The biofilm mode of life : mechanisms and adaptations 9781904933335, 1904933335

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Introduction Staffan Kjelleberg, Kevin C. Marshall, and Michael Givskov

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For more than a century, there has been a strong tradition for studies of microorganisms as planktonic, liquid cultures. his has imprinted the view that bacteria live as unicellular organisms. Although test tube laboratory studies have led to fundamental insights into basic life processes and have unraveled complex intracellular regulatory networks, it is now clear that microbial activity in nature is mainly associated with surfaces and scientists therefore need to turn their attention to studies of the sessile mode of life (Marshall, 1992). Indeed it may be argued that the ability to form surface-associated, structured and cooperative consortia (referred to as biofilms) is one of the most remarkable and ubiquitous characteristics of bacteria (Costerton et al., 1987). From an evolutionary perspective the biofilm lifestyle makes great sense. Sessile consortia of bacteria exhibit increased resistance and elegant adaptive responses to a range of factors that would otherwise negatively impact on their activity. Studies of microbial biofilms have a long history. Antonie van Leeuwenhoek, in the late eighteenth century, marveled at the diversity of the small life forms he observed in his microscope of biofilm scrapings from teeth surfaces. Early observations on surface related phenomena also included those of swarming by Proteus spp. on agar plates used for the detection of pathogenic bacteria (Hauser, 1885). It was seen that the swarming phenomenon allowed the bacteria to effectively colonize the entire surface of the agar plate. In the twentieth century, the development of slimes, or biofilms, on surfaces subjected to water flow was recognized and explored in a variety of habitats and applications. In particular, the effects of biofilms were regularly reported by engineers concerned by alterations in the efficiencies of high-speed vessels, of heat-transfer systems and of flow rates in water reticulation and hydroelectric pipelines. Furthermore, the importance of an increased surface area of microbial populations in the treatment of sewage and industrial wastes was realized at that time. While most studies of biofilms in this regard were carried out by engineers, microbiologists were also conscious of the effects of surfaces on microbial growth in natural habitats, such as soils (Cholodny, 1930) and waters, where the so-called “bottle effect,” with water samples in bottles of varying sizes or with added glass beads, resulted in a rapid increase in microbial numbers because of the increased surface area (Henrici, 1933; Heukelekian and Heller, 1940; Stark et al., 1938; ZoBell, 1937, 1943; ZoBell and Anderson, 1936). he importance and impacts of biofilms in natural ecosystems were also increasingly understood in the latter half of the last century. For example, the prevalence

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of bacterial and algal biofilms on surfaces in natural streams and the predominant effect of such mixed community biofilms on the otherwise pristine water ecosystem were unraveled by Geesey et al. (1977, 1978). Also, biofilm-forming marine bacteria were shown to be the initial colonizers with significant effects on the subsequent development of complex biofouling communities (Wilson, 1955). What were the early observations on mechanisms of attachment and biofilm formation? For a biofilm to develop, micro-organisms must first attain the proximity of the surface in question and then attach to that surface. Micro-organisms may be transported to surfaces in aquatic environments by several means (Marshall, 1986), including sedimentation, fluid dynamic forces, motility guided by chemotaxis and more passive mechanisms such as Brownian motion and cell surface hydrophobicity (Marshall and Cruickshank, 1973). he first definitive microscopic studies of bacteria in the vicinity of a solid surface was undertaken by ZoBell (1943), who noted that attachment was probably a two-phase process consisting of a primary, but reversible, attraction to the surface followed by a later firm, irreversible, adhesion. ZoBell (1943) also speculated on the possibility of polymer production by the bacteria being involved in the firm adhesion phase. Later, Marshall et al. (1971) published a seminal paper detailing the behavior of marine motile bacteria at solid surfaces and laid the foundation for a mechanistic physico-chemical explanation of the initial stages of reversible and irreversible attachment and hence colonization of surfaces by bacteria. Marshall and co-workers addressed why bacteria which possess a net negative charge are not repelled from like-charged surfaces in natural habitats. As a bacterium approaches a solid surface, the energy of interaction that occurs between the interacting electrical double layers around such surfaces was explained in terms of the Derjaguin-Landau and VerweyOverbeek (or DLVO) theory (Marshall, 1976). At low electrolyte concentrations the thickness of the double layers of counter ions increases resulting in a total repulsion between the interaction surfaces, whereas at high electrolyte concentrations the double layers are compressed exposing a secondary attraction zone, resulting from London–van der Waals attraction forces, at some distance from the surface. It is this zone that accounts for the reversible attraction of bacteria to a surface. Marshall et al. (1971) also demonstrated the need for the production by the bacterium of very fine extracellular polymeric fibrils to bridge the repulsion barrier and, hence, ensure irreversible adhesion to the surface, at a primary attraction zone in close proximity to the surface, observations subsequently confirmed by several investigators (e.g. Marshall and Cruickshank, 1973, and Fletcher and Floodgate, 1973). Further details and our current understanding of the orientation, motility and the shift from reversible to irreversible attachment by surface localized bacterial cells are provided in the chapter by MacEachran and O’Toole in this volume. While other factors, such as cell-surface hydrophobicity (Marshall and Cruickshank, 1973), the nature of the conditioning films on surfaces (Baier, 1980) surface active components for altered viscosity, temporary adhesion and translational motion (Humphrey et al., 1979), and extracellular matrix components (Pamp et al., this volume) contribute to bacterial adhesion processes in natural habitats, the discovery of the mechanisms of reversible and irreversible adhesion as the key steps in early bacterial colonization, laid the foundation for modern molecular biofilm research, as detailed in this monograph, for a range of organisms and habitats. Also, genetic analysis substantiated the important role of

Introduction

surface appendages, such as fimbriae and flagella, in specific adhesin–receptor mediated irreversible binding (Marshall, 1984). he current volume presents recent progress in our understanding of specific mechanisms that underpin the biofilm mode of life. his includes, in addition to areas highlighted above, studies on interactions among biofilm cells at a variety of levels; metabolic interactions, cell to cell communication and signal transmission across cells and species (see chapters by Yarwood, Wood and Bentley, Atkinson et al., Kolenbrander et al., Dow et al., and Eberl et al., this volume). Moreover, interactions based on competition and predation across species and domains have enjoyed strong interest in recent years as described in the chapters by Hogan, Matz and Eberl et al. Modern molecular techniques in combination with confocal laser scanning microscopy have also contributed greatly to our understanding of biofilm community structure and composition (see chapters by Kjelleberg and Givskov, and Kolenbrander et al.) as well as mechanisms of biofilm dissolution (see Chapter 9). Genomic based studies of such biofilm specific traits are rapidly unraveling much novel information on both monospecies and mixed species communities. Interestingly modern molecular microbiology increasingly adopts approaches developed in microbial ecology to better understand the sessile life style of bacteria in particular microbial habitats, including those of the human body. he recent surge in studies of biofilms is driven not only by our desire to explore the basic mechanisms that mediate bacterial colonization on surfaces, but also the realization that bacterial evolution very effectively allows for resistance to develop and hence makes almost any control measure we develop ineffective. Hence, like a century ago we still face bacterial supremacy, a fact that urges us to focus on more applied aspects of the discipline with new ideas and approaches generated from our basic knowledge about “life on surfaces” to win the battle against unwanted bacteria. he updated aspects of biofilm research reported in this volume should encourage interested researchers to integrate these new concepts in attempts to develop newer innovative approaches in studies of the behavior and functions of sessile bacteria. Acknowledgments We wish to thank the authors for providing their first class contributions to this book, Hugh Griffin and staff at Horizon Scientific Press, and Adam Abdool and Evi Fuary at the Centre for Biofilm and Bio-Innovation for their excellent contribution and support in preparing this volume. References Baier, R.E. (1980). Substrate influence on adhesion of microorganisms and their resultant new surface properties. In: Bitton, G., and Marshall, K.C. Eds. Adsorption of Microorganisms to Surfaces, WileyInterscience, New York, pp. 59–104. Costerton, J.W., Cheng, K.J., Geesey, G.G., Ladd, T.I., Nickel, J.C., Dasgupta, M., and Marrie, T.J. (1987). Bacterial biofilms in nature and disease. Annu. Rev. Microbiol. 41, 435–464. Cholodny, N. (1930). Über eine neue Methode sur Untersuchung der Bodenflora. Arch. Mikrobiol. 1, 620–652. Fletcher, M., and Floodgate, G.D. (1973). An electron microscope demonstration of an acidic polysaccharide involved in the adhesion of a marine bacterium to solid surfaces. J. Gen. Microbiol. 74, 325–334.

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Geesey, G.G., Richardson, W.T., Yeomans, H.G., Irvin, R.T., and Costerton, J.W. (1977). Microscopic examination of natural sessile bacterial populations from an alpine stream. Can. J. Microbiol. 23, 1733–1736. Geesey, G.G., Mutch, R., Costerton, J.W., and Green, R.B. (1978). Sessile bacteria, an important component of the microbial population in small mountain streams. Limnol. Oceanogr. 23, 1214–1223. Henrici, A.T. (1933). Studies of freshwater bacteria. 1. A direct microscopic technique. J. Bacteriol. 25, 277–287. Hauser, G. (1885). Über Fäulnisbakterien und deren Beziehung zur Septicämie. F.G.W. Vogel, Leipzig. Heukelekian, H., and Heller, A. (1940). Relation between food concentration and surface for bacterial growth. J. Bacteriol. 40, 547–558. Humphrey, B.A., Dickson, M.R., and Marshall, K.C. (1979). Physiochemical and in situ observations on the adhesion of gliding bacteria to surfaces. Arch. Microbiol. 120, 231–238 Marshall, K.C. (1976). Interfaces in Microbial Ecology (Cambridge, MA: Harvard University Press). Marshall, K.C. (1984). Microbial Adhesion and Aggregation (Berlin: Springer-Verlag). Marshall, K.C. (1986). Adsorption and adhesion processes in microbial growth at interfaces. Adv. Colloid Interface Sci. 25, 59–86. Marshall, K.C. (1992). Planktonic versus sessile life of prokaryotes. In he Prokaryotes. A Handbook on the Biology of Bacteria. Ecophysiology, Isolation, Identification, Applications. 2nd Edn. (A. Balows, H.G. Trüper, M. Dworkin, W. Harder and K.H. Schleifer, eds), pp. 262–275 (Berlin: SpringerVerlag). Marshall, K.C., and Cruickshank, R.H. (1973). Cell surface hydrophobicity and the orientation of certain bacteria at interfaces. Arch. Mikrobiol. 91, 29–40. Marshall, K.C., Stout, R., and Mitchell, R (1971). Mechanism of the initial events in the sorption of marine bacteria to surfaces. J. Gen. Microbiol. 68, 337–348. Stark, W.H., Stadler, J., and McCoy, E. (1938). Some factors affecting the bacterial population of freshwater lakes. J. Bacteriol. 36, 653–654. Wilson, D P 1955. he role of micro-organisms in the settlement of Ophelia biocornis Savigny. J. Mar. Biol. Ass. UK 34, 531–543. ZoBell, C.E. (1937). he influence of solid surfaces upon the physiological activities of bacteria in sea water. J. Bacteriol. 33, 86. ZoBell, C.E. (1943). he effect of solid surfaces upon bacterial activity. J. Bacteriol. 46, 39–56. ZoBell, C.E., and Anderson, D.Q. (1936). Observations on the multiplication of bacteria in different volumes of sea water and the influence of oxygen tension and solid surfaces. Biol. Bull. (Woods Hole) 71, 324–342. Stark, W.H., Stadler, J., and McCoy, E. (1938). Some factors affecting the bacterial population of freshwater lakes. J. Bacteriol. 36: 653–654. Wilson D.P. (1955). he role of micro-organisms in the settlement of Ophelia biocornis Savigny. J. Mar. Biol. Ass. UK 34, 531–543. ZoBell, C.E. (1937). he influence of solid surfaces upon the physiological activities of bacteria in sea water. J. Bacteriol. 33: 86. ZoBell, C.E. (1943). he effect of solid surfaces upon bacterial activity. J. Bacteriol. 46: 39–56. ZoBell, C.E., and Anderson, D.Q. (1936). Observations on the multiplication of bacteria in different volumes of sea water and the influence of oxygen tension and solid surfaces. Biol. Bull. (Woods Hole) 71: 324–342.

The Bioilm Mode of Life Staffan Kjelleberg and Michael Givskov

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Abstract Recent advances in studies of biofilm systems have generated a wealth of novel information on multicellular prokaryotic biology and have established models for the formation of biofilms and the biology of their lifecycles. As a prelude to the subsequent chapters in this volume, this introductory article is aimed at identifying the contextual scientific and experimental framework for contemporary biofilm research programs, and addresses the strengths and weaknesses of some of the current key biofilm models. We will discuss whether or not a unique biofilm specific gene expression underpins our observations on biofilm structure and biology. Further, we will highlight the limitations inherent to current genetic and physiological analyses of bacterial biofilms, including the strengths and weaknesses of the molecular toolbox and the biofilm assays commonly employed. Moreover, the extent by which multiple parallel pathways of biofilm formation exist will be addressed, with reference also to applications for novel control strategies based on contemporary advances in studies of bacterial biofilms. he chapter will conclude by discussing the relevance of a consensus view of bacterial biofilm formation and biology. Bioilm organization and differentiation Bacterial biofilms are multicellular consortia in which cells are embedded in an extracellular matrix at close proximity to one another. Such consortia are generally studied to assess particular properties of biofilms attached to solid surfaces, but they occur also as multicellular aggregates, flocs and granules suspended in the aqueous phase in many habitats. Biofilms of these kinds can be accommodated by single species, but in most cases in natural as well as artificial systems, they are mixed species consortia, ranging from phylogenetically highly diverse communities ( Juretschko et al., 2002; Lyautey et al., 2005; Taylor et al., 2004) to those in which one or a restricted number of species will be numerically and functionally dominant (Tujula et al., 2006). Intriguingly, whether biofilms consist of one or a mixture of a range of species, they display similarities with respect to architectural features and behavioral responses. For example, the microcolony structure observed in established mature biofilms is strikingly similar across mono- and multispecies biofilms, across different habitats, as well as for different organismal levels (Lawrence et al., 1991; O’Toole et al., 2000; Sauer et al., 2002). he latter applies to the similarities found between the multicellular consortia of well studied

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model bacterial systems, those of highly differentiated multicellular prokaryotes such as streptomycetes and myxobacteria, and colonial or sessile marine invertebrates. Moreover, strong similarities in behavioral responses and functional features have been reported for mono- vs. multispecies biofilms, as well as across organismal levels on the evolutionary scale. For example, it is now appreciated that biofilms have unique differentiation events that generate subpopulations of cells with apparently defined tasks and hence sharing of labor in the biofilm population (Allesen-Holm et al., 2006; Webb et al., 2003). hese events include different cells that build the different parts of the mushroom-like, or equivalent structures, of biofilms, and the generation of specialized dispersal cells or propagules for colonization of new surfaces (Caldwell et al., 2000; Labbate et al., 2004; Mai-Prochnow et al., 2006). he advances recently made in our understanding of biofilms systems whether well defined or of more complexity, derive in the main from a series of seminal studies on a restricted set of monospecies biofilm (Ghannoum and O’Toole G, 2004). he experimental approaches have preferentially been focusing on high resolution genetic and microscopic techniques. hese studies have laid the foundation for our current models of bacterial biofilms, their formation and life cycle, and they have subsequently been extrapolated and assessed for more complex systems (e.g. Auschill et al., 2001; Hope et al., 2002). Specifically, it is now appreciated that biofilm formation can be defined as a process that consists of defined stages. Commonly, we recognize that these stages include: reversible and irreversible attachment; surface motility and initiation of microcolony (or alternative structural arrangements) formation; maturation, ageing and differentiation of microcolonies; and, finally, biofilm dissolution and generation of specialized dispersal cells. It is further appreciated from the research reported on our currently best defined model systems that these stages and structural rearrangements include physiological characteristics and phenotypic responses apparently supportive of suggestions for the presence of a unique biofilm biology. Intuitively, the appearance of morphological arrangements uniquely observed for biofilm consortia, and the development of traits allowing for remarkable resilience against antimicrobials (Fux et al., 2004; Hall-Stoodley et al., 2004) or removal of biofilm cells by the predatory actions of protozoans and phagocytes (Bjarnsholt et al., 2005; Matz et al., 2005; Queck et al., 2006), to name a few characteristics commonly observed for biofilms, would support the notion of a biology different from that of planktonically living bacteria. In further support of this, recent reports on transcriptomic and proteomic analysis of biofilm vs. free-living cells of different monospecies systems appear to lay a strong foundation for the existence of genes and proteins uniquely expressed by biofilm cells (Hume et al., 2004; Sauer, 2003). For example, swarming, or coordinated population migration on surfaces, involves a defined differentiation event with distinct temporal stages in certain species (Eberl et al., 1999; Hume et al., 2004; Shapiro, 1988; Shapiro, 1998). Following the observation of apparent program-like responses in this and other specific multicellular surface dependent responses, it is logical to anticipate the involvement of a development program of which biofilm specific genes are expressed in a defined order reflective of a biofilm life cycle. In the following sections we will address whether a unique program, and hence biofilm specific gene expression, underpins the structural and functional features discussed above,

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and whether the interpretations made from contemporary molecular analyses of biofilm genetics and physiology accurately reflect the biology of these communities. he implications of our analysis of currently available data sets on biofilm gene expression and seemingly similar behavioral responses of biofilms or multicelluar consortia of different organisms will also be explored. In addition, we will discuss whether multiple, parallel, pathways of biofilm formation and development of specific traits exists not only for different taxa, but also within a species. he implications of such parallel pathways for biofilm research and biofilm control measures will also be assessed. Finally, we will address strengths and limitations with current biofilm assays and conclude by discussing whether or not there should be a consensus view on how bacterial biofilms form and function. Bioilm program and speciic gene expression Recent years have witnessed detailed mechanistic studies of biofilm formation in a range of species, including E. coli, Vibrio cholera, Staphylococcus aureus, and Serratia marcescens (Davey and O’Toole, 2000; Ghannoum and O’Toole G, 2004; Labbate et al., 2004). For in-depth studies, one of the preferred model organisms is Pseudomonas aeruginosa. here are several good reasons for the many reports on biofilm biology in this organism. First, a number of molecular tools are available for this organism including commercial DNA arrays for the study of genome wide expression. Second, the genus Pseudomonas has simple growth requirements and tolerance to a wide range of temperatures (4°C to 42°C), significantly facilitating experimental efforts. hird, the organism represents a highly diverse and ecologically significant group of bacteria. Fourth, P. aeruginosa is also the most common bacterium found in nosocomial and life-threatening infections of immune compromised patients for which the development of effective treatments are desirable. he bacterium causes a variety of acute and chronic infections (as listed by Van Delden and Iglewski, 1998) and it is particularly known for its involvement in chronic infections of the respiratory pathways including, diffuse panbronchiolitis, bronchiectasia and cystic fibrosis (CF) (Frederiksen et al., 1997; Hoiby, 1974; Kobayashi, 2005; Koch and Hoiby, 1993). Much of what we know about P. aeruginosa biofilm formation derives from studies of monospecies biofilms established in flow cells. A number of specific features are required for the type of biofilm development we see in these flow cells: (1) Attachment to the surface involving specific adhesive proteins, (2) cell to cell binding involving proteins, extra-cellular DNA and polysaccharides in order for the cells to resist the hydrodynamic forces, and (3) cell motility to enable the cells to crawl on the surface (Ghannoum and O’Toole G, 2004). When the biofilm is fully developed, the bacteria are less likely to be displaced by physical processes including shear forces. Moreover, they are better able to withstand competition by invading organisms and they are more resilient to predation challenges (in nature or by host immune responses). In fact, biofilm cells show a remarkable tolerance to a variety of antimicrobial measures as compared to their, planktonic, growing, counterparts (Bjarnsholt et al., 2005; Drenkard and Ausubel, 2002; Fux et al., 2004; Hentzer et al., 2003; Lewis, 2001). Most standard antibiotics will kill only minor parts of the population (Hentzer et al., 2003b), and similar difficulties in eliminating biofilm bacteria have been documented for the host defense systems (Anderson et al., 2003).

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he characteristics detailed above have prompted researchers to suggest the existence of a specific, developmental, program which would operate to ensure proper biofilm formation and differentiation, and account for the tolerance phenotype characteristic for all biofilms. Such proposals are also born out of the formation of characteristic biofilm architectures, including the mushroom-like structures frequently observed in glucose irrigated flow cell biofilms. Recent analysis based on transcriptomics revealed that the bulk of P. aeruginosa biofilm cells even at the early stages express genes in a pattern that is reminiscent of gene expression seen in early stationary phase of planktonic cells (Hentzer et al., 2005). his would in part explain the elevated tolerance to antibiotics since many drugs are relatively ineffective against slow or non-growing stationary cells. In addition, quorum sensing (QS) regulated gene expression also contributes to biofilm tolerance. Davies et al. (1998) demonstrated that a QS deficient lasI mutant of P. aeruginosa formed biofilms that were more susceptible to biocides. Likewise, biofilms formed by a lasR, rhlR double mutant of P. aeruginosa is much more prone to killing by tobramycin and hydrogen peroxide (Bjarnsholt et al., 2005), as well as ciprofloxacin and ceftizidime (our unpublished results) and kanamycin treatment, shear forces, protozoan grazing and neutrophil phagocytosis, than biofilms formed by a wild-type counter part (Allesen-Holm et al., 2006; Barraud et al., 2006; Bjarnsholt et al., 2005; Davies et al., 1998; Hentzer et al., 2002; Hentzer et al., 2003; Matz et al., 2004; Matz et al., 2005; Queck et al., 2006; Rasmussen et al., 2005; Shih and Huang, 2002) suggestive of biofilm specific QS controlled genes. Remarkably, we have observed that the majority of QS regulated genes are expressed even at the early stages of biofilm formation. However, the results generated by different research groups have failed to consistently identify a specific biofilm regulon. More generally, the genes classified by the different research groups as differentially expressed by biofilm cells (as compared to planktonic cultures present at different stages of the growth cycle) do not reveal the presence of a specific program operating in P. aeruginosa during conditions of sessile growth and biofilm formation. his failure to detect common themes in gene expression profiles in different reports on P. aeruginosa biofilms leaves us with the possibility that biofilm development in this organism is mainly governed by adaptive responses. Such a conclusion is strengthened by the finding that biofilm development is associated with the expression of genes required for various physiological life styles (Hentzer et al., 2005). For example, mature P. aeruginosa biofilms express a large number of genes for anaerobic respiration and iron metabolism (Hentzer et al., 2005). We submit that although the experimental conditions would differ in different experiments, the existence of a specific biofilm program would always require a core set of key genes to be expressed, regardless of the experimental conditions. To date, we conclude that transcriptomic studies of P. aeruginosa biofilms have not delivered such an outcome. Clearly, the field of biofilm transcriptomics is at an early stage. Hence, the lack of expression of a consistent set of biofilm genes in P. aeruginosa does not necessarily translate to other biofilm forming bacteria. In fact, development of fruiting bodies in Myxococcus spp. biofilms (which are reminiscent of the mushroom structures and the stationary phase physiological response in P. aeruginosa biofilms) is controlled by a genetically programmed series of events regulated by signal molecules (Sager and Kaiser, 1994). Intriguingly, many of the features assigned to Myxococcus xanthus development, such as the structure of fruiting bodies, role of signal molecules, and type IV pili dependency appear to have strong

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similarities with analogous features and events in P. aeruginosa, and it is possible that further more in depth studies will reveal the presence of a program also in P. aeruginosa (Lux et al., 2004; O’Toole, 2003). While our understanding of gene expression by biofilm cells of other bacterial species has also increased significantly in recent years, the studies of differential gene expression in these organisms do not yet allow for comparative measures of the existence of genes specific for the biofilm mode of life. Hence, the generality of the findings for P. aeruginosa remains to be addressed. With respect to the expression of the entire set of QS genes by early biofilms, it is still likely that the QS system in P. aeruginosa regulates gene expression in a similar fashion in both biofilm and planktonic cells. It may be proposed that QS is not required for biofilm formation per se, and that the QS mediated communication between the cells is a consequence of increased cell density and growth physiology. Again, comparative studies in other bacteria species for which the QS system plays a key role for biofilm development and function, such as S. marcescens and V. cholerae, will provide much needed information on this issue. he failure, in analyses of P. aeruginosa biofilm and planktonic cells, to detect common themes in biofilm gene expression profiles, may also be caused by the severe limitations of current transcriptomic research. hese refer to the poor sensitivity (108–109 cells are required) of transcriptomic analysis and the fact that it only provides us with an average view of the phenomenon of how cells behave in compartmentalized three dimensional biofilms. Application of confocal laser scanning microscopy has shown that in vitro biofilms of several bacterial species often display elaborate multicellular structures separated by channels and void spaces (Allesen-Holm et al., 2006; Lawrence et al., 1991). For example, P. aeruginosa differentiates into two distinct subpopulations, a process that has implications for the antibiotic resistance of biofilms. One of these subpopulations is fully susceptible to the drug (the antimicrobial peptide colistin) whereas the motile, migrating, subpopulation found on the top of the microcolonies in mature biofilms is orders of magnitude more resistant to the peptide ( Janus et al., 2006). he formation of the mushroom-shaped structures in glucose-grown P. aeruginosa biofilms occurs in a sequential process involving the non-motile bacterial subpopulation that forms the stalks and a migrating bacterial subpopulation which subsequently forms the mushroom caps via a process which requires type IV pili (Allesen-Holm et al., 2006). Recent detailed examination of the dispersal cells of P. aeruginosa and S. marcescens biofilms also provides clear evidence for different subpopulations of cells, as well as significant phenotypic variation among dispersal cells (Boles et al., 2004; Drenkard and Ausubel, 2002; Koh et al., 2006; Mai-Prochnow et al., 2006). he data suggest that motile dispersal cells are different from the cells of the microcolony hollow structure remaining after the biofilm dissolution event and that the dispersal cells in turn are stable genotypes, following an increased frequency of mutational events in the mature microcolony. his leads to a broad variation of phenotypes pertinent for the optimal colonization of new surfaces, during a range of conditions. Hence, transcriptomic analysis of the entire biofilm population clearly will not offer information specific to biofilm subpopulations. Additional examples of compartmentalized populations of biofilm cells include the recently emerging evidence that the biosurfactant rhamnolipid is preferentially produced

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by subpopulations present in the stalks of the mushroom structures (Lequette and Greenberg, 2005). Similarly, the generation of the matrix component of extracellular DNA is primarily confined to the stalks of the mushroom-shaped structures in P. aeruginosa biofilms (Allesen-Holm et al., 2006). Intriguingly, both these QS controlled processes involve distinct subpopulations. By first impression this contradicts the view of QS as a method for pooling the activities of the entire population. However, spatial limitations that will contribute to increased local cell concentration taken together with the observation by van Delden and Iglewski (1998) who reported induction of QS regulated genes independent of cell density but in response to nutrient unavailability and gradients, might contribute to local divergence in QS regulated gene expression. Bioilm development In support of the apparent lack of a specific core set of genes characteristic of the biofilm mode of life, it is possible that biofilm formation may proceed through multiple pathways of development. First, different species appear to acquire similar structural and functional endpoints during biofilm formation, including the different stages of microcolony formation, matrix embedded mature biofilms, and tolerance to antimicrobial agent or other stress factors. his has been well documented for P. aeruginosa, E. coli and V. cholerae (Davey and O’Toole, 2000; Ghannoum and O’Toole G, 2004) perhaps the best studied model organisms for elucidating the detailed stages of biofilm formation. However, these species may or may not employ cell surface structures such as pili, flagella, LPS, and exopolymeric substances at specific stages. Second, the use of different pathways for biofilm formation and function occur also within a species. Perhaps the best example is the development of tobramycin resistance in P. aeruginosa. Mah et al. (2003) recently reported on the presence of a specific gene product which inactivates tobramycin by binding to the drug in P. aeruginosa PA14. he locus identified, ndvB, is required for the synthesis of periplasmic glucans. hese periplasmic glucans interact physically with drugs and therefore might prevent antibiotics such as gentamicin, ciprofloxacin, chloramphenicol and ofloxacin from reaching their sites of action by sequestering the antimicrobial agents in the periplasm. However, this gene is not QS regulated neither does the gene product contribute significantly to tobramycin tolerance in the sequenced P. aeruginosa PAO1 strain. Nevertheless, also P. aeruginosa PAO1 tolerates the exposure to high levels of tobramycin without loss of viability during biofilm formation (Bjarnsholt et al., 2005). his illustrates the notion of multiple converging pathways for biofilm development. Importantly, this is pertinent to some of the hallmarks, i.e. tolerance to antimicrobials and stress, of the biofilm mode of life biofilm. Further examples of parallel pathways for development of common biofilm characteristics include different phenotypic characteristics across a range of clinical P. aeruginosa isolates during biofilm formation and dispersal. Although exopolysaccharides encoded by psl, pel and the alg genes (Evans and Linker, 1973; Friedman and Kolter, 2004; Vasseur et al., 2005) have been reported to be critical matrix components where a carbohydraterich matrix component appears to be sufficient for mature biofilm formation, Nemoto et al. (2003) found that mature biofilms formed by four independent clinical P. aeruginosa isolates could be dissolved by DNase treatment, suggesting that extracellular DNA is the primary cell-to-cell interconnecting compound in mature biofilms. Murakawa (1973a,

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1973b) reported on the chemical composition of “slimes” from 20 clinical P. aeruginosa isolates and found that slimes from 18 strains consisted primarily of DNA, while two strains with a mucoid phenotype produced slimes composed primarily of polyuronic acid, an important component of alginate. Also Wozniak et al., (2003) demonstrated that alginate was not the important matrix component of non-mucoid PAO1 and P14. Additionally, a recent study in our laboratories assessed the role of filamentous phage in inducing differentiation, autolysis of a subpopulation of cells in mature microcolonies, and subsequent biofilm dissolution through the release of specialized dispersal cells. his study followed the initial demonstration that filamentous phage Pf4 in P. aeruginosa PAO1, as well as some clinical isolates, is involved in mediating active dispersal from biofilms (Webb et al., 2003). Our recent study showed no phage induction in some isolates and variation in the timing of phage induction, and hence lack of correlation between phage appearance and autolysis and dispersal in other isolates (unpublished results). It is also clear that different filamentous phage, and hence different genes, are induced during biofilm formation in the different clinical P. aeruginosa strains (Mooij et al., 2005). A final example of parallel pathways for development of biofilm traits is the use of cell surface associated adhesins for the same outcome. For example, Staphylococcus epidermidis produces a polysaccharide intercellular adhesin (PIA) which is involved in biofilm accumulation. However, infections of PIA-negative strains are not uncommon, suggesting the existence of PIA-independent biofilm accumulation mechanisms (Rohde et al., 2005). he examples above are in favor of a model in which various environmental conditions select for different parallel pathways by which biofilm formation and dispersal proceed across different species, as well as across strains within a given species. In these cases, the structural and functional endpoints of the biofilms are similar at the level of characteristics used to describe the biofilm life cycle. Furthermore, we suggest that the capacity of forming a multicellular biofilm consortium from planktonic cells developed through parallel evolutionary processes and at different times. While several observations appear to support this view, there are also examples of the use of very similar systems and genes for biofilm formation in different bacteria. For example, within the genus Serratia, different species employ the QS system for biofilm formation, however, the organization and function of these systems vary (Givskov et al., 1996; Givskov et al., 1998; Horng et al., 2002) possibly suggestive of a common ancestral QS system which has been acquired by different species through lateral gene transfer. Implications of different bioilm processes he notion that multiple parallel processes for biofilm formation and dissolution are employed by not only different species of bacteria but also a different strain within a species, has implications for our understanding of a range of bacterial colonization events. For example, an immediate outcome of our ability to detail the various stages in biofilm formation and hence colonization of host surfaces by bacterial pathogens, is the development of novel biofilm control strategies. Novel and effective biofilm control require an in depth understanding of biofilm processes. One biofilm control measure has been suggested to be based on interference of bacterial QS systems (Givskov et al., 1996; Hentzer et al., 2002; Ren et al., 2001). For example, a number of pathogens appear to control much of their

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virulence arsenal by means of QS that relay on extracellular signal molecules (Dunny and Winans, 1999). Most diseases in which QS regulation is expected to play an important role originate from infections caused by opportunistic pathogens. he sessile biofilm mode of growth is recognized in all implant related infections and with every foreign medical equipment inserted into a patient there is a potential risk that a bacterial biofilm will form over time on that artificial surface and that the infection develops into a chronic state (Hentzer et al., 2003). Combined with the involvement of QS in biofilm tolerance to various antimicrobial measures and modulation of both the cellular and adaptive immune systems, it is feasible to propose that QS signaling and interference with important host defense mechanisms afford the bacteria with a mechanism to strategically cause disease. A drug capable of blocking signaling will be expected to attenuate bacterial pathogenicity and cause the biofilm cells of the infectious organism to become susceptible to the host defense and consequently be cleared from the host. Further, an important and appealing aspect is that antibiofilm measures directed against QS systems interfere with non-essential phenotypes and therefore are not expected to create a similar harsh selection pressure for development of resistance as seen with conventional antibiotics. Furthermore, being non-toxic they are not expected to eliminate communities of helpful and beneficial bacteria present in the host. Obviously, QS interference with the aim to remove or prevent biofilms constitutes a powerful tool for biofilm control on all surfaces at which bacteria form and maintain biofilms in a QS dependent fashion. Hence, the application of QS blockers is expected to have wide utility. In the case of P. aeruginosa, the majority of animal model experiments strongly suggests that QS plays an important role in the ability of this bacterium to cause disease but the direct involvement in for example the CF pulmonary infection is still largely based on circumstantial evidence (Hardman et al., 1999; Middleton et al., 2002). Given the possibility that biofilms may be formed by multiple pathways (see above), the lack of clear evidence for a role of QS systems is not surprising. However, it is likely that the environment in the lung tissue constitutes a rather defined selection pressure and hence allows for one or a limited number only of parallel pathways for biofilm formation by the infecting organisms. While such a scenario clearly calls for further studies, the results obtained from the animal models referred to above nevertheless justifies the development of pharmacological relevant QS blockers as an important constituent of future chemotherapies for treatments of a variety of P. aeruginosa based infections. Finally, an increasing number of clinical isolates show multiple resistances to conventional antibiotics which only speak in favor of speeding up the pace in this particular field (Oliver et al., 2000). he use of QS inhibitors to inhibit biofilm formation by Gram positive staphylococci and streptococci, has also been argued as an attractive novel therapeutic tool. However, the use of peptide blockers or inhibitors requires careful investigation in every specific pathogen and type of infection, as a consequence of alternate biofilm processes. Moreover, according to Yarwood (this volume) the signal used can be varied, and the responses to the different signals are complex in staphylococci. For example, peptide signaling via the Agr system may facilitate early biofilm and prevent late biofilm formation. Hence, signal inhibitors may prevent the acute phase of infection but facilitate chronic biofilm based infections.

