80 D Yeast st rain 3 ~ a Yeast strain 4 .,C 70 E E., 60 .,"' so j 01 ~ 40 J .,C ~ ., 30 1 Q. 20 j

10 a l E+0B 1E+07 1E+06 l E+0S 1E+04 1E +03 1E+02 WmpR Seawa ter

Density (cells cm·')

Fig. 4.8 The attachment(% surface cover) (mean ± SE, n = 5) of marine fungi after 48 h incubation with P. tunicata biofilms established at different densities on plastic Petri dishes. Sterile culture medium (I 0% VNSS) served as a negative control and P. tunicata FM3 and WmpR mutant biofilms establi shed at 106 cells cm -2 were the positive controls.

112 100

90 1

80 .... C 70 Cl) E Cl) 60 , ;; .... D Cytophaga fu cico/a Cl) u, D Pseudoalteromonas gracilis Cl) 50 Cl D Alleromonas Sf' ,a .... D Pseudoalleromonas tunicata C 40 Cl) ..V Cl) II. 30 ,

20 ,

10 ' 0 1------~---- ~--~~ M1ITill 1E+08 1E+07 1E+06 1E+05 1E+04 1E+03 1E+02 Seawater

Density (cells cm-2 )

Fig. 4.9 The attachment (% surface cover) (mean ± SE, n = 5) of marine bacteria after 48 h incubation with R.gallaeciensis biofilms established at different densities on plastic Petri dishes. Sterile culture medium (lO¾YNSS) served as a control.

113 90

70 < ... ~ 60 E (II so E(II Ill (II C, f 40 C (II ..u : 30

1E+08 1E+07 1E+06 l E+OS 1E+04 1E+03 1E+02 Seawater

Density (cells cm"2 )

Fig. 4.10 The settlement (%) (mean ± SE, n = 5) of spores of Ulva australis after 5 days of incubation with R. gallaeciensis biofilms established at different densities on U. australis discs. Axenic algal discs cultured in sterile seawater served as the control.

114 100

D Cytophaga fucicola 80 D Pseudoalteromonas gracilis

.., 70 OA/teromonas sp . C Ill D Pseudoalteromonas tunicata ~ 60 E Ill

Ill"' 50 ' Cl ...Ill ~ 40 ...u Ill c. 30

0 1------lE+OB 1E+07 1E+06 lE+OS 1E+04 1E+03 1E+02 Seawater

Density (cells cm-2 )

Fig. 4.11 The attachment (% surface cover) (mean ± SE, n = 5) of marine bacteria after 48 h incubation with R. gallaeciensis biofilms established at different densities on U. australis di scs. Axenic algal di scs incubated in sterile seawater served as a negative control.

115 100 0 Yeast strain 1 0 Yeast strain 2 0Yeast strain 3 .I. 80 1 OYeast strain 4 l .... 70 1 C QI .I ' E QI 60 i E QI , Ill QI so I en I ....n, ~ 40 , ...u QI ll. 30 j

, I I ~

1E+08 1E+07 1E+06 lE+0S 1E+04 1E+03 1E+02 W3Gfp Seawater

Density (cells cm-2 )

Fig. 4.12 The attachment(% surface cover) (mean ± SE, n = 5) of marine fungi after 48 h incubation with P. tunicata biofilms established at different densities on U. australis discs.Axenic algal discs incubated in sterile seawater served as a negative control and P. tunicata FM3 and WmpR mutant biofilms established at 106 cells cm -2 were the positive controls.

116 4.3.1.3 Fungal colonisation

A range of densities of P. tunicata was able to inhibit fungal growth of the three yeast

strains (Yl,Y2 and Y3). P. tunicata was inhibitory at densities as low as 102 - 103 cells per cm-2 (Fig. 4.8) suggesting that the anti-fungal compound is effective at very low concentrations. Y4, which is a filamentous fungus, was not inhibited by P. tunicata biofilms and mycelia extended over the mirocolonies without any hindrance. In contrast, P. tunicata antifungal mutant (FM3) did not affect the attachment of any of the fungal strains on the control biofilm. The results are very similar to the study conducted by Franks (2005) who observed that after 24 hours of growth, P. tunicata fully inhibited the attachment of Yl to a glass flow cell surface, whereas the antifungal mutant, FM3, had no inhibitory effect. Furthermore, P. tunicata was able to invade and disrupt an already established biofilm of Yl (Franks, 2005). Purification of the broad spectrum antifungal component has shown it to be composed of two novel tambjamine based compounds (Franks, 2005). In contrast R. gallaeciensis had no antifungal activity at any of the densities tested and no reports of inhibition of fungi by Roseobacter spp. were found in the literature.

4.3.1.4 Effects of bacterial biofilms on attachment of bacteria

P. tunicata biofilms did not impact on the settlement of other marine bacteria and there were no differences in attachment, regardless of the density of P. tunicata biofilms. The settlement assay for bacteria indicated that biofilms of P. tunicata were unable to prevent other marine strains from settling and attaching onto the surface (Data not shown). This was unexpected as in competition experiments conducted in a flow through system, P. tunicata was able to outcompete and remove these same strains. In glass flow cells, competing strains formed separate microcolonies with very little interaction of cells from different species (Rao et al., 2005). However, in the bacterial settlement assay, in a batch system, the cells formed mixed microcolonies.

Biofilms of R. gallaeciensis were very effective in preventing the settlement of other bacterial strains, including P. tunicata. Densities as low as 103 cells cm-2 were able to prevent growth of marine bacteria (Fig. 4.9). This result is similar to that of the 117 competition studies conducted in glass flow cells (Rao et al., 2005). Roseobacter gallaeciensis strains have been found to display strong in vitro antagonism (Ruiz-Ponte et al., 1998; Brinkhoff et al., 2004; Hjelm et al., 2004) and this has been attributed to the presence of peptides (Ruiz-Ponte et al., 1998) and tropodithietic acid (Brinkhoff et al., 2004), suggesting that at least two inhibitory compounds may be produced.

4.3.2 Effects of biofilms on the surface of CJ. australis

Major differences in the ability of biofilms of bacteria to deter fouling were observed when assays were carried out on the surface of U australis algal thallus rather than on plastic surfaces. Polysiphonia and Ulva spores were not inhibited by P. tunicata biofilms on the algal surface. However, Ulva spore settlement on R. gallaeciensis biofilms on U australis surface, mimicked the trends observed with polystyrene plates - Ulva spores were stimulated by R. gallaeciensis biofilms and the degree of stimulation roughly correlated with the densities of cells established on the algal surface (Fig. 4.10). It is unclear as to why biofilms on the surface of the algae would enhance Ulva spore settlement. Cues produced by the bacteria may be enhanced on the surface of the algae.

It was further observed that there were no differences between larval settlement on biofilm covered algae as compared to axenic algal tissue. Biofilms of both P. tunicata and R. gallaeciensis were unable to deter B. neretina larval settlement. There was also no inhibition of larval settlement by the natural community established on U australis. Based on the assumption that there are inhibitory bacteria on Ulva, the effectiveness of a natural biofilm in inhibiting larval settlement was explored. It was found that settlement of larvae on algal tissue was generally poor and there were no differences between natural community discs and axenic algae. Other studies have indicated that the post-settlement success or failure of Bugula neretina is largely dependent on the structural complexity of its substrata (Walters and Wethey, 1996). Most bryozoans require a substratum that provides firm support for attachment and many prefer surfaces that have a smooth or glossy finish (Ryland, 2001 ). It is possible that algal discs, suspended in seawater, as prepared in this study, did not provide surfaces that were sufficiently firm or smooth to allow for attachment by larvae.

118 When anti-fungal assays were conducted on the surface of U. australis, P. tunicata had an inhibitory effect on Yl, Y2 and Y3 reflecting assays conducted on polystyrene surfaces. In a study by Franks (Franks, 2005), which explored the effect of P. tunicata biofilms established on U. australis and their effects on a natural marine fungal community, it was found that the fungi on the surface of U. australis were no longer detectable by DGGE after 48 hours, indicating the removal or the inhibition of the natural mixed fungal community (Franks, 2005). It is not known what densities of cells were established on the surface in that study, but the biofilm was very effective at inhibiting fungi. The results presented here show that cell densities as low as 103 -104 cm-2 P. tunicata cells on the plant surface were inhibitory against fungi (Fig. 4.11 ).

In contrast to observations that P. tunicata was not able to inhibit attachment of marine bacteria in Petri dishes, it was found that biofilms of P. tunicata established on U. australis were able to prevent settlement and attachment of other marine isolates. However, this does not seem to be an inhibitory effect, because the control biofilm of P. tunicata AlpP mutant was also able to deter attachment of the marine strains. Results described in Chapter 3 suggested that pre-established biofilms could deplete resources on the surface of U. australis, and make it more difficult for invading bacteria to sequester themselves in the existing biofilm community. Such phenomena have been shown in higher plants, where pre-established biofilms deplete a large percentage of the carbon source, leading to pre-emptive exclusion of secondary colonisers (Wilson and Lindow, 1994 ). Similarly to the results in Petri dishes, biofilms of R. gallaeciensis at densities as low as 104-105 cells cm-2 on U. australis, were effective at preventing attachment of other marine bacteria (Fig. 4.12).

For negative settlement cues to be ecologically relevant they must be present in the habitat that fouling organisms recruit to, and they must be present in quantities that deter fouling organisms (Williamson et al., 2000). However, in some cases, settlement in the field does not correlate with laboratory settlement observations (Cameron and Schroeter, 1980; Thompson et al., 1998). Results presented here demonstrate that low densities of cells in biofilms established in Petri dishes are inhibitory to settlement by various fouling organisms. However, assays conducted on algal discs are difficult to interpret. To my knowledge, there are no reports of settlement assays being conducted on living algal tissue, and it is likely that several factors influence the outcome of the 119 assays. Colonisation studies (Chapter 3), indicate that environmental factors that influence nutrient accumulation on the surface, and in tum microbial colonisation, are complex. Thus the reasons for differences on the two surfaces are unclear but could to be related to the expression of inhibitors and plant mediated factors, both subject to substrate availability.

4.3.3 Density and antifouling activity.

Skovhus and co-workers showed that P. tunicata is present in very low numbers in the natural environment (Skovhus et al., 2004). These researchers estimated that the absolute abundance of the antifouling subgroup (P. tunicata and P. ulvae) was in the range of lx 103 cells cm-2 which implies that P. tunicata was present at less than 1 x 103 cells cm-2. The results presented here confirm that low cell densities P. tunicata are effective at inhibiting settlement of antifouling organisms. Holmstrom and co-workers demonstrated that eight out of ten tested Pseudoalteromonas sp. contained at least one of the four tested antifouling properties: growth inhibition of bacteria and fungi and settlement inhibition of algal spores and invertebrate larvae (Holmstrom et al., 2002). It is likely that the antifouling protection of U. australis consists of several surface associated Pseudoalteromonas sp. and other inhibitory bacteria such as R. gallaeciensis working as an antifouling consortium, rather than one individual species of P. tunicata. Roseobacter spp. are numerous in the marine environment and using CARD-FISH it has been shown that they comprise 12% of the bacterial epiphytic community on U. australis (Tujula, N., Crocetti, G., Dahllof, I., Holmstrom, C., Kjelleberg, S. unpublished data). It is proposed that both of the bacteria examined in the present study, participate in the antifouling defence, with R. gallaeciensis being more effective at inhibiting larvae and bacteria, whilst P. tunicata is more effective against algal spores and fungi. Mixed communities need both P. tunicata and R. gallaeciensis to inhibit a diverse fouling community. Hugget (2005) showed that a wide range of bacteria from various genera induce the same settlement response in sea urchin larvae, suggesting a redundancy in the function of bacteria on the surface of coralline algae (Huggett, 2005). Similarly, results reported here indicated that R. gallaeciensis and P. tunicata provide complementary benefits to the host, with R. gallaeciensis being more effective at inhibiting larvae and bacteria, whilst P. tunicata is more effective for algal spores and

120 fungi. Thus P. tunicata and R. gallaeciensis are quite possibly present in sufficient quantities on the plant to inhibit fouling organisms.

4.4 CONCLUSIONS

Bacteria that produce inhibitory compounds on the surface of marine algae are thought to contribute to the defence of the host plant against colonisation by fouling organisms. However this hypothesis had not been tested using living plants in situ and the number of bacterial cells necessary to defend against fouling on the plant surface was not known. The present study demonstrated that low cell densities of both P. tunicata and R. gallaeciensis were effective in preventing settlement and attachment of diverse fouling organisms in Petri dishes. It is likely that R. gallaeciensis and P. tunicata provide complementary benefits to the host, with R. gallaeciensis being more effective at inhibiting larvae and bacteria, whilst P. tunicata is more effective for algal spores and fungi. This strongly supports the hypothesis that P. tunicata and R. gallaeciensis can play a role in the defence against fouling on U. australis at ecologically relevant densities.

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126

Chapter Five

Effects of introducing inhibitory bacteria into mixed species marine biofilms

5.1 INTRODUCTION

In the marine environment all surfaces become rapidly fouled leading to the formation of a complex biofouling community. The process is initiated by the formation of an organic "conditioning" film that occurs on clean surfaces within minutes of being immersed in seawater. The organic film facilitates the attachment of bacteria which divide rapidly and form a biofilm. In nature, colonisation of habitats by mixtures of bacterial populations is the rule rather than the exception (Marsh and Bowden, 2000).

As microbial communities establish and mature, they are often subjected to challenge with planktonic cells. Whether or not an immigrant organism can survive and grow within the community will depend on its ability to displace, compete or co-operate effectively with the resident population (Marsh and Bowden, 2000). This ability will change as a function of time as the biofilm becomes physiologically and nutritionally more heterogenous (McBain et al., 2000). With the progression of time, the diversity of species within naturally occurring biofilm communities increases, until competitive interactions lead to the selection of a few successful partnerships, or consortium phenotypes (Jackson et al., 2001). Successful partnerships of cooperating organisms can lead to increased persistence under stressful conditions (Cowan et al., 2000). Occasionally, cooperative interactions can lead to synergistic biofilm formation by strains that are unable to form a biofilm alone. Benefits accruing from synergistic interactions include communal protection from inhibitory bacteria, production of essential growth factors, and modification of the local environment and the protection of sensitive species by the inactivation of inhibitors (Brook, 1989). Eventually a mature biofilm community achieves a pseudo-steady state, where sequestration of planktonic cells and the growth and division of attached cells are counterbalanced by losses through dispersal and cell death (Willcock et al., 1997).

127 Once established, biofilm communities serve as a focus for the attachment and growth of a range of fouling organisms such as diatoms, invertebrate larvae and algal spores. Attachment of macrofoulers adds further complexity to the community, which continues to develop through both the growth of populations within the biofilm and additional recruitment from the water column. Biofouling communities can also develop on living surfaces, with subsequent detrimental effects on the host organism. However, the extent of biofouling on marine organisms is markedly less than that on inanimate structures and it is evident that algae can regulate bacterial colonisation of their surfaces. Marine algae have evolved a range of defence mechanisms against fouling and it has been suggested that the green alga Ulva australis, (previously known as Ulva lactuca) depends on inhibitory bacteria to protect it against fouling (Holmstrom et al., 1992; Egan et al., 2002a). There is also increasing evidence of beneficial associations between surface bacteria and their hosts (Gil-Tumess and Fenical, 1992; Barbieri et al., 2001; Dobretsov and Qian, 2002; Dobretsov and Qian, 2004) suggesting that bacterial biofilms may contribute to the fouling defence of their hosts.

The best studied of the inhibitory bacteria on U australis are Pseudoalteromonas tunicata and Roseobacter gallaeciensis which have been shown to display strong antibacterial activity (Ruiz-Ponte et al., 1999; Brinkhoff et al., 2004; Hjelm et al., 2004; Mai-Prochnow et al., 2004; Rao et al., 2005). It has been demonstrated that P. tunicata and R. gallaeciencsis display antibacterial activity against monospecies biofilms under laboratory conditions (Chapter 2) but the effect of P. tunicata and R. gallaeciensis on either synergistic mixed species bacterial biofilms or complex seawater and epiphytic communities has not been explored. It was also found that monospecies biofilms of P. tunicata and R. gallaeciensis on Petri dishes can inhibit the settlement of algal spores and invertebrate larvae, as well as attachment of fungi and bacteria at ecologically relevant numbers of bacteria (Chapter 4). However, the integration of inhibitory marine bacteria into a synergistic mixed species biofilm or an epiphytic community established on algal surfaces remains largely unstudied.

128 The aims of the work described in this chapter were to determine if:

1) ecologically relevant densities of P. tunicata alter the diversity of a natural seawater community on inanimate surfaces. 2) P. tunicata or R. gallaciensis could colonise and impact on an epiphytic community on U. australis 3) P. tunicata or R. gallaeciensis can colonise synergistic mixed species biofilms on both inanimate and living surfaces.

5.2 MATERIALS AND METHODS

5.2.1 Establishment of a natural seawater community on glass slides

In order to asses whether ecologically relevant densities of P. tunicata are able to alter a natural seawater community on inanimate surfaces, a natural seawater community was allowed to develop on glass microscope slides immersed in seawater for two weeks, before being invaded with different densities of P. tunicata. These experiments were done in collaboration with Torben Skovhus of the University of Aarhus, Denmark.

Glass slides were arranged in open microscope slide storage boxes filled with seawater and incubated on a shaker at 40 rpm at 14°C and light intensities of 20 µE/m2/s on a 12 h photoperiod. Seawater for the experiment was collected from Aarhus Bay, Denmark. After a two week period, slides were transferred from the slide storage boxes to three seawater tanks containing 1200 ml fresh seawater. Each of the three tanks had 12 slides immersed in it. GFP-labelled P. tunicata cells were prepared as described previously (Chapter 3), and added at in situ densities (9.7 x 104 cells per ml (Skovhus et al., 2004) to Tank 2. Cell densities of P. tunicata added to Tank 3 were approximately 1,000 fold higher (7.8 x 107 cells per ml). Tank 1 was a control that did not contain P. tunicata. Slides in three tanks were further incubated for a week at the conditions described above. Of the twelve slides in each seawater tank, nine slides were used for DNA extractions (each DNA extraction was from three pooled slides) and three were used for staining and microscopy observations.