The Bioilm Mode of Life

Methodologies for bioilm analyses While the last few years have witnessed much progress in our understanding of the means by which bacteria form biofilms on different surfaces and in different habitats, it is also clear that the methodologies employed to study biofilm populations and the rather intricate differentiation processes that appear to underpin biofilm development are unsatisfactory. In fact it has been argued that the progress has been limited in several aspects of biofilm research. his concern speaks to the range of approaches used to study biofilms, i.e. including biofilm assays, such as current batch and flow cell systems, and transcriptomic and proteomic analyses yielding information on average activities rather than assessments of discrete activities reflective of the compartmentalized nature of three dimensional biofilms. Activity of speciic regulatory systems here is clearly a need for the development of several assays for detection of specific traits in biofilms, such as improved assays for the detection of specific regulatory elements. For example, the typical screen for QS blocker activity involves a gene fusion of a QS regulated promoter to a reporter gene. In the presence of exogenously added signal molecules, the reporter gene is activated to express its corresponding phenotype. If in addition to AHL an exogenous blocker is also present, expression of the reporters is reduced or switched-off. Rasmussen et al. (2005) developed an alternative system in which growth of the screeningbacterium indicates the presence of QS blocker activity. Briefly, by genetic manipulation QS regulates expression of a detrimental gene product, however presence of a QS blocker compound blocks expression of the “killer” gene, rescues the host cells and enables growth. Performed as an agar based diffusion assay it enables a quick verification of the inhibitory as well as toxic properties of a test compound or extract. his novel paradigm for bioassays can be applied not only to QS systems and their inhibition, but extends into a multitude of key regulatory systems of direct value for biofilm research. Detection of differential gene expression in bioilms In order to fully capitalize on novel high resolution and specific assays, we require a much enhanced understanding of the differential gene expression that occurs in biofilm communities. However, this is not easily achieved at present. he current “omic” technologies are limited in regard to resolution. While mechanical tools, such as micromanipulators exist to sample individual cells or biofilm compartments, the sample size is not sufficient to support current analyses by transcriptomic or proteomic tools. Ideally, the sensitivity of such tools needs to be improved by at least six orders of magnitude, allowing for analysis of a population of ideally less than 1000 cells. While studies of biofilms have improved significantly using methods for detection of individual cells, by means of for example micromanipulator assisted analysis, GFP tagging, CARD FISH, confocal scanning laser microscopy, and the separation of cells by flow cytometry and cell sorting, current “omic” technologies require a return to sample preparation which includes entire biofilms. Ideally, our experimental toolbox should include the analysis of genome-wide expression by single cells. If the latter scenario can be applied, a range of experiments addressing central questions on gene expression in bacterial biofilms and their individual cells upon interactions

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with hosts as well as other organisms in the environment, including protozoan grazers, may be conducted. Such high resolution methods might enable the detection of key responses in individual cells, reflective of the true interaction between cells and their hosts or predators. Currently, our measurements only depict the average signal or response from all cells in the population which may reflect the majority of cells that do not respond to the presence of the recognition events mediated by the eukaryotic opponent. If analysis of genome-wide expression by single cells can be accomplished we will be in the position to identify relevant gene responders which in turn can be used in the construction of appropriate bioscreens. Such screens would be ideal for the identification and design of interfering chemical compounds. Bioilm assays here are several examples of in vitro biofilm devices, including Robbin’s device (Kharazmi et al., 1999), spinning disk biofilms (Charlton et al., 2001), run through flow cells (Moller et al., 1998) and microtiter dish based batch systems (often referred to as static biofilms). he former three systems produce biofilms irrigated with fresh medium and therefore constitutes biofilms that experience a continuous flow supplemented with fresh nutrients. he latter system generates biofilms that have exhausted important nutrient components at the end of an over night incubation. he key features of this system are that numerous biofilms can be handled at any given time, and does not require time consuming sterilization and setup procedures allowing it to be used as a high through put system for biofilm analysis. Importantly, it provides a basis for a rapid screening procedure of biofilm mutants. For example, a library of random mutants generated by transposon mutagenesis can and has been readily screened for the expression of adhesive properties (Kulasekara et al., 2005), biomass development and biofilm forming capacity (O’Toole and Kolter, 1998; Watnick and Kolter, 1999), as well as extracellular matrix composition (Friedman and Kolter 2004a, 2004b). Essentially, detection of any phenotype that can be processed by a microtiter reader can be handled by this system and mutants can be isolated in pure cultures. Hence, to a limited extent the batch based microtiter analysis can be used to study functional aspects of the biofilm linked to a certain genotype. On the other hand, the system is incompatible with confocal scanning laser microscopy based investigations, the preferred methodology to reveal structural aspects of biofilms. Structural biofilm analyses require the use of irrigated biofilm systems. he irrigated biofilm flow systems are counterparts of chemostats normally employed for studying physiological steady state conditions in planktonic prokaryotes. An important distinction, however, is that the irrigated and flow through cell systems allow for studying the development over time and space of the biofilm. his includes non-destructive studies of temporal and spatial expression of selected genes—as limited by the range of fluorescent reporters available—biofilm structure, and the complete life cycle of biofilm formation and dispersal. Importantly, several of the findings on unique behavioral responses by biofilm cells and subpopulations derive from observations made using irrigated biofilm flow systems e.g. (Davey and O’Toole, 2000). Such information can not be generated using static microtiter batch systems. Moreover, other studies of biofilm characteristics such as toler-

The Bioilm Mode of Life

ance to antimicrobial measures require the use of one of the irrigated biofilm systems for reliable data acquisition (unpublished observations). Genomics he field of genomics is currently experiencing enormous progress in sequencing technology and strategies (Margulies et al., 2005; Shendure et al., 2005; Goldberg et al., 2006) that will open exciting opportunities for biofilm research. In fact, we expect that sequencing of individual cells, populations or natural communities will become an indispensable tool in biofilm research. An important, recent observation in biofilm research has been that a large number of stable, genetic variants are generated during the process of biofilm formation and dispersal; a process that ensures a next generation of cells with better fitness for environmental adaptation (Boles et al., 2004; Mai-Prochnow et al., 2006; see also section “Biofilm program and specific gene expression” above). Applying whole-genome sequencing on entire dispersal populations would help to quickly and effectively identify the mutational processes (such as reduced mismatch repair or increased transposition) that cause the generation of variants. Furthermore, information about the positional bias of mutations in certain genes or genomic regions will point towards selection of specific genotypes in the process from initial attachment to final dispersal. his in turn can enhance our understanding of the evolutionary forces of the biofilm cycle. Recent studies of the genomes of P. aeruginosa CF strains isolated from patients at early and late stages of the disease have provided a strong case for the power of such genomic-based approaches (Smith et al., 2006). Comparing the “early” and the “late” stage derived genomes showed that the strains undergo numerous genetic adaptations during the disease progression. Interestingly, virulence factors such as the QS regulator LasR and the multi-drug efflux pump MexZ are selected against during the chronic phase of the disease. his highlights the notion that advanced CF strains are quite different from “wild-type” P. aeruginosa and might offer new strategies for treatment. he study by Smith et al. (2006) also shows that genome analysis can generate results that are initially counterintuitive to our current understanding of a biological system, while also challenging us to define new hypotheses for the observations made. With the dramatic reduction of sequencing cost expected in the near future, we will see genomics becoming increasingly integrated into other defined, experimental systems of biofilm research. High-throughput DNA sequencing is now well established as a powerful tool to explore and describe the diversity of natural, microbial communities in a field termed environmental genomics or metagenomics (Handelsman, 2004; Riesenfeld et al., 2004; Venter et al., 2004; Tyson et al., 2004; DeLong et al., 2006). In fact the first large-scale environmental sequencing effort was applied to a biofilm community from an acid mine drainage system (Tyson et al., 2004). he relatively low complexity of this microbial system (less then 10 species) allowed for the reconstruction of nearly complete genomes for the populations of the two most-abundant species, Leptospirillum group II and Ferroplasma type II. While the Leptospirillum group II showed very little nucleotide polymorphism, the Ferroplasma type II exhibited extensive homologous recombination resulting in a complex, mosaic genome for its populations. his study illustrates that natural biofilm populations can undergo significant genetic diversification yet the degree of variation might depend on

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the organism itself or the specific evolutionary pressure experienced. It is also tempting to speculate that these observations reflect cell differentiation required for the adaptation to complex, environmental conditions or the biofilm lifestyle itself. Clearly, there is a need to expand these studies to other naturally occurring biofilm systems in order to obtain a better and more refined picture of the genetic diversification and its underlying mechanisms. Should we expect there to be a consensus view on bioilms? Contemporary biofilm research has the potential to reveal novel prokaryotic biology. While there may not be one specific biofilm program, as discussed above, there are certainly a range of observations that suggest the elucidation of prokaryotic biology not previously assigned to planktonic cells or cell cultures. Moreover, we expect that the field of biofilm research will continue to deliver novel information on prokaryotic biology and hence greatly improve our understanding of how biofilm consortia in the environment drive biogeochemical cycles and other key ecological process, as well as unravel disease mechanisms used by bacterial pathogens. Given these predictions, and the fact that several reports on biofilm diversity and heterogeneity have been published in recent years, it is surprising that an almost consensus view on bacterial biofilm biology appear to prevail today. his rather narrow model possibly derives from the focus on mushroom-like structures. It would appear that many bacterial species have the capacity to form such structures in vitro, should appropriate nutrient conditions be provided, and such architectures have become a focal point in biofilm research. In fact, this has lead to a widely held view that biofilm research is the science of mushroom-like structures as studied by CLSM. However, it is not clear that such structures are common in natural habitats. For example, biofilms on living surfaces studied in the authors’ laboratories, e.g. those of the surface of lung tissue, chronically infected wounds and marine sessile organisms, do not display mushroom-like structures. Rather, these biofilms exist as microcolonies (Rao et al., 2006) and in some instances filamentous biofilms (Labbate et al., 2004). Also, other naturally occurring biofilms such as those in the phyllosphere ( Jacques et al., 2005; Morris et al., 1998), the oral cavity (Auschill et al., 2001; Hope et al., 2002) and the rhizosphere (Timmusk et al., 2005; Walker et al., 2004), form aggregates or microcolonies rather than mushrooms. Hence, many current studies appear to be directed at a phenotype that may be of limited relevance in natural habitats. It may be argued that this prevailing consensus view of biofilms has directed the field into a much too narrow research path. Contributing to this situation is the lack of technologies which allow for more detailed analysis of the activity and interactions of single cells, or subpopulations, beyond the current microscopy based descriptions of biofilm structures, as discussed in sections above. What should biofilm research also address and what do we expect will be the focal points in this research area in the near future? Several issues may be considered in this regard. We assume that the driving force for cells to form multicellular arrangements relates to bacterial survival strategies in hostile environments. Hence, studies of the mechanisms by which cells form a multicellular sanctuary or a fortress, as have been the descriptions used for biofilms in which cells are protected from environmental conditions or engage in competitive interactions with other organisms, will be crucial to explore differences between planktonic and sessile prokaryotic lifestyles and identify elements key to the ecological suc-

The Bioilm Mode of Life

cess of biofilms. Further, a much improved understanding of the genes and pathways the cells need to call on for achieving the shift from a planktonic to a true multicellular life style will not only disclose targets for biofilm control but likely also reveal evolutionary traits common to prokaryotes and eukaryotes. With respect to both structural stability and tolerance of biofilms, the exact composition and role of the extracellular matrix in which biofilm cells are embedded should be explored in considerably more detail than is hitherto the case. It is likely that such studies will reveal structural arrangements, akin to the lattice recently reported for Staphylococcus aureus, following the dissolution of mature biofilms (Costerton, J.W. personal communication), or the proposed biofilm backbone characteristic of filamentous phage fibrils embedding P. aeruginosa cells (Webb et al., 2004). hese arrangements are likely to represent novel “nanostructures,” potentially contributing also to rapid advances in the nanosciences. Related to the observation of a network of phage filaments around these cells are current reports on predominantly DNA containing, rather than other macromolecules, extracellular biofilm matrices (Allesen-Holm et al., 2006). Obviously, the implications of DNA as the biofilm matrix are profound as discussed by Pamp et al., this volume. Finally, perhaps the most rapid and ecologically relevant contribution to our understanding of biofilms will derive from precise mechanistic based studies of increasingly complex multispecies biofilms. It is possible that only the inclusion of high resolution genetic based analyses of defined mixed species biofilms, representing a broad range of habitats, will deliver truly meaningful information of prokaryotic multicellular communities. his speaks to their intrinsic biology as well as their ecosystem functions. It may be argued that in depth analyses of relevant mixed species biofilm communities are also required to fully understand a range of bacterially mediated diseases. As discussed in the previous section, current rapid progress in whole bacterial community sequencing and functional metagenomics may make such studies a reality within a very short period of time. We suggest that the apparently prevailing consensus view on bacterial biofilms may not reflect the organization and function of most biofilms, neither in in vitro situations nor in their natural habitats. We also submit that the current consensus view is not helpful in allowing us to embrace the taxonomically and functionally diverse biofilms that constitute the predominant prokaryotic biomass in host and environmental settings. Acknowledgments We wish to express our sincere thanks to our colleagues at the Centre for Marine Biofouling and Bio-Innovation at UNSW and BioCentrum at DTU for their contributions to the topics discussed in this chapter, and Anne Mai-Prochnow and Torsten homas for their help in preparing the chapter. References Allesen-Holm, M., Bundvig Barken, K., Yang, L.K.M., Webb, J.S., Kjelleberg, S., Molin, S., Givskov, M., and Tolker-Nielsen, T. (2006). A characterization of DNA release in Pseudomonas aeruginosa cultures and biofilms. Mol. Microbiol. 59, 1114–1128. Anderson, G.G., Palermo, J.J., Schilling, J.D., Roth, R., Heuser, J., and Hultgren, S.J. (2003). Intracellular bacterial biofilm-like pods in urinary tract infections. Science 301, 105–107.

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Auschill, T.M., Arweiler, N.B., Netuschil, L., Brecx, M., Reich, E., Sculean, A., and Artweiler, N.B. (2001). Spatial distribution of vital and dead microorganisms in dental biofilms. Arch. Oral Biol. 46, 471–476. Barraud, N., Hassett, D.J., Hwang, S., Kjelleberg , S., and Webb, J.S. (2006). Involvement of nitric oxide in biofilm dispersal of Pseudomonas aeruginosa. J. Bacteriol. 188, 7344–7353. Bjarnsholt, T., Jensen, P.O., Burmolle, M., Hentzer, M., Haagensen, J.A., Hougen, H.P., Calum, H., Madsen, K.G., Moser, C., Molin, S., et al. (2005). Pseudomonas aeruginosa tolerance to tobramycin, hydrogen peroxide and polymorphonuclear leukocytes is quorum-sensing dependent. Microbiology 151, 373–383. Boles, B.R., hoendel, M., and Singh, P.K. (2004). Self-generated diversity produces “insurance effects” in biofilm communities. Proc. Natl. Acad. Sci. USA 101, 16630–16635. Caldwell, D.E., Colwell, R.R., Harayama, S., Kjelleberg, S., Givskov, M., Molin, S., Nielsen, A.T., Heydorn, A., Tolker-Nielsen, T., Sternberg, C. (2000). Crystal ball: leading scientists in the field of environmental microbiology consider the technical and conceptual developments that they believe will drive innovative research during the first years of the new millennium. Environ. Microbiol. 2, 3–10. Charlton, T., Givskov, M., de Nys, R., Andersen, J.B., Hentzer, M., Rice, S.A., and Kjelleberg, S. (2001). Genetic and chemical tools for investigating signalling processes in biofilms. In: Methods in Enzymology, R.J. Doyle, ed. (San Diego, CA: Academic Press), pp. 108–128. Davey, M.E., and O’Toole, G.A. (2000). Microbial biofilms: from ecology to molecular genetics. Microbiol. Mol. Biol. Rev. 64, 847–867. Davies, D.G., Parsek, M.R., Pearson, J.P., Iglewski, B.H., Costerton, J.W., and Greenberg, E.P. (1998). he involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280, 295–298. DeLong, E.F., Preston, C.M., Mincer, T., Rich, V., Hallam, S.J., Frigaard, N.U., Martinez, A., Sullivan, M.B., Edwards, R., Brito, B.R., Chisholm, S.W., and Karl, D.M. (2006). Community genomics among stratified microbial assemblages in the ocean’s interior. Science 311, 496–503. Drenkard, E., and Ausubel, F.M. (2002). Pseudomonas biofilm formation and antibiotic resistance are linked to phenotypic variation. Nature 416, 740–743. Dunny, G.M., and Winans, S.C. (1999). Cell–Cell Signaling in Bacteria (Washington DC: ASM Press). Eberl, L., Molin, S., and Givskov, M. (1999). Surface motility of Serratia liquefaciens MG1. J. Bacteriol. 181, 1703–1712. Evans, L.R., and Linker, A. (1973). Production and characterization of the slime polysaccharide of Pseudomonas aeruginosa. J. Bacteriol. 116, 915–924. Frederiksen, B., Koch, C., and Hoiby, N. (1997). Antibiotic treatment of initial colonization with Pseudomonas aeruginosa postpones chronic infection and prevents deterioration of pulmonary function in cystic fibrosis. Pediatr. Pulmonol. 23, 330–335. Friedman, L., and Kolter, R. (2004a). Two genetic loci produce distinct carbohydrate-rich structural components of the Pseudomonas aeruginosa biofilm matrix. J. Bacteriol. 186, 4457–4465. Friedman L., and Kolter R. (2004b). Genes involved in matrix formation in Pseudomonas aeruginosa PA14 biofilms. Mol. Microbiol. 51, 675–90. Fux, C.A., Wilson, S., and Stoodley, P. (2004). Detachment characteristics and oxacillin resistance of Staphylococcus aureus biofilm emboli in an in vitro catheter infection model. J. Bacteriol. 186, 4486– 4491. Ghannoum, M.A., and O’Toole G, A. (2004). Microbial biofilms (Washington, DC: ASM Press). Givskov, M., de Nys, R., Manefield, M., Gram, L., Maximilien, R., Eberl, L., Molin, S., Steinberg, P.D., and Kjelleberg, S. (1996). Eukaryotic interference with homoserine lactone-mediated prokaryotic signalling. J. Bacteriol. 178, 6618–6622. Givskov, M., Ostling, J., Eberl, L., Lindum, P.W., Christensen, A.B., Christiansen, G., Molin, S., and Kjelleberg, S. (1998). Two separate regulatory systems participate in control of swarming motility of Serratia liquefaciens MG1. J. Bacteriol. 180, 742–745. Goldberg, S.M., Johnson, J., Busam, D., Feldblyum, T., Ferriera, S., Friedman, R., Halpern, A., Khouri, H., Kravitz, S.A., Lauro, F.M., Li, K., Rogers, Y.H., Strausberg, R., Sutton, G., Tallon, L., homas, T., Venter, E., Frazier, M., and Venter, J.C. (2006). A Sanger/Pyrosequencing hybrid approach for generation of high quality draft assemblies of marine microbial genomes. Proc. Natl. Acad. Sci. USA 103 (30), 11240–45. Hall-Stoodley, L., Costerton, J.W., and Stoodley, P. (2004). Bacterial biofilms: from the natural environment to infectious diseases. Nat. Rev. Microbiol. 2, 95–108.

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Handelsman, J. (2004). Metagenomics: application of genomics to uncultured microorganisms. Microbiol. Mol. Biol. Rev. 68 (4), 669–85. Hardman, A., Rodgers, H.C., Middleton, B., Cámara, M., Knox, A.J., and Williams, P. (1999). Detection of N-acylhomoserine lactones in CF sputum—a new marker of inflammation in cystic fibrosis. Pediat. Pulmonol. (suppl) 19, 274. Hentzer, M., Eberl, L., and Givskov, M. (2005). Transcriptome Analysis of Pseudomonas aeruginosa: anaerobic respiration and iron limitation. Biofilms 2, 37–61. Hentzer, M., Riedel, K., Rasmussen, T.B., Heydorn, A., Andersen, J.B., Parsek, M.R., Rice, S.A., Eberl, L., Molin, S., Hoiby, N., et al. (2002). Inhibition of quorum sensing in Pseudomonas aeruginosa biofilm bacteria by a halogenated furanone compound. Microbiology 148, 87–102. Hentzer, M., Wu, H., Andersen, J.B., Riedel, K., Rasmussen, T.B., Bagge, N., Kumar, N., Schembri, M.A., Song, Z., Kristoffersen, P., et al. (2003). Attenuation of Pseudomonas aeruginosa virulence by quorum sensing inhibitors. EMBO J. 22, 3803–3815. Hoiby, N. (1974). Epidemiological investigations of the respiratory tract bacteriology in patients with cystic fibrosis. Acta Pathol. Microbiol. Scand. [B] Microbiol. Immunol. 82, 541–550. Hope, C.K., Clements, D., and Wilson, M. (2002). Determining the spatial distribution of viable and nonviable bacteria in hydrated microcosm dental plaques by viability profiling. J. Appl. Microbiol. 93, 448–455. Horng, Y.T., Deng, S.C., Daykin, M., Soo, P.C., Wei, J.R., Luh, K.T., Ho, S.W., Swift, S., Lai, H.C., and Williams, P. (2002). he LuxR family protein SpnR functions as a negative regulator of N-acylhomoserine lactone-dependent quorum sensing in Serratia marcescens. Mol. Microbiol. 45, 1655–1671. Hume, E.B.H., Baveja, J., Muir, B., Schubert, T.L., Kumar, N., Kjelleberg, S., Griesser, H.J., hissen, H., Read, R., Poole-Warren, L.A. (2004). he control of Staphylococcus epidermidis biofilm formation and in vivo infection rates by covalently bound furanones. Biomaterials 25, 5023–5030. Jacques, M.A., Josi, K., Darrasse, A., and Samson, R. (2005). Xanthomonas axonopodis pv. phaseoli var. fuscans is aggregated in stable biofilm population sizes in the phyllosphere of field-grown beans. Appl. Environ. Microbiol. 71, 2008–2015. Janus, A., Haagensen, J., Klausen, M., Ernst, R.K., Miller, S.I., Tolker-Nielsen, T., and Molin S. (2006). Differentiation and distribution of colistin/SDS tolerant cells in Pseudomonas aeruginosa flow-cell biofilms. J. Bacteriol. 189, 28–37. Juretschko, S., Loy, A., Lehner, A., and Wagner, M. (2002). he microbial community composition of a nitrifying-denitrifying activated sludge from an industrial sewage treatment plant analyzed by the full-cycle rRNA approach. Syst. Appl. Microbiol. 25, 84–99. Kharazmi, A., Giwercman, B., and Hoiby, N. (1999). Robbins device in biofilm research. Methods Enzymol. 310, 207–215. Kobayashi, H. (2005). Airway biofilms: implications for pathogenesis and therapy of respiratory tract infections. Treat. Respir. Med. 4, 241–253. Koch, C., and Hoiby, N. (1993). Pathogenesis of cystic fibrosis. Lancet 341, 1065–1069. Koh, K.S., Lam, K.W., Alhede, M., Kjelleberg, S., and Rice, S.A. (2006). Phenotypic diversification and adaptation in S. marcescens MG1 microcolony biofilm morphotype. J. Bacteriol. 189, 119–130. Kulasekara, H.D., Ventre, I., Kulasekara, B.R., Lazdunski, A., Filloux, A., and Lory, S. (2005). A novel two-component system controls the expression of Pseudomonas aeruginosa fimbrial cup genes. Mol. Microbiol. 55, 368–380. Labbate, M., Queck, S.Y., Koh, K.S., Rice, S.A., Givskov, M., and Kjelleberg, S. (2004). Quorum sensingcontrolled biofilm development in Serratia liquefaciens MG1. J. Bacteriol. 186, 692–698. Lawrence, J.R., Korber, D.R., Hoyle, B.D., Costerton, J.W., and Caldwell, D.E. (1991). Optical sectioning of microbial biofilms. J. Bacteriol. 173, 6558–6567. Lequette, Y., and Greenberg E.P. (2005). Timing and localization of rhamnolipid synthesis gene expression in Pseudomonas aeruginosa biofilms. J. Bacteriol. 187, 37–44. Lewis, K. (2001). Riddle of biofilm resistance. Antimicrob. Agents Chemother. 45, 999–1007. Lux, R., Li, Y., Lu, A., and Shi, W. (2004). Detailed three-dimensional analysis of structural features of Myxococcus xanthus fruiting bodies using confocal laser scanning microscopy. Biofilms 1, 293–303. Lyautey, E., Jackson, C.R., Cayrou, J., Rols, J.L., and Garabetian, F. (2005). Bacterial community succession in natural river biofilm assemblages. Microb. Ecol. 50, 589–601. Mah, T.F., Pitts, B., Pellock, B., Walker, G.C., Stewart, P.S., and O’Toole, G.A. (2003). A genetic basis for Pseudomonas aeruginosa biofilm antibiotic resistance. Nature 426, 306–310.

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Mai-Prochnow, A., Webb, J.S., Ferrari, B.C., and Kjelleberg, S. (2006). Ecological advantages of autolysis during the development and dispersal of Pseudoalteromonas tunicata biofilms. Appl. Environ. Microbiol. 72, 5414–5420. Margulies, M., Egholm, M., Altman, W.E., Attiya, S., Bader, J.S., Bemben, L.A., Berka, J., Braverman, M.S., Chen, Y.J., Chen, Z., Dewell, S.B., Du, L., Fierro, J.M., Gomes, X.V., Godwin, B.C., He, W., Helgesen, S., Ho, C.H., Irzyk, G.P., Jando, S.C., Alenquer, M.L., Jarvie, T.P., Jirage, K.B., Kim, J.B., Knight, J.R., Lanza, J.R., Leamon, J.H., Lefkowitz, S.M., Lei, M., Li, J., Lohman, K.L., Lu, H., Makhijani, V.B., McDade, K.E., McKenna, M.P., Myers, E.W., Nickerson, E., Nobile, J.R., Plant, R., Puc, B.P., Ronan, M.T., Roth, G.T., Sarkis, G.J., Simons, J.F., Simpson, J.W., Srinivasan, M., Tartaro, K.R., Tomasz, A., Vogt, K.A., Volkmer, G.A., Wang, S.H., Wang, Y., Weiner, M.P., Yu, P., Begley, R.F., and Rothberg, J.M. (2005). Genome sequencing in microfabricated high-density picolitre reactors. Nature 437 (7057), 376–80. Matz, C., Deines, P., Boenigk, J., Arndt, H., Eberl, L., Kjelleberg, S., and Jurgens, K. (2004). Impact of violacein-producing bacteria on survival and feeding of bacterivorous nanoflagellates. Appl. Environ. Microbiol. 70, 1593–1599. Matz, C., McDougald, D., Moreno, A.M., Yung, P.Y., Yildiz, F.H., and Kjelleberg, S. (2005). Biofilm formation and phenotypic variation enhance predation-driven persistence of Vibrio cholerae. Proc. Natl. Acad. Sci. USA 102, 16819–16824. Middleton, B., Rodgers, H.C., Cámara, M., Knox, A.J., Williams, P., and Hardman, A. (2002). Direct detection of N-acylhomoserine lactones in cystic fibrosis sputum. FEMS Microbiol. Lett. 207, 1–7. Moller, S., Sternberg, C., Andersen, J.B., Christensen, B.B., Ramos, J.L., Givskov, M., and Molin, S. (1998). In situ gene expression in mixed-culture biofilms: evidence of metabolic interactions between community members. Appl. Environ. Microbiol. 64, 721–732. Mooij, M.J., Llamos, M., Vandenbroucke-Grauls, C., Savelkoul, P.H., and Bitter, W. (2005). Bacteriophages and small-colony variants in clinical isolates of Pseudomonas aeruginosa. Pseudomonas 10th International Congress, August 2005, Marseille, France. Morris, C.E., Monier, J.M., and Jacques, M.A. (1998). A technique To quantify the population size and composition of the biofilm component in communities of bacteria in the phyllosphere. Appl. Environ. Microbiol. 64, 4789–4795. Murakawa, T. (1973a). Slime production by Pseudomonas aeruginosa. III. Purification of slime and its physicochemical properties. Jpn. J. Microbiol. 17, 273–281. Murakawa, T. (1973b). Slime production by Pseudomonas aeruginosa. IV. Chemical analysis of two varieties of slime produced by Pseudomonas aeruginosa. Jpn. J. Microbiol. 17, 513–520. Nemoto, K., Hirota, K., Murakami, K., Taniguti, K., Murata, H., Viducic, D., and Miyake, Y. (2003). Effect of Varidase (streptodornase) on biofilm formed by Pseudomonas aeruginosa. Chemotherapy 49, 121–125. O’Toole, G., Kaplan, H.B., and Kolter, R. (2000). Biofilm formation as microbial development. Annu. Rev. Microbiol. 54, 49–79. O’Toole, G.A. (2003). To build a biofilm. J. Bacteriol. 185, 2687–2689. O’Toole, G.A., and Kolter, R. (1998). Initiation of biofilm formation in Pseudomonas fluorescens WCS365 proceeds via multiple, convergent signalling pathways: a genetic analysis. Mol. Microbiol. 28, 449– 461. Oliver, A., Canton, R., Campo, P., Baquero, F., and Blazquez, J. (2000). High frequency of hypermutable Pseudomonas aeruginosa in cystic fibrosis lung infection. Science 288, 1251–1254. Queck, S.Y., Weitere, M., Moreno, A.M., Rice, S.A., and Kjelleberg, S. (2006). he role of quorum sensing mediated developmental traits in the resistance of Serratia marcescens biofilms against protozoan grazing. Environ. Microbiol. 8 1017–1025. Rao, D., Webb, J.S., and Kjelleberg, S. (2006). Microbial colonisation and competition on the marine alga Ulva australis. Appl. Environ. Microbiol. 72, 5547–5555 Rasmussen, T.B., Bjarnsholt, T., Skindersoe, M.E., Hentzer, M., Kristoffersen, P., Kote, M., Nielsen, J., Eberl, L., and Givskov, M. (2005). Screening for quorum-sensing inhibitors (QSI) by use of a novel genetic system, the QSI selector. J. Bacteriol. 187, 1799–1814. Ren, D., Sims, J.J., and Wood, T.K. (2001). Inhibition of biofilm formation and swarming of Escherichia coli by (5Z)-4-bromo-5-(bromomethylene)-3-butyl-2 (5H)-furanone. Environ. Microbiol. 3, 731–736. Riesenfeld, C., Schloss, P., and Handelsman, J. (2004). Metagenomics: genomic analysis of microbial communities. Annu. Rev. Genet. 38, 525–52.

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Rohde, H., Burdelski, C., Bartscht, K., Hussain, M., Buck, F., Horstkotte, M.A., Knobloch, J.K., Heilmann, C., Herrmann, M., and Mack, D. (2005). Induction of Staphylococcus epidermidis biofilm formation via proteolytic processing of the accumulation-associated protein by staphylococcal and host proteases. Mol. Microbiol. 55, 1883–1895. Sager, B., and Kaiser, D. (1994). Intercellular C-signaling and the traveling waves of Myxococcus. Genes Dev. 8, 2793–2804. Sauer, K. (2003). he genomics and proteomics of biofilm formation. Genome Biol. 4, 219. Sauer, K., Camper, A.K., Ehrlich, G.D., Costerton, J.W., and Davies, D.G. (2002). Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J. Bacteriol. 184, 1140–1154. Shapiro, J.A. (1988). Bacteria as multicellular organisms. Sci. Am. 256, 82–89. Shapiro, J.A. (1998). hinking about bacterial populations as multicellular organisms. Annu. Rev. Microbiol. 52, 81–104. Shendure, J., Porreca, G., Reppas, N.B., X., L., McCutcheson, J.P., Rosenbaum, A.M., Wang, M.D., Zhang, K., Mitra, R.D., and Church, G.M. (2005). Accurate multiplex colony sequencing of an evolved bacterial genome. Science 309, 1728–32. Shih, P.C., and Huang, C.T. (2002). Effects of quorum-sensing deficiency on Pseudomonas aeruginosa biofilm formation and antibiotic resistance. J. Antimicrob. Chemother. 49, 309–314. Smith, E.E., Buckley, D.G., Wu, Z., Saenphimmachak, C., Hoffman, L.R., D’Argenio, D.A., Miller, S.I., Ramsey, B.W., Speert, D.P., Moskowitz, S.M., Burns, J.L., Kaul, R., and Olson, M.V. (2006). Genetic adaptation by Pseudomonas aeruginosa to the airways of cystic fibrosis patients. Proc. Natl. Acad. Sci. USA 103 (22), 8487–92. Taylor, M.W., Schupp, P.J., Dahllof, I., Kjelleberg, S., and Steinberg, P.D. (2004). Host specificity in marine sponge-associated bacteria, and potential implications for marine microbial diversity. Environ. Microbiol. 6, 121–130. Timmusk, S., Grantcharova, N., and Wagner, E.G. (2005). Paenibacillus polymyxa invades plant roots and forms biofilms. Appl. Environ. Microbiol. 71, 7292–7300. Tujula, N.A., Holmstrom, C., Mussmann, M., Amann, R., Kjelleberg, S., and Crocetti, G.R. (2006). A CARD-FISH protocol for the identification and enumeration of epiphytic bacteria on marine algae. J. Microbiol. Methods 65, 604–607. Tyson, G., Chapman, J.P.H., Allen, E., Ram, R., Richardson, P., Solovyev, V., Rubin, E., Rokhsar, D., and Banfield, J. (2004). Community structure and metabolism through reconstruction of microbial genomes from the environment. Nature 428 (6978), 37–43. van Delden, C., and Iglewski, B.H. (1998). Cell-to-cell signaling and Pseudomonas aeruginosa infections. Emerg. Infect. Dis. 4, 551–560. Vasseur, P., Vallet-Gely, I., Soscia, C., Genin, S., and Filloux, A. (2005). he pel genes of the Pseudomonas aeruginosa PAK strain are involved at early and late stages of biofilm formation. Microbiology 151, 985–997. Venter, J., Remington, K., Heidelberg, J.F., Halpern, A.L., Rusch, D., Eisen, J.A., Wu, D., Paulsen, I., Nelson, K.E., Nelson, W., Fouts, D.E., Levy, S., Knap, A.H., Lomas, M.W., Nealson, K., White, O., Peterson, J., Hoffman, J., Parsons, R., Baden-Tillson, H., Pfannkoch, C., Rogers, Y.H., and Smith, H.O. (2004). Environmental genome shotgun sequencing of the Sargasso Sea. Science 304, 66–74. Walker, T.S., Bais, H.P., Deziel, E., Schweizer, H.P., Rahme, L.G., Fall, R., and Vivanco, J.M. (2004). Pseudomonas aeruginosa-plant root interactions. Pathogenicity, biofilm formation, and root exudation. Plant Physiol. 134, 320–331. Watnick, P.I., and Kolter, R. (1999). Steps in the development of a Vibrio cholerae El Tor biofilm. Mol. Microbiol. 34, 586–595. Webb, J.S., Givskov, M., and Kjelleberg, S. (2003). Bacterial biofilms: prokaryotic adventures in multicellularity. Curr. Opin. Microbiol. 6, 578–585. Webb, J.S., Lau, M., and Kjelleberg, S. (2004). Bacteriophage and phenotypic variation in Pseudomonas aeruginosa biofilm development. J. Bacteriol. 186, 8066–8073. Wozniak, D.J., Wyckoff, T.J., Starkey, M., Keyser, R., Azadi, P., O’Toole, G.A., and Parsek, M.R. (2003). Alginate is not a significant component of the extracellular polysaccharide matrix of PA14 and PAO1 Pseudomonas aeruginosa biofilms. Proc. Natl. Acad. Sci. USA 100, 7907–7912.