129 5.2.2 Biofilm sampling and DNA extraction of the natural seawater community

on glass surfaces

After one week, biofilm material was removed from both sides of the glass slides using sterile razor blades. The material was transferred directly into DNA extraction tubes. The total area of sampled biofilm for one DNA extraction was 120 cm2 (pooled from three slides), which provided enough DNA for PCR amplification. Slides used for DNA extractions and microscopy were removed from seawater tanks according to a table of random numbers. Colour and structure of the biofilms were observed by microscopy prior to DNA extraction. Nucleic acids from the biofilm samples were extracted using the FastDNA Spin Kit for Soil (Qbiogene, Carlsbad, CA), according to the manufacturer's instructions. The concentration of the extracted DNA was measured fluorometrically (TD-700 fluorometer; Turner Design, Sunnyvale, CA). DNA extracts (1-3 ng per PCR tube) were diluted 1: 10 in Sigma molecular biology-grade water (Sigma, Saint Louis, MO) prior to DNA amplification to avoid high levels of PCR inhibitors in the DNA extract.

5.2.3 Diversity of the natural seawater community on glass slides

The Pseudoalteromonas specific Polymerase Chain Reaction- Denaturant Gradient Gel Electrophoresis (PCR-DGGE) protocol was based on the primers Eub341F-GC and Psalt815R and the running conditions were as described by Skovhus et al., (2004). The eubacterial specific PCR-DGGE protocol was based on the primers Eub341F-GC and Univ907RC. The reaction mixture for the PCR amplification contained: 36.5 µl dH2O (Sigma), 5 µl buffer (l00mM Tris-HCI, 750 mM KCl, 15 mM MgC12, pH 8.8), 5 µl of l0X dNTP mix (final concentration of 125 µM for each dNTP; Invitrogen, Leek, The Netherlands), 1 µl of each primer (50 pmol/µl), 0.5 µl Taq DNA polymerase (5,000 U/ml; Amersham Biosciences, Uppsala, Sweden), and 1 µl of template DNA (diluted 1: 10). PCR amplification was carried out in a DNA thermocycler (PT-200 Peltier thermal cycler; MJ Research). The PCR consisted of an initial denaturation for 60 s at 93°C followed by 25 cycles with each cycle consisting of 30 s denaturing at 92°C, 60 s annealing at 57°C and 45 s (adding 1 s/cycle) extension at 72°C. The reaction was completed by a 5 min final extension step at 72°C. PCR products were separated by DGGE analysis according to their melting behaviour. The D-GENE DGGE system from Bio-Rad was used with an 8% acrylamide gel and a denaturating gradient ranging

130 from 30 to 70% ( 100% denaturant was 7 M urea and 40% vol/vol formamide ). The DGGE protocol was optimised to run at a constant voltage of 100 V at 60°C for 15 h for maximum band separation. For both DGGE approaches, the banding patterns were analysed with the Plot Profile tool in the Scion Image software package for presence and absence of DGGE-bands (Scion Beta version 4.0.2; http://www.scioncorp.com). In each of the two DGGE approaches, PCR amplified P. tunicata DNA was used in a DGGE-ladder to verify if P. tunicata was present in the biofilm samples at densities high enough to be detected on the DGGE-gels.

5.2.4 Microscopy of the natural seawater community on glass slides

One slide from each tank was used to investigate if GFP-labelled P. tunicata cells were present in the seawater biofilms. A further two slides from each tank were used to investigate biofilm morphology, structure and sloughing activity under epifluorescence microscopy. Biofilm biomass was removed from one side of the slide, stained with SybrGreen (Molecular Probes) (diluted 1:50 in sterile NSS) and visualised under a microscope (Axiovert 200M ApoTome; Carl Zeiss, Jena, Germany).

5.2.5 Epiphytic communities on U. australis

In order to assess if P. tunicata and R. gallaeciensis could colonise and impact on an epiphytic community on U australis, algal discs were pre-incubated in natural seawater for 6 days to allow a complex epiphytic community to develop on the surface. Although untreated algal tissue had a pre-existing biofilm community on its surface, excised discs were incubated in seawater for 6 days to allow for further growth and development of the epiphytic community such that biofilms with a thickness of 10-15 µm were established. Discs were excised from thallus tissue of U australis as described previously (Chapter 3) and incubated in Falcon 24 well plates on a shaker at

60 rpm at 25°C (16 h photoperiod at 20 µEm- 2s- 1).

Bacterial strains were isolated from the surface of U. australis as described previously (Chapter 2). P. tunicata, R. gallaeciensis and all mutants used for colonising the epiphytic community were GFP-labelled (Chapter 2). P. tunicata AlpP mutant (Mai Prochnow et al, 2004) and WmpR mutant (W3) strains, that did not produce any of the antifouling compounds (Egan et al., 2000), served as the controls. Bacteria were

131 cultured for 24 h at 25°C in VNSS broth for preparation of inoculae and bacteria were applied by immersing the discs in a suspension of either P. tunicata or R. gallaeciensis. Overnight cultures of bacteria were inoculated onto U. australis at densities of either 106cells mr' or l 08cells mr' in filtered seawater and incubated for 3 hours without shaking at room temperature. Inoculated discs were rinsed three times with filtered seawater and transferred to sterile seawater in Falcon 24 well plates. The discs were then rinsed twice in filtered seawater and transferred to fresh Falcon 24 well plates containing 2 ml of filtered seawater. The plates were incubated on a shaker at 60 rpm at

25°C ( 16 h photoperiod at 20 µEm- 2). The epiphytic community was visualised by staining with acridine orange. Attachment and colonisation of GFP-labelled P. tunicata and R. gallaeciensis within the community was visualised by observing unstained biofilm under a confocal microscope. The time points taken were: 1, 2, 4 and 8 days following inoculation. At each sampling time, five discs were randomly selected and a total of 12 random fields of view visualised for each disc.

5.2.6 Synergistic mixed species biofilms established on glass surfaces

In order to determine if members of the epiphytic community on U. austra/is were capable of synergistic biofilm formation, and whether such a synergistic mixed species biofilm would be more resistant to invasion by P. tunicata and R. gallaeciensis, strains isolated from the algae (Chapter 2) were mixed in different combinations of three or four strains and screened for synergistic interactions.

Experiments with synergistic mixed species biofilms were conducted in collaboration with Mette Burm0lle of Copenhagen University, Denmark. Of the different strains isolated from U. australis (Chapter 2), apart from P. tunicata and R. gallaeciensis, seventeen survived repeated subculturing on VNSS plates. The seventeen strains were mixed in combinations of three or four strains and screened for synergistic interactions. Mixtures of strains formed biofilms in 96- well microtitre plates and four particular strains were found to interact synergistically in biofilms. The four strains identified by the use of 16S rDNA sequencing, were found to be Microbacterium sp., Shewanella pacifica, Acinetobacter /owffii and Dokdonia donghaensis. A mixture of these strains was shown to form a better biofilm than each of the strains individually, and biofilm formation was found to increase by 167%. The synergistic mixed species biofilm was

132 more resistant to hydrogen peroxide and tetracycline compared to that of single species biofilms. Therefore, the mixed species biofilm was tested for resistance to colonisation by P. tunicata and R. gallaeciensis and that was compared to single species biofilms.

Biofilms were grown in continuous-culture flow cells (channel dimensions lx4 x 40 mm) at room temperature as previously described (Moller et al., 1998). Monospecies biofilms were established by inoculating channels with 1 ml overnight cultures of Microbacterium sp., Shewanella pacifica, Acinetobacter lowffii and Dokdonia donghaensis. To cultivate mixed biofilms, flow chambers were inoculated with 250 µl of stationary phase culture of each strain resulting in a total volume of 1 ml. Cultures were incubated without flow for 1 h at room temperature. Cultures were adjusted so that biofilms were established with a flow rate of 150 µl min· 1 (Biofilms were grown in 20% VNSS broth (Marden et al., 1985). Monoculture biofilms and mixed species biofilms were allowed to pre-establish for 2 h. Pre-formed biofilms (both monospecies and mixed species) were inoculated with - 10 6 cells of GFP-labelled P. tunicata and the flow stopped for 1 h. After the resumption of flow, the biofilm was monitored at regular intervals for a period of 3 days. Biofilms were stained with Syto 59 diluted to 3 µl mr1 in biofilm media. Biofilms were visualised with a confocal laser scanning microscope (CLSM) (Olympus) using fluorescein isothiocyanate and tetramethyl rhodamine isocyanate optical filters. Cells were examined for red and green fluorescence. Percentage surface coverage was calculated using image analysis (ImageJ) and percentage inhibition was calculated. Experiments were repeated in three separate rounds with three independent flow cells running in parallel.