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Do Not Fear Commitment: The Initial Transition to a Surface Lifestyle by Pseudomonads

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Daniel P. MacEachran and George A. O’Toole

Abstract his chapter aims to describe the early stages of biofilm formation, particularly on abiotic surfaces, by focusing on Pseudomonas aeruginosa. Specifically, we will dissect the early steps in the establishment of a multicellular community: (i) translocation to the surface from a free-swimming planktonic lifestyle, (ii) initial or reversible attachment, and finally (iii) irreversible attachment. We will also compare the mechanisms used by P. aeruginosa to its related fluorescent pseudomonad cousins, Pseudomonas fluorescens and Pseudomonas putida. We argue that, for pseudomonads, irreversible attachment is the first committed step in the transition to a biofilm lifestyle. Introduction It has been understood for some time that the majority of bacteria exist in nature attached to a substratum (see Eberl et al. and Hogan, this volume), and more recently it has been suggested that a subset of bacterial infections are the result of surface attached multicellular bacterial communities (Costerton et al., 1999; Vinh and Embil, 2005). One can readily envision the myriad of advantages afforded to a bacterium attached to surface. For example, nutrients are typically most abundant within relative proximity to a substratum (Paul and Clark, 1989). Furthermore, bacteria growing in a biofilm gain protection from protozoan and bacterial predator grazing, phage infection and a variety of other environmental insults (Matz et al., 2004; Patel, 2005; Picioreanu et al., 2000; Sutherland et al., 2004; Matz, this volume). Relatively recently, molecular and genetic techniques have been applied to the study of this phenomenon. Several model organisms for the study of biofilm formation have emerged and among the most studied microbes is the ubiquitous Gram-negative bacilli Pseudomonas aeruginosa. P. aeruginosa has been previously noted for its ability to thrive in a vast array of environments ranging from the rhizosphere to medical facilities (Bloemberg and Lugtenberg, 2001; Rahme et al., 1995; Stoodley et al., 2005). his ubiquitous nature is partially due to the relatively large coding capacity of the P. aeruginosa genome allowing for both metabolic plasticity as well as the ability to rapidly sense the environment and alter gene expression accordingly, which is attributed to this microbe’s myriad of two-component regulatory systems (Stover et al., 2000). here is growing evidence that the ability to attach to surfaces and form complex multicellular communities is also a major factor contributing

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to the metabolically diverse nature of P. aeruginosa. It is likely that these features are interrelated, thereby increasing the fitness of this bacterium. he growing attention regarding P. aeruginosa and its ability to form biofilms has begun to reveal that the formation of these communities follows a fairly defined series of steps and may be analogous the developmental pathways previously reserved for more complex organisms (O’Toole et al., 2000). A look back he phenomenon we currently refer to as biofilm formation has been described in the literature for some time. his should come as little surprise since biofilms are observable to the naked eye. Indeed, when Anthony van Leeuwenhoek first took scrapings from between his teeth and observed them under his crude microscope he observed small “animalcules” embedded in an organic substance (van Leeuwenhoek, 1683). Today we know that these scrapings were oral communities rich in a variety of bacteria, including Streptococcus spp., embedded in a polymeric matrix secreted by the microbes (Kolenbrander et al., 1999; Kolenbrander et al., this volume). With the improvement in microscopy over the next several centuries came further study of bacteria and their ability to attach to surfaces. Work carried out by several early pioneers of biofilm formation, including Henrici, Zobell, Anderson, Waksman and many others, demonstrated through a variety of techniques that bacteria from a range of environmental niches were capable of attaching to a surface and proliferating (Henrici, 1933; Waksman and Vartiovaara, 1938; Zobell and Allen, 1935; Zobell and Anderson, 1936; Zobell, 1937, 1943). Work by Zobell in 1943 demonstrated not only that a large number of bacteria in seawater were capable of attaching to a surface and preferred to do so, but that this attachment seemed to consist of two distinct stages (Zobell, 1943). He noted that when glass slides were incubated with the bacteria for a short period of time the attached bacteria could easily be washed off. However, if the slides were left in the seawater for somewhat more extended periods of time the bacteria were much more difficult to remove through simple washing. he concept that bacterial attachment to a surface consists of two stages, one in which the bacterium is weakly attached to the surface followed by a more secure form of attachment, is further supported in work by Marshall and colleagues in 1971 (Marshall et al., 1971). heir work using both Achromobacter and Pseudomonas demonstrated that both bacteria undergo an immediate reversible attachment followed by a time-dependent irreversible attachment. Furthermore, their studies showed that the Pseudomonas species used in these studies attached via the pole of the cell and could rotate freely about their pole during the reversible stage of attachment (Figure 3.1). his rotation ceased once the bacterium had made the switch to irreversible attachment. Work by Lawrence and colleagues demonstrated that there were two distinct stages of early adhesion in Pseudomonas fluorescens (Lawrence, 1987). Utilizing several different microscopic methods they observed that the bacteria attached initially by one of their poles wherein the bacteria were capable of rotating freely around this pole. his polar attachment was observed to be reversible as some bacteria were seen to re-enter the planktonic phase. Following reversible attachment, the bacteria switched to a more secure interaction with the surface by attaching along their longitudinal axis, followed by microcolony formation

Pseudomonad Transition to a Surface Lifestyle

Figure 3.1 Initial attachment to a substratum. Pseudomonads translocate to the surface and adhere via a pole. Which pole, and which part of the cell directly contacts the surface, is unknown. Several possibilities are illustrated here. Bacteria subsequently adhere via the long axis of the cell in a process known as irreversible attachment. Irreversible attachment is the irst committed step in the transition to a bioilm lifestyle.

and subsequent biofilm maturation. From these early findings the models of adhesion and biofilm formation arose: bacteria swim to a surface whereupon they initiate attachment through polar, reversible attachment, followed by irreversible attachment, which we will argue below, is the first committed stage of biofilm formation. Translocation to the surface One can readily imagine that before a bacterium can attach to a substratum, it must first locate the surface and be capable of translocating to that surface. Approaching the surface may be more difficult than it appears on its face because in order to attach, a bacterium must not only move towards the substratum, but also be capable of overcoming the repulsive forces found at this liquid-surface interface. Both the substratum and bacterial surfaces are often of a similar charge, thereby resulting in electrostatic repulsion between the microbe and the surface to which it may attach. Hydrophobic interactions can also have an impact on initial attachment as these forces can act either as a repulsive or attractive force thereby altering rates of attachment (An and Friedman, 1998; Bos et al., 1999). Bacteria can overcome repulsive forces as they approach the surface in several ways. he bacterium may move passively towards the surface, subject to the will of gravity, Brownian motion or the simple flow of the liquid environment within which it finds itself. Alternatively, the bacterium can actively translocate to the surface using a variety of swimming appendages. As discussed in greater detail below, there are a large number of flagellated bacteria that form robust biofilms, including P. aeruginosa. However, many bacteria lacking any form of swimming organelle are quite capable of forming robust biofilms, as is the case for Staphylococcus aureus (Caiazza and O’Toole, 2003; Cramton et al., 1999;

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Marrie and Costerton, 1984), suggesting that for these organisms, active translocation to the surface during biofilm formation is dispensable. P. aeruginosa expresses a single peritrichous flagellum and exhibits robust swimming motility under most conditions. Several studies have implicated a role for swimming motility in biofilm formation. One of the earliest studies of the genetics of biofilm formation found that P. aeruginosa mutants lacking a functional flagellum were incapable of forming a biofilm in minimal medium supplemented with glucose and amino acids under static conditions (O’Toole and Kolter, 1998b). In these studies, pseudomonads were inoculated into wells of a 96-well microtiter dish containing minimal glucose plus amino acids medium and incubated under static conditions for up to 24 hours. Following incubation, the individual wells were washed with water and stained with crystal violet, followed by subsequent solubilization of the stain with ethanol to quantify the attachment. Mutant strains lacking a flagellum demonstrated a severe defect in attachment in this system. Microscopic studies confirmed the staining data (O’Toole and Kolter, 1998b). Work by Ramsey and Whiteley (2004) further supported this notion, only these investigators demonstrated a role for the flagellum in a somewhat more dynamic environment that also incorporated shear stress. In this study, the author’s biofilm assay was performed as described above, utilizing a similar minimal medium containing glucose and amino acids, but they also added glass beads to the wells. Rather than incubating the assay plates under static conditions, the microtiter dishes were shaken at high speed, thus the addition of the glass beads allowed for both aeration of the culture and provided a source of shear stress. his model further emphasized the role for flagella in biofilm formation by demonstrating that the cell appendage is not only important in a static environment but also in environments subjected to shear stress. Several genetic studies of soil pseudomonads also demonstrated that a functional flagellum is required for attachment and subsequent biofilm formation on both biotic and abiotic surfaces (De Weger et al., 1987; DeFlaun et al., 1994). How might the translocation of bacteria to a substratum to initiate reversible attachment be regulated? his precise question has not been directly addressed, however because the regulation of flagellar assembly and function has been studied in detail, and is quite complex (Soutourina and Bertin, 2003), this entire regulatory cascade could be included in a discussion of the regulation of biofilm formation. One confounding issue regarding interpreting the role of the flagellum in biofilm formation relates to the fact that this structure has two potential roles: as a propeller used to move the cells through a liquid environment and an adhesive appendage (Arora et al., 1998; Landry et al., 2006; Lillehoj et al., 2002). Recent work has allowed us to begin to address each of these roles individually. Using a bioinformatic approach, Toutain et al. (2005) and Doyle et al. (2004) identified two pairs of genes with sequence similarity to motAB of E. coli in the P. aeruginosa genome. MotAB comprises the stator, the stationary component of the flagellar motor. Both groups showed that in regards to swimming motility, the two sets of motor genes, motAB and motCD, were redundant—mutations in either stator alone revealed no measurable effect on swimming (Doyle et al., 2004; Toutain et al., 2005). Interestingly, in a more recent study, Toutain and O’Toole (unpublished) showed that mutations affecting either set of stators resulted in a significant reduction in biofilm formation despite the absence of any discernable defect in swimming motility. Disruption of both

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sets of stators further exacerbated this biofilm formation defect under static conditions, thus demonstrating that not only is the presence of a flagellum required, but it must also be functional to initiate biofilm formation under these conditions (Toutain and O’Toole, unpublished). Taken together, the lines of evidence above certainly support the notion that for a flagellated Gram-negative bacterium such as P. aeruginosa, actively translocating to the surface facilitates attachment to a greater degree than relying on a passive mechanism, and that flagellar-mediated motility plays a key role in early biofilm formation. Interestingly, there is very little evidence suggesting a role for chemotaxis in biofilm formation. Most studies have focused primarily on the classic che mediated chemotactic pathways, leaving room for less or uncharacterized chemotactic mechanisms. Work by Pratt and Kolter (1998) demonstrated that in E. coli disruptions in the che chemotaxis system did not result in a loss of biofilm formation in a static assay. Similarly, large genetic screens for biofilm mutants in P. aeruginosa and P. fluorescens did not yield any strains with mutations in the chemotaxis gene cluster that is most similar to E. coli (O’Toole and Kolter, 1998a, 1998b). here are several lines of apparently contradictory data concerning the role of flagellar motility in biofilm formation. Microscopic studies by Klausen et al. (2003) demonstrated that under flowing conditions, in medium containing citrate as the sole carbon source, P. aeruginosa flagellar mutants were quite capable of forming biofilms albeit with gross morphological differences from those formed by the wild-type bacterium. Similar results were observed for strains lacking either or both flagellar stators (Toutain and O’Toole, unpublished data). hese results seem at odds with those outlined above, however several differences should be noted. First, the biofilm formed by the wild-type bacterium when grown on citrate is significantly different morphologically from that formed in medium supplemented with glucose, and furthermore, biofilms formed under static conditions are morphologically distinct from those formed under flowing conditions. Furthermore, there is a significant difference in the time frame of these two assays—typically the static biofilm assays are incubated for 8 hours while those experiments using the flow conditions were incubated for 24 hours or more. Finally, it is also important to consider that the flagellum also plays a role in surface-associated swarming motility. How flagellar-mediated swarming contributes to biofilm formation is still an open question. Taken together, we believe that the data presented here can be interpreted to mean that there is a more active role for flagellar-mediated motility in the formation of a biofilm than simply getting the bacterial cell to the surface. Reversible attachment Once the bacterium has overcome the various repulsive forces found at the surface/liquid interface, it must attach to the substratum. As is quite evident from several early studies, this attachment consists of two distinct steps: an initial and reversible attachment defined by a relatively unstable, polar adhesion event and the ability of the bacterium to rapidly return to the liquid environment, followed by a more stable irreversible attachment via the long axis of the cell (Lawrence et al., 1987; Marshall et al., 1971; Zobell, 1943). While this phenomenon has been noted in the literature for several decades, in depth investigation of the molecular mechanism of reversible and irreversible attachment has been lacking.

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Recent microscopic analysis of reversible attachment has suggested that the bacterium is indeed attached in a polar manner and may be attached via its flagellum (Caiazza and O’Toole, 2004; Sauer et al., 2002) as reversibly attached bacteria appear to rotate about their polar axis while attached to the substratum. Direct proof that the flagellum is indeed responsible for this initial attachment has been elusive thus far as strains lacking these structures mutants are also impacted in their ability to translocate to the surface as previously mentioned. It has been postulated by a variety of groups that during the initial attachment stage the bacterium may be sensing the local environment of the surface prior to committing to the surface (Caiazza and O’Toole, 2004; Hinsa et al., 2003; Lawrence et al., 1987; Marshall et al., 1971; Zobell, 1943). his is a fairly sensical assumption as this affords the bacterium the ability to either return to the liquid environment, in the case of an unsuitable environment, or further commit if the surface and the local environs are indeed suitable to promote initial biofilm formation and maturation. If the transition from reversible to irreversible attachment is actively controlled, there must presumably be set cues leading to commitment, however, few if any of these cues have been identified but several candidates have been proposed (Marshall et al., 1971). hese attachment signals may include sensing carbon availability, carbon nitrogen ratios, presence of other bacteria or the surface itself. Work by Otto and Silhavey (2002) demonstrated that the cpx signaling pathway was not only required for surface adhesion but also played a key role in signaling upon adhesion of E. coli to a surface under certain conditions. he cpx system had previously been shown to play a role in sensing perturbations and stresses within the cell envelope (Raivio and Silhavy, 1997). hese experiments utilized a lacZ reporter system that served as a marker for induction of the cpx system. Upon adhesion to the surface a cpx-lacZ reporter system was induced and reached maximal expression 1 hour following attachment, while planktonic bacteria did not show this same level of induction. he model put forth would suggest that upon contact with the surface there is some form of perturbation of the cell envelope due to contact with the surface, the cpx system interprets this contact and relays the information resulting in increased expression of adhesins and subsequent biofilm formation (Raivio, 2005). Little is still known concerning the signals that mediate the decision to return to the planktonic stage or further commit to the surface. It seems likely that the bacterium is capable of receiving and acting upon various environmental cues, whether they be metabolic as would be the case for nutrient conditions, or mechanical such as sensing the surface. How do pseudomonads mediate polar attachment to the surface? While a fundamentally important question, very little is known regarding this point. As described above, flagellar motility is thought to be important in pseudomonads for their translocation to the substratum. It has been postulated that the flagellar filament also serves as an adhesin. Furthermore, several lines of evidence demonstrate that very defined portions of the flagellum, and perhaps not the organelle in its entirety, play a key role in attachment of P. aeruginosa to various surfaces. Arora et al. (1998) demonstrated in P. aeruginosa that disruption of fliD, which codes for the flagellar cap protein at the tip of the flagellum, resulted in an inability to attach to mucin, while strains incapable of producing flagellin, the structural subunit of flagella could still attach to mucin. Further work by Landry et al. (2006) demonstrated that under flow conditions fliD mutants were capable of attaching to a glass surface

Pseudomonad Transition to a Surface Lifestyle

at a rate comparable to the wild-type bacteria. However, if the surface was coated with mucin, the fliD mutant was significantly attenuated in attachment while the flgK mutant, which lacks the flagellar filament, showed a dramatic decrease in attachment under both conditions when compared to the wild-type bacteria (Landry et al., 2006). hese data suggest that in addition to its role in motility, the flagellum, through FliD, mediates contact with some types of surfaces. However, the observation that flagellar mutants do indeed eventually attach and form a biofilm suggests that bacteria have other adhesive molecules on their surface to promote cell-surface interactions. One obvious mechanism for promoting reversible attachment would be an outer membrane protein (OMP) that functions primarily as an adhesin, or alternatively, an OMP that may be a part of, for example, a secretion system with a secondary role in cell-surface attachment. Several OMP have been implicated as adhesins (Hinsa and O’Toole, 2004), including OprF in P. aeruginosa, which has been shown to play a role in anaerobic biofilm formation and adhesion to epithelial cells (Azghani et al., 2002; Yoon et al., 2002). Another potential adhesin is the LecB lectin, which is localized to the OM of P. aeruginosa (Tielker et al., 2005). Mutating this fucose-binding protein results in a severe biofilm formation defect, however it is not clear where in the biofilm formation pathway this block occurs (Tielker et al., 2005). he lack of any single, clearly defined adhesin required for reversible attachment might indicate multiple, redundant adhesins for attachment to a wide range of surfaces. Alternatively, there may be multiple adhesins whose primary function is to bind to one particular surface but with enough cross-specificity to make it difficult to identify these proteins via typical genetic approaches. he identification of adhesins required for polar interaction of bacteria during biofilm formation is an area that warrants further study. It is also possible that LPS plays a direct or indirect role in reversible attachment. LPS structure appears to play a role in adhesion of pseudomonads, as mutants lacking O-polysaccharide are decreased in their ability to attach to a variety of surfaces. Work by DeFlaun and colleagues on the soil bacterium Burkholderia cepacia G4, a close relative of P. aeruginosa, identified a biofilm-defective mutant utilizing a sand column assay (DeFlaun et al., 1999). It was found that this mutant strain, ENV435, demonstrated significant differences in the lipopolysaccharide O-antigen compared to the wild type suggesting that, indeed, lipopolysaccharide composition plays a role in attachment to abiotic surfaces (DeFlaun et al., 1999). A role for LPS O-antigen in attachment was not entirely novel amongst pseudomonads as work by Dekkers and colleagues in 1998 had demonstrated that mutants of P. fluorescens WCS365 defective in O-antigen synthesis were defective for colonization of roots (Dekkers et al., 1998). Because of the pleiotropic effects changes in LPS are likely to cause, it is not currently possible to develop any firm mechanistic models to explain why altering the O-antigen results in a biofilm defect. Irreversible attachment: commitment to the surface As stated previously there are a number of different studies that have described the phenomenon of reversible and irreversible attachment in a variety of different bacterial systems. Teleologically, one might postulate that bacteria would develop the ability to initially attach to a surface, sense the suitability of the local environment, and either commit to the surface via irreversible attachment or to re-enter the planktonic stage and seek out a more suitable

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niche. his choice to “stay or go” is best typified by the observable change in orientation of the bacterium to the surface, from a polarly attached cell capable of rotating freely about its axis to a longitudinally attached cell that is firmly bound to the surface. Work by Caiazza and O’Toole (2004) provided some of the first insight into the molecular mechanisms underlying irreversible attachment to a surface. hese investigators demonstrated that a cytoplasmic protein of unknown function in P. aeruginosa, designated SadB, was required for the transition from reversible to irreversible attachment. Furthermore, the sadB mutant was severely defective for biofilm formation under static conditions irrespective of the carbon source present. his study further demonstrated that the biofilm formation defect of a sadB mutant was also observable under flowing conditions. Upon close inspection of micrographs and time-lapse images of sadB mutants in a flow cell, no obvious defect in reversible attachment was observed, a majority of the sadB mutant bacteria were bound by their poles and rotated freely about the attached axis of the cell. In contrast, most wild-type bacteria under similar conditions were irreversibly attached via their long axis (Caiazza and O’Toole, 2004). How SadB mediates irreversible attachment is still not known. Interestingly, the gene coding for SadB is conserved amongst the pseudomonads begging the question of whether this system is integral for biofilm formation in all members of this genus. Levels of SadB protein, as assayed by Western blot, are upregulated in strains carrying mutations in rpoN, which codes for an alternative sigma factor that regulates a large number of genes including those required for flagellar-mediated motility, and fleR, which is involved in regulation of flagellar biosynthesis (Caiazza and O’Toole, 2004). hese data suggest that the regulation of irreversible attachment is co-regulated with flagellar-mediated motility and thus reversible attachment events. Studies in P. fluorescens have identified another series of components required for the switch from reversible to irreversible attachment that utilizes a large adhesin, termed LapA, which weakly associates with the bacterial cell envelope, and a predicted ABC transporter encoded by the lapEBC genes (Hinsa et al., 2003). Mutations in lapBCE are still capable of making LapA, but this protein is localized only to the cytoplasm and is not found outside of the cell. hese data suggest that the LapBCE system is required for the secretion of the LapA adhesion. Hinsa (2003) identified the Lap system through a genetic screen to identify genes required for biofilm formation in a static assay. Mutations in the Lap system resulted in bacteria, that when observed microscopically, were incapable of progressing past the reversible stage of adhesion. Adjacent to the lapABCE cluster is lapD, which is shown to also be required for irreversible attachment in P. fluorescens WCS363 and P. putida (Gjermansen et al., 2005; Hinsa and O’Toole, 2006). LapD is an inner membrane protein, and strains lacking LapD have a significant decrease in the amount of LapA found associated with the cell surface (Hinsa and O’Toole, 2006). herefore, it is possible that LapD participates in the secretion of LapA or its association with the outer membrane. Interestingly, LapA is not found in P. aeruginosa and a mutation in the ABC transporter of this microbe with the greatest sequence similarity to LapBCE has no biofilm defect on abiotic surfaces (Hinsa and

Pseudomonad Transition to a Surface Lifestyle

O’Toole, unpublished data), suggesting that these related organisms, while both capable of irreversible attachment, commit to the surface via distinct mechanisms. Espinosa-Urgel and colleagues identified several genes critical for attachment of P. putida to corn seeds, and furthermore several of the mutants from this screen were deficient in attaching to abiotic surfaces as well. Of the eight genes identified, four of them encode proteins predicted to localize to the cell envelope suggesting that one or more of them may act as an adhesin during attachment (Espinosa-Urgel et al., 2000). Furthermore, one of the proteins identified has sequence similarity to LapA, suggesting that this adhesin is also involved in irreversible attachment (Hinsa et al., 2003). Recent work by Overhage and colleagues demonstrated that the pslA gene, a member of the exopolysaccharide synthesis psl regulon in P. aeruginosa, played a role in initial attachment—approximately a third as many pslA mutants were attached to PVC as the wild type in minimal media supplemented with glucose (Overhage et al., 2005). Furthermore, they demonstrated, using a PpslA-gfp transcriptional fusion, that the pslA gene was induced upon attachment to the surface (Overhage et al., 2005). hese data are quite interesting, because in addition to the often-mentioned role of exopolysaccharide in the stabilization of the mature biofilm structure (Friedman and Kolter, 2004a, 2004b; Jackson et al., 2004; Matsukawa and Greenberg, 2004; Vasseur et al., 2005; Pamp et al., this volume), this works suggests that exopolysaccharide may also play a role in early events in biofilm formation, such as irreversible attachment. As mentioned above, the process of reversible attachment allows an individual bacterial cell to sample the surface, integrate the appropriate local environmental signals and either release from the surface or trigger the transition to irreversible attachment. Because irreversible attachment appears to be the first stable interaction with the substratum, we proposed that this step is the “first committed” step in biofilm formation. In part, the choice of the phrase “committed step,” typically used within the context of metabolic pathways, was chosen to convey the idea that the initial step in surface commitment is part of a larger set of behaviors of which P. aeruginosa is capable. It is possible that P. aeruginosa, subsequent to reversible attachment to a surface, is able to initiate other surface behaviors, such as twitching motility or swarming in addition to biofilm formation. A greater understanding of the early steps in biofilm formation is needed to understand this surface behavior in the greater context of the lifestyle of P. aeruginosa. Genomics and proteomics While the switch from reversible to irreversible attachment is evident when observed microscopically, is this transition accompanied by changes in gene or protein expression in the cell? Recently, several studies have sought to determine the extent of gene expression changes during early stages of biofilm formation. Generally, the results of genomic or proteomic studies have demonstrated a profound difference in gene expression in cells attached to a surface compared with those grown in the liquid environment, and in one study, approximately 25% of all proteins were upregulated in 1 day old biofilms as compared to planktonic cultures (Sauer et al., 2002; Sauer, 2003; Southey-Pillig et al., 2005). Sauer and colleagues also tracked the changes in protein expression over several stages of biofilm formation, breaking the pathway into three distinct stages: maturation 1 and maturation 2 and

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dispersion (Southey-Pillig et al., 2005). his work compared the proteome of P. aeruginosa during these three stages as well as during planktonic growth and was able to demonstrate that a large number of proteins were differentially regulated during these stages and that a distinct set of proteins is expressed at each stage in biofilm maturation (Southey-Pillig et al., 2005). Work by Waite and colleagues explored the transcriptional profile of P. aeruginosa during logarithmic and stationary growth, and four different points in biofilm formation: 8, 14, 24, and 48 hours post inoculation, by growing P. aeruginosa on nitrocellulose membranes placed on LB agar (Waite et al., 2005). he percentage of the genome differentially regulated depended upon the comparison made. For example, by 8 hours 0.8% of the genome was differentially regulated in the model biofilm used in this study compared to a log-phase planktonic culture, but close to 10% of the genes were differentially regulated when an 8 hour biofilm was compared to a stationary phase culture grown in the same medium (Waite et al., 2005). However, the significance of these changes and whether they are a direct or indirect consequence of growing in a biofilm-like environment is unclear. Few transcriptome- or proteome-based studies have been conducted that have focused on the switch from reversible to irreversible attachment. One study that has tackled this issue head-on comes from work in Vibrio cholerae. Moorthy and Watnick (2004) used genetic studies to indicate that there were differences between planktonic bacteria and bacteria which had formed a monolayer. We believe that this monolayer stage may be thought of as the functional equivalent of irreversibly attached bacteria. Furthermore, these investigators showed that V. cholerae could be locked in a monolayer stage by growing them in a medium that lacked monosaccharides (Moorthy and Watnick, 2004), thus providing an excellent tool for specifically accessing differences between planktonic and irreversibly attached bacteria. Moorthy and Watnick (2005) used microarrays to analyze differences between planktonic- and monolayer-grown bacteria and found 150 genes were differentially regulated (91 upregulated and 59 downregulated). hese data suggest that even the early transition to irreversible attachment results in changes in the biology of the microbe compared to their planktonic counterparts. Summary While the majority of prokaryotic research has focused on studying planktonic bacteria grown in liquid media, it is evident that the majority of bacteria exist in nature attached to a substratum in what we now refer to as biofilms. his field has gained a fair amount of attention in recent years leaving one with the impression that it is a relatively new area of research, and while there have been many recent advances in our understanding of this process, history has proven that much of the early groundwork in the field performed in the 1930s and 40s is on the mark. Zobell’s early recognition that there were two types of attachment, an immediate yet unstable attachment and a more secure, time-dependent attachment has been repeatedly confirmed in a variety of different organisms. With the advent of modern molecular genetic and biochemical techniques, this evidence has been augmented further with the identification of adhesins and regulatory proteins that regulate the switch from reversible to irreversible attachment. he large number of genes involved in the switch from reversible to irreversible attachment suggests that this is an important stage

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in biofilm formation. In fact, we argue that this is the first committed step in the transition from a planktonic to biofilm lifestyle. References An, Y.H., and Friedman, R.J. (1998). Concise review of mechanisms of bacterial adhesion to biomaterial surfaces. J. Biomed. Mater. Res. 43, 338–348. Arora, S.K., Ritchings, B.W., Almira, E.C., Lory, S., and Ramphal, R. (1998). he Pseudomonas aeruginosa flagellar cap protein, FliD, is responsible for mucin adhesion. Infect. Immun. 66, 1000–1007. Azghani, A.O., Idell, S., Bains, M., and Hancock, R.E. (2002). Pseudomonas aeruginosa outer membrane protein F is an adhesin in bacterial binding to lung epithelial cells in culture. Microb. Pathog. 33, 109–114. Bloemberg, G.V., and Lugtenberg, B.J. (2001). Molecular basis of plant growth promotion and biocontrol by rhizobacteria. Curr. Opin. Plant Biol. 4, 343–350. Bos, R., van der Mei, H.C., and Busscher, H.J. (1999). Physico-chemistry of initial microbial adhesive interactions—its mechanisms and methods for study. FEMS Microbiol. Rev. 23, 179–230. Caiazza, N.C., and O’Toole, G.A. (2003). Alpha-toxin is required for biofilm formation by Staphylococcus aureus. J. Bacteriol. 185, 3214–3217. Caiazza, N.C., and O’Toole, G.A. (2004). SadB is required for the transition from reversible to irreversible attachment during biofilm formation by Pseudomonas aeruginosa PA14. J. Bacteriol. 186, 4476–4485. Costerton, J.W., Stewart, P.S., and Greenberg, E.P. (1999). Bacterial biofilms, a common cause of persistent infections. Science 284, 1318–1322. Cramton, S.E., Gerke, C., Schnell, N.F., Nicols, W.W., and Gotz, F. (1999). he intercellular adhesion (ica) locus is present in Staphylococcus aureus and is required for biofilm formation. Infect. Immun. 67, 5427–5433. De Weger, L.A., van der Vlught, C.I.M., Wij es, A.H.M., Bakker, P.A.H.M., Schippers, B., and Lugtenberg, B. (1987). Flagella of a plant-growth-stimulating Pseudomonas fluorescens are required for colonization of potato roots. J. Bacteriol. 169, 2769–2773. DeFlaun, M., F., Marshall, B.M., Kulle, E.-P., and Levy, S.B. (1994). Tn5 insertion mutants of Pseudomonas fluorescens defective in adhesion to soil and seeds. Appl. Environ. Microbiol. 60, 2637–2642. DeFlaun, M.F., Oppenheimer, S.R., Streger, S., Condee, C.W., and Fletcher, M. (1999). Alterations in adhesion, transport, and membrane characteristics in an adhesion-deficient pseudomonad. Appl. Environ. Microbiol. 65, 759–765. Dekkers, L.C., van der Bij, A.J., Mulders, I.H.M., Phoelich, C.C., Wentwoord, R.A.R., Glandorf, D.C.M., Wijffelman, C.A., and Lugtenberg, B.J.J. (1998). Role of the O-antigen of lipopolysaccharide, and possible roles of growth rate and of NADH, ubiquinone oxidoreductase (nuo) in competitive tomato roottip colonization by Pseudomonas fluorescens WCS365. Mol. Plant-Microbe Interact. 11, 763–771. Doyle, T.B., Hawkins, A.C., and McCarter, L.L. (2004). he complex flagellar torque generator of Pseudomonas aeruginosa. J. Bacteriol. 186, 6341–6350. Espinosa-Urgel, M., Salido, A., and Ramos, J.L. (2000). Genetic analysis of functions involved in adhesion of Pseudomonas putida to seeds. J. Bacteriol. 182, 2363–2369. Friedman, L., and Kolter, R. (2004a). Two genetic loci produce distinct carbohydrate-rich structural components of the Pseudomonas aeruginosa biofilm matrix. J. Bacteriol. 186, 4457–4465. Friedman, L., and Kolter, R. (2004b). Genes involved in matrix formation in Pseudomonas aeruginosa PA14 biofilms. Mol. Microbiol. 51, 675–690. Gjermansen, M., Ragas, P., Sternberg, C., Molin, S., and Tolker-Nielsen, T. (2005). Characterization of starvation-induced dispersion in Pseudomonas putida biofilms. Environ. Microbiol. 7, 894–906. Henrici, A.T. (1933). Studies of freshwater bacteria. I. a direct microscopic technique. J. Bacteriol. 25, 277–287. Hinsa, S.M., Espinosa-Urgel, M., Ramos, J.L., and O’Toole, G.A. (2003). Transition from reversible to irreversible attachment during biofilm formation by Pseudomonas fluorescens WCS365 requires an ABC transporter and a large secreted protein. Mol. Microbiol. 49, 905–918. Hinsa, S.M., and O’Toole, G.A. (2004). Mechanisms of Adhesion By Pseudomonads. In: Pseudomonas. Vol. 1. Ramus, J.-L. (ed). New York, NY, Kluwer Academic/Plenum Publishers, pp. 699–720. Hinsa, S.M., and O’Toole, G.A. (2006). Biofilm formation by Pseudomonas fluorescens WCS365: a role for LapD. Microbiology 152, 1375–1383.