5.2. 7 Synergistic mixed species biofilms on U. australis

To determine if the synergistic mixed species biofilm established on U. australis was able to resist colonisation by inhibitory bacteria, mixed species biofilms were established on axenic surfaces of the algae and inoculated with either P. tunicata or R. gallaeciensis.

Axenic discs were prepared from U. australis axenic as described previously (Chapter 3). Axenic algal discs were cultured in 2 ml of sterile seawater and agitated at room

133 temperature at 60 rpm. All experiments were done in triplicate, and five individual discs were randomly sampled at each sampling time as described previously.

Overnight cultures of strains Microbacterium sp., Shewanella pacifica, Acinetobacter lowfjii and Dokdonia donghaensis were inoculated onto algal discs at densities of 106cells mr' in filtered seawater and incubated for 3 h without shaking at room temperature. Mixed species biofilms were established by inoculating a mixture of the four strains. Inoculated discs were rinsed three times with filtered seawater and transferred to sterile seawater in Falcon 24 well plates and incubated on a shaker at 60 rpm at 25°C (16 h photoperiod) for 8 h. Mixed species biofilms on the surface of U. australis were inoculated with either GFP-labelled P. tunicata or R. gallaeciensis and incubated for 3 h without shaking at room temperature. Discs were rinsed three times with filtered seawater and incubated on a shaker as described earlier. The mixed species biofilm was visualised by staining with acridine orange. Attachment and colonisation of GFP-labelled P. tunicata and R. gallaeciensis within the mixed biofilm was visualised by observing unstained biofilm under a confocal microscope. The time points taken were 24, 48, 72 and 96 h after the inoculation of P. tunicata or R. gallaeciensis.

5.3 RESULTS

5.3.1 Diversity of the natural seawater community on glass surfaces

The first aim was to determine if ecologically relevant densities of P. tunicata could alter a natural seawater community on glass surfaces. In order to achieve this aim, natural seawater communities were allowed to establish on glass slides and then invaded with P. tunicata at in situ densities (Tank 2) and densities a thousand fold higher (Tank 3).

The diversity of both the eubacterial community and of Pseudoalteromonas species was studied using two different PCR-DGGE protocols. The number of DGGE-bands (which is an indication of species richness) was lowest in the control in Tank 1 (8-13 bands), slightly higher in Tank 2 ("low addition") (11-16 bands) and highest in Tank 3 ("high addition") (16-24 bands). The species richness was significantly different

134 between the three treatments (ANOVA, p<0.03), where Tank 1 and 2 did not differ from each other, but were both significantly different from Tank 3 (SNK-test, p<0.1 ). Four bands were present at the same location in the lanes of all the analysed slides and may therefore be assigned to represent seawater biofilm generalists (Fig. 5.1 ). P. tunicata was added in a DOGE-ladder located between the lanes for each of the treatments, but no bands matching with the P. tunicata band in the DOGE-ladder were observed in any of the treatments.

Pseudoalteromonas diversity within replicates of the same treatment was more similar than the diversity observed between different treatments inoculated with different densities of P. tunicata (Fig. 5.2). The number of DOGE-bands was highest in samples from the "low addition tank" (5 bands) and lowest in samples from the control and the "high addition tank" (both with 3 bands). By matching the P. tunicata band in the DOGE-ladder with each of the treatments, the P. tunicata DGGE-band in both treatments with P. tunicata addition was identified (Fig.5.2). The P. tunicata band in the samples from the "high addition tank" had higher intensity than in samples from the "low addition tank", which reflected the number of P. tunicata cells added to the two tanks (Fig. 5.2).

5.3.2 Morphology of the natural seawater community on glass surfaces

There were distinct differences in the appearance of the slides from different treatments. Visually, the control biofilms appeared white, the low addition biofilms were light yellow in colour and the biofilm with the high dose of added P. tunicata appeared yellow, presumably because of the presence of GFP-labelled P. tunicata cells. Also the strength of attachment of the biofilm to the glass surface differed between treatments. The biofilms in the tank to which had been added high amounts of P. tunicata were easily removed from the glass surface. The control biofilms were solid and sticky in appearance. The biofilms from the tank to which had been added small amounts of P. tunicata contained both well-defined biofilm threads and slime and were not easily removed from the surfaces. However, when the slides were observed under the microscope there were no major differences between the treatments or the control.

135 5.3.3 Introducing P. tunicata and R. gal/aeciensis into an epiphytic community on U. australis

The second aim was to determine if P. tunicata or R. gallaeciensis could colonise and impact on an epiphytic community on U. australis. To achieve this aim, a complex epiphytic community was allowed to establish on U. australis discs before being colonised with P. tunicata or R. gallaeciensis at two different densities.

When P. tunicata was inoculated at densities of 106cells mr', it did not attach very well and the few attached cells did not persist on the surface. Even at higher densities of 108cells mr'P. tunicata did not colonise effectively and had no discemable effect on biofilm structure. There were no significant differences between the community invaded with P. tunicata (Fig. 5.3A) or P. tunicata AlpP mutant (Fig. 5.3B). There was also no difference in the epiphytic community inoculated with P. tunicata as compared to the control community that had not been inoculated with any bacteria (Fig. 5.30).

In contrast, R. gallaeciensis was able to colonise at both cell densities. However, colonisation was more effective at higher densities and R. gallaeciensis was able to invade the epiphytic community and totally dominate it within 8 days (Fig 5.3C).

5.3.4 Mixed species biofilms established on glass surfaces.

The third aim was to determine if P. tunicata or R. gallaeciensis could colonise a synergistic mixed species biofilm both on inanimate and animate surfaces. To achieve this aim, 17 strains of epiphytic bacteria from U. australis were screened in various combinations for their ability to form a better biofilm as a mixture, rather than individually. The possibility that certain combinations of bacteria within the epiphytic community may be able to form a synergistic mixed species biofilm that may allow it to resist invasion by P. tunicata or R. gallaeciensis was explored.

It was seen that the synergistic mixed species biofilm formed by Microbacterium sp., Shewanella pacifica, Acinetobacter lowffii and Dokdonia donghaensis isolates was indeed able to resist invasion by P. tunicata as evidenced by Fig. 5.4. However, R. gallaeciensis was able to invade and dominate the synergistic mixed species biofilm (Fig 5.5).

136 45% P. tunicata low Control P. tunicata high

• •

. . .- ._ . .. -

•

11 15 16 8 13 13 16 22 24

60 %

Fig. 5.1 Denaturing Gradient Gel Electrophoresis (DGGE) profiles of eubacterial communities in the "low addition", "high addition" and "control" tanks. Profiles were obtained with the primers Eub34 I F-GC and Univ907RC. Numbers indicate the species richness in each lane. The gradient was 30-70% (visible 45-60%). Asterisks indicate bands that could be found at the same location in all nine lanes.

137 42% 3 3 3 5 5 5 3 3 3 .,_,,._ 1-.W

• I .. ,.

P. tunicata P. tunicata Control low high 48%

Fig. 5.2 DGGE profiles of Pseudoalteromonas communities in the "low addition", "high addition" and "control" tanks. Profiles were obtained with the primers Eub341F GC and Psalt8 l 5R. Numbers indicate the species richness in each lane. The gradient was 35-55% (visible 42-48%). The asterisk indicates that the band belongs to P. tunicata, as verified with a DGGE-ladder containing P. tunicata amplified DNA.

138 A

Day l Day 2 Day4 Day 8

Fig. 5.3 Colonisation of an epiphytic community by P. tunicata and R. gallaeciensis. Non-axenic V. australis was allowed to develop an epiphytic community for 6 days before being inoculated with bacterial strains. (A) P. tunicata inoculated at 108cells m1-1 colonises an epiphytic community poorly and does not persist beyond a week. Image is of unstained algal tissue, and cells visible are GFP-labelled P. tunicata, as well as some autofluouresent algal cells. (B) The P. tunicata AlpP mutant strain is unable to colonise the epiphytic community and image is the epiphytic community stained with acridine orange. (C) GFP-labelled R. gallaeciensis inoculated at 108cells mr1 is able to invade an epiphytic community and dominate by day 8. (D) The control epiphytic community in the absence of inoculated bacteria. Image is of epiphytic community stained with acridine orange. Images are representative of 3 biofilm experiments and were taken at 600X magnification.

139 100

C: 0 'iii 60 I'll > .E QI Cl I'll 50 C: -QI ...u QI a. 40

30 ----+--- All species --+- M. phyl/ospaerae --+- S. Japonica 20 -x- D. donghaensis ----t,-A. lwoffi

0 0 ------~ ----~------r ------, 4 16 24 44 Time (hours)

Fig. 5.4 Bacterial invasion of one- and four-species biofilm by Pseudoalteromonas tunicata. Biofilms composed of one or four strains of the epiphytic isolates, Microbacterium phyllosphaerae, Shewanella japonica, Dokdonia donghaensis and Acinetobacter lwoffii, were established in glass flow cells inoculated with equal total cell densities. After 2 h of growth in the presence of media flow, the antibacterial protein-producing Pseudoalteromonas tunicata was introduced to the biofilms. This strain constitutively expressed green fluorescent protein. At various time points, the fraction of the surface covered by P. tunicata-biofilm was determined by staining of the biofilm cells followed by confocal laser scanning microscopy, image analysis, and comparisons to corresponding biofilms not subjected to P. tunicata invasion. This is presented as "percentage of invasion by P. tunicata". Bars represent means±standard errors of eight replicates.

140 100

---o--- All species C 60 ---- M. phyl/osphaerae 0 'iii --S. japonica ftl > --- D. donghaensis .5 -llE- A. lwoffi QI 50 Cl ftl C -QI u ... 40 QI a.

0 4 16 24 44 Time (hours)

Fig. 5.5 Bacterial invasion of one- and four-species biofilm by R. gallaeciensis. Biofilms composed of one or four strains of the epiphytic isolates, Microbacterium phyllosphaerae, Shewanella japonica, Dokdonia donghaensis and Acinetobacter lwoffii, were established in glass flow cells inoculated with equal total cell densities. After 2 h of growth in the presence of media flow, R. gallaeciensis was introduced to the biofilms. At various time points, the fraction of the surface covered by R. gallaeciensis -biofilm was determined by staining of the biofilm cells followed by confocal laser scanning microscopy, image analysis, and comparisons to corresponding biofilms not subjected to R. gallaeciensis invasion. This is presented as "percentage of invasion by P. gallaeciensis". Bars represent means±standard errors of eight replicates.

141 5.3.5 Synergistic mixed species biofilms on U. australis.

In order to examine the synergistic effects of mixed species biofilm formation in an ecologically relevant setting, experiments were repeated on the surface of axenic U australis discs. Results obtained here mimicked that of the flow cells, with R. gallaeciensis dominating the mixed species biofilm and P. tunicata unable to colonise the mixed species biofilm. As mixed species biofilms took longer to establish on algal tissue incubated in filtered seawater, compared to those in flow cells, experiments were extended up to 96 hours. Despite these experiments being conducted over a longer time period, results were strikingly similar for the two surfaces.

5.4 DISCUSSION

Although mixed species microbial communities are common m the marme environment, relatively little is known about their resistance to colonisation and invasion by inhibitory bacteria. Evidence presented here suggests that epiphytic bacteria have evolved to exploit the benefits of a community lifestyle, with some of the component species interacting cooperatively to enhance resistance to extracellular inhibitory compounds produced by inhibitory bacteria. The present study demonstrates that although P. tunicata is able to colonise a natural seawater community on glass surfaces at in situ concentrations and change biofilm structure, it is unable to colonise an epiphytic community on U australis. A synergistic mixed species biofilm is able to resist the invasion of P. tunicata when compared to biofilms formed by individual strains. In contrast R. gallaeciensis is able to invade a synergistic mixed species biofilm and an epiphytic community on U australis.

5.4.1 Natural seawater community on glass surfaces

It was observed that the addition of high concentrations of P. tunicata to the natural seawater community on glass slides resulted in an increased species richness in the community. This may be due to the production and release of the antibacterial protein AlpP, (James et al, 1996), which may result in cell lysis and release of cellular material to the biofilm community. The increased level of available nutrients might increase the number of free available niches and thereby stimulate bacterial diversity as seen in the

142 "high addition" treatment (Fig.5.1). Alternatively, the release of the antibacterial protein may result in the removal of some dominant members of the microbial community, paving the way for an invasion by opportunistic bacteria, which would lead to an increase in diversity.

Not surprisingly, no increase in Pseudoalteromonas species richness was observed for the "high addition" tank as compared to the control (Fig. 5.2). It has been shown previously that P. tunicata displays antibacterial activity against other Pseudoalteromonas species when present in high cell densities (Holmstrom et al., 2002). It has also been shown that the AlpP produced by P. tunicata has autotoxic effects (James et al., 1996) and causes cell death and detachment in dense and mature P. tunicata biofilms (Mai-Prochnow et al, 2004) Results presented in this study support these findings, where high concentrations of P. tunicata cells appeared to enhance detachment of biofilm cells which resulted in an altered biofilm community as compared to the control biofilms (Fig. 5.1).

Unexpectedly, there was also an increase in diversity in the community that had P. tunicata added at "in situ" densities. At such low densities of cells, the antibacterial protein would be expected to have minimal effects on cell lysis and nutrient release. However, an increase from 11 to 14 DGGE-bands was observed as compared to the control treatment. Microscopy showed that there were no differences in biofilm structure between the control and low addition tank slides. Also no dispersal of cells was observed, resulting in no major community changes. Results from DGGE revealed that Pseudoalteromonas species richness, however, was higher in biofilms from the "low addition" treatment compared to the control and "high addition" treatment (Fig. 5.2). It is possible that P. tunicata at in situ densities stimulates the attachment of other surface living Pseudoalteromonas species from the seawater and thereby strengthens the antifouling properties of the biofilm towards subsequent macro fouling and grazing. In support of this conclusion, other reports provide evidence that eukaryotic fouling protection is caused by the action of mixed species biofilms on, for example soft corals (Dobretsov and Qian, 2004), squid egg capsules (Barbieri et al, 2001), Ulva reticulata (Dobretsov and Qian, 2002) and U lactuca (Holmstrom et al., 1996; Egan et al., 2000; Rao et al, 2005). These biofilm communities were all found to contain members of the genus Pseudoalteromonas that might act as regulators of the protective 143 biofilm community. Likewise, earlier studies suggested that the expression of antibactenal activity is important for maintaining microbial activity and diversity within microhabitats (Defreitas and Fredrickson, 1978).

5.4.2 Epiphytic community on U. australis

Major differences in the ability of P. tunicata to colonise and invade a complex community were observed when inoculation was conducted on an epiphytic community on U. australis, as compared to a natural seawater community on glass slides. P. tunicata did not colonise U. australis at in situ cell densities, and even when inoculated at high densities to give a final density of 108 cells mr1, P. tunicata cells did not persist on the surface, and therefore had minimal effects on the epiphytic community. As described above, P. tunicata appears to require a natural planktonic seawater community to colonise and establish itself on the plant surface (Chapter 3), but is unable to integrate into an established epiphytic community.

It is also likely that a stable epiphytic community on U. australis may have evolved resistance to invasion. It has been proposed that as the composition of a community becomes stable over time and reaches a climax (the "climax community"), it is better able to withstand environmental perturbations (Alexander, 1971). One of the oldest generalisations about community resistance to invasions is that more diverse communities are more resistant to invasion (Elton, l 958). A mechanism proposed to underlie this relationship is the complementarity effect, in which greater phenotypic diversity in communities enhances resource utilisation or positive interactions among species, thereby leaving fewer resources for invaders (Wilsey and Polley, 2002). It is possible that positive interactions have evolved among a number of the strains within the epiphytic community on U. australis which would make them more resistant to invasion by P. tunicata. Furthermore, previous studies (Chapter 3) suggest that the surface of the algae is nutrient limited, so that nutrient resources are likely to be depleted by the initial colonists of the algal surface (Lindow and Andersen, 1996), making it potentially more difficult for P. tunicata to establish itself. A second mechanism to explain community resistance to invasion is the selection effect whereby communities with higher diversity have a greater probability of including a species with a strong effect on the invader (Wilsey and Polley, 2002). It is conceivable that the

144 presence of inhibitory bacteria such as R. gallaeciensis within the natural epiphytic community on U. australis would result in P. tunicata not being able to integrate into the community. Thus compared to the natural seawater community on glass slides, the epiphytic community on U. australis may have evolved resistance to invasion.

However, the resistance offered by a stable epiphytic community on U. australis and possible nutrient constraints appear not to be an impediment to R. gallaeciensis which is able to invade the established epiphytic community and become the dominant species. R. gallaeciensis with its strong antibacterial activity, coupled with its nutritional versatility (Ruiz-Ponte et al., 1998) is able to dominate the epiphytic community. One possibility is that the success of R. gallaeciensis may also be attributed to its production of signalling molecules as recent studies from our laboratory have shown that R. gal/aeciensis produces N-acyl-homoserine lactones (AHLs) suggesting that it is capable of quorum sensing (Case R.; Low, A.; Kjelleberg, S. unpublished). To date there is no evidence that P. tunicata produces signalling molecules (Franks, 2005). Signalling molecules have been shown to exert not only intra-species control but also interspecies control on growth and expression of specific phenotypes, such as the interference of inhibitor production in other bacterial strains (Egland et al., 2004 ). Thus signalling molecules have the ability to alter composition of bacterial communities.