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Jackson, K.D., Starkey, M., Kremer, S., Parsek, M.R., and Wozniak, D.J. (2004). Identification of psl, a locus encoding a potential exopolysaccharide that Is essential for Pseudomonas aeruginosa PAO1 biofilm formation. J. Bacteriol. 186, 4466–4475. Klausen, M., Heydorn, A., Ragas, P., Lambertsen, L., Aaes-Jorgensen, A., Molin, S., and Tolker-Nielsen, T. (2003). Biofilm formation by Pseudomonas aeruginosa wild type, flagella and type IV pili mutants. Mol. Microbiol. 48, 1511–1524. Kolenbrander, P.E., Andersen, R.N., Kazmerzak, K., Wu, R., and Palmer, R.J., Jr. (1999). Spatial organization of oral bacteria in biofilms. Methods Enzymol. 310, 322–332. Landry, R.M., An, D., Hupp, J.T., Singh, P.K., and Parsek, M.R. (2006). Mucin-Pseudomonas aeruginosa interactions promote biofilm formation and antibiotic resistance. Mol. Microbiol. 59, 142–151. Lawrence, J.R., Delaquis, P.J., Korber, D.R., and Caldwell, D.E. (1987). Behavior of Pseudomonas fluorescens within the hydrodynamic boundary layers of surface microenvironments. Microb. Ecol. 14, 1–14. Lillehoj, E.P., Kim, B.T., and Kim, K.C. (2002). Identification of Pseudomonas aeruginosa flagellin as an adhesin for Muc1 mucin. Am. J. Physiol. Lung Cell. Mol. Physiol. 282, L751–756. Marrie, T.J., and Costerton, J.W. (1984). Scanning and transmission electron microscopy of in situ bacterial colonization of intravenous and intraarterial catheters. J. Clin. Microbiol. 19, 687–693. Marshall, K.C., Stout, R., and Mitchell, R. (1971). Mechanism of the initial events in the sorption of marine bacteria to surfaces. J. Gen. Microbiol. 68, 337–348. Matsukawa, M., and Greenberg, E.P. (2004). Putative exopolysaccharide synthesis genes influence Pseudomonas aeruginosa biofilm development. J. Bacteriol. 186, 4449–4456. Matz, C., Bergfeld, T., Rice, S.A., and Kjelleberg, S. (2004). Microcolonies, quorum sensing and cytotoxicity determine the survival of Pseudomonas aeruginosa biofilms exposed to protozoan grazing. Environ. Microbiol. 6, 218–226. Moorthy, S., and Watnick, P.I. (2004). Genetic evidence that the Vibrio cholerae monolayer is a distinct stage in biofilm development. Mol. Microbiol. 52, 573–587. Moorthy, S., and Watnick, P.I. (2005). Identification of novel stage-specific genetic requirements through whole genome transcription profiling of Vibrio cholerae biofilm development. Mol. Microbiol. 57, 1623–1635. O’Toole, G.A., and Kolter, R. (1998a). Initiation of biofilm formation in Pseudomonas fluorescens WCS365 proceeds via multiple, convergent signalling pathways, a genetic analysis. Mol. Microbiol. 28, 449–461. O’Toole, G.A., and Kolter, R. (1998b). Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol. Microbiol. 30, 295–304. O’Toole, G.A., Kaplan, H., and Kolter, R. (2000). Biofilm formation as microbial development. Annu. Rev. Microbiol. 54, 49–79. Otto, K., and Silhavy, T.J. (2002). Surface sensing and adhesion of Escherichia coli controlled by the Cpxsignaling pathway. Proc. Natl. Acad. Sci. USA 99, 2287–2292. Overhage, J., Schemionek, M., Webb, J.S., and Rehm, B.H. (2005). Expression of the psl operon in Pseudomonas aeruginosa PAO1 biofilms: PslA performs an essential function in biofilm formation. Appl. Environ. Microbiol. 71, 4407–4413. Patel, R. (2005). Biofilms and antimicrobial resistance. Clin. Orthop. Relat. Res. 437, 41–47. Paul, E.A., and Clark, F.E. (1989). Soil Microbiology and Biochemistry. San Diego, CA, Academic Press. Picioreanu, C., Van Loosdrecht, M.C., and Heijnen, J.J. (2000). Effect of diffusive and convective substrate transport on biofilm structure formation, a two-dimensional modeling study. Biotechnol. Bioeng. 69, 504–515. Pratt, L.A., and Kolter, R. (1998). Genetic analysis of Escherichia coli biofilm formation, roles of flagella, motility, chemotaxis and type I pili. Mol. Microbiol. 30, 285–293. Rahme, L.G., Stevens, E.J., Wolfort, S.F., Shao, J., Tompkins, R.G., and Ausubel, F.M. (1995). Common virulence factors for bacterial pathogenicity in plants and animals. Science 268, 1899–1902. Raivio, T.L., and Silhavy, T.J. (1997). Transduction of envelope stress in Escherichia coli by the Cpx twocomponent system. J. Bacteriol. 179, 7724–7733. Raivio, T.L. (2005). Envelope stress responses and Gram-negative bacterial pathogenesis. Mol. Microbiol. 56, 1119–1128. Ramsey, M.M., and Whiteley, M. (2004). Pseudomonas aeruginosa attachment and biofilm development in dynamic environments. Mol. Microbiol. 53, 1075–1087.

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Sauer, K., Camper, A.K., Ehrlich, G.D., Costerton, J.W., and Davies, D.G. (2002). Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J. Bacteriol. 184, 1140–1154. Sauer, K. (2003). he genomics and proteomics of biofilm formation. Genome Biol. 4, 219. Southey-Pillig, C.J., Davies, D.G., and Sauer, K. (2005). Characterization of temporal protein production in Pseudomonas aeruginosa biofilms. J. Bacteriol. 187, 8114–8126. Soutourina, O.A., and Bertin, P.N. (2003). Regulation cascade of flagellar expression in Gram-negative bacteria. FEMS Microbiol. Rev. 27, 505–523. Stoodley, P., Kathju, S., Hu, F.Z., Erdos, G., Levenson, J.E., Mehta, N., Dice, B., Johnson, S., Hall-Stoodley, L., Nistico, L., Sotereanos, N., Sewecke, J., Post, J.C., and Ehrlich, G.D. (2005). Molecular and imaging techniques for bacterial biofilms in joint arthroplasty infections. Clin. Orthop. Relat. Res. 31–40. Stover, C.K., Pham, X.Q., Erwin, A.L., Mizoguchi, S.D., Warrener, P., Hickey, M.J., Brinkman, F.S., Hufnagle, W.O., Kowalik, D.J., Lagrou, M., Garber, R.L., Goltry, L., Tolentino, E., WestbrockWadman, S., Yuan, Y., Brody, L.L., Coulter, S.N., Folger, K.R., Kas, A., Larbig, K., Lim, R., Smith, K., Spencer, D., Wong, G.K., Wu, Z., and Paulsen, I.T. (2000). Complete genome sequence of Pseudomonas aeruginosa PA01, an opportunistic pathogen. Nature 406, 959–964. Sutherland, I.W., Hughes, K.A., Skillman, L.C., and Tait, K. (2004). he interaction of phage and biofilms. FEMS Microbiol. Lett. 232, 1–6. Tielker, D., Hacker, S., Loris, R., Strathmann, M., Wingender, J., Wilhelm, S., Rosenau, F., and Jaeger, K.E. (2005). Pseudomonas aeruginosa lectin LecB is located in the outer membrane and is involved in biofilm formation. Microbiology 151, 1313–1323. Toutain, C.M., Zegans, M.E., and O’Toole, G.A. (2005). Evidence for two flagellar stators and their role in the motility of Pseudomonas aeruginosa. J. Bacteriol. 187, 771–777. van Leeuwenhoek, A. (1683). An abstract of a Letter from Antonie van Leeuwenhoek, About Animals in the scurf of the Teeth. Vol. 14. London, Philosophical Transactions of the Royal Society of London, pp. 568–574. Vasseur, P., Vallet-Gely, I., Soscia, C., Genin, S., and Filloux, A. (2005). he pel genes of the Pseudomonas aeruginosa PAK strain are involved at early and late stages of biofilm formation. Microbiology 151, 985–997. Vinh, D.C., and Embil, J.M. (2005). Device-related infections, a review. J. Long Term Eff. Med. Implants 15, 467–488. Waite, R.D., Papakonstantinopoulou, A., Littler, E., and Curtis, M.A. (2005). Transcriptome analysis of Pseudomonas aeruginosa growth, comparison of gene expression in planktonic cultures and developing and mature biofilms. J. Bacteriol. 187, 6571–6576. Waksman, S.A., and Vartiovaara, U. (1938). he adsorption of bacteria by marine bottom. Biol. Bull. 74, 56–63. Yoon, S.S., Hennigan, R.F., Hilliard, G.M., Ochsner, U.A., Parvatiyar, K., Kamani, M.C., Allen, H.L., DeKievit, T.R., Gardner, P.R., Schwab, U., Rowe, J.J., Iglewski, B.H., McDermott, T.R., Mason, R.P., Wozniak, D.J., Hancock, R.E., Parsek, M.R., Noah, T.L., Boucher, R.C., and Hassett, D.J. (2002). Pseudomonas aeruginosa anaerobic respiration in biofilms. Relationships to cystic fibrosis pathogenesis. Dev. Cell 3, 593–603. Zobell, C.E., and Allen, E.C. (1935). he significance of marine bacteria in the fouling of submerged surfaces. J. Bacteriol. 29, 239–251. Zobell, C.E., and Anderson, D.Q. (1936). Observations on the multiplication of bacteria in different volumes of stored sea water and the influence of oxygen tension and solid surfaces. Biol. Bull. 71, 324–342. Zobell, C.E. (1937). he influence of solid surfaces upon the physiological activities of bacteria in sea water. J. Bacteriol. 33, 86. Zobell, C.E. (1943). he effects of solid surfaces upon bacterial activity. J. Bacteriol. 46, 39–56.

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The Bioilm Matrix: A Sticky Framework Sünje Johanna Pamp, Morten Gjermansen, and Tim Tolker-Nielsen

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Abstract he extracellular matrix of structured microbial communities constitutes the framework that holds the component cells together. Although the presence of cell-to-cell interconnecting matrices appears to be a common feature of structured microbial communities, there is a remarkable diversity in the composition of these matrices. Compounds such as polysaccharides, fimbriae, mating pili, and extracellular DNA can all function as extracellular matrix components. In the present chapter we provide examples of the diversity of biofilm matrices. Introduction he formation and maintenance of structured microbial communities critically depends on the presence of extracellular substances that constitute cell-to-cell interconnecting matrices. Proliferation of sessile bacteria and the production of cell-to-cell interconnecting extracellular compounds lead to the formation of microbial biofilms. he extracellular matrix that surrounds the bacteria in biofilms is believed to offer protection against various adverse factors including protozoan predation in environmental settings (Matz and Kjelleberg, 2005) and host immune responses in medical settings (Costerton et al., 1999). he production of the cell-to-cell interconnecting components in biofilms apparently is a cost each bacterium pays in order to contribute to the social activity of creating a protective biofilm domicile. Although the presence of a cell-to-cell interconnecting matrix appears to be a common feature of microbial biofilms, there is a remarkable diversity in the composition of these matrices. Many kinds of exopolymers, e.g. polysaccharide, protein, and DNA may function as biofilm matrix material. In addition to these exopolymers, outer membrane proteins and a variety of cell appendages such as fimbriae, pili, and flagella may also function as part of the biofilm matrix. he components of the biofilm matrix are usually, but not always, produced by the bacteria themselves. In the present chapter we present examples of polysaccharides, proteins, and extracellular DNA as matrix components in biofilms. It appears that the expression of polysaccharides and proteins of the extracellular matrix in many cases is regulated through proteins which contain GGDEF and/or EAL domains. hese proteins control intracellular levels of the cyclic dinucleotide c-di-GMP through diguanylate cyclase and phosphodiesterase

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activity. c-di-GMP in turn acts as a second messenger and affects matrix production and the adhesiveness of the bacteria. For a thorough description of the regulatory mechanism involving GGDEF and EAL domain proteins the reader is referred to Dow et al., this volume. he generation of extracellular DNA is evidently in many cases regulated by means of quorum-sensing which is a mechanism that enable bacteria to monitor their cell population density through the extracellular accumulation of signaling molecules. For thorough descriptions of quorum-sensing in Gram-negative and Gram-positive bacteria the reader is referred to Atkinson et al., and Yarwood, this volume. Polysaccharides as matrix components in bioilms Extracellular polysaccharides are usually very important parts of biofilm matrices. he chemical composition and physical properties of the polysaccharides in biofilm matrices can vary greatly due to the type of monomer units, the kind of glycosidic linkages (e.g. B-1,4, B-1,3 or A-1,6), and the occurrence of different organic and inorganic substitutions. Here we address five types of polysaccharides that have been shown to play important roles in biofilm formation by a number of different bacterial species: cellulose, PNAG/PIA, PEL, PSL and VPS. Cellulose Cellulose is the most abundant polysaccharide in nature and is produced by both plants and bacteria. Bacterial cellulose biogenesis and the role of cellulose in biofilm formation has been described for a number of bacterial species including Gluconacteobacter xylinus (formerly called Acetobacter xylinum) Sarcina ventriculi, Agrobacterium tumefaciens, Rhizobium leguminosarum, Escherichia coli, Salmonella spp., and Pseudomonas fluorescence (Ausmees et al., 1999; Deinema and Zevenhuizen, 1971; Matthysse et al., 1995; Napoli et al., 1975; Ross et. al., 1991; Zogaj et al., 2001; Spiers et al., 2003). However, comparative sequence analyses indicate that many other bacteria including Vibrio, Yersinia, and Burkholderia species have the capacity to synthesize cellulose. In the present chapter we will focus on G. xylinus, Salmonella spp., and P. fluorescence as model organisms. When G. xylinus is grown in vitro in a static broth culture it forms a thick cellulose-containing biofilm at the air/liquid interface (Schramm and Hestrin, 1954). In nature G. xylinus has predominantly been found on decaying fruits, where a cellulose-matrix surrounding the bacteria may protect against competitors and lethal effects of UV light. he native cellulose is synthesized as a long polymeric chain composed of B-1,4-linked D-Glucose units by a multimeric enzyme complex, which is located in the cytoplasmic membrane of G. xylinus. A single row of pore-like structures on the outer membrane along the longitudinal axis of the rod-shaped bacterium, called linear terminal-complexes, has been identified to be the site of extrusion of the native cellulose chain (Brown et al., 1976; Kimura et al., 2001). Several of these single glucan chains coalesce via van der Waals forces and hydrogen bonds to form crystalline microfibrils. An entangled mesh of these microfibrils produces a gelatinous structure which constitutes the major component of the biofilm matrix. In G. xylinus and other cellulose-synthesizing bacteria, the genetic elements coding for cellulose synthesis are located in an operon consisting of four genes which are generally designated bcsA, bcsB, bcsC and bcsD (bacterial cellulose synthesis). he polymerization of

The Bioilm Matrix

single monomer units of UDP-glucose to cellulose polymer is catalyzed by the cellulose synthase BcsA. Several transmembrane domains anchor the cellulose synthase BcsA in the cytoplasmic membrane. In spatial proximity with the BcsA enzyme the BcsB protein can be found. his protein regulates the activity of cellulose synthesis via binding of the second messenger c-di-GMP (Ross et al., 1987). It has been proposed that a controlled release of c-di-GMP from BcsB to the BcsA enzyme may activate the cellulase synthase allosterically. Recent bioinformatic investigations suggest the localization of a binding site of c-di-GMP, called the PilZ domain, at the C-terminal end of the BcsA cellulose synthase (Amikam and Galperin, 2006). he functions of BcsC and BcsD have not been described yet. However, sequence analysis suggests that BcsC might be involved in pore formation through the cell wall, whereas BcsD might have an effect on cellulose crystallization (Saxena et al., 1994; Wong et al., 1990). Synthesis of cellulose by enterobacteria such as Salmonella spp. and E. coli has been associated with the ability to form biofilms on abiotic surfaces, such as glass and polystyrene (Garcia et al., 2004; Römling et al., 2000; Zogaj et al., 2001). High-level production of cellulose and proteinaceous curli fimbria by these organisms result in the formation of wrinkly or rough colonies on agar plates. If the bacteria are grown on rich agar medium (of low osmolarity), containing the diazo dye Congo Red, which binds to cellulose and curli fimbriae, they will form characteristic colonies which appear red, dry and rough (Hammar et al,. 1995; Römling et al., 2000; Zogaj et al., 2001). Regulation of the biosynthesis of cellulose and curli fimbriae evidently occurs through a complex coordinated pathway. he CsgD protein, a member of the LuxR superfamily of transcriptional regulators, is believed to regulate the synthesis of curli fimbria directly via transcriptional activation of the curli biosynthesis operon csgDEFG-csgBAC (Römling et al., 2000; Gerstel et al., 2003). In addition, CsgD stimulates the synthesis of cellulose indirectly via transcriptional activation of AdrA (Römling et al., 2000; Zogaj et al., 2001). AdrA is a two-domain protein consisting of an N-terminal MASE2 domain (Nikolskaya et al., 2003) and a C-terminal GGDEF domain, capable of producing c-di-GMP (Simm et al., 2004). High concentrations of c-di-GMP have been shown to induce the cellulose biosynthesis operon (yhjRQbcsABZC-bcsEFG) and thereby the excretion of cellulose from Salmonella enterica (Zogaj et al., 2001; Solano et al., 2002). Recent data indicate that in addition to AdrA, other GGDEF-domain containing proteins of Salmonella spp., are involved in modulating the level of the c-di-GMP, and thus biofilm formation (Garcia et al., 2004; Kader et al., 2006). Cellulose and curli fimbriae together form a highly inert matrix, which account for the strength and integrity of biofilms formed under various conditions by these enterobacteria (Garcia et al., 2004; Römling et al., 2000; Zogaj et al., 2001). When P. fluorescens was grown for extended periods in static liquid cultures, which contained numerous niches, spontaneous variants that colonized the different niches arose at high frequencies (Rainey and Travisano, 1998). One group of these variants, which colonized the air/liquid interface of the static cultures by forming a robust biofilm-pellicle, was termed “the wrinkly spreader” because the bacteria formed distinct wrinkled colonies on agar plates (Rainey and Travisano, 1998). his highly aggregative phenotype of the wrinkly spreader has been linked to the production of an acetylated form of cellulose and a proteinaceous fimbrial like attachment factor (Spiers et al., 2003; Spiers and Rainey 2005).

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Complementation experiments suggested that the cellulose fibers could interact with the lipopolysaccharide (LPS) of neighboring cells and that this interaction in conjunction with the proteinaceous attachment factor was responsible for the strength and integrity of the pellicle (Spiers and Rainey 2005). Studies of P. fluorescens in the spatially heterogeneous environment of a static culture have also shown that cheater variants, which benefit from the production of matrix compounds by neighboring bacteria without contributing themselves, arise at high frequencies in the biofilm pellicle (Rainey and Rainey, 2003). he frequency of cheaters in a biofilm in a given system is limited to the point where their presence damages the integrity of the biofilm too much. In general, the emergence of cheaters is the key problem for the evolution of microbial cooperation in biofilms. However, the predominantly clonal structure of the microcolonies in biofilms, combined with the re-establishment of them by single cells, acts as a purification mechanism to get rid of cheaters, and promotes the evolution of cooperation in biofilms (Kreft, 2004). Production of the acetylated cellulose polymer in P. fluorescens was shown to be encoded by genes in the 10-gene wss operon, and on the basis of homology with the G. xylinus bcs genes the WssB, WssC and WssE proteins were identified as the bacterial cellulose synthase subunits (Spiers et al., 2002; Spiers et al., 2003). he WssG, WssH and WssI proteins are unique to the P. fluorescens cellulose operon, but their homology to the AlgF, AlgI and AlgJ proteins of P. aeruginosa (Franklin and Ohman 1996) suggests a role in the acetylation of the polymer. he production of the cellulosic polymer has been demonstrated to enhance the fitness of P. fluorescens during colonization of the rhizosphere (Gal et al., 2003). he wrinkly spreader phenotype was found to be regulated by the GGDEF domain protein WspR, which consists of a C-terminal CheY-like receiver domain and an N-terminal GGDEF output domain. Mutagenesis showed that the activity of WspR was dependent on phosphorylation of the protein via gene products encoded by the wsp operon, and that the GGDEF domain was essential for the regulation of cellulose production. (Spiers et al., 2003; Goymer et al., 2006). PNAG/PIA he role of poly-N-acetylglucosamine as matrix component in biofilms has been extensively studied in two closely related Gram-positive bacteria; the coagulase-negative organism Staphylococcus epidermidis which synthesizes polysaccharide intercellular adhesin (PIA), and the coagulase-positive organism Staphylococcus aureus which synthesizes poly-Nacetylglucosamine (PNAG). More recently it has been found that PNAG/PIA-like polysaccharides are also synthesized by Gram-negative bacteria such as E. coli, Yersinia pestis and Actinobacillus sp. In addition, comparative genome sequence analysis has revealed that homologues of the poly-N-acetylglucosamine biosynthesis genes are present in many bacterial species including Pseudomonas fluorescence, Bordetella pertussis, Ralstonia solanacearum, and Lactococcus lactis. Here we will turn our attention on PNAG/PIA synthesized by S. epidermidis, S. aureus, E. coli and Y. pestis. S. aureus and S. epidermidis are frequently found as harmless inhabitants of the mucosal nasal passages or the normal skin flora of humans. However, these organisms are opportunistic pathogens and they are increasingly found to be the cause of invasive, and chronic, medical device-associated infections. hese implant-associated infections are difficult to

The Bioilm Matrix

eradicate and it is believed that the biofilm mode of growth is responsible for the inherent tolerance towards host immune responses and antimicrobial treatment. Biofilm development by staphylococci is multifactorial. Many extracellular or surface-bound polymers have been identified to play a role in cell-to-cell adhesion and binding to biotic and abiotic surfaces, but in general PNAG/PIA is believed to be the most important biofilm matrix component for the staphylococci (Caiazza and O’Toole, 2003; Foster and Höök, 1998; Gross et al., 2001; Götz, 2002; Vuong et al., 2000). he polysaccharides PIA and PNAG synthesized by S. epidermidis and S. aureus, are chemically and immunologically closely related. PIA and PNAG are both homopolysaccharides, composed of unbranched long polymeric chains of D-glucosamine uniformly linked together by A-1,6 glucosidic bonds, but they differ in chain length and modification (Mack et al., 1996; Maira-Litran et al., 2002). he wheat germ agglutinin lectin (WGA) was found to bind to N-acetylglucosamine produced by staphylococci ( Jäger et al., 2005; Sanford et al., 1995; Sanford et al., 1996; homas et al., 1997). Figure 4.1 visualizes the

Figure 4.1 Confocal laser scanning micrographs showing bacterial cells (stained with Syto 9) and PNAG exopolysaccharide (stained with WGA-TMR) in a bioilm formed by Staphylococcus aureus SJ235 (a clinical isolate obtained from a cystic ibrosis patient). Staphylococcal cells were cultivated in a continuous low chamber setup perfused with minimal medium. After 24 hours of cultivation, the cells had formed a single-layer bioilm on the glass surface (A), and PNAG could be visualized (B). An overlay of the two images demonstrates that most cells were surrounded by PNAG exopolysaccharide (C). After 3 days of cultivation, a bioilm with microcolonies containing staphylococci and PNAG had formed as visualized by the top-down shadow projection with two lanking section images (D) and with the 3D image projection (E).

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presence of the extracellular polysaccharide in a staphylococcal biofilm using fluorescently labeled WGA lectin and confocal laser scanning microscopy. he genetic elements coding for the biosynthesis of poly-N-acetylglucosamine are located in an operon consisting of the four genes icaA, icaD, icaB and icaC (intercellular adhesin). A gene, icaR, located upstream of icaADBC and transcribed divergently, encodes a transcriptional repressor of the icaADBC-operon (Conlon et al., 2002a; Jefferson et al., 2004; Cramton et al., 1999; Heilmann et al., 1996; Gerke et al., 1998). IcaA, IcaC and IcaD are located in the cellular membrane, whereas IcaB can be found extracellularly. he polymerization of single monomer units of UDP-N-acetylglucosamine to polymer chains of B-1,6-N-acetylglucosamine is catalyzed by the N-acetylglucosaminyl-transferase IcaA. Co-expression of icaA with icaD increased the enzyme activity significantly, indicating a supportive function of IcaD for IcaA enzyme activity. Oligomers of N-acetylglucosamine produced by IcaAD only reach a maximal length of 20 residues. Expression of long Nacetylglucosaminyl chains requires co-expression of icaA and icaD with icaC (Gerke et al., 1998). TcaR (Teicoplanin associated regulator), a protein with sequence homology to MarR-like transcriptional regulators, was shown to influence transcription of icaADBC negatively in S. aureus ( Jefferson et al., 2004). SarA (staphylococcal accessory regulator A), a global regulator for expression of virulence factors in S. aureus, was shown to upregulate transcription of icaADBC in S. aureus as well as S. epidermidis (Beenken et al., 2003; Tormoet al., 2005b; Valle et al., 2003). Staphylococcal cells lacking SarA showed reduced binding to abiotic surfaces such as polystyrene or glass in static as well as continuous flow systems (Beenken et al., 2003; Tormo et al., 2005b; Valle et al., 2003). Quorum sensing in staphylococci (encoded by the accessory gene regulator (agr) locus) was shown to influence biofilm formation of both S. aureus and S. epidermidis, but apparently through an ica-independent pathway (Vuong et al., 2000; Vuong et al., 2003; Yarwood et al., 2004). he alternative sigma factor B (SB) is part of a complex regulatory network that regulates basic cellular processes as well as virulence factor expression in S. aureus and possibly also in S. epidermidis (Kies et al., 2001; Novick, 2003; Pane-Farre et al., 2006; Ziebandt et al., 2001). Previously SB was found to regulate biofilm formation of an S. aureus mucosal isolate (Rachid et al., 2000a), but more recent data indicate that SB might play a minor role in regulating biofilm development of S. aureus as complete deletion of sigB did not significantly affect PNAG production (Valle et al., 2003). In contrast, a positive, although indirect, influence of SB on PIA regulation seems to be established for S. epidermis. Deletion of either SB itself or its positive regulator rsbU was shown to decrease production of PIA and therefore lead to a biofilm-negative phenotype in a microtiter tray assay (Knobloch et al., 2001; Knobloch et al., 2004). he biofilm-negative phenotype could be restored by addition of ethanol resulting in downregulation of icaR and increased synthesis of PIA (Knobloch et al., 2001; Knobloch et al., 2004), indicating the presence of an additional regulatory element in the pathway that regulates PIA synthesis. It has been widely observed that biofilm formation of staphylococci is enhanced in the presence of additional NaCl, glucose, and ethanol. While the glucose- and NaCl-dependent biofilm formation in S. aureus seems to be regulated independently of the ica-locus, enhanced biofilm formation of S. epidermidis in the presence of NaCl or ethanol is regulated via transcriptional induction of icaADBC (Conlon et al., 2002a; Conlon et al., 2002b; Lim

The Bioilm Matrix

et al., 2004). Addition of glucose seems to affect the biosynthesis of PIA in S. epidermidis positively, but at a stage subsequent to icaADBC transcription (Mack et al., 1992; Dobinsky et al., 2003). Of therapeutic relevance is the observation that some antibiotics are able to induce PIA production in S. epidermidis. Subinhibitory concentrations of tetracycline and quinuprisin-dalfopristin were found to induce transcription of the ica-operon and promote biofilm formation in a microtiter tray assay (Rachid et al., 2000b). E. coli is, in addition to cellulose, colanic acid and capsular polysaccharides, able to synthesize the PNAG/PIA-like polysaccharide PGA (poly-N-acetyl-glucosamine). Disruption of the pga locus in E. coli was found to decrease biofilm formation significantly in a polystyrene microtiter tray assay (Blattner et al., 1997; Wang et al., 2004). he PGA exopolymer produced by E. coli consists of unbranched poly-B-1,6-N-acetylglucosamine, contains less than 3% non-N-acetylated glucosaminyl moieties and does not possess any major substitutions (Wang et al., 2004). he genetic elements coding for the biosynthesis of PGA are located in an operon consisting of the four genes pgaABCD (formerly called ycdSRQP) (Blattner et al., 1997; Wang et al., 2004). he low G+C content of the pgABCD locus in E. coli (44% versus 51%) suggests that these genes were horizontally transferred (Wang et al., 2004). PgaC is an N-glycosyltransferase which is predicted to be anchored in the membrane by two N-terminal and three C-terminal transmembrane domains. PgaB is assumed to be a lipoprotein and shares sequence similarities with the staphylococcal protein IcaB (Wang et al., 2004). he protein sequence of PgaA indicates that it is located in the outer membrane, suggesting that it might participate in mediating translocation of PGA to the cell surface. PgaD is supposed to be a small inner membrane protein with two N-terminal membrane-spanning domains (Wang et al., 2004). he pgaABCD operon in E. coli was shown to be under negative control of CsrA (global storage regulator A). Disruption of csrA in E. coli led to a significant increase in biofilm formation in microtiter trays and on a glass surfaces ( Jackson et al., 2002). CsrA is a small RNA-binding protein which is a central effector molecule in the global regulatory system Csr, which controls bacterial gene transcription on the post-transcriptional level (Romeo, 1998). CsrA is known to be involved in regulating central metabolic pathways, such as glycogen synthesis and catabolism, gluconeogenesis, and glycolysis (Sabnis et al., 1995; Yang et al., 1996). CsrA binds to the pgaA mRNA in competition with the ribosome and prevents its translation by destabilizing the transcript (Romeo, 1998). Six putative CsrA binding sites in the pgaA mRNA leader have been identified (Wang et al., 2005). CsrB and CsrC, two small RNAs (sRNAs), have been found to antagonize the regulatory effects of CsrA. hese sRNAs have repeat sequences and were shown to bind several CsrA proteins, thereby sequestering CsrA from its target mRNA leader. his mechanism leads to positive modulation of the transcription of pgaABCD and therefore PGA synthesis (Weilbacher et al., 2003). Recent research indicates that sRNAs in general may be key regulators of virulence factors and adaptive processes in Gram-negative as well as Gram-positive pathogens (Romby et al., 2006). he hmsHFRS (hemin storage) genes of Y. pestis share similarities to the icaADBC genes of staphylococci (Darby et al., 2002; Pendrak and Perry, 1993; Perry et al., 1990), and were shown to be required for biofilm formation by Y. pestis in a flow-chamber system ( Jarrett et al., 2004). he hmsHFRS operon is located on a 102 kb large mobile high-patho-

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genicity island (HPI) which has been found in Y. pestis and Y. pseudotuberculosis (Buchrieser et al., 1999; Fetherston et al., 1992). Sequence analysis predicts that HmsH is located in the outer membrane of Y. pestis, whereas HmsF is supposed to be a lipoprotein (Lillard et al., 1997). HmsR has about 39% identity to the staphylococcal N-glycosyltransferase IcaA, and HmsS is predicted to be a small inner membrane protein with two N-terminal membrane-spanning domains (Darby et al., 2002; Lillard et al., 1997). Evidence has been presented that expression of the hsmHFRS locus is regulated in a temperature dependent fashion via the HmsT and HmsP proteins, which contain a GGDEF and an EAL domain, respectively (Kirillina et al., 2004). Biofilm formation allows Y. pestis to colonize the proventricular valve within fleas (Darby et al., 2002; Hinnebusch et al., 1996; Jarrett et al., 2004). Colonization of the proventricular valve, which separates the midgut from the esophagus, causes physical blockage of the flea, and efficient transmission of Y. pestis, the etiological agent of plague, to humans and rodents is facilitated when a blocked flea attempts to feed (Darby et al., 2002; Hinnebusch et al., 1996; Jarrett et al., 2004). PEL and PSL P. aeruginosa can, dependent on the strain and growth conditions, produce at least three different polysaccharides: alginate, PEL and PSL. Besides these, two additional gene clusters which putatively are involved in polysaccharide biosynthesis (PA1381-PA1398 and PA3552-PA3558) have been identified in the chromosome of the reference strain P. aeruginosa PAO1. Mucoid forms of P. aeruginosa that over-express the alg genes (alginate biosynthesis, PA3540-PA3551) are primarily found in infected lungs of cystic fibrosis patients (Govan and Deretic, 1996). However, most P. aeruginosa strains are non-mucoid, and alginate was found not to be a significant component of the extracellular matrix of nonmucoid laboratory strains (Wozniak et al., 2003). In the present chapter we will describe PEL and PSL as examples of exopolysaccharides in the extracellular matrix of P. aeruginosa biofilms. While PEL and PSL both seem to be branched heteropolysaccharides, the main component of PEL is glucose, whereas PSL has a high content of mannose (Friedman and Kolter, 2004a; Friedman and Kolter, 2004b). High-level expression of PEL and PSL in P. aeruginosa was shown to lead to the formation of wrinkly colonies on agar plates, and synthesis of PEL was shown to enable P. aeruginosa to form pellicle-biofilm at the air/liquid interface of broth cultures (Friedman and Kolter, 2004a; Friedman and Kolter, 2004b). Proteinaceous cup fimbriae appear to participate together with PEL in P. aeruginosa biofilm formation under some conditions (Friedman and Kolter, 2004a), similar to the cellulose and curli fimbriae-containing matrix of Salmonella sp. biofilms described above. Studies using a static attachment assay provided evidence that PSL is important in the early stages of P. aeruginosa biofilm development, whereas the synthesis of PEL seems to be important in later stages of biofilm development (Friedman and Kolter, 2004b; Vasseur et al., 2005). In a continuous culture flow-chamber set-up, a P. aeruginosa strain deficient in PSL production was found to be impaired in biofilm formation ( Jackson et al., 2004; Matsukawa and Greenberg, 2004), supporting a role of PSL in the early stages of biofilm development.

The Bioilm Matrix

he genetic elements encoding the biosynthesis of PEL are organized in a gene cluster which consists of seven open reading frames termed pelA-G (pellicle formation; PA3058PA3064), spanning a 12.2 kb region of the P. aeruginosa genome. Although the exact function of the gene products is not described yet, sequence analysis revealed that these proteins contain domains, which are present in proteins involved in polysaccharide processing in other organisms. For example, PelF shares sequence homology with group IV glycosyl transferases, PelA has similarity to endo A-1,4 polygalactosaminidase, and PelE contains domains which resembles those of sucrose synthases. In addition, PelG was suggested to be a PST-family protein, whose members are involved in translocating glycolipid precursors through the membrane. Many of the proteins contain transmembrane domains, indicating their final location in the membrane (Friedman and Kolter, 2004a; Vasseur et al., 2005). he psl (polysaccharide synthesis locus) gene cluster contains the 15 co-transcribed open reading frames pslA-O (PA2231-PA2245). Sequence analysis of the first 11 predicted gene products revealed homologies to proteins involved in polysaccharide biosynthesis. he putative gene products PslF, PslH and PslI share sequence homologies with group I family glycosyltransferases, whereas PslA is similar to sugar transferases and PslD might be involved in polysaccharide transport. Since PslA, PslJ and PslK are predicted to contain 6, 11 and 12 transmembrane domains respectively, they are most likely located in the cellular membrane (Friedman and Kolter, 2004b; Jackson et al., 2004; Matsukawa and Greenberg, 2004) Recent data indicate that synthesis of the polysaccharide matrix in P. aeruginosa biofilms is regulated via intracellular levels of c-di-GMP. Transcription of the pel and psl loci were found to be regulated through the wsp chemosensory system (wspABCDEFR, PA3708-PA3702), of which the two gene products, WspR and WspF contain the catalytic GGDEF and EAL domains respectively. Whereas high levels of c-di-GMP were found to stimulate transcription of the pel and psl loci and induce biofilm formation, low intracellular c-di-GMP levels were found to decrease biofilm formation in a microtiter tray assay and in a flow-chamber system (Hickman et al., 2005). VPS Vibrio cholerae produces the exopolysaccharide VPS (Vibrio polysaccharide) which causes the formation of wrinkly (rugose) colonies on agar plates and has been shown to have an important role in biofilm formation on solid surfaces and at liquid–air interfaces (CasperLindley and Yildiz, 2004; Wai et al., 1998; Watnick and Kolter, 1999; Yildiz et al., 2001; Yildiz and Schoolnik, 1999). he bacteria in V. cholerae biofilms have been shown to exhibit an enhanced survival in chlorinated water, and an elevated tolerance towards osmotic, acid and oxidative stresses compared to their planktonic counterparts (Morris et al., 1996; Wai et al., 1998; Yildiz and Schoolnik, 1999). he polysaccharide produced by V. cholerae contains mainly glucose and galactose as monomeric sugar components, and in addition smaller amounts of N-acetylglucosamine and mannose (Yildiz and Schoolnik, 1999). he genetic elements coding for the biosynthesis of VPS are organized in two gene clusters, vspI (vpsA-K, VC0917-VC0927) and vspII (vpsL-Q, VC09334-VC0939), that encompass a 30.7 kb region on the large chromosome of V. cholera. Sequence analysis revealed that the gene products of the vps gene cluster share sequence similarities with

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proteins involved in exopolysaccharide synthesis and polysaccharide modification in other bacterial species, among these glycosyl transferase, UDP-glucose dehydrogenase, glycosyl1-phosphate transferase and NDP-N-acetyl-D-galactosaminuronic acid dehydrogenase (Yildiz and Schoolnik, 1999). In vitro analysis demonstrated that transcription of the gene clusters vpsI and vpsII are regulated positively by the two response regulators VpsR and VpsT. Knock-out mutants in either vpsR or vpsT or both genes displayed reduced biofilm formation in a polyvinyl microtiter tray assay and in a static borosilicate glass slide assay compared to the rugose isogenic wild-type strain. VpsR belongs to the NtrC subclass of transcriptional regulators, whereas VspT shares sequence homologies with the transcriptional regulators CsgD and AgfD of E. coli and Salmonella sp. (Casper-Lindley and Yildiz, 2004; Yildiz et al., 2001). Evidence has been provided that transcription of the vps genes in V. cholerae is regulated through the second messenger c-di-GMP (Rashid et al., 2003; Tischler and Camilli, 2004). Whole genome transcriptional studies of bacteria which formed smooth and rugose colonies, respectively, showed that five genes which code for proteins containing GGDEF and/or EAL domains were differentially expressed in the two colonial phenotypes (Yildiz et al., 2004). More detailed analysis demonstrated that the GGDEF/EAL domain proteins CdgC (cyclic diguanylate), RocS (regulation of cell signaling) and MbaA (maintenance of biofilm architecture) all regulate vps transcription and V. cholerae biofilm formation negatively via the VpsR regulator (Bomchil et al., 2003; Lim et al., 2006; Rashid et al., 2003). In addition, high intracellular levels of the EAL domain-containing protein AcgA was shown to decrease V. cholerae biofilm formation, whereas high intracellular levels of the GGDEF domain-containing protein AcgB was shown to increase V. cholerae biofilm formation in a microtiter tray assay. Whether regulation of V. cholerae biofilm formation via AcgA and AcgB is dependent on the vps genes remains to be investigated (Kovacikova et al., 2005). Several environmental stimuli and complex regulatory pathways are involved in the aquatic and intestinal life cycles of V. cholera. Along with the synthesis of the cholera toxin (CT) and toxin co-regulated pili (TCP), the synthesis of the extracellular VPS matrix seems to play a major role in the life cycle and pathogenesis of V. cholera. It appears that quorum sensing negatively regulates biofilm formation as well as CT and TCP expression in V. cholera. At low cell density, the transcriptional regulator LuxO (together with the alternative sigma factor S54) activates expression of four small regulatory RNAs. hese sRNAs together with Hfq repress HapR (homologue to LuxR of V. harveyi) expression by destabilizing the hapR mRNA. In the absence of HapR the genes vps, ctx (cholera toxin) and tcp (toxin co-regulated pilus) are expressed. At high cell density, LuxO is inactive and therefore repression of hapR is relieved, and HapR negatively regulates biofilm formation as well as ctx and tcp expression (Hammer and Bassler, 2003; Jobling and Holmes, 1997; Lenz et al., 2004; Vance et al., 2003; Zhu and Mekalanos, 2003; Zhu et al., 2002). Recent investigations indicate that the biofilm growth mode of V. cholera might play an important role in the transmission of the diarrheal disease cholera (Zhu and Mekalanos, 2003; Faruque et al., 2006). V. cholera cells have been found to exist as aggregates (biofilms) of partially dormant cells in surface waters (Faruque et al., 2006). Within these aquatic environments the biofilm mode of growth might increase survival of V. cholera, for instance against grazing by protozoa as has been shown by Matz et al. (2005). Upon oral ingestion,

The Bioilm Matrix

these biofilms might furthermore be protected against acid stress in the gastric area. In the intestinal environment single cells might detach from the biofilm, due to high levels of quorum-sensing signal. In the intestinal environment with low quorum-sensing levels, expression of CT and TCP is enhanced and colonization of the intestinal sites induced. Again, high cell densities lead to high levels of quorum-sensing signal and repression of VPS, CT and TCP but induction of proteases, which might facilitate detachment of cell aggregates and single cells from the intestinal sites and exit from the host (Zhu and Mekalanos, 2003). In agreement, single free-swimming cells and small aggregates have been observed in stools from cholera patients (Faruque et al., 2006). Proteins as matrix components in bioilms Genetic and microscopic approaches have provided information about proteins that play roles as cell-to-cell interconnecting factors in the course of biofilm formation by different bacterial species. In the present chapter we present examples from two different groups: multimeric cell appendages and surface proteins. Multimeric cell appendages Large multimeric cellular appendages such as flagella, fimbriae, and pili typically consist of numerous major structural protein components and several auxiliary proteins. In many bacterial species flagella play a role in the initial phase of biofilm formation under some conditions (e.g. O’Toole and Kolter 1998a, b; Klausen et al., 2003b; Watnick and Kolter, 1999), and they may also play a role in the later phases of structural biofilm development (Klausen et al., 2003b; Yamada et al., 2005). It appears that flagellum-driven motility can promote initial biofilm formation by facilitating transport of the bacteria to a surface (Gilbert et al., 1993). In addition, evidence has been provided that flagella can act as both cell-to-surface adhesins and cell-to-cell adhesins (O’Toole and Kolter 1998a; Yamada et al., 2005). Type IV pili are used by a number of bacteria to perform surface associated motility (Mattick, 2002). In addition type IV pili have been shown to mediate adhesion to both abiotic and biotic surfaces under some conditions (Giltner et al., 2006; Mattick, 2002; Sheth et al., 1994; Schweizer et al., 1998; O’Toole and Kolter 1998a). In vitro studies revealed that type IV pili of P. aeruginosa display specificity towards asialo-GM1 and asialo-GM2 on host cell surfaces (Hahn, 1997; Gupta et al., 1994; Ramphal et al., 1991). Recently type IV pili of Neisseria gonorrhoeae and P. aeruginosa were shown to bind with high affinity to DNA (Aas et al., 2002; van Schaik et al., 2005), and because extracellular DNA has been shown to be part of the extracellular matrix material in P. aeruginosa biofilms (Whitchurch, et al., 2002; Nemoto et al., 2003; Matsukawa and Greenberg, 2004; Allesen-Holm et al., 2006), type IV pili might act as crosslinkers between the cells and the extracellular DNA matrix. Sauer and Camper (2001) presented evidence that expression of the major structural component of type IV pili, PilA, is downregulated in the initial stages of biofilm formation, but upregulated in the later stages of biofilm development. Type 1 fimbriae-like organelles are encoded by various members of the enterobacteria and they are believed to play an important role during pathogenesis of some of these organisms (Abraham et al., 1988). Each bacterial cell typically carries 100–500 type 1 fimbriae

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on its surface (Hahn et al., 2002). Type 1 fimbriae consist primarily of the structural protein FimA, but several auxiliary proteins are necessary for transport and assembly of the structural proteins (Klemm 1984; Klemm and Christiansen 1990; Klemm 1992). A minor component of type 1 fimbriae is the mannose specific adhesin FimH which is responsible for the ability of type 1 fimbriae to bind to eukaryotic cells (Krogfelt et al., 1990). Evidence has been presented that type 1 fimbriae play a role in the formation of E. coli biofilms at the air–liquid interface in static liquid cultures (Duguid et al., 1966; Pratt and Kolter 1998). he formation of these pellicles was inhibited by addition of mannose derivatives, suggesting that the process was dependent on the adhesive functions of the FimH adhesin (Old and Duguid, 1970; Harris et al., 1990). Type 1 fimbriae have also been associated with increased biofilm formation by E. coli in flow-chamber systems (Schembri and Klemm 2001). hese observations suggest that type 1 fimbriae, besides mediating binding to eukaryotic cell surfaces, may also act as matrix components in biofilms. Mutations in genes encoding P. aeruginosa cell appendages termed Cup fimbria were shown to affect the ability of the cells to attach to surfaces in microtiter trays (Vallet et al., 2001). Evidence is accruing that Cup fimbriae in addition to their role in initial biofilm formation also play a role as cell-to-cell interconnecting compounds in mature biofilms. he sadARS genes code for a putative sensor histidine kinase and two response regulators, and mutations in any of these genes were shown to result in the formation of P. aeruginosa biofilms with an altered mature structure (Kuchma et al., 2005). In another study the sadARS genes (termed rocARS) were shown to regulate transcription of the CupB and CupC fimbriae through the action of the positive regulator RocA, and the negative regulator RocR, which contain an EAL domain (Kulasekara et al., 2005). D’Argenio and colleagues demonstrated that expression of CupA fimbriae was necessary for the formation of wrinkly colonies by a P. aeruginosa mutant, and that the GGDEF domain protein WspR was required for this phenotype (D’Argenio et al., 2002). It appears that the presence of conjugative pili may promote biofilm formation in E. coli. Ghigo (2001) observed that strains harboring and expressing conjugative plasmids displayed an increase in biofilm formation on Pyrex slides placed in microfermentors. In a flow-chamber system the presence of conjugative pili on the surface of E. coli K-12 cells was shown to be the critical biofilm matrix component whereas other known E. coli biofilm formation factors like Ag43 and type 1 fimbria were dispensable (Reisner et al., 2003). Even minor changes of the conjugative pili structure, such as those conferred by a deletion of the traX gene, resulted in either the formation of biofilms with altered spatial structure, or in a decrease in biofilm formation (Reisner et al., 2003). Curli were first identified in E. coli but subsequent studies have shown that they are also produced by Salmonella, Citrobacter and Enterobacter species (Prigent-Combart et al., 2000; Zogaj et al., 2003). Curli are thin amyloid-like structures protruding from the cell surface as a tangled amorphous matrix, and they may function as both cell-to-surface and cell-tocell adhesins (see Figure 4.2) (Vidal et al., 1998; Römling et al., 1998; Prigent-Combart et al., 2000). he CsgA protein is the primary structural component of curli and the CsgB protein is an important minor structural unit. Polymerization of curli occurs outside the cell in a process referred to as extracellular nucleation. his nucleation is dependant of the CsgB protein and mutant studies have shown that mixing of csgA and csgB mutants can

The Bioilm Matrix

Figure 4.2 Scanning electron micrographs showing curli-mediated adherence of E. coli cells to a surface (A) and to each other (B). Adapted from Environ. Microbiol. 2:450–464 with permission from Blackwell Publishing.

result in the precipitation of curli subunits on the surface of the csgA mutant, suggesting that the CsgA protein is freely diffusible in the extracellular surroundings. In addition to binding to abiotic surfaces and stimulating biofilm formation by mediating cell-to-cell adherence, curli have been demonstrated to interact with host cell exopolymers such as fibronectin (Olsen et al., 1989; Austin et al., 1998). Evidence has been provided that curli together with cellulose play a role for biofilm formation by members of the enterobacteria such as E. coli, Enterobacter and Citrobacter (Zogaj et al., 2003; Bokranz et al., 2005). In E. coli and S. enterica it has been shown that the coordinated expression of curli and cellulose is regulated through the transcriptional regulator CsgD and the GGDEF domain protein AdrA as described above (Prigent-Combaret et al., 2001; Römling et al., 2000; Garcia et al., 2004; Kader et al., 2006). Surface proteins In E. coli a group of surface proteins (e.g. Ag43, AIDA and TibA) termed self associating autotransporters or SAAT has been identified (Diderichsen 1980; Benz and Schmidt 1992; Lindenthal and Elsinghorst, 1999; Klemm et al., 2006). hese proteins all have sequence similarities and share common features such as promotion of cell aggregation and biofilm formation (Henderson et al., 1997; Danese et al., 2000). Cell-to-cell interconnection mediated by these surface proteins occurs by virtue of there self-recognizing nature (Klemm et al., 2004; Henderson et al., 1997). he TasA protein of Bacillus subtilis was recently recognized as a major component of the extracellular matrix surrounding B. subtilis cells during formation of biofilms at the liquid-air interface (Branda et al., 2006). Based on genetic and biochemical evidence it was demonstrated that biofilm formation, in addition to the TasA protein, required an exopolysaccharide component encoded by the epsA-O operon, and that the absence of both of these components led to the abolishment of B. subtilis biofilm formation (Branda et al., 2001; Branda et al., 2006). Inactivation of either component alone resulted in a residual biofilm matrix and extracellular complementation was possible through mixing of the two mutant

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strains. Evidence has been provided that the eps and tasA genes are coregulated through the B. subtilis regulator SinR (Chu et al., 2006). he exact function of TasA in the extracellular matrix of B. subtilis remains unknown, but in addition to its role in biofilm formation it appears to have broad-spectrum antibacterial activity (Stover and Driks, 1999), and seems to be part of the spore coat assembly and germination (Serrano et al., 1999). Lectins are characterized by affinity towards carbohydrate residues on host cell surfaces, but evidence has been provided that some lectins recognize carbohydrates in extracellular biofilm matrices and thereby promote cell-to-cell interconnection. he P. aeruginosa fucose specific LecB lectin has recently been demonstrated to be important for the development of flow-chamber grown biofilms (Loris et al., 2003; Tielker et al., 2005). Cell fractionation experiments suggested that LecB was exported and bound to the outer-membrane through interaction with fucose containing ligands (Tielker et al., 2005). Staining of P. aeruginosa biofilms with fluorescently labeled LecB protein confirmed the presence of lectin binding sites in these structures (Tielker et al., 2005). he galactophilic lectin, LecA, has also recently been shown to have a role in biofilm development by P. aeruginosa (Diggle et al., 2006). he search for factors involved in biofilm formation has revealed the widespread existence of a large group of high molecular weight surface proteins that share limited sequence homology but are characterized by extensive repeat structures (Lasa and Penadés, 2005). Although the repetition of domains is central to the grouping of these diverse proteins, this might only reflect a general requirement for such surface proteins to protrude from the bacterial surface. Repetitive protein sequences could reflect a number of structural units necessary for obtaining protrusion from the surface and could otherwise be unrelated to the biological function of the individual proteins of the group. In addition to the repetition of domains, these proteins share a number of functional characteristics such as promotion of cell aggregation, surface adhesion and biofilm formation. he protein family includes the biofilm-associated protein (Bap) of Staphylococcus aureus, the large adhesion protein (LapA) of P. fluorescens and P. putida, the biofilm associated protein (BapA) of Salmonella enterica, the enterococcal surface protein (Esp) of Enterococcus faecalis, and the AdhA adhesin of Burkholderia cenocepacia. In addition to these experimentally characterized proteins a large number of similar proteins are found in the genome databases of both environmental and medically relevant bacteria. he Bap protein was first described in a study with an S. aureus bovine mastitis isolate, and was found to be essential for biofilm formation by this organism (Cucarella et al., 2001). Bap was shown to promote both primary attachment to abiotic surfaces and cell-to-cell adhesion. In addition, evidence was presented that deletion of the bap gene was linked to a decreased accumulation of the major staphylococcal exopolysaccharide, PIA. Bap appears to be almost universally conserved in biofilm forming S. aureus isolates, and has also been linked to the pathogenesis of bovine isolates (Cucarella et al., 2004). Close homologs of Bap have been found in numerous other staphylococcal species among these S. epidermidis (Tormo et al., 2005a). Mutations in the bapA gene of S. enterica were shown to abolish biofilm formation, but the defect could be rescued by overexpression of curli (Latasa et al., 2005). he bapA gene is located next to genes encoding a putative type 1 transport system and the BapA protein

The Bioilm Matrix

might be exported to the surface in a process similar to that identified for the LapA protein of P. fluorescens (described below) (Lasa and Penadés, 2005). he expression of the BapA protein has been proposed to be co-regulated with the other two major components of the S. enterica matrix, curli and cellulose, through the CsgD regulatory protein (Latasa et al., 2005). In the case of the Esp protein of E. faecalis it was shown by in-frame deletions that only the non-repetitive N-terminal domain was required for enhancement of biofilm formation (Toledo-Arana et al., 2001; Tendolkar et al., 2004), and that a fusion between the N-terminal region and a heterologous anchoring protein was sufficient to restore biofilm formation (Tendolkar et al., 2005). Expression of esp in Lactococcus lactis or Enterobacter facium did not enhance biofilm formation, suggesting that Esp stimulates biofilm formation by E. faecalis through interaction with another component of E. faecalis such as a surface protein or an exopolymer (Tendolkar et al., 2005). In addition, an esp mutant was found to have decreased cell surface hydrophobicity compared to its isogenic wild type (Tendolkar et al., 2005); a phenomenon which has also been observed for other members of the Bap family. he connection between expression of Bap-like proteins and increased cell surface hydrophobicity is somewhat surprising since a high serine/threonine content and a low theoretical pI of these proteins suggest that they would be highly charged and soluble in an aqueous environment. his apparent paradox might be related to an interaction of Bap-like proteins with other matrix components such as polysaccharides. LapA (large adhesion protein A) was first identified by Espinosa-Urgel et al. (2000) in a study aimed at determining factors of P. putida which are involved in adhesion to corn seed. Among a number of mus (mutants unattached to seed) mutants one (mus-24) displayed a very severe corn adhesion defect. his mutant was also tested for adhesion to abiotic surfaces and was found to be severely defective in adhesion to polystyrene, polypropylene and borosilicate glass in both minimal and rich media. Co-inoculation of the wild type and the mus-24 strain in part rescued the adhesion defect of the mus-24 strain, suggesting that the adhesion factor was secreted or that coaggregation with the wild type occurred. A gene almost identical to the mus-24 gene was identified in a transposon mutant screen for attachment defective mutants in P. fluorescens by O’Toole and Kolter (1998a). he P. fluorescens protein was further characterized by Hinsa et al. (2003) who designated the protein LapA. he LapA protein was reported to have sequence similarities to the CshA adhesin of the oral bacterium Streptococcus gordonii (Hinsa et al., 2003). CshA has been shown to be essential for colonization of the oral cavity and to participate in coaggregation of S. gordonii with other oral bacteria (McNab et al., 1996). LapA is transported to the bacterial surface via an ABC transport system which is encoded by the lapEBC genes, and is analogous to the type 1 transporter associated with transport of the BapA protein of S. enterica. he LapE protein has been identified in earlier work by Buell and Anderson (1992) referred to as the AggA protein, and was shown to be necessary for adhesion of P. putida to plant roots. Since a lapE mutant does not display surface located LapA it seems likely that the phenotypes described by Buell and Anderson (1992) for the aggA mutant also applies to a lapA mutant. Further work indicated that the plant receptor might consist of carbohydrate moieties (Buell et al., 1993) suggesting that LapA might also facilitate adhesion to carbohydrate residues. he involvement of LapA in adhesion to both abiotic

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and biotic surfaces suggests a function as a general adhesion. So far, no conditions that rescue the biofilm formation defect of a lapA mutant have been reported, suggesting an important role in biofilm formation of P. putida and P. fluorescens under diverse conditions. Our unpublished experiments with P. putida suggest that LapA not only acts as a surface adhesin but also plays a role during the later stages of biofilm formation as a cell-to-cell adhesin. he GGDEF and EAL domain containing protein LapD appears to be necessary for the activity of the LapA protein, possibly through a mechanism involving transport of LapA to the cell surface (Hinsa and O’Toole, 2006). he Bap-type protein of Burkholderia cenocepacia, AdhA, was identified as an adhesin mediating binding to cytokeratin 13 filaments, which are expressed on the apical surface of injured tracheobronchial epithelial cells (Urban et al., 2005). It was found that in conjunction with cable pili, the AdhA protein was required for strong binding to the epithelial cells and for migration across the epithelium surface. Huber et al. (2002) identified mutants in B. cenocepacia that were impaired in biofilm formation in both microtiter trays and flowchambers. hese mutants (termed m13 and m15) had insertions in a gene that showed homology to the Bap protein of S. aureus, and surface protein extracts from them showed that they were missing a 22 kDa protein. he mutants obtained by Huber et al. (2002) were shown to have decreased surface hydrophobicity compared to that of the wild type. However, because the Huber et al. (2002) report does not contain sequence information no conclusive correlation between the studies in the two B. cenocephacia strains can be made. In Gram-positive bacteria a large group of proteins termed MSCRAMM proteins (microbial surface components that recognize adhesive matrix molecules) has been described (Patti et al., 1994). he large family of MSCRAMM proteins share many of the characteristics of the Bap-type protein family, but the functions have mostly been demonstrated in relation to adhesion to host factors such as fibronectin-, fibrinogen-, collagen-, and heparin-related polysaccharides, although this does not rule out a function in biofilm matrices as well. Extracellular DNA as matrix component in bioilms Because most, if not all, bacterial populations are accompanied by extracellular DNA (e.g. Lorenz and Wackernagel, 1994), and because most bacterial species bind to DNA (e.g. Dubnau, 1999), it appears that extracellular DNA may serve as a cell-to-cell interconnecting compound in many different biofilms. On top of a basal level of DNA release it appears that many bacteria, especially those that are able to develop natural competence, possess a specific DNA-release program. A correlation between DNA release and competence development has been established in many different bacteria including Streptococcus pneumoniae (Steinmoen et al., 2002), Bacillus subtilis (Lorenz et al., 1991), Acinetobacter calcoaceticus (Palmen and Hellingwerf, 1995), Neisseria gonorrhoeae (Dillard and Seifert, 2001), and Pseudomonas stutzeri (Stewart et al., 1983). In all these cases DNA release and competence development was shown to occur in late-log phase cultures, and in some of the cases it has been documented that competence development is regulated through a quorum-sensing mechanism (e.g. Pestova et al., 1996; Magnuson et al., 1994). Since many kinds of polymers can function as biofilm matrix material, it is difficult to conceive that bacteria should release large amounts of costly information material solely with the purpose of stabilizing biofilms.

The Bioilm Matrix

It is possible, that bacteria release DNA both in order to exchange genetic material, and in order to form and stabilize biofilms. he relatively long-lasting physical proximity of bacteria in biofilms enable the constituent cells to establish long term relationships with each other, and evidence has been presented that biofilms are optimal environments for transformation-based gene transfer (e.g. Li et al., 2001; Wang et al., 2002; Hendrickx et al., 2003). In the present chapter we limit the description of extracellular DNA in biofilms to P. aeruginosa as an example from the Proteobacteria and Streptococcus species as examples from the Gram-positive bacteria. Extracellular DNA and bioilm formation by P. aeruginosa Evidence for a role of extracellular DNA as cell-to-cell interconnecting compound in P. aeruginosa biofilms has been presented both for the P. aeruginosa PAO1 reference strain and for clinical P. aeruginosa isolates (Whitchurch et al., 2002; Nemoto et al., 2003). P. aeruginosa PAO1 biofilm formation in the wells of microtiter plates was attenuated by the presence of DNase I, and biofilm formation by P. aeruginosa PAO1 in flow-chambers was almost absent when the flow-chambers were perfused with medium containing DNase I (Whitchurch et al., 2002). In addition, young P. aeruginosa PAO1 biofilms, which had been grown in flow-chambers perfused with DNase-free medium, were dispersed rapidly after addition of DNase I to the flowing medium, whereas older P. aeruginosa PAO1 biofilms were not dispersed by DNase I treatment, suggesting that other components than extracellular DNA stabilizes older P. aeruginosa PAO1 biofilms (Whitchurch et al., 2002). Matsukawa and Greenberg (2004) investigated the composition of the extracellular matrix of mature P. aeruginosa PAO1 biofilms, and found that extracellular DNA by far was the most abundant polymer, although exopolysaccharide encoded by the psl genes appeared to be the most critical structural matrix component. In contrast to the finding that extracellular DNA is not the primary cell-to-cell interconnecting compound in mature P. aeruginosa PAO1 biofilms, Nemoto et al. (2003) found that mature biofilms formed by four different clinical P. aeruginosa isolates could be dispersed by DNase treatment, suggesting that extracellular DNA is the critical matrix component in mature biofilms formed by these P. aeruginosa strains. Long before biofilms became a central research area Murakawa (1973a,b) conducted a study to characterize extracellular “slime” produced by P. aeruginosa. he chemical composition of slimes from 20 clinical P. aeruginosa isolates was investigated, and it was found that slimes from 18 strains consisted primarily of DNA, while two strains with a mucoid phenotype produced slimes composed primarily of polyuronic acid (which most likely was alginate). Figure 4.3 visualizes the extracellular DNA matrix in a flowchamber-grown P. aeruginosa PAO1 biofilm. Evidence has been presented that P. aeruginosa is capable of producing an extracellular DNase (Allesen-Holm et al., 2006) which might have a role during biofilm dispersal processes. PCR and Southern analysis have suggested that the extracellular DNA released from P. aeruginosa in biofilms and planktonic cultures is similar to whole-genome DNA (Steinberger and Holden, 2005; Allesen-Holm et al., 2006). In agreement, it has been shown that different chromosomal genes, including his+, leu+, and trp+, could be transferred by transformation of CaCl2-treated P. aeruginosa cells with extracellular DNA at the

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Figure 4.3 Horizontal confocal laser scanning microscope sections in a 2-day-old DDAOstained bioilm formed by Gfp-tagged P. aeruginosa PAO1. The images show the luorescent bacteria (A), the luorescent extracellular DNA (B), and an overlay of the two (C). Reproduced from Mol. Microbiol. 59:1114–1128 with permission from Blackwell Publishing.

same frequencies as when transformation was done with an equivalent amount of purified intracellular DNA (Hara et al., 1981; Muto and Goto, 1986). A basal level of extracellular DNA present in P. aeruginosa PAO1 biofilms is evidently generated via a pathway which is not linked to quorum-sensing, whereas the generation of large amounts of extracellular DNA in P. aeruginosa biofilms evidently depends on the las, rhl and pqs quorum sensing systems (Allesen-Holm et al., 2006). he increased level of extracellular DNA in P. aeruginosa wild-type biofilms in comparison to P. aeruginosa lasIrhlI biofilms appears to be linked to quorum-sensing via a mechanism that results in lysis of a small subpopulation of the cells (Allesen-Holm et al., 2006). In support of a role of quorumsensing in cell lysis, D’Argenio et al. (2002) reported that mutants which overproduce the Pseudomonas quinolone signal (PQS) displayed high levels of autolysis, whereas mutants which could not produce PQS did not show autolysis. In addition, Heurlier et al. (2005) presented evidence that P. aeruginosa quorum-sensing mutants, unlike the wild type, did not undergo cell lysis in stationary phase cultures. Quinolone compounds have previously been shown to induce prophages in bacteria (Phillips et al., 1987; Froshauer et al., 1996), and recent studies by Webb et al. (2003) and Hentzer et al. (2004) have suggested that quorum-sensing regulated DNA release might be linked to phage induction in biofilms. In support of a role of phage-mediated cell lysis in DNA release a P. aeruginosa fliMpilA mutant, which was reported not to undergo phage-mediated cell lysis (Webb et al., 2003), showed a defect in DNA release (Allesen-Holm et al., 2006). However, membrane vesicles produced by P. aeruginosa might also have a role in DNA release. P. aeruginosa releases membrane vesicles which have bacteriolytic effects and contain DNA (Kadurugamuwa and Beveridge, 1996; Renelli et al., 2004). Extracellular DNA might be released either from vesicles that eventually lyse, or through the bacteriolytic activity of the vesicles which might lyse a small subpopulation of the P. aeruginosa cells. Recently it was shown that PQS is necessary for vesicle formation in P. aeruginosa (Mashburn and Whiteley, 2005), and evidence was presented that type IV pili and flagella are necessary for quorum sensing in P. aeruginosa (Hassett, 2005). he involvement of PQS, type IV pili, and flagella in DNA release, therefore, could be consistent with a role of vesicles in the generation of extracellular DNA in P. aeruginosa biofilms.

The Bioilm Matrix

he extracellular DNA appears to be organized in distinct patterns in P. aeruginosa biofilms (Allesen-Holm et al., 2006). In 4-day-old flow-chamber-grown P. aeruginosa biofilms, which contain mushroom-shaped structures, the extracellular DNA was located primarily in the stalk-portion of the mushroom-shaped structures with the highest concentration in the outer parts of the stalks forming a border between the stalk-subpopulation and the cap-subpopulation (Allesen-Holm et al., 2006). he finding that biofilms formed by wildtype P. aeruginosa contained the highest concentration of extracellular DNA in the stalkportion of the mushroom-shaped structures is in agreement with a study showing that the expression of lasI and rhlI in P. aeruginosa biofilms was highest in the portion of the biofilm closest to the substratum (DeKievit et al., 2001). In addition it was shown that synthesis of rhamnolipid, an established quorum-sensing regulated process (Ochsner and Reiser, 1995; Pearson et al., 1997), occurs primarily in the stalks of the mushroom-shaped structures in P. aeruginosa biofilms (Lequette and Greenberg, 2005). Evidence has been presented that the formation of the mushroom-shaped structures in glucose-grown P. aeruginosa biofilms occurs in a sequential process involving a non-motile bacterial subpopulation that forms the stalks by growth in certain foci of the biofilm, and a migrating bacterial subpopulation which subsequently forms the mushroom caps via a process that requires type IV pili (Klausen et al., 2003a). It is currently not understood how the migration of the motile cells is coordinated so that they form mushroom caps. However, because type IV pili bind to DNA (Aas et al., 2002; van Schaik et al., 2005), it is tempting to speculate that the high concentration of extracellular DNA on the outer parts of the mushroom stalks might cause accumulation of the migrating bacteria, which in combination with bacterial growth, might result in the formation of the mushroom caps. In agreement with this suggestion, type IV pili-mediated migration of Myxobacteria during fruiting body formation has been shown to depend on the presence of exopolymer material (Lu et al., 2005). he extracellular DNA in P. aeruginosa biofilms appears to have a stabilizing effect, as mature P. aeruginosa PAO1 biofilms which were pre-treated with DNase I were more susceptible to SDS treatment than biofilms which were not pre-treated with DNase I (Allesen-Holm et al., 2006). P. aeruginosa colonizes the lungs of cystic fibrosis (CF) patients and is a major cause of lung deterioration, health decline, and death of these patients (Høiby, 2002). Several studies have shown that P. aeruginosa forms biofilms in the CF lung (e.g. Lam et al., 1980; Baltimore et al., 1989; Worlitzsch et al., 2002; Høiby, 2002), and the biofilm mode of growth is considered the major reason that these bacteria can not be eradicated by host defenses or antibiotic treatment (Costerton et al., 1999). CF lungs evidently contain large amounts of extracellular DNA from necrotized neutrophils (Lethem et al., 1990), and evidence has been presented that extracellular actin-DNA filaments can provide a matrix for biofilm formation by P. aeruginosa (Walker et al., 2005). In another study the presence of extracellular DNA was shown to be important for P. aeruginosa biofilm formation in artificial CF sputum medium (Sriramulu et al., 2005). In addition to a role of extracellular DNA, it was reported that biofilm formation in artificial CF sputum medium depended on the presence of amino acids (Sriramulu et al., 2005). Evidence has been presented that P. aeruginosa, in part due to the presence of a high level of aromatic amino acids, produces large amounts of PQS when it is present in CF lungs (Collier et al., 2002; Palmer et al., 2005). Because PQS evidently plays a role in DNA release from P. aeruginosa (Allesen-Holm et al., 2006),

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PQS-mediated release of DNA from the bacteria might play a role in biofilm formation in the CF lung. In further support of this possibility, the amino acid content of CF sputum has been shown to correlate with the severity of the disease (homas et al., 2000). Extracellular DNA and bioilm formation by streptococci Evidence that extracellular DNA plays a role in biofilm formation by streptococci is accumulating steadily. Competence mutants of S. mutans and S. gordonii were shown to be attenuated in biofilm formation (Loo et al., 2000; Li et al., 2002; Yoshida et al., 2002), and because these competence mutants were also deficient in generating extracellular DNA, it is possible that the biofilm formation defect was caused by a lack of extracellular DNA. Accordingly, the presence of DNase I was subsequently shown to attenuate biofilm formation by S. mutans and S. intermedius wild-type strains in the wells of microtiter plates (Petersen et al., 2004; 2005). Addition of exogenous quorum-sensing signal molecules to S. intermedius cultures was shown to promote biofilm formation, and simultaneous treatment with DNase I was shown to reverse the effect, suggesting that extracellular DNA was responsible for the increase in biofilm formation upon addition of signal molecules (Petersen et al., 2004). Among the genes regulated by quorum-sensing in streptococci are those required for DNA binding and uptake of extracellular DNA. Evidence has been presented, that proteins in streptococci which are necessary for binding and uptake of extracellular DNA play a role in biofilm formation. Petersen et al. (2005) reported that a comGB mutant of S. mutans, which is deficient in DNA binding but unaffected in quorumsensing signaling, showed reduced biofilm formation. In the presence of DNase I, biofilm formation by the S. mutans wild type was reduced to a level similar to that displayed by the comGB mutant. he comGB mutant was not impaired in DNA-release as growth in the presence of quorum-sensing signaling molecules promoted DNA-release from both the wild type and the comGB mutant. he addition of exogenous quorum-sensing signaling molecules to S. mutans wild-type cultures was shown to promote DNA-release and biofilm formation, and the simultaneous addition of DNase I reversed the effect, emphasizing the importance of extracellular DNA in the biofilm formation process (Petersen et al., 2005). Moreover, it was shown that addition of exogenous quorum-sensing signaling molecules to cultures of S. mutans comX, comE, and comD competence mutants did not promote DNArelease and biofilm formation. Evidence has been presented that S. pneumoniae is capable of producing an extracellular DNase (Moscoso and Claverys, 2004) which might have a role during biofilm dispersal processes. he extracellular DNA in streptococcal populations appears to be generated via lysis of a subpopulation of the cells (e.g. Steinmoen et al., 2002; 2003; Moscoso and Claverys, 2004; Shibata et al., 2005), and should therefore be similar to whole genome DNA. Release of DNA in S. pneumoniae populations was shown to involve cell lysis via the cell-wall hydrolases LytA, LytC, and CbpD (Steinmoen et al., 2003; Moscoso and Claverys, 2004; Guiral et al., 2005). Evidence has been provided that competent S. pneumoniae cells trigger lysis of S. pneumoniae sibling cells that are non-competent because they respond slower to the quorum-sensing signaling molecules (Steinmoen et al., 2003; Moscoso and Claverys, 2004; Guiral et al., 2005). he phenomenon evidently involves a system consisting of a bacteriocin (CibAB), its immunity factor (CibC), and the cell wall hydrolases (Guiral et al., 2005).