5.4.3 Invasion of synergistic mixed species biofilms

The synergistic mixed species biofilm established on glass surfaces was resistant to invasion by P. tunicata both on artificial and living surfaces. It is well established that bacteria in biofilms are more tolerant of antimicrobial agents, but this effect can be enhanced in mixed species biofilms. Neighbouring cells of a different species can produce neutralising enzymes that protect inherently susceptible organisms from inhibitors (Brook, 1989). Increased resistance to P. tunicata may also reflect changes in mixed species biofilm matrix, which might influence the permeability of the antibacterial protein. It is possible that interactions between the different matrix polymers might result in a more viscous matrix. Increased matrix viscosity was suggested to explain the enhanced resistance to disinfection of mixed species biofilms of Enterobacter agglomerans and Klebsiella pneumoniae (Skillman et al., 1999). Furthermore, rheological interactions between polysaccharides from Pseudomonas

145 cepacia and P. aeruginosa have also been shown to decrease the diffusion and antimicrobial activity of antibiotics (Allison and Matthews, 1992).

The fact that individually the target strains examined were almost eliminated from dual species biofilms with P. tunicata, suggests that they benefited from being part of the multi-species biofilm. Protective effects of one species on another within mixed biofilms have been described in the literature (Whiteley et al., 2001; Leriche et al., 2003). Generally it has been shown that within communities, strongly interacting species limit the invasion possibilities of most similar species (Case, 1990). Even a superior invading competitor is repelled by strongly interacting species (Miller et al., 2002) which are able to maintain population integrity and stability.

However, as noted above, possible changes in the mixed species biofilm matrix did not lead to increased resistance to the inhibitors produced by R. gallaeciensis. Diffusion of antibacterial proteins produced by R. gallaeciensis may not be hampered by increased matrix viscosity in mixed species biofilms. Similar to results from flow cell studies, R. gallaeciensis was able to invade and dominate the mixed species biofilm on the surface of the algae.

5.5 CONCLUSIONS

Microbial communities are ubiquitous in nature, and it has been suggested that the environmental heterogeneity generated within biofilm communities provides a form of "biological insurance" that can safeguard the microbial community in the face of adverse conditions (Marsh, 2005). It is shown here that a natural seawater community established on glass slides is stable and able to resist invasion by P. tunicata at in situ cell densities. It was only when cells were inoculated at densities which were a thousand fold higher than the in situ density of P. tunicata, that significant changes in the bacterial community structure (increased diversity) were observed, implying that a stable natural seawater community is able to resist perturbations.

Although P. tunicata at high densities is able to change the structure of a natural seawater community on inanimate surfaces, it is unable to invade a synergistic mixed

146 species biofilm in glass flow cells. Given that microbial communities are made up of a combination of different consortia, it is likely that certain assemblages of bacteria within the community would have evolved strategies to persist under hostile conditions. A synergistic mixed species biofilm comprising a combination of four specific strains isolated from U. australis was found to resist invasion by P. tunicata. Bacterial species appear to gain fitness advantages from residing in synergistic mixed species biofilms compared to single species biofilms, and such relationships may have evolved as local pockets of resistance in a largely susceptible community on U. australis. These assemblages may have developed in response to certain dominant bacteria such as P. tunicata. However, mixed species biofilms in flow cells did succumb to R. gallaeciensis.

On the plant surface, P. tunicata has no effect on the epiphytic community, possibly because fewer resources remain available for new colonists and the likelihood of selection for inhibitory bacteria within the community preventing the establishment of invaders. In contrast R. gallaeciensis, at high cell densities, is able to invade and dominate both the epiphytic community and synergistic mixed species biofilms. This has implications in biofouling control, as R. gallaeciensis has the potential to control the growth of natural marine communities.

147 5.6 REFERENCES

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Allison, D.G., and Matthews, M.J. (1992) Effect of polysaccharide interactions on antibiotic susceptibility of Pseudomonas aeruginosa. J Appl. Bacterial. 73: 484-488.

Barbieri, E., Paster, B.J., Hughes, D., Zurek, L., Moser, D.P., Teske, A., and Sogin, M.L. (2001) Phylogenetic characterization of epibiotic bacteria in the accessory nidamental gland and egg capsules of the squid Loligo peali (Cephalopoda: Loliginidae). Environ. Microbial. 3: 151-167.

Brook, I. (1989) Direct and indirect pathogenecity of beta-lactamase producing bacteria in mixed infections in children. Crit. Rev. Microbial. 16: 161-180.

Case, T.J. (1990) Invasion resistance arises in strongly interacting species-rich model competition communities. Proc. Natl. Acad. Sci. USA 87: 9610-9614.

Cowan, S.E., Gilbert, E., Liepmann, D., and Keasling, J.D. (2000) Commensal interactions in a dual-species biofilm exposed to mixed organic compounds. Appl. Environ. Microbial. 66: 4481-4485.

Defreitas, M.J., and Fredrickson, A.G. (1978) Inhibition as a Factor in Maintenance of Diversity of Microbial Ecosystems. J Gen. Microbial. 106: 307-320.

Dobretsov, S., and Qian, P.Y. (2004) The role of epibotic bacteria from the surface of the soft coral Dendronephthya sp in the inhibition of larval settlement. J. Exp. Mar. Biol. Ecol. 299: 35-50.

Dobretsov, S.V., and Qian, P.Y. (2002) Effect of bacteria associated with the green alga Ulva reticulata on marine micro- and macrofouling. Biofouling 18: 217-228.

Egan, S., James, S., Holmstrom, C., and Kjelleberg, S. (2002a) Correlation between pigmentation and antifouling compounds produced by Pseudoalteromonas tunicata. Environ. Microbial. 4: 433-442.

Egland, P.G., Palmer, R.J., and Kolenbrander, P.E. (2004) Interspecies communication in Streptococcus gordonii-Veillonella atypica biofilms: Signaling in flow conditions requires juxtaposition. Proc. Natl. A cad. Sci. USA 101: 16917-16922.

148 Elton, C.S. (1958) The Ecology of Invasion by Plants and Animals. London: Chapman and Hall.

Franks, A. (2005) Jdentfication and characterisation of an antifungal comound produced by the marine bacterium Pseudoalteromonas tunicata. PhD Thesis: University of New South Wales, Sydney, Australia.

Gil-Turness, M.S., and Fenical, W. (1992) Embryos of Homarus americanus are protected by epibiotic bacteria. Bio. Bull. 182: 105-108.

Holmstrom, C., Egan, S., Franks, A., McCloy, S., and Kjelleberg, S. (2002) Antifouling activities expressed by marine surface associated Pseudoalteromonas species. FEMS Microbiol. Ecol. 41: 47-58.

Jackson, C.R., Churchill, P.F., and Roden, E.E. (2001) Successional changes in bacterial assemblage structure during epilithic biofilm development. Ecology 82: 555- 566.

James, S., Holmstrom, C., and Kjelleberg, S. (1996) Purification and characterisation of a novel antibacterial protein from the marine bacterium D2. Appl. Environ. Microbiol. 62: 2783-2788.

Lindow, S.E., and Andersen, G.L. (1996) Influence of immigration on epiphytic bacterial populations on navel orange leaves. Appl. Environ. Microbiol. 62: 2978-2987.

149 Marsh, P.O. (2005) Dental plaque: biological significance of a biofilm and community life-style . .f. Clin. Periodontol. 32: 7-15.

Marsh, P.O., and Bowden, G.H.W. (2000) Microbial community interactions in biofilms. Cambridge: Cambridge University Press.

McBain, A.J., Allison, D., and Gilbert, P. (2000) Emerging strategies for the chemical treatment of microbial biofilms. Biotechnol. Genet. Eng. 17: 267-279.

Miller, T.E., Kneitel, J.M., and Bums, J.H. (2002) Effect of community structure on invasion success and rate. Ecology 83: 898-905.

Moller, S., Sternberg, C., Andersen, J.B., Christensen, B.B., Ramos, J.L., Givskov, M., and Molin, S. (1998) In situ gene expression of mixed-culture biofilms: evidence of metabolic interactions between community members. Appl. Environ. Microbiol. 64: 721-732.

Rao, D., Webb, J.S., and Kjelleberg, S. (2005) Competitive interactions in mixed species biofilms containing the marine bacterium Pseudoalteromonas tunicata. Appl. Environ. Microbiol. 71: 1729-1736.

Ruiz-Ponte, C., Samain, J.F., Sanchez, J.L., and Nicolas, J.L. (1999) The benefit of a Roseobacter species on the survival of scallop larvae. Mar. Biotechnol. 1: 52-59.

Skillman, L.C., Sutherland, I.W., and Jones, M.V. (1999) The role of exopolysaccharides in dual species biofilm development. J. Appl. Microbiol. 85: 13s- 18s.

Whiteley, M., Ott, J.R., Weaver, E.A., and McLean, R.J.C. (2001) Effects of community composition and growth rate on aquifer biofilm bacteria and their susceptibility to betadine disinfection. Environ. Microbiol. 3: 43-52.

Willcock, L., Holah, J., Allison, D.G., and Gilbert, P. (1997) Population dynamics in steady-state biofilms: effects of growth environment upon dispersal. Cardiff: Bioline Press.