The Bioilm Matrix

Competent cells are immune to the bacteriocin, presumably because they also produce the immunity factor, but the bacteriocin induces lysis of non-competent cells via a process that depends on the cell-wall hydrolases. A similar bacteriocin-based system appears to operate during DNA release from S. mutans populations (Kreth et al., 2005; van der Ploeg, 2005) and from S. sanguis populations (Schlegel and Slade, 1973). Because biofilms most often contain numerous microenvironments, the streptococcal DNA-release mechanism described above will most likely lead to a stratified distribution of extracellular DNA in biofilms formed by streptococci. However, the investigation of structural biofilm development by the streptococci is not as advanced as in the case of P. aeruginosa, and at present the spatial organization of the extracellular DNA in streptococcal biofilms has not been investigated. Competence-triggered DNA-release from streptococci has been proposed to ensure coordination in time and space between DNA-release and uptake, thus favoring genetic exchange (e.g. Steinmoen et al., 2002; 2003). However, the finding that DNA-release in S. pneumoniae cultures continued a long time after competence had disappeared suggested that genetic exchange is not the only purpose of competence-triggered cell lysis (Moscoso and Claverys, 2004). Competence-triggered lysis of streptococcal cells could be important for the release of virulence factors such as pneumolysin, (lipo-) teichoic acid, and DNA (Guiral et al., 2005). Because streptococci often are involved in biofilm-related infections such as those occurring in middle ears or lungs (e.g. S. pneumoniae) or on teeth (e.g. S. mutans), it is possible that the extracellular DNA plays a role in stabilizing medically relevant streptococcal biofilms. In agreement with this possibility the comB, comD, lytA, and cbpD genes have all been implicated in the virulence of S. pneumoniae ( Jedrzejas, 2001; Bartilson et al., 2001; Lau et al., 2001; Hava et al., 2003). Concluding remarks he extracellular matrix is arguably the most critical component of biofilms as it constitutes the framework that holds the component cells together. Indeed, it may be argued that the difference between a planktonic bacterium and a biofilm bacterium basically is that the biofilm bacterium has upregulated its adhesiveness and produces one or more biofilm matrix components. On top of that of course, the micro-environmental conditions prevailing in the different parts of a biofilm during the different stages of biofilm development leads to the expression of distinct sets of genes in time and space. Many different compounds may function as extracellular biofilm matrix component. he emerging picture is that almost anything that can interconnect bacteria may function as a matrix component. Recent research in the field has provided some surprises. For example it has been found that mating pili and extracellular DNA can function as biofilm matrix components. By promoting biofilm formation, mating pili and extracellular DNA also create optimal environments for gene transfer via conjugation or transformation. In this way efficient gene transfer may be both a consequence and a cause of biofilm development. A single bacterial species can produce several different biofilm matrix components. Usually not all of the biofilm matrix components are expressed during biofilm formation in a particular environment, but it is anticipated that the capacity of bacteria to produce dif-

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ferent biofilm matrix components allows colonization of different niches through different biofilm development pathways. Although the matrix components used for biofilm development are diverse and vary amongst bacterial species, and in response to environmental cues, there may be common features underlying these factors. Recent work indicates that one common denominator to bacterial adhesiveness and biofilm matrix production may be regulatory proteins which contain GGDEF and/or EAL domains. hrough diguanylate cyclase or phosphodiesterase activity proteins with GGDEF or EAL domains control intracellular levels of c-di-GMP which acts as a second messenger and affects matrix production and the adhesiveness of the bacteria. It appears that in many cases, exemplified in this chapter by S. enterica, P. fluorescens, P. putida, and P. aeruginosa, GGDEF/EAL domain proteins regulate production (or transport) of both polysaccharide and protein components of the biofilm matrix. Biofilms may be very dynamic and contain migrating bacterial subpopulations, and it appears that the biofilm matrix may serve as a framework that the bacteria can migrate on. Evidence is emerging that exopolymer material is necessary for coordinated bacterial migration during structural development in P. aeruginosa biofilms and in myxobacterial biofilms. Continued research in the field will improve our understanding of the composition of extracellular matrices in biofilms formed by particular species under particular conditions, and will provide knowledge about the regulation of bacterial adhesiveness and matrix production and the transition between planktonic and biofilm lifestyles. Interference with the production of biofilm matrix components, or with the physical integrity of the biofilm matrix, are obvious therapeutic strategies for combating biofilm-based persistent infections. References Aas, F.E., Wolfgang, M., Frye, S., Dunham, S., Lovold, C., and Koomey, M. (2002). Competence for natural transformation in Neisseria gonorrhoeae, components of DNA binding and uptake linked to type IV pilus expression. Mol. Microbiol. 46, 749–760. Abraham, S.N., Sun, D., Dale, J.B., and Beachey, E.H. (1988). Conservation of the D-mannose-adhesion protein among type 1 fimbriated members of the family Enterobacteriaceae. Nature 336, 682–684. Allesen-Holm, M., Barken, K.B., Yang, L., Klausen, M., Webb, J.S., Kjelleberg, S., Molin, S., Givskov, M., and Tolker-Nielsen, T. (2006). A characterization of DNA-release in Pseudomonas aeruginosa cultures and biofilms. Mol. Microbiol. 59, 1114–1128. Amikam, D., and Galperin, M.Y. (2006). PilZ domain is part of the bacterial c-di-GMP binding protein. Bioinformatics 22, 3–6. Ausmees, N., Jonsson, H., Hoglund, S., Ljunggren, H., and Lindberg M. (1999). Structural and putative regulatory genes involved in cellulose synthesis in Rhizobium leguminosarum bv. trifolii. Microbiology 145, 1253–62. Austin, J.W., Sanders, G., Kay, W.W., and Collinson, S.K. (1998). hin aggregative fimbriae enhance Salmonella enteritidis biofilm formation. FEMS Microbiol. Lett. 162, 295–301. Bartilson, M., Marra, A., Christine, J., Asundi, J.S., Schneider, W.P., and Hromockyj, A.E. (2001). Differential fluorescence induction reveals Streptococcus pneumoniae loci regulated by competence stimulatory peptide. Mol. Microbiol. 39, 126–135. Baltimore, R.S., Christie, C.D.C., and Walker Smith, G.J. (1989). Immunohistopathologic localization of Pseudomonas aeruginosa in lungs from patients with cystic fibrosis. Am. Rev. Respir. Dis. 140, 1650–1661. Beenken, K.E., Blevins, J.S., and Smeltzer, M.S. (2003). Mutation of sarA in Staphylococcus aureus limits biofilm formation. Infect. Immun. 71, 4206–11.

The Bioilm Matrix

Benz, I., and Schmidt, M.A. (1992). Isolation and serologic characterization of AIDA-I, the adhesin mediating the diffuse adherence phenotype of the diarrhea-associated Escherichia coli strain 2787 (O126, H27). Infect. Immun. 60, 13–18. Blattner, F.R., Plunkett, G., Bloch, C.A., Perna, N.T., Burland, V., Riley, M., Collado-Vides, J., Glasner, J.D., Rode, C.K., Mayhew, G.F., Gregor, J., Davis, N.W., Kirkpatrick, H.A., Goeden, M.A., Rose, D.J., Mau, B., and Shao, Y. (1997). he complete genome sequence of Escherichia coli K-12. Science 277, 1453–74. Bokranz, W., Wang, X., Tschape, H., and Römling, U. (2005). Expression of cellulose and curli fimbriae by Escherichia coli isolated from the gastrointestinal tract. J. Med. Microbiol. 54, 1171–1182. Branda, S.S. Gonzalez-Pastor, J.E., Ben-Yehuda, S., Losick, R., and Kolter, R. (2001). Fruiting body formation by Bacillus subtilis. Proc. Natl. Acad. Sci. USA 98, 11621–11626. Branda, S.S., Chu, F., Kearns, D.B., Losick, R., and Kolter, R. (2006). A major protein component of the Bacillus subtilis biofilm matrix. Mol. Microbiol. 59, 1229–1238. Bomchil, N., Watnick, P., and Kolter, R. (2003). Identification and characterization of a Vibrio cholerae gene, mbaA, involved in maintenance of biofilm architecture. J. Bacteriol. 185, 1384–90. Brown, R.M., Willison, J.H., and Richardson, C.L. (1976). Cellulose biosynthesis in Acetobacter xylinum, visualization of the site of synthesis and direct measurement of the in vivo process. Proc. Natl. Acad. Sci. USA 73, 4565–9. Buchrieser, C., Rusniok, C., Frangeul, L., Couve, E., Billault, A., Kunst, F., Carniel, E., and Glaser, P. (1999). he 102-kilobase pgm locus of Yersinia pestis, sequence analysis and comparison of selected regions among different Yersinia pestis and Yersinia pseudotuberculosis strains. Infect. Immun. 67, 4851–61. Buell, C.R., and Anderson, A.J. (1992). Genetic analysis of the aggA locus involved in agglutination and adherence of Pseudomonas putida, a beneficial fluorescent pseudomonad. Mol. Plant Microbe Interact. 5, 154–162. Buell, C.R., Whetton, R., Tari, P., and Anderson, A.J. (1993). Characterization of cell surface properties in agglutinable and nonagglutinable mutants of Pseudomonas putida. Can. J. Microbiol. 39, 787–794. Caiazza, N.C., and O’Toole, G.A. (2003). Alpha-toxin is required for biofilm formation by Staphylococcus aureus. J. Bacteriol. 185, 3214–7. Casper-Lindley, C., and Yildiz, F.H. (2004). VpsT is a transcriptional regulator required for expression of vps biosynthesis genes and the development of rugose colonial morphology in Vibrio cholerae O1 El Tor. J. Bacteriol. 186, 1574–8. Chu, F., Kearns, D.B., Branda, S.S., Kolter, R., and Losick, R. (2006). Targets of the master regulator of biofilm formation in Bacillus subtilis. Mol. Microbiol. 59, 1216–1228. Collier, D.N., Anderson, L., McKnight, S.L., Noah, T.L., Knowles, M., Boucher, R., Schwab, U., Gilligan, P., and Pesci E.C. (2002). A bacterial cell to cell signal in the lungs of cystic fibrosis patients. FEMS Microbiol. Lett. 215, 41–46. Conlon, K.M., Humphreys, H., and O’Gara, J.P. (2002a). icaR encodes a transcriptional repressor involved in environmental regulation of ica operon expression and biofilm formation in Staphylococcus epidermidis. J. Bacteriol. 184, 4400–8. Conlon, K.M., Humphreys, H., and O’Gara, J.P. (2002b). Regulation of icaR gene expression in Staphylococcus epidermidis. FEMS Microbiol. Lett. 216, 171–7. Costerton, J.W., Stewart, P.S., and Greenberg, E.P. (1999). Bacterial biofilms, a common cause of persistent infections Science 284, 1318–1322. Cramton, S.E., Gerke, C., Schnell, N.F., Nichols, W.W., and Götz F. (1999). he intercellular adhesion (ica) locus is present in Staphylococcus aureus and is required for biofilm formation. Infect. Immun. 67, 5427–33. Cucarella, C., Solano, C., Valle, J., Amorena, B., Lasa, I., and Penades, J.R. (2001). Bap, a Staphylococcus aureus surface protein involved in biofilm formation. J. Bacteriol. 183, 2888–2896. Cucarella, C., Tormo, M.A., Ubeda, C., Trotonda, M.P., Monzon, M., Peris, C., Amorena, B., Lasa, I., and Penades, J.R. (2004). Role of biofilm-associated protein bap in the pathogenesis of bovine Staphylococcus aureus. Infect. Immun. 72, 2177–2185. Danese, P.N., Pratt, L.A., Dove, S.L., and Kolter, R. (2000). he outer membrane protein, antigen 43, mediates cell-to-cell interactions within Escherichia coli biofilms. Mol. Microbiol. 37, 424–432. Darby, C., Hsu, J.W., Ghori, N., and Falkow, S. (2002). Caenorhabditis elegans, plague bacteria biofilm blocks food intake. Nature 417, 243–4.

|

59

60

|

Pamp et al.

D’Argenio, D.A., Calfee, M.W., Rainey, P.B., and Pesci, E.C. (2002). Autolysis and autoaggregation in Pseudomonas aeruginosa colony morphology mutants. J. Bacteriol. 184, 6481–6489. Deinema, M.H., and Zevenhuizen, L.P. (1971). Formation of cellulose fibrils by gram-negative bacteria and their role in bacterial flocculation. Arch. Mikrobiol. 78, 42–51. DeKievit, T.R., Gillis, R., Marx, S., Brown, C., and Iglewski, B.H. (2001). Quorum-sensing genes in Pseudomonas aeruginosa biofilms, their role and expression patterns. Appl. Environ. Microbiol. 67, 1865–1873. Diderichsen, B. (1980). flu, a metastable gene controlling surface properties of Escherichia coli. J. Bacteriol. 141, 858–67. Diggle, S.P., Stacey, R.E., Dodd, C., Cámara, M., Williams, P., and Winzer, K. (2006). he galactophilic lectin, LecA, contributes to biofilm development in Pseudomonas aeruginosa. Environ. Microbiol. 8, 1095–1104. Dillard, J.P., and Seifert, H.S. (2001). A variable genetic island specific for Neisseria gonorrhoeae is involved in providing DNA for natural transformation and is found more often in disseminated infection isolates. Mol. Microbiol. 41, 263–77. Dobinsky, S., Kiel, K., Rohde, H., Bartscht, K., Knobloch, J.K., Horstkotte, M.A., and Mack, D. (2003). Glucose-related dissociation between icaADBC transcription and biofilm expression by Staphylococcus epidermidis, evidence for an additional factor required for polysaccharide intercellular adhesin synthesis. J. Bacteriol. 185, 2879–86. Dubnau, D. (1999). DNA uptake in bacteria. Annu. Rev. Microbiol. 53, 217–244. Duguid, J.P., Anderson E. S., and Campbell I. (1966). Fimbriae and adhesive properties in Salmonellae. J. Pathol. Bacteriol. 92, 107–138. Espinosa-Urgel, M., Salido, A., and Ramos, J.L. (2000). Genetic analysis of functions involved in adhesion of Pseudomonas putida to seeds. J. Bacteriol. 182, 2363–2369. Faruque, S.M., Biswas, K., Udden, S.M., Ahmad, Q.S., Sack, D.A., Nair, G.B., and Mekalanos, J.J. (2006). Transmissibility of cholera, in vivo-formed biofilms and their relationship to infectivity and persistence in the environment. Proc. Natl. Acad. Sci. USA 103, 6350–5 Fetherston, J.D., Schuetze, P., and Perry, R.D. (1992). Loss of the pigmentation phenotype in Yersinia pestis is due to the spontaneous deletion of 102 kb of chromosomal DNA which is flanked by a repetitive element. Mol. Microbiol. 6, 2693–704. Foster, T.J., and Höök M. (1998). Surface protein adhesins of Staphylococcus aureus. Trends Microbiol. 6, 484–8. Franklin, M.J., and Ohman, D.E. (1996). Identification of algI and algJ in the Pseudomonas aeruginosa alginate biosynthetic gene cluster which are required for alginate O acetylation. J. Bacteriol. 178, 2186– 2195. Friedman, L., and Kolter, R. (2004a). Genes involved in matrix formation in Pseudomonas aeruginosa PA14 biofilms. Mol. Microbiol. 51, 675–90. Friedman, L., and Kolter, R. (2004b). Two genetic loci produce distinct carbohydrate-rich structural components of the Pseudomonas aeruginosa biofilm matrix. J. Bacteriol. 186, 4457–65. Froshauer, S., Silvia, A.M., Chidambaram, M., Sharma, B., and Weinstock, G.M. (1996). Sensitization of bacteria to danofloxacin by temperate prophages. Antimicrob. Agents Chemother. 40, 1561–1563. Gal, M., Preston, G.M., Massey, R.C., Spiers, A.J., and Rainey, P.B. (2003). Genes encoding a cellulosic polymer contribute toward the ecological success of Pseudomonas fluorescens SBW25 on plant surfaces. Mol. Ecol. 12, 3109–3121. Garcia, B., Latasa, C., Solano, C., Garcia-del Portillo, F., Gamazo, C., and Lasa, I. (2004). Role of the GGDEF protein family in Salmonella cellulose biosynthesis and biofilm formation. Mol. Microbiol. 54, 264–277. Gerke, C., Kraft, A., Sussmuth, R., Schweitzer, O., and Götz, F. (1998). Characterization of the Nacetylglucosaminyltransferase activity involved in the biosynthesis of the Staphylococcus epidermidis polysaccharide intercellular adhesin. J. Biol. Chem. 273, 18586–93. Gerstel, U., and Römling, U. (2003). he csgD promoter, a control unit for biofilm formation in Salmonella typhimurium. Res. Microbiol. 154, 659–67 Ghigo, J.M. (2001). Natural conjugative plasmids induce bacterial biofilm development. Nature 412, 442–5. Gilbert, P., Evans, D.J., and Brown, M.R.W. (1993). Formation and dispersal of bacterial biofilms in vivo and in situ. J. Appl. Bacteriol. 74, 67S-78S.

The Bioilm Matrix

Giltner, C.L., van Schaik, E.J., Audette, G.F., Kao, D., Hodges, R.S., Hassett, D.J., and Irvin, R.T. (2006). he Pseudomonas aeruginosa type IV pilin receptor binding domain functions as an adhesin for both biotic and abiotic surfaces. Mol. Microbiol. 59, 1083–1096. Götz F. (2002). Staphylococcus and biofilms. Mol. Microbiol. 43, 1367–78. Govan, J.R., and Deretic, V. (1996). Microbial pathogenesis in cystic fibrosis, mucoid Pseudomonas aeruginosa and Burkholderia cepacia. Microbiol. Rev. 60, 539–74. Goymer, P., Kahn, S.G., Malone, J.G., Gehrig, S.M., Spiers, A.J., and Rainey, P.B. (2006). Adaptive divergence in experimental populations of Pseudomonas fluorescens. II. Role of the GGDEF regulator WspR in evolution and development of the wrinkly spreader phenotype. Genetics 173, 515–526. Gross, M., Cramton, S.E., Götz, F., and Peschel, A. (2001). Key role of teichoic acid net charge in Staphylococcus aureus colonization of artificial surfaces. Infect. Immun. 69, 3423–6. Guiral S., Mitchell T.J, Martin B., Claverys J.P. (2005). Competence-programmed predation of noncompetent cells in the human pathogen Streptococcus pneumoniae, genetic requirements. Proc. Natl. Acad. Sci. USA 102, 8710–8715. Gupta, S.K., Berk, R.S., Masinick, S., and Hazlett, L.D. (1994). Pili and lipopolysaccharide of Pseudomonas aeruginosa bind to the glycolipid asialo GM1. Infect. Immun. 62, 4572–4579 Hahn, H.P. (1997). he type-4 pilus is the major virulence-associated adhesin of Pseudomonas aeruginosa—a review. Gene 192, 99–108. Hahn, E., Wild, P., Hermanns, U., Sebbel, P., Glockshuber, R., Haner, M., Taschner, N., Burkhard, P., Aebi, U., and Muller, S.A. (2002). Exploring the 3D molecular architecture of Escherichia coli type 1 pili. J. Mol. Biol. 323, 845–857. Hammar, M., Arnqvist, A., Bian, Z., Olsen, A., and Normark S. (1995). Expression of two csg operons is required for production of fibronectin- and congo red-binding curli polymers in Escherichia coli K-12. Mol. Microbiol. 18, 661–70. Hammer, B.K., and Bassler, B.L. (2003). Quorum sensing controls biofilm formation in Vibrio cholerae. Mol. Microbiol. 50, 101–4. Hara, T., Aumayr, A., and Ueda, S. (1981). Genetic transformation of Pseudomonas aeruginosa with extracellular DNA. J. Gen. Appl. Microbiol. 27, 109–114. Harris, S.L., Elliott, D.A., Blake, M.C., Must, L.M., Messenger, M., and Orndorff, P.E. (1990). Isolation and characterization of mutants with lesions affecting pellicle formation and erythrocyte agglutination by type 1 piliated Escherichia coli. J. Bacteriol. 172, 6411–6418. Hassett, D.J. (2005). Tails, hairs, and speech in Pseudomonas aeruginosa. Pseudomonas, 10th international congress. Marseille, France, August 27–31, 2005 Oral presentation and Abstract S19. Hava, D.L., LeMieux, J., and Camilli, A. (2003). From nose to lung, the regulation behind Streptococcus pneumoniae virulence factors. Mol. Microbiol. 50, 1103–1110. Heilmann, C., Schweitzer, O., Gerke, C., Vanittanakom, N., Mack, D., and Götz, F. (1996). Molecular basis of intercellular adhesion in the biofilm-forming Staphylococcus epidermidis. Mol. Microbiol. 20, 1083–91. Henderson, I.R., Meehan, M., and Owen, P. (1997). Antigen 43, a phase-variable bipartite outer membrane protein, determines colony morphology and autoaggregation in Escherichia coli K-12. FEMS Microbiol. Lett. 149, 115–120. Hendrickx, L., Hausner, M., and Wuertz, S. (2003). Natural genetic transformation in monoculture Acinetobacter sp. BD413 biofilms. Appl. Environ. Microbiol. 69, 1721–1727. Hentzer, M., Eberl, L., and Givskov, M. (2004). Quorum sensing in Biofilms, Gossip in the slime world? In: Microbial Biofilms, eds M. Ghannoum and G. O’Toole, ASM Press, Washington, D.C. Heurlier, K., Denervaud, V., Haenni, M., Guy, L., Krishnapillai, V., and Haas, D. (2005). Quorum-sensing-negative (lasR) mutants of Pseudomonas aeruginosa avoid cell lysis and death. J. Bacteriol. 187, 4875–4883. Hickman, J.W., Tifrea, D.F., and Harwood, C.S. (2005). A chemosensory system that regulates biofilm formation through modulation of cyclic diguanylate levels. Proc. Natl. Acad. Sci. USA 102, 14422–7. Hinnebusch, B.J., Perry, R.D., and Schwan, T.G. (1996). Role of the Yersinia pestis hemin storage (hms) locus in the transmission of plague by fleas. Science 273, 367–70. Hinsa, S.M., and O’Toole, G.A. (2006). Biofilm formation by Pseudomonas fluorescens WCS365, a role for LapD. Microbiology 152, 1375–1383.

|

61

62

|

Pamp et al.

Hinsa, S.M., Espinosa-Urgel, M., Ramos, J.L., and O’Toole, G.A. (2003). Transition from reversible to irreversible attachment during biofilm formation by Pseudomonas fluorescens WCS365 requires an ABC transporter and a large secreted protein. Mol. Microbiol. 49, 905–918. Huber, B., Riedel, K., Kothe, M., Givskov, M., Molin, S., and Eberl, L. (2002). Genetic analysis of functions involved in the late stages of biofilm development in Burkholderia cepacia H111. Mol. Microbiol. 46, 411–426. Høiby N. (2002). Understanding bacterial biofilms in patients with cystic fibrosis, current and innovative approaches to potential therapies. J. Cyst. Fibros. 1, 249–254. Jackson, K.D., Starkey, M., Kremer, S., Parsek, M.R., and Wozniak, D.J. (2004). Identification of psl, a locus encoding a potential exopolysaccharide that is essential for Pseudomonas aeruginosa PAO1 biofilm formation. J. Bacteriol. 186, 4466–75. Jackson, D.W., Suzuki, K., Oakford, L., Simecka, J.W., Hart, M.E., and Romeo, T. (2002). Biofilm formation and dispersal under the influence of the global regulator CsrA of Escherichia coli. J. Bacteriol. 184, 290–301. Jäger, S., Mack, D., Rohde, H., Horstkotte, M.A., and Knobloch, JK. (2005). Disintegration of Staphylococcus epidermidis biofilms under glucose-limiting conditions depends on the activity of the alternative sigma factor sigmaB. Appl. Environ. Microbiol. 71, 5577–81. Jedrzejas, M.J. (2001). Pneumococcal virulence factors, structure and function. Microbiol. Mol. Biol. Rev. 65, 187–207. Jarrett, C.O., Deak, E., Isherwood, K.E., Oyston, P.C., Fischer, E.R., Whitney, A.R., Kobayashi, S.D., DeLeo, F.R., and Hinnebusch, B.J. (2004). Transmission of Yersinia pestis from an infectious biofilm in the flea vector. J. Infect. Dis. 190, 783–92. Jefferson, K.K., Pier, D.B., Goldmann, D.A., and Pier, G.B. (2004). he teicoplanin-associated locus regulator (TcaR) and the intercellular adhesion locus regulator (IcaR) are transcriptional inhibitors of the ica locus in Staphylococcus aureus. J. Bacteriol. 186, 2449–56. Jobling, M.G., and Holmes, R.K. (1997). Characterization of HapR, a positive regulator of the Vibrio cholerae HA/protease gene hap, and its identification as a functional homologue of the Vibrio harveyi luxR gene. Mol. Microbiol. 26, 1023–34. Kader, A., Simm, R., Gerstel, U., Morr, M., and Romling, U. (2006). Hierarchical involvement of various GGDEF domain proteins in rdar morphotype development of Salmonella enterica serovar Typhimurium. Mol. Microbiol. 60, 602–616. Kadurugamuwa, J.L., and Beveridge, T.J. (1996). Bacteriolytic effect of membrane vesicles from Pseudomonas aeruginosa on other bacteria including pathogens, conceptually new antibiotics. J. Bacteriol. 178, 2767–2774. Kimura, S., Chen, H.P., Saxena, I.M., Brown, R.M., and Itoh, T. (2001). Localization of c-di-GMP-binding protein with the linear terminal complexes of Acetobacter xylinum. J. Bacteriol. 183, 5668–74. Kies, S., Otto, M., Vuong, C., and Götz F. (2001). Identification of the sigB operon in Staphylococcus epidermidis, construction and characterization of a sigB deletion mutant. Infect. Immun. 69, 7933–6. Kirillina, O., Fetherston, J.D., Bobrov, A.G., Abney, J., and Perry, R.D. (2004). HmsP, a putative phosphodiesterase, and HmsT, a putative diguanylate cyclase, control Hms-dependent biofilm formation in Yersinia pestis. Mol. Microbiol. 54, 75–88. Klausen, M., Aaes-Jorgensen, A., Molin, S., and Tolker-Nielsen, T. (2003a). Involvement of bacterial migration in the development of complex multicellular structures in Pseudomonas aeruginosa biofilms. Mol. Microbiol. 50, 61–68. Klausen, M., Heydorn, A, Ragas, P, Lambertsen, L., Aaes-Jørgensen, A., Molin, S., and Tolker-Nielsen, T. (2003b). Biofilm formation by Pseudomonas aeruginosa wild-type, flagella, and type IV pili mutants. Mol. Microbiol. 48, 1511–1524. Klemm, P. (1984). he mA gene encoding the type-1 fimbrial subunit of Escherichia coli. Nucleotide sequence and primary structure of the protein. Eur. J. Biochem. 143, 395–399. Klemm, P. (1992). FimC, a chaperone-like periplasmic protein of Escherichia coli involved in biogenesis of type 1 fimbriae. Res. Microbiol. 143, 831–838. Klemm, P., and G. Christiansen (1990). he mD gene required for cell surface localization of Escherichia coli type 1 fimbriae. Mol. Gen. Genet. 220, 334–338. Klemm, P., Hjerrild, L., Gjermansen, M., and Schembri, M.A. (2004). Structure-function analysis of the self-recognizing Antigen 43 autotransporter protein from Escherichia coli. Mol. Microbiol. 51, 283–296.

The Bioilm Matrix

Klemm, P., Vejborg, R.M., Sherlock, O. (2006). Self-associating autotransporters, SAATs, Functional and structural similarities. Int. J. Med. Microbiol. (In press). Knobloch, J.K., Bartscht, K., Sabottke, A., Rohde, H., Feucht, H.H., and Mack, D. (2001). Biofilm formation by Staphylococcus epidermidis depends on functional RsbU, an activator of the sigB operon, differential activation mechanisms due to ethanol and salt stress. J. Bacteriol. 183, 2624–33. Knobloch, J.K., Jager, S., Horstkotte, M.A., Rohde, H., and Mack, D. (2004). RsbU-dependent regulation of Staphylococcus epidermidis biofilm formation is mediated via the alternative sigma factor sigmaB by repression of the negative regulator gene icaR. Infect. Immun. 72, 3838–48. Kovacikova, G., Lin, W., and Skorupski, K. (2005). Dual regulation of genes involved in acetoin biosynthesis and motility/biofilm formation by the virulence activator AphA and the acetate-responsive LysR-type regulator AlsR in Vibrio cholerae. Mol. Microbiol. 57, 420–33. Kreft, J.U. (2004). Biofilms promote altruism. Microbiology 150, 2751–2760. Kreth, J., Merritt, J., Shi, W., and Qi, F. (2005). Co-ordinated bacteriocin production and competence development, a possible mechanism for taking up DNA from neighbouring species. Mol. Microbiol. 57, 392–404. Krogfelt, K.A., Bergmans H., and Klemm P. (1990). Direct evidence that the FimH protein is the mannose-specific adhesin of Escherichia coli type 1 fimbriae. Infect. Immun. 58, 1995–1998. Kuchma, S.L., Connolly, J.P., and O’Toole, G.A. (2005). A three-component regulatory system regulates biofilm maturation and type III secretion in Pseudomonas aeruginosa. J. Bacteriol. 187, 1441–1454. Kulasekara, H.D., Ventre, I., Kulasekara, B.R., Lazdunski, A., Filloux, A., and Lory, S. (2005). A novel two-component system controls the expression of Pseudomonas aeruginosa fimbrial cup genes. Mol. Microbiol. 55, 368–380. Lam, J., Chan, R., Lam, K., and Costerton, J.W. (1980). Production of mucoid microcolonies by Pseudomonas aeruginosa within infected lungs in cystic fibrosis. Infect. Immun. 28, 546–556. Lasa, I., and Penadés, J.R. (2005). Bap, a family of surface proteins involved in biofilm formation. Res. Microbiol. 157, 99–107. Latasa, C., Roux, A., Toledo-Arana, A., Ghigo, J.M., Gamazo, C., Penades, J.R., and Lasa, I. (2005). BapA, a large secreted protein required for biofilm formation and host colonization of Salmonella enterica serovar Enteritidis. Mol. Microbiol. 58, 1322–1339. Lau GW, Haataja S, Lonetto M, Kensit SE, Marra A, Bryant AP, McDevitt D, Morrison DA, Holden DW. (2001). A functional genomic analysis of type 3 Streptococcus pneumoniae virulence. Mol. Microbiol. 40, 555–571. Lenz, D.H., Mok, K.C., Lilley, B.N., Kulkarni, R.V., Wingreen, N.S., and Bassler, B.L. (2004). he small RNA chaperone Hfq and multiple small RNAs control quorum sensing in Vibrio harveyi and Vibrio cholerae. Cell 118, 69–82. Lethem MI, James SL, Marriott C, Burke JF. (1990). he origin of DNA associated with mucus glycoproteins in cystic fibrosis sputum. Eur. Respir. J. 3, 19–23. Lequette, Y., and Greenberg, E.P. (2005). Timing and localization of rhamnolipid synthesis gene expression in Pseudomonas aeruginosa biofilms. J. Bacteriol. 187, 37–44. Li, Y.-H., Lau, P.C.Y., Lee, J.H., Ellen, R.P., and Cvitkovitch, D.G. (2001). Natural genetic transformation of Streptococcus mutans growing in biofilms. J. Bacteriol. 183, 897–908. Li Y-H, Tang N, Aspiras MB, Lau PC, Lee JH, Ellen RP, Cvitkovitch DG. (2002). A quorum-sensing signaling system essential for genetic competence in Streptococcus mutans is involved in biofilm formation. J Bacteriol. 184, 2699–2708. Lillard, J.W., Fetherston, J.D., Pedersen, L., Pendrak, M.L., and Perry, R.D. (1997). Sequence and genetic analysis of the hemin storage (hms) system of Yersinia pestis. Gene 193, 13–21. Lim, B., Beyhan, S., Meir, J., Yildiz, F.H. (2006). Cyclic-diGMP signal transduction systems in Vibrio cholerae, modulation of rugosity and biofilm formation. Mol. Microbiol. 60, 331–48. Lim, Y., Jana, M., Luong, T.T., and Lee, C.Y. (2004). Control of glucose- and NaCl-induced biofilm formation by rbf in Staphylococcus aureus. J. Bacteriol. 186, 722–9. Lindenthal, C., and Elsinghorst, E.A. (1999). Identification of a glycoprotein produced by enterotoxigenic Escherichia coli. Infect. Immun. 67, 4084–4091. Loo, C.Y., Corliss, D.A., and Ganeshkumar, N. (2000). Streptococcus gordonii biofilm formation, identification of genes that code for biofilm phenotypes. J. Bacteriol. 182, 1374–1382. Lorenz, M.G., Gerjets, D., and Wackernagel, W. (1991). Release of transforming plasmid and Chromosomal DNA from two cultured soil bacteria. Arch. Microbiol. 156, 319–326.

|

63

64

|

Pamp et al.

Lorenz, M.G., and Wackernagel, W. (1994). Bacterial gene transfer by natural genetic transformation in the environment. Microbiol. Rev. 58, 563–602. Loris, R., Tielker, D., Jaeger, K.E., and Wyns, L. (2003). Structural basis of carbohydrate recognition by the lectin LecB from Pseudomonas aeruginosa. J. Mol. Biol. 331, 861–870. Lu, A., Cho, K., Black, W.P., Duan, X.Y., Lux, R., Yang, Z., Kaplan, H.B., Zusman, D.R., and Shi, W. (2005). Exopolysaccharide biosynthesis genes required for social motility in Myxococcus xanthus. Mol Microbiol. 55, 206–220. Mack, D., Fischer, W., Krokotsch, A., Leopold, K., Hartmann, R., Egge, H., and Laufs, R. (1996). he intercellular adhesin involved in biofilm accumulation of Staphylococcus epidermidis is a linear beta-1, 6-linked glucosaminoglycan, purification and structural analysis. J. Bacteriol. 178, 175–83. Mack, D., Siemssen, N., and Laufs, R. (1992). Parallel induction by glucose of adherence and a polysaccharide antigen specific for plastic-adherent Staphylococcus epidermidis, evidence for functional relation to intercellular adhesion. Infect. Immun. 60, 2048–57. Magnuson, R., Solomon, J., and Grossman AD. (1994). Biochemical and genetic characterization of a competence pheromone from B. subtilis. Cell. 77, 207–216. Maira-Litran, T., Kropec, A., Abeygunawardana, C., Joyce, J., Mark, G. 3rd., Goldmann, D.A., and Pier, G.B. (2002). Immunochemical properties of the staphylococcal poly-N-acetylglucosamine surface polysaccharide. Infect. Immun. 70, 4433–40. Mashburn, L.M., and Whiteley, M. (2005). Membrane vesicles traffic signals and facilitate group activities in a prokaryote. Nature. 437, 422–425. Matsukawa, M., and Greenberg, E.P. (2004). Putative exopolysaccharide synthesis genes influence Pseudomonas aeruginosa biofilm development J. Bacteriol. 186, 4449–4456. Matthysse, A.G., White, S., and Lightfoot, R. (1995). Genes required for cellulose synthesis in Agrobacterium tumefaciens. J. Bacteriol. 177, 1069–75. Mattick, J.S. (2002). Type IV pili and twitching motility. Annu. Rev. Microbiol. 56, 289–314. Matz, C., McDougald, D., Moreno, A.M., Yung, P.Y., Yildiz, F.H., and Kjelleberg, S. (2005). Biofilm formation and phenotypic variation enhance predation-driven persistence of Vibrio cholerae. Proc. Natl. Acad. Sci. USA 102, 16819–24. Matz, C., and Kjelleberg, S. (2005). Off the hook—how bacteria survive protozoan grazing. Trends Microbiol. 13, 302–307. McNab, R., Holmes, A.R., Clarke, J.M., Tannock, G.W., and Jenkinson, H.F. (1996). Cell surface polypeptide CshA mediates binding of Streptococcus gordonii to other oral bacteria and to immobilized fibronectin. Infect. Immun. 64, 4204–4210. Morris, J.G. Jr, Sztein, M.B., Rice, E.W., Nataro, J.P., Losonsky, G.A., Panigrahi, P., Tacket, C.O., and Johnson JA. (1996). Vibrio cholerae O1 can assume a chlorine-resistant rugose survival form that is virulent for humans. J. Infect. Dis. 174, 1364–8. Moscoso, M., and Claverys, J.P. (2004). Release of DNA into the medium by competent Streptococcus pneumoniae, kinetics, mechanism and stability of the liberated DNA. Mol. Microbiol. 54, 783–794. Murakawa, T. (1973a). Slime production by Pseudomonas aeruginosa. III. Purification of slime and its physicochemical properties. Jpn. J. Microbiol. 17, 273–281. Murakawa, T. (1973b). Slime production by Pseudomonas aeruginosa. IV. Chemical analysis of two varieties of slime produced by Pseudomonas aeruginosa. Jpn. J. Microbiol. 17, 513–20. Muto, Y., and Goto, S. (1986). Transformation by extracellular DNA produced by Pseudomonas aeruginosa. Microbiol. Immunol. 30, 621–628. Napoli, C., Dazzo, F., and Hubbell, D. (1975). Production of cellulose microfibrils by Rhizobium. Appl. Microbiol. 30, 123–31. Nemoto, K., Hirota, K., Murakami, K., Taniguti, K., Murata, H., Viducic, D., and Miyake, Y. (2003). Effect of Varidase (streptodornase) on biofilm formed by Pseudomonas aeruginosa. Chemotherapy 49, 121–125. Nikolskaya, A.N., Mulkidjanian, A.Y., Beech, I.B., and Galperin, M.Y. (2003). MASE1 and MASE2, two novel integral membrane sensory domains. J Mol. Microbiol. Biotechnol. 5, 11–6. Novick, R.P. (2003). Autoinduction and signal transduction in the regulation of staphylococcal virulence. Mol. Microbiol. 48, 1429–49. Ochsner, U.A., and Reiser, J. (1995). Autoinducer-mediated regulation of rhamnolipid biosurfactant synthesis in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 92, 6424–8.

The Bioilm Matrix

Old, D.C., and Duguid J. P. (1970). Selective outgrowth of fimbriate bacteria in static liquid medium. J. Bacteriol. 103, 447–456. Olsen, A., Jonsson, A., and Normark, S. (1989). Fibronectin binding mediated by a novel class of surface organelles on Escherichia coli. Nature 338, 652–655. O’Toole, G.A., and Kolter, R. (1998a). Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol. Microbiol. 30, 295–304. O’Toole, G.A., and Kolter, R. (1998b). Initiation of biofilm formation in Pseudomonas fluorescens WCS365 proceeds via multiple, convergent signalling pathways, a genetic analysis. Mol. Microbiol. 28, 449–461. Palmen, R., and Hellingwerf, K.J. (1995). Acinetobacter calcoaceticus liberates chromosomal DNA during induction of competence by cell lysis. Curr. Microbiol. 30, 7–10. Palmer, K.L., Mashburn, L.M., Singh, P.K., and Whiteley, M. (2005). Cystic fibrosis sputum supports growth and cues key aspects of Pseudomonas aeruginosa physiology. J. Bacteriol. 187, 5267–77. Pane-Farre, J., Jonas, B., Forstner, K., Engelmann, S., and Hecker M. (2006). he sigma (B) regulon in Staphylococcus aureus and its regulation. Int. J. Med. Microbiol. 296, 237–258. Patti, J.M., Allen, B.L., McGavin, M.J., and Höök, M. (1994). MSCRAMM-mediated adherence of microorganisms to host tissues. Annu. Rev. Microbiol. 48, 585–617. Pearson, J.P., Pesci, E.C., and Iglewski B. H. (1997). Roles of Pseudomonas aeruginosa las and rhl quorum-sensing systems in control of elastase and rhamnolipid biosynthesis genes. J. Bacteriol. 179, 5756–5767. Pendrak, M.L., and Perry, R.D. (1993). Proteins essential for expression of the Hms+ phenotype of Yersinia pestis. Mol. Microbiol. 8, 857–64. Perry, R.D., Pendrak, M.L., and Schuetze, P. (1990). Identification and cloning of a hemin storage locus involved in the pigmentation phenotype of Yersinia pestis. J. Bacteriol. 172, 5929–37. Pestova, E.V., Havarstein, L.S., and Morrison, D.A. (1996). Regulation of competence for genetic transformation in Streptococcus pneumoniae by an auto-induced peptide pheromone and a two-component regulatory system. Mol. Microbiol. 21, 853–862. Petersen, F.C., Tao, L., Scheie, A.A. (2005). DNA binding-uptake system, a link between cell-to-cell communication and biofilm formation. J. Bacteriol. 187, 4392–4400. Petersen, F.C., Pecharki, D., and Scheie, A.A. (2004). Biofilm mode of growth of Streptococcus intermedius favored by a competence-stimulating signaling peptide. J. Bacteriol. 186, 6327–6331. Phillips, I., Culebras, E., Moreno, F., and Baquero, F. (1987). Induction of the SOS response by new 4-quinolones. J. Antimicrob. Chemother. 20, 631–638. van der Ploeg, J.R. (2005). Regulation of bacteriocin production in Streptococcus mutans by the quorumsensing system required for development of genetic competence. J. Bacteriol. 187, 3980–3989. Pratt, L.A., and Kolter, R. (1998). Genetic analysis of Escherichia coli biofilm formation, roles of flagella, motility, chemotaxis and type I pili. Mol. Microbiol. 30, 285–293. Prigent-Combaret, C., Brombacher, E., Vidal, O., Ambert, A., Lejeune, P., Landini, P., Dorel, C. (2001). Complex regulatory network controls initial adhesion and biofilm formation in Escherichia coli via regulation of the csgD gene. J. Bacteriol. 183, 7213–7223. Prigent-Combaret, C., Prensier, G., Le hi, T.T., Vidal, O., Lejeune, P., Dorel, C. (2000). Developmental pathway for biofilm formation in curli-producing Escherichia coli strains, role of flagella, curli and colanic acid. Environ. Microbiol. 2, 450–464. Rachid, S., Ohlsen, K., Wallner, U., Hacker, J., Hecker, M., and Ziebuhr W. (2000a). Alternative transcription factor sigma (B) is involved in regulation of biofilm expression in a Staphylococcus aureus mucosal isolate. J. Bacteriol. 182, 6824–6. Rachid, S., Ohlsen, K., Witte, W., Hacker, J., and Ziebuhr W. (2000b). Effect of subinhibitory antibiotic concentrations on polysaccharide intercellular adhesin expression in biofilm-forming Staphylococcus epidermidis. Antimicrob. Agents Chemother. 44, 3357–63. Rainey P.B, and Rainey K. (2003). Evolution of cooperation and conflict in experimental bacterial populations. Nature. 425, 72–74. Rainey, P.B., and Travisano, M. (1998). Adaptive radiation in a heterogeneous environment. Nature 394, 69–72. Ramphal, R., Carnoy, C., Fievre, S., Michalski, J.C., Houdret, N., Lamblin, G., Strecker, G., and Roussel, P. (1991). Pseudomonas aeruginosa recognizes carbohydrate chains containing type 1 (Gal beta 1–3GlcNAc) or type 2 (Gal beta 1–4GlcNAc) disaccharide units. Infect. Immun. 59, 700–4.

|

65

66

|

Pamp et al.

Rashid, M.H., Rajanna, C., Ali, A., and Karaolis, D.K. (2003). Identification of genes involved in the switch between the smooth and rugose phenotypes of Vibrio cholerae. FEMS Microbiol. Lett. 227, 113–9. Reisner, A., Haagensen, J.A., Schembri, M.A., Zechner, E.L., Molin, S. (2003). Development and maturation of Escherichia coli K-12 biofilms. Mol. Microbiol. 48, 933–946 Renelli, M., Matias, V., Lo, R.Y., and Beveridge, T.J. (2004). DNA-containing membrane vesicles of Pseudomonas aeruginosa PAO1 and their genetic transformation potential. Microbiology. 150, 2161–2169. Romby, P., Vandenesch, F., and Wagner, E.G. (2006). he role of RNAs in the regulation of virulence-gene expression. Curr. Opin. Microbiol. 9, 229–36. Romeo, T. (1998). Global regulation by the small RNA-binding protein CsrA and the non-coding RNA molecule CsrB. Mol. Microbiol. 29, 1321–30. Römling, U., Sierralta, W.D., Eriksson, K., and Normark, S. (1998). Multicellular and aggregative behaviour of Salmonella typhimurium strains is controlled by mutations in the agfD promoter. Mol. Microbiol. 28, 249–264. Römling, U., Rohde, M., Olsen, A., Normark, S, and Reinkoster, J. (2000). AgfD, the checkpoint of multicellular and aggregative behaviour in Salmonella typhimurium regulates at least two independent pathways. Mol. Microbiol. 36, 10–23. Ross, P., Mayer, R., and Benziman, M. (1991). Cellulose biosynthesis and function in bacteria. Microbiol. Rev. 55, 35–58. Ross, P., Weinhouse, H., Aloni, Y., Michaeli, D., Weinberger-Ohana, P., Mayer, R. Braun, S., de Vroom, E., van der Marel, G.A., van Boom, J.H., and Benziman, M. (1987). Regulation of cellulose synthesis in Acetobacter xylinum by cyclic di-guanylic acid. Nature 325, 279–281 Sabnis, N.A., Yang, H., and Romeo, T. (1995). Pleiotropic regulation of central carbohydrate metabolism in Escherichia coli via the gene csrA. J. Biol. Chem. 270, 29096–104. Sanford, B.A., homas, V.L., Mattingly, S.J., Ramsay, M.A., and Miller, M.M. (1995). Lectin-biotin assay for slime present in in situ biofilm produced by Staphylococcus epidermidis using transmission electron microscopy (TEM). J. Ind. Microbiol. 15, 156–61. Sanford, B.A., de Feijter, A.W., Wade, M.H., and homas, V.L. (1996). A dual fluorescence technique for visualization of Staphylococcus epidermidis biofilm using scanning confocal laser microscopy. J. Ind. Microbiol. 16, 48–56. Sauer, K., and Camper, A.K. (2001). Characterization of phenotypic changes in Pseudomonas putida in response to surface-associated growth. J. Bacteriol. 183, 6579–6589. Saxena, I.M., Kudlicka, K., Okuda, K., and Brown, R.M. Jr. (1994). Characterization of genes in the cellulose-synthesizing operon (acs operon) of Acetobacter xylinum, implications for cellulose crystallization. J. Bacteriol. 176, 5735–52. van Schaik, E.J., Giltner, C.L., Audette, G.F., Keizer, D.W., Bautista, D.L., Slupsky, C.M., Sykes, B.D., and Irvin., R.T. (2005). DNA Binding, a novel function of Pseudomonas aeruginosa type IV pili. J. Bacteriol. 187, 1455–1464. Schembri, M.A., and Klemm, P.. (2001). Biofilm formation in a hydrodynamic environment by novel FimH variants and ramifications for virulence. Infect. Immun. 69, 1322–1328. Schlegel, R., and Slade, H.D. (1973). Properties of a Streptococcus sanguis (group H) bacteriocin and its separation from the competence factor of transformation. J. Bacteriol. 115, 655–661. Schramm, M., and Hestrin, S. (1954). Factors affecting production of cellulose at the air/liquid interface of a culture of Acetobacter xylinum. J. Gen. Microbiol. 11, 123–129. Schweizer, F., Jiao, H., Hindsgual, O., Wong, W., and Irvin, R.T. (1998). Interaction between the pili of Pseudomonas aeruginosa PAK and its carbohydrate receptor beta-D-GalNAc (1m4)beta-D-Gal. Can. J. Microbiol. 44, 307–311. Serrano, M., Zilhao, R., Ricca, E., Ozin, A.J., Moran, C.P. Jr, and Henriques, A.O. (1999). A Bacillus subtilis secreted protein with a role in endospore coat assembly and function. J. Bacteriol. 181, 3632–3643. Sheth, H.B., Lee, K.K., Wong, W.Y., Srivastava, G., Hindsgaul, O., Hodges, R.S., Paranchych, W., and Irvin, R.T. (1994). he pili of Pseudomonas aeruginosa strains PAK and PAO bind specifically to the carbohydrate sequence beta GalNAc (1–4)beta Gal found in glycosphingolipids asialo-GM1 and asialo-GM2. Mol. Microbiol. 11, 715–723. Shibata Y, Kawada M, Nakano Y, Toyoshima K, Yamashita Y. (2005). Identification and characterization of an autolysin-encoding gene of Streptococcus mutans. Infect. Immun. 73, 3512–3520.

The Bioilm Matrix

Shih, P.-C., and Huang, C.-T. (2002). Effects of quorum-sensing deficiency on Pseudomonas aeruginosa biofilm formation and antibiotic resistance. J. Antimicrob. Chemother. 49, 309–314. Simm, R., Morr, M., Kader, A., Nimtz, M., and Römling, U. (2004). GGDEF and EAL domains inversely regulate cyclic di-GMP levels and transition from sessility to motility. Mol. Microbiol. 53, 1123–34. Solano, C., Garcia, B., Valle, J., Berasain, C., Ghigo, J.M., Gamazo, C., and Lasa I. (2002). Genetic analysis of Salmonella enteritidis biofilm formation, critical role of cellulose. Mol. Microbiol. 43, 793–808. Spiers, A.J., Bohannon, J., Gehrig, S.M., and Rainey, P.B. (2003). Biofilm formation at the air-liquid interface by the Pseudomonas fluorescens SBW25 wrinkly spreader requires an acetylated form of cellulose. Mol. Microbiol. 50, 15–27. Spiers, A.J., Kahn, S.G., Bohannon, J., Travisano, M., and Rainey, P.B. (2002). Adaptive divergence in experimental populations of Pseudomonas fluorescens. I. Genetic and phenotypic bases of wrinkly spreader fitness. Genetics 161, 33–46. Spiers, A.J., and Rainey, P.B. (2005). he Pseudomonas fluorescens SBW25 wrinkly spreader biofilm requires attachment factor, cellulose fibre and LPS interactions to maintain strength and integrity. Microbiology 151, 2829–2839 Sriramulu DD, Lunsdorf H, Lam JS, Römling U. (2005). Microcolony formation, a novel biofilm model of Pseudomonas aeruginosa for the cystic fibrosis lung. J. Med. Microbiol. 54, 667–676. Steinberger, R.E., and Holden, P.A. (2005). Extracellular DNA in single- and multiple-species unsaturated biofilms. Appl. Environ. Microbiol. 71, 5404–5410. Steinmoen, H., Knutsen, E., and Håvarstein, L.S. (2002). Induction of natural competence in Streptococcus pneumoniae triggers lysis and DNA release from a subfraction of the cell population. Proc. Natl. Acad. Sci. USA 99, 7681–7686. Steinmoen, H., Teigen, A., and Havarstein, L.S. (2003). Competence-induced cells of Streptococcus pneumoniae lyse competence-deficient cells of the same strain during cocultivation. J. Bacteriol. 185, 7176–7183. Stewart, G.J., Carlson, C.A., and Ingraham, J.L. (1983). Evidence for an active role of donor cells in natural transformation of Pseudomonas stutzeri. J. Bacteriol. 156, 30–35. Stover, A.G., and Driks, A. (1999). Secretion, localization, and antibacterial activity of TasA, a Bacillus subtilis spore-associated protein. J. Bacteriol. 181, 1664–1672. Tendolkar, P.M., Baghdayan, A.S., and Shankar, N. (2005). he N-terminal domain of enterococcal surface protein, Esp, is sufficient for Esp-mediated biofilm enhancement in Enterococcus faecalis. J. Bacteriol. 187, 6213–6222. Tendolkar, P.M., Baghdayan, A.S., Gilmore, M.S., and Shankar, N. (2004). Enterococcal surface protein, Esp, enhances biofilm formation by Enterococcus faecalis. Infect. Immun. 72, 6032–6039. homas, S.R., Ray, A., Hodson, M.E., and Pitt, T.L. (2000). Increased sputum amino acid concentrations and auxotrophy of Pseudomonas aeruginosa in severe cystic fibrosis lung disease. horax. 55, 795–797. homas, V.L., Sanford, B.A., Moreno, R., and Ramsay, M.A. (1997). Enzyme-linked lectinsorbent assay measures N-acetyl-D-glucosamine in matrix of biofilm produced by Staphylococcus epidermidis. Curr. Microbiol. 35, 249–54. Tielker, D., Hacker, S., Loris, R., Strathmann, M., Wingender, J., Wilhelm, S., Rosenau, F., and Jaeger, K.E. (2005). Pseudomonas aeruginosa lectin LecB is located in the outer membrane and is involved in biofilm formation. Microbiology 151, 1313–1323. Tischler, A.D., and Camilli, A. (2004). Cyclic diguanylate (c-di-GMP) regulates Vibrio cholerae biofilm formation. Mol. Microbiol. 53, 857–69. Toledo-Arana, A., Valle, J., Solano, C., Arrizubieta, M.J., Cucarella, C., Lamata, M., Amorena, B., Leiva, J., Penades, J.R., and Lasa, I. (2001). he enterococcal surface protein, Esp, is involved in Enterococcus faecalis biofilm formation. Appl. Environ. Microbiol. 67, 4538–4545. Tormo, M.A., Knecht, E., Götz, F., Lasa, I., and Penades, J.R. (2005a). Bap-dependent biofilm formation by pathogenic species of Staphylococcus, evidence of horizontal gene transfer? Microbiology 151, 2465–2475. Tormo, M.A., Marti, M., Valle, J., Manna, A.C., Cheung, A.L., Lasa, I., and Penades, J.R. (2005b). SarA is an essential positive regulator of Staphylococcus epidermidis biofilm development. J. Bacteriol. 187, 2348–56. Urban, T.A., Goldberg, J.B., Forstner, J.F., and Sajjan, U.S. (2005). Cable pili and the 22-kilodalton adhesin are required for Burkholderia cenocepacia binding to and transmigration across the squamous epithelium. Infect. Immun. 73, 5426–5437.

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Valle, J., Toledo-Arana, A., Berasain, C., Ghigo, J.M., Amorena, B., Penades, J.R., and Lasa, I. (2003). SarA and not sigmaB is essential for biofilm development by Staphylococcus aureus. Mol. Microbiol. 48, 1075–87. Vallet, I., Olson, J.W., Lory, S., Lazdunski, A., Filloux, A. (2001). he chaperone/usher pathways of Pseudomonas aeruginosa, identification of fimbrial gene clusters (cup) and their involvement in biofilm formation. Proc. Natl. Acad. Sci. USA 98, 6911–6916. Vance, R.E., Zhu, J., and Mekalanos, J.J. (2003). A constitutively active variant of the quorum-sensing regulator LuxO affects protease production and biofilm formation in Vibrio cholerae. Infect. Immun. 71, 2571–6. Vasseur, P., Vallet-Gely, I., Soscia, C., Genin, S., and Filloux, A. (2005). he pel genes of the Pseudomonas aeruginosa PAK strain are involved at early and late stages of biofilm formation. Microbiology 151, 985–97. Vidal, O., R. Longin, C. Prigent-Combaret, C. Dorel, M. Hooreman, and P. Lejeune. (1998). Isolation of an Escherichia coli K-12 mutant strain able to form biofilms on inert surfaces, involvement of a new ompR allele that increases curli expression. J. Bacteriol. 180, 2442–9. Vuong, C., Gerke, C., Somerville, G.A., Fischer, E.R., and Otto, M. (2003). Quorum-sensing control of biofilm factors in Staphylococcus epidermidis. J. Infect. Dis. 188, 706–18. Vuong, C., Saenz, H.L., Götz, F., and Otto, M. (2000). Impact of the agr quorum-sensing system on adherence to polystyrene in Staphylococcus aureus. J. Infect. Dis. 182, 1688–93. Wai, S.N., Mizunoe, Y., Takade, A., Kawabata, S.I., and Yoshida, S.I. (1998). Vibrio cholerae O1 strain TSI-4 produces the exopolysaccharide materials that determine colony morphology, stress resistance, and biofilm formation. Appl. Environ. Microbiol. 64, 3648–55. Walker, T.S., Tomlin, K.L., Worthen, G.S., Poch, K.R., Lieber, J.G., Saavedra, M.T., Fessler, M.B., Malcolm, K.C., Vasil, M.L., and Nick, J.A. (2005). Enhanced Pseudomonas aeruginosa biofilm development mediated by human neutrophils. Infect. Immun. 73, 3693–3701. Wang, B.Y., Chi, B., and Kuramitsu, H.K. (2002). Genetic exchange between Treponema denticola and Streptococcus gordonii in biofilms. Oral Microbiol. Immunol. 2, 108–112. Wang, X., Dubey, A.K., Suzuki, K., Baker, C.S., Babitzke, P., and Romeo, T. (2005). CsrA post-transcriptionally represses pgaABCD, responsible for synthesis of a biofilm polysaccharide adhesin of Escherichia coli. Mol. Microbiol. 56, 1648–63. Wang, X., Preston, J.F. 3rd and Romeo, T. (2004). he pgaABCD locus of Escherichia coli promotes the synthesis of a polysaccharide adhesin required for biofilm formation. J. Bacteriol. 186, 2724–34. Watnick, P.I., and Kolter, R. (1999). Steps in the development of a Vibrio cholerae El Tor biofilm. Mol. Microbiol. 34, 586–95. Webb, J.S., hompson, L.S., James, S., Charlton, T., Tolker-Nielsen, T., Koch, B., Givskov, M., and Kjelleberg, S. (2003). Cell Death in Pseudomonas aeruginosa Biofilm Development. J. Bacteriol. 185, 4585–4592. Weilbacher, T., Suzuki, K., Dubey, A.K., Wang, X., Gudapaty, S., Morozov, I., Baker, C.S., Georgellis, D., Babitzke, P., and Romeo, T. (2003). A novel sRNA component of the carbon storage regulatory system of Escherichia coli. Mol. Microbiol. 48, 657–70. Whitchurch, C.B., Tolker-Nielsen, T., Ragas, P.C., and Mattick, J.S. (2002). Extracellular DNA is required for bacterial biofilm formation. Science. 295, 1487. Wong, H.C., Fear, A.L., Calhoon, R.D., Eichinger, G.H., Mayer, R., Amikam, D., Benziman, M., Gelfand, D.H., Meade, J.H., Emerick, A.W., et al. (1990). Genetic organization of the cellulose synthase operon in Acetobacter xylinum. Proc. Natl. Acad. Sci. USA 87, 8130–4. Worlitzsch D, Tarran R, Ulrich M, Schwab U, Cekici A, Meyer KC, Birrer P, Bellon G, Berger J, Weiss T, Botzenhart K, Yankaskas JR, Randell S, Boucher RC, Döring G. (2002). Effects of reduced mucus oxygen concentration in airway Pseudomonas infections of cystic fibrosis patients. J. Clin. Invest. 109, 317–325. Wozniak, D.J., Wyckoff, T.J., Starkey, M., Keyser, R., Azadi, P., O’Toole, G.A., Parsek, M.R. (2003). Alginate is not a significant component of the extracellular polysaccharide matrix of PA14 and PAO1 Pseudomonas aeruginosa biofilms. Proc. Natl. Acad. Sci. USA 100, 7907–12. Yamada, M., Ikegami, A., and Kuramitsu, H.K. (2005). Synergistic biofilm formation by Treponema denticola and Porphyromonas gingivalis. FEMS Microbiol. Lett. 250, 271–277. Yang, H., Liu, M.Y., and Romeo, T. (1996). Coordinate genetic regulation of glycogen catabolism and biosynthesis in Escherichia coli via the CsrA gene product. J. Bacteriol. 178, 1012–7.

The Bioilm Matrix

Yarwood, J.M., Bartels, D.J., Volper, E.M., and Greenberg, E.P. (2004). Quorum sensing in Staphylococcus aureus biofilms. J. Bacteriol. 186, 1838–50. Yildiz, F.H., Dolganov, NA., and Schoolnik, G.K. (2001). VpsR, a member of the response regulators of the two-component regulatory systems, is required for expression of vps biosynthesis genes and EPS (ETr)-associated phenotypes in Vibrio cholerae O1 El Tor. J. Bacteriol. 183, 1716–26. Yildiz, F.H., Liu, X.S., Heydorn, A., and Schoolnik, G.K. (2004). Molecular analysis of rugosity in a Vibrio cholerae O1 El Tor phase variant. Mol. Microbiol. 53, 497–515. Yildiz, F.H., and Schoolnik, G.K. (1999). Vibrio cholerae O1 El Tor, identification of a gene cluster required for the rugose colony type, exopolysaccharide production, chlorine resistance, and biofilm formation. Proc. Natl. Acad. Sci. USA 96, 4028–33. Yoshida A, Kuramitsu HK. (2002). Multiple Streptococcus mutans Genes Are Involved in Biofilm Formation. Appl. Environ. Microbiol. 68, 6283–6291. Ziebandt, A.K., Weber, H., Rudolph, J., Schmid, R., Höper, D., Engelmann, S., and Hecker M. (2001). Extracellular proteins of Staphylococcus aureus and the role of SarA and sigma B. Proteomics 1, 480–93. Zogaj, X., Bokranz, W., Nimtz, M., and Romling, U. (2003). Production of cellulose and curli fimbriae by members of the family Enterobacteriaceae isolated from the human gastrointestinal tract. Infect. Immun. 71, 4151–4158. Zogaj, X., Nimtz, M., Rohde, M., Bokranz, W., and Römling U. (2001). he multicellular morphotypes of Salmonella typhimurium and Escherichia coli produce cellulose as the second component of the extracellular matrix. Mol. Microbiol. 39, 1452–63. Zhu, J., and Mekalanos, J.J. (2003). Quorum sensing-dependent biofilms enhance colonization in Vibrio cholerae. Dev. Cell 5, 647–56. Zhu, J., Miller, M.B., Vance, R.E., Dziejman, M., Bassler, B.L., and Mekalanos J.J. (2002). Quorum-sensing regulators control virulence gene expression in Vibrio cholerae. Proc. Natl. Acad. Sci. USA 99, 3129–34.