Wilsey, B.J., and Polley, H.W. (2002) Reductions in grassland species evenness increase dicot seedling invasion and spittle bug infestation. Ecol. Lett. 5: 676-684.

150

Chapter Six

General Discussion

The work presented in this thesis explored the hypothesis that epiphytic bacteria on the green alga Ulva australis protect the plant surface against fouling. In order to understand the processes that determine the composition of epiphytic communities, interactions were studied at three levels: (i) between the host plant and epiphytic bacterial communities (Chapter 3), (ii) interactions within epiphytic bacterial communities (Chapters 2, 3 and 5) and (iii) betw~en bacterial biofilms and micro- and macro-fouling organisms (Chapter 4). The principal microorganisms chosen for the study of the colonisation ecology of U. australis were the inhibitory bacteria P. tunicata and R. gallaeciensis and results presented here suggest that these bacteria may play a role in the defence against fouling organisms.

6.1 HOST-BACTERIAL INTERACTIONS

The factors that influence microbial colonisation and the outcome of competitive interactions on U. australis are likely to have a profound impact on the antifouling and inhibitory activity of the epiphytic community. The results presented in Chapter 3 revealed fundamental differences in the way P. tunicata and R. gallaeciensis colonised the surface of U. australis. Whilst R. gallaeciensis is capable of colonising U. australis under a range of conditions, colonisation by P. tunicata is enhanced by high cell densities, presence of cellobiose in the pre-culture, inoculation in the dark and interactions with a natural seawater community which promote the ability of this organism to attach and persist on the surface of the algae.

A spatial heterogeneity of nutrients on the plant surface is suggested by the distribution of microcolonies across the surface of the alga. Observations of vast expanses of uncolonised areas interspersed with clumps of bacteria, suggest "oases" of relatively abundant nutrient leakage in a largely oligotrophic environment. Unlike higher plants

151 where certain regions of the leaves accumulate more nutrients, the thallus surface of U australis is morphologically undifferentiated and it is not known why certain areas may harbour more nutrients or are more conducive for attachment and growth of epiphytic bacteria. Distribution of nutrients has implications for microbial colonisation and competitive interaction on algae, as it seems to significantly limit microbial interactions and leads to coexistence of competing strains. Spatial separation of microcolonies provides refuges where strains can persist out of reach of their respective "nemeses". Thus environmental factors, such as the distribution and abundance of nutrients are likely to have a profound impact on the composition of the epiphytic community and consequently the antifouling activity of host surfaces. It suggests that the antifouling activity of epiphytic bacteria can be enhanced by the alga, through alteration of the nutritional status of the algal surface.

A nutrient limited surface on U australis is a possible explanation for the requirement of a mixture of strains by P. tunicata for colonisation of the algal surface. P. tunicata inoculated in a natural seawater community may be able to overcome nutrient limitations imposed by the external macro-environment by creating, through its metabolites, a range of microenvironments that enable the survival and growth of P. tunicata (Schreiber et al., 2005). The underlying mechanisms of cooperative biofilm formation are the basis of ongoing experiments at the Centre for Marine Biofouling and Bio-innovation.

The extent and nature of microcolony formation have implications for survival during microbial colonisation of marine algal surfaces. Microcolony formation allows for enhanced ability to scavenge nutrients and also results in persistence of bacterial strains during competition within biofilms. Moreover, the aggregation of cells encased within an EPS matrix may provide protection against periodic desiccation. Thus, microcolonies may represent an adaptive strategy that promotes persistence under stressful conditions.

Microcolony formation may also be a mechanism for promoting diversity on algal surfaces. Clustering of cells in close proximity would maximise opportunities for gene transfer to take place within microcolonies. Although plasmid transfer in algal epiphytic communities remains unstudied, given that terrestrial epiphytic bacterial species exhibit 152 remarkably high rates of gene transfer (Lilley et al., 1996; Bjorklof et al., 2000) and the high levels of plasmid transfer detected in marine bacterial communities (Dahlberg et al., 1998), the potential for gene transfer on algal surfaces seems high. As bacterial communities on algae undergo substantial changes during a growing season (Fisher et al, 2002), it is likely that there is extensive mixing of genes within these communities. Thus, microcolonies on U australis may also mediate gene transfer and enhance diversity.

The success of R. gallaeciensis in colonising the plant surface may be attributed to several factors, and these include the production of signalling molecules, possible attraction of bacterial cells to plant derived cues and the likely production of plant hormones. Although the role of cell-cell signalling in colonisation of algal surfaces is not known, it is feasible that quorum sensing contributes to attachment and microcolony formation in Roseobacter gallaeciensis. Recent studies from our laboratory have shown that R. gallaeciensis produces N-acyl-homoserine lactones (AHLs) suggesting that it is capable of quorum sensing (Case, R., Low, A., Kjelleberg, S., unpublished data). Identification of the phenotypes under AHL regulation would be a crucial next step and further study is required to establish whether AHLs play a role in mediating interactions between algae and bacteria. It is also conceivable that algal cues may serve as an attractant LO R. gallaeciensis and DMSP is a likely candidate for such a cue. Several members of the Roseobacter clade are attracted to DMSP released by dinoflagellates (Miller and Belas, 2004). While there is no information on DMSP-utilizing bacterial epiphytes of macroalgae, there is evidence that fungi involved in the decay of cordgrass (Spartina alternaria) are capable of using cytosolic DMSP (Bacic et al., 1998), and data suggests coevolution of the DMSP-utilizing fungi and the DMSP producing plant. Furthermore, it has been proposed that the substrate for the sulphur based inhibitory compounds produced by R. gallaeciensis (thiotropocin and tropodithietic acid), is DMSP (Bruhn et al, 2005). The production of plant hormones such as indole acetic acid (IAA) may also confer epiphytic fitness to Roseobacter gallaeciensis. It has been shown in higher terrestrial plants that not only does epiphytic bacterial secretion of IAA down-regulate genes involved in plant defence responses, making it easier for bacteria to establish themselves on the surface (Yamada, 1993), it also modifies the microhabitat of epiphytic bacteria by increasing nutrient leakage from plant cells (Brandl and Lindow, 1998). It is not known if R. gallaeciensis produces 153 IAA, but given that several members of this group do (Ashen and Goff, 2000), it may be another mechanism it has at its disposal for the effective colonisation of algal surfaces. Thus, colonisation of U. australis by R. gallaeciensis is probably multifaceted, involving a combination of inhibitory compounds, quorum sensing molecules, the ability to utilise a wide range of carbon sources and to respond to the presence of cues such as DMSP and the ability to secrete plant hormones such as IAA.

Mutualisms have traditionally been viewed as obligate, coevolved interactions uniformly benefiting both partners (Thomson, 2003), but this definition has broadened in more recent years, to include interactions in which the benefits exceed the costs for both participants (Hay et al., 2004). Furthermore, mutualisms are now considered to be dynamic in nature and could change from mutual to neutral to antagonistic, depending on the environmental setting. Based on this broader definition of mutualism, results described in this study suggest that U. australis shares a mutualistic relationship with P. tunicata and R. gallaeciensis. The host provides a preferred surface for bacterial growth and the bacteria chemically defend this resource, enhancing host fitness. In return, the host provides the microbes with nutrients and a predictable environment. It is likely that U. australis produces compounds which allow it to interact with its epiphytic microbial community. As cross-talk between host and symbiont is presumably required for the maintenance of the symbiosis (Sara et al., 1998), it would be interesting to explore the role of signalling molecules in structuring bacterial communities on algae.

6.2 INTERACTIONS WITHIN BIOFILM COMMUNITIES

Although there have been some investigations on algal-bacterial interactions in marine systems, bacterial-bacterial interactions in algal epiphytic communities remain largely unstudied. Experiments conducted in flow cells demonstrated that during competitive biofilm formation P. tunicata and R gallaeciensis were dominant against other marine strains. This dominance could be attributed to the production of inhibitory compounds. Epiphytic bacteria live in a highly competitive environment where space and access to nutrients are limited and the production of inhibitors may provide fitness benefits under a wide range of conditions (Durrett and Levin, 1997). It was found that the P. tunicata AlpP mutant was less competitive in certain situations, suggesting that this compound

154 provided a competitive advantage for P. tunicata. R. gallaeciensis was resistant to the antibacterial protein produced by P. tunicata, and it is likely that other members of the epiphytic community have developed resistance mechanisms in response to inhibitory compounds produced by P. tunicata and R. gallaeciensis. The ability to resist such toxic effects, may be a prerequisite to survival in the habitat. However, the distribution or occurrence of resistant strains in the epiphytic community is not known and mechanisms of resistance are relatively unexplored in the marine environment.

Studies of competitive biofilm formation in glass flow cells also suggested that microcolony formation may enhance the ability of some organisms to compete against P. tunicata and persist within the flow cell reactor. It seems that high densities of cells within microcolonies allow for enhanced persistence during co-culture with a superior competitor and improve the competitiveness of an otherwise poor competitor. Although the formation of well-defined microcolonies m a pre-established biofilm appears to buy more time for strains competing with P. tunicata, the superior competitor eventually overtakes strains. Furthermore, microcolonies were not always a guarantee against invasion, and pre-established microcolonies of P. tunicata always succumbed to R. gallaeciensis. R. gallaeciensis produces inhibitory compounds which are small (Ruiz-Ponte et al., 1998; Brinkhoff et al., 2004) and likely to diffuse easily, therefore microcolonies do not appear to present a barrier to these compounds. The large antibacterial protein produced by P. tunicata may diffuse slowly and have a delayed response on strains that establish large microcolonies. The shielding effects of large numbers of cells coupled with the reduced permeability of EPS mkay result in protective effects of large microcolonies (Wilson and Lindow, 1994).

Competitive interactions among epiphytic bacteria may lead to a predominance of inhibitory strains on U. australis, which in tum may benefit the plant in its defence against fouling. Interestingly, competitive interactions may actually promote diversity in the natural environment. The impact of a single interaction on diversity would be small, but competitive interactions between several inhibitory bacteria would potentially provide a network of interlacing competitive interactions and therefore have a significant cumulative effect on diversity (Riley, 1998). Diverse communities have long been equated with stability (Elton, 1958), so this may have implications for the stability of epiphytic populations on U. australis. Thus, an "arms race" between 155 interacting organisms can promote genetic and phenotypic diversity, suggesting that competitive interactions may lead to an increased stability of epiphytic communities. Clone library data studies demonstrate that although there are spatial and temporal changes in epiphytic community composition on U. australis, a subpopulation of the community remains unchanged (Tujula, N., Crochetti, G., Holmstrom, C., Dahllof, I., and Kjelleberg, S. unpublished data) indicating that the epiphytic community may indeed be stable.

In most natural environments biofilms will consist of a mixture of organisms which form complex communities. Diversity of complex communities has been well studied, and results indicate that the level of microbial diversity observed in marine epibiotic communities is much greater than had been anticipated (Weidner et al., 1996; Fisher et al., 1998; Gillan et al., 1998; Polz et al., 1999). Relatively little is known however, of the synergistic or antagonistic interactions within complex communities. With the exception of studies on oral biofilms, few studies have specifically addressed interactions within multi-species biofilms. Understanding the inter-relationships within bacterial communities, particularly as they adapt to changes, is likely to open up new avenues for their control, and give insight into their resistance to invasion.

The impact of invasion by P. tunicata and R. gallaeciensis on an epiphytic community on U. australis was described in Chapter 5. It was found that R. gallaeciensis was an aggressive coloniser and invaded and dominated an epiphytic community. This was not an unexpected finding, as R. gallaeciensis has been shown to have strong antibacterial activity on both inanimate (Chapter 2, 4) and animate surfaces (Chapter 3). However, it would have been useful to conduct some molecular analysis of the community to get an indication of the change in microbial diversity after the introduction of R. gallaeciensis. In addition to the production of inhibitory compounds, production of signalling molecules by this bacterium may enhance its ability to colonise and dominate mixed communities. Recent work suggests that production of inhibitory compounds in a R. gallaeciensis strain is regulated in a quorum dependent manner (Bruhn et al., 2005). Given the potential significance of signalling molecules, coupled to the known interdependencies among biofilm populations, modulation of signalling could provide an important means of control of community diversity and structure. A full

156 understanding of the role of cell-cell signalling in epiphytic microbial communities would be a major goal for research in the future.

Epiphytic bacterial strains isolated from U. australis were screened for synergistic interactions and four isolates were found to interact synergistically in biofilms. Synergistic mixed species biofilms were more resistant to invasion by P. tunicata but succumb to R. gallaeciensis. Mechanisms for synergistic interactions are not known, but one possibility is coaggregation of cells. Similar to bacteria that grow in the oral cavity, it is possible that epiphytic bacteria have coevolved surface molecules for specific binding to each other. Interspecies binding could also be due to synergistic gelation of extracellular polymers leading to the formation of a stable microconsortium (Sutherland, 1985). Thus a strongly interacting consortium of isolates may be able to resist invasion by certain inhibitory bacteria.

6.3 INTERACTIONS BETWEEN BACTERIA AND FOULING ORGANISMS

Cell densities of P. tunicata and R. gallaeciensis required to deter settlement and attachment of fouling organisms were determined, and it was seen that surprisingly low densities of cells within biofilms of these bacteria were capable of antifouling effects

(Chapter 4). P. tunicata biofilms with densities in the range of 103 - 105 cells mr1 were inhibitory to algal spores, invertebrate larvae and fungi, whereas R. gallaeciensis was inhibitory at 103 - 104 cells mr1 for bacteria and larvae.

Although the role of bacterial biofilms in mediating the settlement of invertebrate larvae or algal spores is now well established, the prevention of settlement by bacterial cell densities as low as those used in the present study have not previously been reported. Furthermore, results described in this study demonstrate that low density biofilms of P. tunicata and R. gallaeciensis are also inhibitory to the attachment and growth of marine bacteria and fungi, indicating a broad spectrum of activity against fouling organisms. Although it has been reported that P. tunicata is found in very low numbers on the surface of U. australis (< lx 103 cells cm-2) (Skovhus et al., 2004), P. tunicata exhibits anti-algal and anti-fungal activity at such low densities. Furthermore, it is possible that R. gallaeciensis is present in larger numbers, and it is likely that

157 together P. tunicata and R. gallaeciensis are present in sufficient quantities on the plant to inhibit fouling organisms.

Recent studies on induction of morphogenesis of sea urchin larvae on both artificial and algal surfaces show that a wide range of bacteria from various genera induce the same settlement response, suggesting a redundancy in the function of bacteria on the surface of coralline algae (Huggett et al., 2005). A diverse range of bacteria induced settlement, with each performing a similar function. Similarly, results reported here indicated rhat R. gallaeciensis and P. tunicata provide complementary benefits to the host, with R. gallaeciensis being more effective at inhibiting larvae and bacteria, whilst P. tunicata is more effective for algal spores and fungi. There is now a growing realisation that the presence of multiple mutualists on a host is the rule rather than the exception (Stanton, 2003). It had been shown earlier that even when mutualists provide redundant benefits, mutualist diversity may serve as a buffer against environmental changes. For example, reef building corals harbour more than one genotype of symbiotic dinoflagellates, and evidence suggests that different genotypes perform better under different light conditions and are differentially susceptible to coral bleaching (Knowlton and Rohwer, 2003). Thus increased mutualist diversity appears to enhance host persistence.

Outcomes of biofouling assays were very different on algal surfaces as compared to Petri dishes. Unlike the case with an inanimate surface, the plant is responsive and dynamic, changing through time with respect to microclimate, exudation of nutrients, inhibitory compounds and volatiles. There appear to be many differences between plants in their natural habitats and plants manipulated in vitro. In order to move beyond the limitations and artefacts of laboratory based assays, it is important to test ecological functions of microbial defences in a field situation. Low densities of bacteria incorporated into a polymer and placed in the field for several weeks, could be examined for the settlement of a mixed assemblage of fouling organisms. For example, this has been done with crude extracts of the alga Delisea pulchra (Denys et al., 1995) and sponge extracts (Henrikson and Pawlik, 1995).

158 6.4 FUTURE DIRECTIONS

There are some issues which have emerged from this work which are worthy of further investigation:

1. The promoter-trapping approach termed in vivo expression technology (IVET) (Rainey and Preston, 2000) would provide an overview of responses as P. tunicata and R. gallaecinesis sense and adapt for epiphytic growth and survival. Bacteria that are found on algal surfaces probably have particular adaptations that allow them to exploit epiphytic environments and it is likely that such adaptive characteristic may have developed at the expense of those promoting survival in other habitats.

2. Genomic studies of P. tunicata and R. gallaeciensis hold the promise of understanding the influence of the environment on microbial activities and combined with approaches such as macro/microarrays would allow one to "interrogate" cells or communities under natural or imposed conditions. Furthermore, genome sequences would facilitate identification of plant inducible genes. The genome sequence for P. tunicata became available in September, 2005 through the Moore Foundation community sequencing programme and R. gallaeciensis is presently being sequenced.

3. Further work would involve making site directed transposon mutants of R. gallaeciensis that do not produce AHLs or inhibitory compounds and ascertain if the mutants have reduced epiphytic fitness. Factors that contribute to epiphytic fitness are just becoming understood in higher plants and it is likely that there may be other traits that contribute to epiphytic fitness. While the approach of constructing mutants altered in individual traits is powerful for evaluating the role of those traits in epiphytic fitness, it may however be restricted to traits that are identifiable in culture.

4. Higher plants respond to pathogens with the synthesis of jasmonic acid (JA) and JA has been reported in several lineages of nonvascular plants including green algae (Fujii et al., 1997). As the same biochemical pathways are triggered when exposed to both pathogens and epiphytic bacteria, it would be useful to determine genes induced in the alga by the presence of P. tunicata and R. gallaeciensis. A comprehensive analysis of induced genes in U. australis might lead to a better understanding of the alga-bacteria interactions.

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