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Cyclic di-GMP as an Intracellular Signal Regulating Bacterial Bioilm Formation

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John M. Dow, Yvonne Fouhy, Jean. Lucey, and Robert P. Ryan

Abstract Cyclic di-GMP is a novel second messenger in bacteria that was first described as an allosteric activator of cellulose synthase in Gluconacetobacter xylinus. It is now established that this nucleotide regulates a range of functions including developmental transitions, aggregative behavior, adhesion, biofilm formation and virulence in diverse bacteria. he level of cyclic di-GMP in bacterial cells is influenced by both synthesis and degradation. he GGDEF protein domain synthesizes cyclic di-GMP, whereas EAL and HD-GYP domains are involved in cyclic di-GMP hydrolysis. Bacterial genomes encode a number of proteins with GGDEF, EAL and HD-GYP domains. he majority of these proteins contain additional signal input domains, suggesting that their activities are responsive to environmental cues. An emerging theme is that high cellular levels of cyclic di-GMP promote biofilm formation and aggregative behavior whereas low cellular levels promote motility. he mechanism(s) by which cyclic di-GMP exerts its effects on these cellular functions is however poorly understood. Introduction he extracellular environment undoubtedly influences many aspects of bacterial behavior including the formation, maturation and dissolution of biofilms. An array of signal transduction systems links the sensing of specific environmental cues to appropriate alterations in bacterial physiology and/or gene expression. In some of these signal transduction mechanism(s), perception of a primary signal alters the level of a second intracellular signal also known as a second messenger. In this chapter we discuss the role of cyclic di-GMP (bis-(3a-5a)-cyclic di-guanosine monophosphate) (Figure 5.1) as a second messenger that is implicated in regulation of processes associated with biofilm formation and communal behavior in diverse bacteria. Cyclic di-GMP was originally described in 1987 as an allosteric regulator of cellulose synthesis in Acetobacter xylinum (now Gluconacetobacter xylinus) (Ross et al., 1987). It was subsequently shown that enzymes from G. xylinus involved in cyclic di-GMP synthesis and turnover contained two protein domains, GGDEF and EAL, of previously unknown function (Tal et al., 1998). Whole genome sequencing has revealed an abundance of GGDEF and EAL domain containing proteins across the majority of bacterial species (both Grampositive and Gram-negative). Most GGDEF/EAL domain proteins contain additional

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Figure 5.1 Structure of the second messenger bis-(3a-5a)-cyclic di-guanosine monophosphate (cyclic di-GMP).

signal input domains, suggesting that their activities are responsive to environmental cues. Importantly, molecular analysis of communal behavior and biofilm formation has uncovered roles for a number of proteins containing GGDEF and/or EAL domains in developmental transitions, aggregative behavior, adhesion, biofilm formation and virulence in a number of bacteria. Taken together these observations indicate that signaling systems involving cyclic di-GMP as a second messenger are of potential importance in the regulation of biofilm formation in many bacteria. In the following sections we review the discovery of cyclic di-GMP and the definition of the biochemical activities of protein domains involved in its synthesis and degradation. We go on to catalogue some proteins with an established role in biofilm or aggregative behavior and to consider the primary environmental cues to which these proteins may respond. Finally we address possible mechanisms for the poorly understood processes by which cyclic di-GMP may exert its influence on bacterial behavior and biofilm formation. he reader is also directed to several recent reviews of this area (D’Argenio and Miller, 2004; Jenal, 2004: Römling et al., 2005). The discovery of cyclic di-GMP through studies of bacterial cellulose biosynthesis he occurrence of cyclic di-GMP in bacteria was first revealed through studies of regulation of cellulose synthesis in Gluconacetobacter xylinus (formerly Acetobacter xylinum) (Ross et al., 1987). In this organism, cyclic di-GMP acts as a reversible and highly specific allosteric activator of cellulose synthase. In vitro the rate of cellulose synthesis is increased by up to 200 fold in the presence of cyclic di-GMP with half maximal activation at 0.35 MM. Original reports suggested that cyclic di-GMP bound to BcsB, the B-subunit of cellulose synthase (Mayer et al., 1991). Subsequent work showed cyclic di-GMP binding at high affinity (KD 20 nM) to a 200 kDa membrane-associated protein or protein complex that had no apparent cellulose synthase activity but that associated with the synthase complex (Weinhouse et al., 1997). It was proposed that this protein introduced an additional level of

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regulation into cellulose biogenesis in vivo, by preventing enzymatic degradation of the effector and by targeting its release to the synthase complex. However the 200 kDa protein(s) was not further characterized. Recent bioinformatic studies (Amikam and Galperin, 2006) have suggested that the true binding site for cyclic di-GMP is a C-terminal domain in the A-subunit of cellulose synthase (BcsA), and propose that the 200 kDa protein comprises either a BcsA dimer or a second form of cellulose synthase in which a single polypeptide contains both subunits (Saxena et al., 1995). his issue of targets for cyclic di-GMP binding will be discussed in detail later. Following the work in G. xylinus, cyclic di-GMP was shown to regulate bacterial cellulose synthesis in the plant pathogen Agrobacterium tumefaciens (Amikam and Benziman, 1989) and plant symbiotic Rhizobium spp. (Ausmees et al., 1999; 2001). What is the role of cellulose synthesis in G. xylinus? Association of cellulose microfibrils in this bacterium leads to the assembly of a multicellular pellicle in static liquid cultures. his allows the organism to colonize a niche that may be inaccessible to other microbes. Furthermore cells within the pellicle stay near the surface, a better-aerated environment with access to nutrients that may be important for the growth of these obligate aerobes (Cook and Colvin, 1980). It has also been suggested that cellulose production may contribute to the attachment of the bacteria to the decaying plant materials on which they grow, providing protection from competitors for the same nutrient source (Williams and Cannon, 1989). Bacterial cellulose production contributes to the initial attachment to plants of symbiotic Rhizobium spp. and the pathogen Agrobacterium tumefaciens (reviewed in Römling, 2002). Identiication of enzymes involved in cyclic di-GMP synthesis and degradation in Gluconacetobacter xylinus Biochemical studies revealed that the level of cyclic di-GMP in G. xylinus were controlled by the opposing action of the enzymes diguanylate cyclase (DGC), which catalyzes its formation from two molecules of GTP and phosphodiesterase A (PDEA) which catalyzes its degradation. In a reverse genetic approach, amino acid sequence information derived from purified DGC and PDEA was used to clone three cdg genes encoding isoforms of DGC and three pdeA genes encoding isoforms of PDEA (Tal et al., 1998). hese genes are organized in three unlinked operons, each containing a pdeA gene immediately upstream of a dgc gene (Tal et al., 1998). Each of the proteins contains a GGDEF domain (formerly known as DUF1, for domain of unknown function) and a C-terminal EAL domain (formerly known as DUF2). he GGDEF and EAL nomenclature relates to conserved amino acid motifs in these domains. In addition to GGDEF and EAL domains each of these proteins has an N-terminal PAS sensory input domain, which typically bind heme and flavin (Taylor and Zhulin, 1999). he GGDEF domain was first identified in PleD, a regulatory protein controlling swarmer-to-stalked-cell transition in Caulobacter crescentus (Hecht and Newton, 1995). he EAL domain was first described in BvgR, a repressor of virulence gene expression in Bordetella pertussis (Merkel et al., 1998a). he findings in G. xylinus were of seminal importance; they linked GGDEF and EAL domains to cyclic di-GMP turnover and gave the first indication that cyclic di-GMP signaling might be widespread in bacteria where it might control functions other than cellulose

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synthesis (Tal et al., 1998). In addition the findings posed a number of questions as to the regulation of cyclic di-GMP levels, in particular what are the specific roles of the GGDEF and EAL domains, and how are the activities of enzymes that have opposite effects on cyclic di-GMP levels regulated so as to give appropriate changes in the cellular level of the nucleotide. he association of GGDEF and EAL domains with cyclic di-GMP turnover was further strengthened by the later demonstration of cyclic di-GMP phosphodiesterase activity of the recombinant PdeA1 protein (Chang et al., 2001). The GGDEF and EAL domains catalyze cyclic di-GMP synthesis and degradation respectively Indirect evidence for the role of the GGDEF domain in cyclic di-GMP synthesis came from in silico studies indicating some structural conservation with the proposed nucleotidebinding loop of eukaryotic adenylyl cyclases (Pei and Grishin, 2001). his suggestion was supported by genetic experiments in which expression of dgc1 from A. xylinum as well as genes encoding proteins with GGDEF but no EAL domain from other bacteria were shown to complement a mutant of Rhizobium for defects in cellulose production (Ausmees et al., 2001). Subsequently it was shown that expression of genes encoding GGDEF proteins could increase the cellular levels of cyclic di-GMP in several bacteria (Paul et al., 2004; Simm et al., 2004; Tischler and Camilli, 2004). Direct evidence for the role of the GGDEF domain has been obtained by biochemical studies of the purified PleD regulatory protein (Paul et al., 2004) and of isolated GGDEF domains (Ryjenkov et al., 2005). In the latter study, the GGDEF domains selected came from proteins from different bacterial phyla (Ryjenkov et al., 2005; Römling et al., 2005). Each of these GGDEF domain proteins converted two molecules of GTP to cyclic di-GMP but had no activity with other nucleotides (Figure 5.2). he conservation of function of GGDEF domains from divergent bacteria has been further demonstrated by reciprocal complementation of hmsT which is involved in biofilm formation in Yersinia pestis and adrA, which is involved in cellulose synthesis in Salmonella enterica serovar typhimurium (Simm et al., 2005). In many of the experiments

Figure 5.2 The role of GGDEF, EAL and HD-GYP domains in the synthesis and degradation of cyclic di-GMP. Synthesis of cyclic di-GMP from two molecules of GTP is catalyzed by the GGDEF domain and is predicted to occur in two steps, with pppGpG as intermediate. Each step releases a molecule of inorganic pyrophosphate. The degradation of cyclic di-GMP to GMP also occurs via a two-step reaction, with the linear dinucleotide pGpG as intermediate. EAL domains characterized thus far characterized catalyze only the irst step, whereas the HDGYP domain catalyzes both steps. Other, perhaps non-speciic, phosphodiesterase enzymes may also convert pGpG to 5aGMP.

Cyclic di-GMP Signaling and Bioilm Formation

described, it was shown by site directed mutagenesis that the conserved GGDEF motif residues were critical for cyclic di-GMP synthesis. Indirect support for the role of the EAL domain in cyclic di-GMP degradation was provided by the demonstration that heterologous expression of genes encoding proteins with EAL but not GGDEF domains could reduce cellular levels of cyclic di-GMP (Simm et al., 2004; Tischler and Camilli 2004; 2005) and conversely that mutation of an EAL domain protein increased cellular cyclic di-GMP levels (Hisert et al., 2005). he EAL domain protein HmsP of Yersinia pestis was shown to possess activity against the model phosphodiesterase substrate bis-(p-nitrophenol) phosphate, an activity that was required for the negative regulation of biofilm formation (Bobrov et al., 2005). Direct biochemical evidence for the role of the EAL domain as a cyclic di-GMP phosphodiesterase has come from studies of the intact proteins as well as isolated EAL domains (Christen et al., 2005, Schmidt et al., 2005). Again in many of these cases mutational analysis indicated the essential role of the conserved EAL residues in enzymatic activity and regulation. he activities of GGDEF and EAL domains in cyclic di-GMP turnover are summarized in Figure 5.2. The biochemical conundrum of GGDEF–EAL domain fusions he definition of the biochemical functions of GGDEF and EAL domains presents a conundrum; what determines the activity of proteins such as the PDEA and DGC from G. xylinus, which contain both domains. One possible resolution to this paradox is that one of the two domains is non-functional (Schmidt et al., 2005). here is evidence that some GGDEF domains are enzymatically inactive and may act in a regulatory capacity. In a GGDEF-EAL domain protein from Caulobacter, binding of GTP to an enzymatically inactive GGDEF domain acts to regulate the activity of the protein in cyclic di-GMP hydrolysis (Christen et al., 2005). he same considerations may also apply to inactive EAL domains (Schmidt et al., 2005). A second resolution of the paradox could be that the proteins can have both activities but switch between states able to synthesize and hydrolyze cyclic diGMP. One possible mechanism could be related to the oligomerization state. Structural analysis of the PleD regulator suggests that the GGDEF domain acts in cyclic di-GMP synthesis as a dimer (Chan et al., 2004) whereas EAL activity is apparently independent of protein oligomerization (Schmidt et al., 2005). Regulation of the oligomerization state of the GGDEF-EAL proteins, perhaps influenced by the sensory input domains, may then serve to determine which activity is expressed. Phylogenetic analysis of the GGDEF/EAL domain proteins in Pseudomonas aeruginosa indicates that GGDEF domains from almost all proteins that do not have an EAL domain are related in a single class (family I) in which the variant GGEEF motif is found (Kulesekara et al., 2006). GGDEF domains from almost all proteins that also contain an EAL domain fall into two further classes, families II and III. his may suggest that the proteins with a GGDEF alone have evolved separately from those in which this domain is linked to an EAL domain. Family III sequences are poorly correlated with the consensus GGDEF sequence, suggesting that they may be enzymatically inactive (Kulesekara et al., 2006).

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The HD-GYP domain is a second cyclic di-GMP phosphodiesterase Bioinformatic studies have suggested that a third domain HD-GYP is also involved in cyclic di-GMP hydrolysis (Galperin et al., 1999; 2001). HD-GYP is a subgroup of the HD superfamily of metal dependent phosphohydrolases. he association of the HD-GYP domain with a CheY-like two-component receiver domain in many bacterial proteomes indicates a role in signaling (Galperin et al., 1999; 2001). A role for HD-GYP in cyclic diGMP hydrolysis was proposed based on an examination of the distribution and numbers of GGDEF, EAL and HD-GYP domains encoded by different bacterial genomes, where several genomes encode proteins with the GGDEF and HD-GYP domains but no EAL domain (Galperin et al., 1999; 2001; Galperin, 2005). In the plant pathogen Xanthomonas campestris, the HD-GYP domain regulator RpfG positively regulates synthesis of extracellular enzyme virulence factor and negatively regulates biofilm formation (Slater et al., 2000; Dow et al., 2003). Expression of genes encoding EAL domain proteins in the X. campestris rpfG mutant restored extracellular enzymes and blocked biofilm formation. In contrast expression of genes encoding a GGDEF domain protein in wild-type X. campestris gave a phenocopy of the rpfG mutant (Ryan et al., 2006). hese indirect observations were consistent with a role for the HD-GYP domain in cyclic di-GMP hydrolysis. his conclusion was supported by biochemical studies that demonstrated that the isolated domain could hydrolyze cyclic di-GMP to GMP via a linear intermediate (Figure 5.2). Mutation of the HD residues comprising the presumed catalytic diad of the HD-GYP domain abolishes both the regulatory activity and enzymatic activity against cyclic di-GMP (Ryan et al., 2006). Bacterial genomes encode multiple proteins with GGDEF, EAL, and HD-GYP domains Large-scale sequencing of bacterial genomes has revealed that GGDEF and EAL domains are highly abundant and widely distributed, although they are not found in archaea (Galperin, 2005). At the time of writing there were over 2400 GGDEF domains and over 1400 EAL domains in the Pfam protein family database. he HD-GYP domain is also widely distributed although slightly less abundant, with over 200 HD-GYP domains in over 70 genomes. Most bacterial genomes encode a number of proteins with these domains. For example, the Pseudomonas aeruginosa PAO1 proteome has 17 predicted proteins with a GGDEF but no EAL domain, 5 proteins with an EAL but no GGDEF domain, 16 proteins with both GGDEF and EAL domains, and 3 proteins with an HD-GYP domain (Galperin, 2005). Many GGDEF, EAL, and HD-GYP domain proteins have associated regulatory/sensory input domains As indicated above, many GGDEF, EAL and HD-GYP domain proteins have additional domains that may directly sense environmental cues (Zhulin et al., 2003). hese domains include PAS, which binds flavin or heme and may sense molecular oxygen or redox potential, GAF, which binds cyclic mononucleotides and other small molecular weight effectors and various membrane-associated or periplasmic domains that may be involved in sensing small

Cyclic di-GMP Signaling and Bioilm Formation

molecules (Table 5.1). As an example, PdeA1 of G. xylinus has a PAS-GAF-GGDEF-EAL domain structure (Tal et al., 1998; Chang et al., 2001). Binding of effectors to the sensory input domain is believed to affect the enzyme activity of the protein. A number of GGDEF, EAL and HD-GYP domain proteins contain a CheY-like REC domain (Table 5.1). hese proteins can be part of either two-component signal transduction systems or of modified chemotaxis systems. In both of these cases the environmental signal is presumably sensed by another element of the system, either a sensory histidine kinase or methyl-accepting chemotaxis protein. Signal transduction involves autophosphorylation of the sensory histidine kinase or of a CheA-like histidine kinase and subsequent phosphotransfer to the REC domain, which alters the activity of the enzymatic domain in cyclic di-GMP synthesis or degradation. Examples of proteins belonging to two component systems are PleD of C. crescentus and RpfG of X. campestris, whereas WspR of P. aeruginosa and P. fluorescens are part of chemotaxis-like signal transduction systems (Aldridge et al., 2003; Paul et al., 2004; Table 5.1 Selected signaling and regulatory domains that are found in association with GGDEF and/or EAL domains in bacterial proteins Domain

Proposed signaling or regulatory function

Cytoplasmic domains REC

CheY-homologous response regulator receiver domain, phosphorylated by sensory or CheA histidine kinases

PAS (PAC)

Oxygen, redox potential and light sensing

GAF

Binding of cyclic nucleotides, other small molecules

HAMP

Signal transduction through conformational changes?

Extracytoplasmic domains PBPb

Bacterial periplasmic substrate binding protein

Reg_Prop

Proteins with multiple Reg_Prop domains may form a beta-propeller structure, unknown function

CHASE

Extracellular/periplasmic sensory domain, also associated with cyclases and histidine kinases

Transmembrane domains 7TMR-DISMED2

Carbohydrate binding?

7TMR-DISM-7TM

Carbohydrate binding?

MHYT

Sensor of oxygen, CO or NO?

MASE1

Membrane-associated sensor

MASE2

Membrane-associated sensor

BLUF

Blue light sensor, FAD-dependent

More detailed description of domains can be found at the Pfam data base at http://pfam.wustl.edu

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Crossman and Dow, 2004; Goymer, 2002; Hickman et al., 2005; Table 5.1). In a number of cases, multiple sensory input domains are found, suggesting complex regulation of individual enzymes in response to a range of environmental cues. Published analyses of the domain organization of all GGDEF/EAL domain proteins from Salmonella typhimurium (Römling, 2002) and P. aeruginosa (Kulesekara et al., 2006) exemplify the diversity of domains within proteins with a proposed role in cyclic di-GMP signaling within a single organism. GGDEF, EAL, and HD-GYP domain proteins with a role in bioilm formation and dispersal Molecular genetic analysis has now implicated a number of proteins with GGDEF, EAL and HD-GYP domains, and by extension cyclic di-GMP, in the regulation of communal bacterial behavior including biofilm formation, aggregation and other multicellular phenotypes (Table 5.2). However it is clear that cyclic di-GMP can also regulate a range of other phenotypes including motility, virulence factor synthesis and cell differentiation (Merkel et al., 1998a,b; Hecht and Newton, 1995; Tischler et al., 2002; Tischler and Camilli, 2004, 2005; Simm et al., 2004; Huang et al., 2003). In a number of these cases, conserved residues within the GGDEF, EAL and HD-GYP domains have been shown to be required for regulation indicating a functional link to the enzymatic activity. Here we address cyclic diGMP regulation of biofilm formation and of the synthesis of biofilm matrix components. For more detailed discussion of the components of the extracellular matrix of biofilms, the reader is directed to Pamp et al., this volume.

Table 5.2 Proteins with GGDEF, EAL, and HD-GYP domains that are implicated in bioilm formation and aggregative behavior in bacteria

Protein

Organism

Domain organization

Regulated phenotype/function

Reference

DGC1,2,3

Gluconacetobacter xylinus

PASGGDEF-EAL

Cellulose synthesis

Tal et al., 1998

PDEA1,2,3

Gluconacetobacter xylinus

PASGGDEF-EAL

Cellulose synthesis

Tal et al., 1998

HmsT

Yersinia pestis

GGDEF

Positive regulation of bioilm formation, aggregation

Jones et al., 1999; Kirillina et al., 2004

HmsP

Yersinia pestis

TM-GGDEFEAL

Negative regulation of bioilm formation, aggregation

Kirillina et al., 2004; Bobrov et al., 2005

PleD

Caulobacter crescentus

CheY-CheY*- Attachment/ GGDEF dispersal; developmental transitions

Hecht and Newton, 1985; Aldridge et al., 2003

Cyclic di-GMP Signaling and Bioilm Formation

Table 5.2 continued

Protein

Organism

Domain organization

Regulated phenotype/function

Reference

WspR

Pseudomonas luorescens

CheYGGDEF

Wrinkled colonies, synthesis of modiied cellulose

Goymer, 2000; Spiers et al., 2003

WspR

Pseudomonas aeruginosa

CheYGGDEF

Wrinkled colonies, bioilm formation, regulation of polysaccharide synthesis operons

D’Argenio et al., 2002; Hickman et al., 2005

CelR2

Rhizobium leguminosarum bv. trifolii

CheYGGDEF

Cellulose production Ausmees et al., 1999

AdrA

Salmonella enterica bv. typhimurium

TM-GGDEF

Rdar colonies

Römling et al., 2000

ScrC

Vibrio parahaemolyticus

GGDEF-EAL

Rugose colonies, capsular polysaccharide synthesis

Boles and McCarter, 2002; Güvener and McCarter, 2003

PvrR

Pseudomonas aeruginosa

CheY-EAL

Repression of autoaggregation, phase variation, virulence

Drenkard and Ausubel, 2002; Kulesekara et al., 2006

MbaA

Vibrio cholerae

TM-HAMPGGDEF-EAL

Bioilm matrix architecture

Bomchil et al., 2003; Karatan et al., 2005

VieA

Vibrio cholerae

CheY-EALHTH

Repression of bioilm formation, activation of cholera toxin production

Tischler et al., 2002; Tischler and Camilli, 2005

RocS

Vibrio cholerae

PASGGDEF-EAL

Rugose colonies

Rashid et al., 2003

RpfG

Xanthomonas campestris

CheY-HDGYP

Repression of bioilm formation, activation of virulence factor synthesis

Slater et al., 2000; Dow et al., 2003

RocR

Pseudomonas aeruginosa

CheY-EVL

Antagonism of expression of cup genes encoding imbrial adhesins

Kulasekara et al., 2005

PPO165

Pseudomonas putida

TM-GGDEFEAL

Bioilm dispersal

Gjermansen et al., 2005

HTH: helix-turn-helix DNA binding domain; TM: transmembrane helix or helices; CheY*: inactive version of the CheY receiver domain.

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As in G. xylinus, the synthesis of cellulose or of an acetylated cellulose derivative has been shown to contribute to the multicellular behavior of a number of bacteria including Escherichia coli, Salmonella enterica bv. typhimurium, Rhizobium leguminosarum and Pseudomonas fluorescens (Ausmees et al., 1999; Zogaj et al., 2001; Solano et al., 2002; Spiers et al., 2003; Römling, 2005). hese multicellular behaviors are apparent through alterations in colony morphology and in some cases through staining by Congo Red (which binds to B-linked glucans like cellulose) to give phenotypes that have been termed rdar (for red, dry and rough), wrinkly spreader and rugose. In each of these cases a GGDEF domain protein has been implicated in positive regulation of the multicellular phenotype, consistent with the earlier work on G. xylinus that showed activation of cellulose synthesis by cyclic di-GMP. he rdar phenotype of Enterobacteriaceae depends not only on cellulose synthesis but also on the expression of thin aggregative fibers termed curli (Römling et al., 2000; Römling, 2005). he two-component regulator CsgD, also called AgfD, regulates transcription of an operon encoding structural components curli and of adrA, a gene that encodes a GGDEF domain protein that is membrane-associated. CsgD has no effect on the expression of genes encoding bacterial cellulose synthesis. he synthesis of the cellulose component of the extracellular matrix is thus presumably responsive to environmental cues at a least two levels; those recognized by the cognate histidine kinase for CsgD, which has not yet been identified, as well as those recognized by AdrA. he wrinkly spreader phenotype of Pseudomonas fluorescens depends upon activation of the CheY-GGDEF domain protein WspR, which is part of a chemotaxis-like sensory transduction system, and the consequent synthesis of cellulose that is modified by acetylation (Goymer, 2002; Spiers et al., 2003). he available evidence suggests that WspR is activated for cyclic di-GMP synthesis by phosphorylation of the CheY domain. Two classes of mutation give rise to the wrinkly spreader phenotype; those in genes encoding other elements of the sensory transduction pathway that result in enhanced or constitutive activation of WspR and those introducing amino acid alterations in WspR giving a “locked-on” conformation (Goymer, 2002). A homologous system is found in Pseudomonas aeruginosa, where mutations in wspF that likely result in constitutive phosphorylation of WspR give rise to aggregation and altered colony morphology (Hickman et al., 2005). here are however some important differences between the two bacteria in the mechanisms underlying alterations in colony morphology and aggregation. P. aeruginosa does not have bcs genes for cellulose biosynthesis. Instead two operons called psl and pel are involved in extracellular polysaccharide production and biofilm formation (Friedman and Kolter, 2004). In the wspF mutant expression of both of these operons is elevated, which may suggest that cyclic di-GMP levels can influence the transcription of genes involved in biofilm polysaccharide formation in P. aeruginosa, in contrast to its (presumed) post-translational effects on cellulose synthesis in P. fluorescens. hese effects on psl and pel gene expression are however only part of a much wider influence of elevated cyclic di-GMP levels on gene transcription; expression levels of at least 560 genes are affected by a wspF deletion (Hickman et al., 2005). Similar broad effects on transcription in response to elevated levels of cyclic di-GMP are seen in Escherichia coli (Méndez-Ortiz et al., 2006). Biofilm formation in P. aeruginosa is also mediated by a family of adherence factors termed CupA, CupB and CupC that are orthologues of the chaperone/usher family of

Cyclic di-GMP Signaling and Bioilm Formation

E. coli adhesins (Vallet et al., 2001). Expression of two of these clusters, cupB and cupC, is regulated by a variant of a classical two-component signal transduction pathway comprising three components RocS1, RocR and RocA1 (Kulasekara et al., 2005). RocS1 is a sensor kinase, RocA1 is a DNA binding response regulator that activates cup genes, and RocR acts as an antagonist of RocA1 activity. RocS1 can phosphorylate both RocR and RocA1. RocR comprises a response regulator receiver domain attached to a variant of the EAL domain in which the EAL motif is replaced by EVL. It is not known whether this variant is still active in hydrolysis of cyclic di-GMP, although the Pseudomonas aeruginosa protein PA3947, which also has the EVL motif, does have activity (Kulesekara et al., 2006). One hypothesis is that binding of cyclic di-GMP to the (inactive) EVL domain modulates the ability of RocR to compete with RocA1 for binding to the RocS1 sensor, thus affecting cup gene expression (Kulasekara et al., 2005). It is also plausible that antagonism of RocS1 is not the only function of RocR and that phosphorylated RocR can be active in regulation of expression of other genes (Kulasekara et al., 2005). A role for cyclic di-GMP in transcription of genes involved in polysaccharide synthesis associated with rugose colony morphology or biofilm formation has been demonstrated in Vibrio spp. In Vibrio cholerae, the CheY-EAL-HTH domain protein VieA acts to negatively regulate transcription of vps genes, required for exopolysaccharide synthesis and biofilm formation (Tischler and Camilli, 2004). hese effects require the cyclic di-GMP phosphodiesterase activity of VieA, as shown by mutations in the EAL motif. In Vibrio parahaemolyticus, mutation of scrC, which encodes a GGDEF-EAL domain protein, leads to elevated levels of capsular polysaccharide production and increased transcription of cpsA, the first gene in an operon directing capsular polysaccharide biosynthesis (Boles and McCarter, 2002). he scrC gene is part of the scrABC operon. Transposon insertions in the cps operon or in an unlinked gene, cpsR, encoding a transcriptional regulator caused reversion of a strain with a polar mutation in scrA to a smooth colony phenotype (Güvener and McCarter, 2003). CpsR is not required for capsular polysaccharide production in srcABC+ strains however. Elevated levels of cyclic di-GMP promote bioilm formation but repress motility and virulence factor synthesis An emerging theme from a number of studies is that high levels of cyclic di-GMP promote biofilm formation and sessility, but low levels promote motility and the synthesis of virulence factors in a range of human, animal and plant pathogens (Figure 5.3). Elevation of the cellular level of cyclic di-GMP by expression of different GGDEF domain proteins promotes biofilm formation and reduces motility in a number of bacterial species including enteric bacteria such as Salmonella spp. and Pseudomonas aeruginosa (Simm et al., 2004). Conversely expression of EAL domain proteins reduces biofilm formation and enhances motility. In Vibrio cholerae, the CheY-EAL-HTH domain VieA positively activates expression of virulence genes toxT, which encodes a transcriptional regulator and ctxAB, which encode cholera toxin (Tischler et al., 2002; Tischler and Camilli, 2005) and negatively influences exopolysaccharide production and biofilm formation (Tischler and Camilli, 2005). A vieA mutant is attenuated for colonization in the infant mouse model (Tischler et al., 2002). All of these effects require the cyclic di-GMP phosphodiesterase activity of VieA

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Figure 5.3 Cyclic di-GMP as a second messenger links the perception of environmental cues to changes in bacterial physiology and behavior. Proteins with GGDEF, EAL or HD-GYP domains have additional sensory input domains that are involved in the perception of diverse environmental cues. Alteration in the activity of the protein in cyclic di-GMP synthesis or hydrolysis occurs upon signal perception and leads to changes the cellular levels of cyclic di-GMP. In general elevated levels of cyclic di-GMP promote bioilm formation and sessility whereas low levels promote motility and virulence factor synthesis. However localized effects on speciic functions may not follow these general rules. Further levels of regulation occur at the level of transcription of the genes encoding the signaling proteins and in their degradation by proteolysis (see text). Figure adapted from Simm et al. (2004).

and are abolished by mutation of the EAL amino acid motif. Similarly the CheY-HD-GYP domain regulator RpfG of the plant pathogen Xanthomonas campestris, which is required for full virulence, positively regulates the synthesis of extracellular enzyme virulence factors and motility, and negatively regulates biofilm formation (Slater et al., 2000; Dow et al., 2003; Crossman and Dow, 2004). Cyclic di-GMP may contribute to bacterial pathogenesis in a number of other pathosystems. he GGDEF domain protein VirC contributes to the virulence of Vibrio anguillarum to fish (Milton et al., 1995). In Bordetella pertussis, activation of virulence factors by the BvgAS two-component system is accompanied by repression of transcription of a further set of genes, which involves the “stand-alone” EAL domain protein BvgR (Merkel and Stibitz, 1995; Merkel et al., 1998a,b). BvgR-mediated regulation of gene expression contributes to respiratory infection of mice (Merkel et al., 1998b). he “stand-alone” EAL domain protein CdgR of Salmonella is required for the bacterium to resist host phagocyte oxidase in vivo and contributes to virulence in mice (Hisert et al., 2005). hese examples illustrate that the effect of cyclic di-GMP on bacterial virulence is not restricted to those circumstances in which high levels promote biofilm formation; the synthesis of some virulence determinants is activated only under low cellular levels of the

Cyclic di-GMP Signaling and Bioilm Formation

nucleotide. It is possible that bacteria within the population in infected tissue may adopt a planktonic, motile lifestyle in which virulence factors are expressed or a sessile lifestyle of residence in aggregates or biofilms, depending on the environmental conditions. he ability to undergo transitions between these physiological states may favor both the spread and persistence of bacterial disease within host tissues. Although work on a number of bacteria has linked increased cyclic di-GMP levels with augmented biofilm formation, this generality is challenged by more recent work on biofilm formation in P. aeruginosa (Hoffman et al., 2005; Kulesekara et al., 2006). Mutation or over-expression of particular genes encoding GGDEF and EAL domain proteins has effects on biofilm formation that are not strictly correlated with the observed or predicted effects on cyclic di-GMP levels. hese findings have been rationalized by suggesting that these signaling elements affect localized cytoplasmic cyclic di-GMP pools which influence specific processes contributing to biofilm formation (Hoffman et al., 2005; Kulesekara et al., 2006). his will be discussed in detail below. Are cyclic di-GMP-dependent systems dedicated to speciic cellular functions? Localization studies of the PleD regulator (CheY-CheY-GGDEF), which influences swarmer to stalk cell transitions and pole development in Caulobacter crescentus (Aldridge et al., 2003), have shown that upon phosphorylation the protein locates to the pole of the cell where the new stalk will be formed (Paul et al., 2004). Phosphorylation also activates the protein for cyclic di-GMP synthesis (Paul et al., 2004). Previously it was shown that the DgcA and PdeA proteins of G. xylinus co-purified with the cellulose synthase (Ross et al., 1987). hese findings have led to the suggestion that the generation of localized pools of cyclic di-GMP by specific components in cyclic di-GMP signaling may activate processes that are determined by co-localizing proteins. In other words, certain cyclic di-GMP synthesis systems are dedicated to specific cellular tasks. An alternative, but not mutually exclusive, view is that a number of signaling systems form a surveillance network to integrate information about various aspects of the cellular environment and to process this information by determining a cellular level of cyclic di-GMP, which may influence bacterial functions. Observations that different GGDEF, EAL or HD-GYP domain proteins have significant roles in regulation of specific bacterial processes under different environmental condition are consistent with the notion of a an environmentally responsive network. his has been reported for the role of GGDEF domain proteins in cellulose synthesis in Salmonella (Römling et al., 2000; Garcia et al., 2004) and is also seen upon examination of the role of HD-GYP, GGDEF and EAL domain proteins in regulation of extracellular enzyme synthesis in Xanthomonas campestris (R.P. Ryan, Y. Fouhy, J. Lucey and J.M. Dow, unpublished). he existence of localized pools of cyclic di-GMP has been proposed to explain the varied effects of mutation of GGDEF and EAL domain proteins on biofilm formation in P. aeruginosa (Hoffman et al., 2005; Kulesekara et al., 2006). In P. aeruginosa PAO1, the EAL domain protein Arr (for aminoglycoside response regulator) is required for biofilm formation in response to subinhibitory concentrations of the antibiotic tobramycin. A variant Arr protein with a mutation in the conserved EAL domain was not able to restore tobramy-

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cin-induced biofilm formation to an arr mutant of P. aeruginosa. his suggests that cyclic di-GMP degradation is required for biofilm formation, which contradicts the consensus view from work a number of bacterial systems. Examination of the effects of mutation of other GGDEF and/or EAL domain proteins in P. aeruginosa PAO1 also reveals a complex relationship to biofilm formation (Hoffman et al., 2005). A comprehensive mutational and over-expression study of the role of all 37 proteins with GGDEF and/or EAL domains in Pseudomonas aeruginosa PA14 has likewise shown that a number of genes with GGDEF or GGDEF/EAL domains affect biofilm formation (Kulesekara et al., 2006). In general the results are consistent with the concept that enhanced cellular levels of cyclic di-GMP promote biofilm formation. here is however no strict correlation; for example mutation of the gene encoding the GGEEF protein PA3343, which is active in cyclic di-GMP synthesis, leads to a hyperbiofilm formation whereas overexpression of PA2879 and PA3343, which both lead to increases in the cellular level of cyclic di-GMP, has no effect on biofilm formation in the wild type (Kulesekara et al., 2006). hese findings in P. aeruginosa have led to the suggestion of localized effects of elements involved in cyclic di-GMP signaling, where synthesis or hydrolysis of the nucleotide is intimately related to its site of action (Hoffman et al., 2005; Kulesekara et al., 2006). By inference, some functions contributing to biofilm formation could be activated by low levels of cyclic di-GMP. However if elements involved in cyclic di-GMP signaling form complexes with proteins that influence biofilm formation, the possibility that loss or overexpression of the signaling component may adversely affect complex assembly and function cannot be overlooked. In this case the function of the GGDEF/EAL domain protein may not be restricted to its action on cyclic di-GMP. It is clear that a great deal more research is needed to resolve the role of individual signaling systems, taking into account issues of cellular localization, interactions with other proteins, and the effects of gene disruption on specific cellular activities. What are the cellular levels of cyclic di-GMP? Knowledge of the cellular levels of cyclic di-GMP is of considerable importance to an overall understanding of the role of the nucleotide in influencing bacterial behavior. Most simply it acts as a guide as to whether effects of cyclic di-GMP seen in in vitro experiments occur at physiologically relevant concentrations. However such measurements could also help to establish the in vivo role of GGDEF-EAL domain fusions, to examine any connection between cyclic di-GMP levels and the temporal expression of particular bacterial genes and to assess the physiological status of bacteria in different environments. he measurement of cyclic di-GMP in bacterial cells presents considerable technical difficulties. It is found in only very low concentrations in most bacteria and may bind to cellular proteins thereby affecting extraction. A number of laboratories have used two-dimensional thin layer chromatography to separate labeled nucleotides extracted from growing cells in 32P-labelled inorganic phosphate. he methodology is of low throughput however and is presumably restricted for use in specific growth conditions. MALDI-TOF and/or liquid chromatography coupled to mass spectrometric analyses are methods of choice, offering highly sensitive and accurate determinations from bacteria under a range of growth conditions. Available data show that ectopic elevation of cyclic di-GMP through the expres-

Cyclic di-GMP Signaling and Bioilm Formation

sion of GGDEF domain proteins can lead to levels of the nucleotide that are over 200 MM (Simm et al., 2004). In contrast, Hisert et al. (2005) determined that in Salmonella, each cell has eight molecules of cyclic di-GMP, which if the nucleotide were freely distributed would put the cellular concentration in the pM range. here are proposals that localized pools of cyclic di-GMP may occur (see above), so the relationship between the concentration that is locally active and that measured as the cytoplasmic concentration is unclear. In the cellulose-producing G. xylinus, the levels of cyclic di-GMP are estimated to be in the range of 5 to 10 MM. However free cyclic di-GMP may account for only 10% of total of intracellular pool, since the nucleotide is believed to bind to a number of cellular proteins. he level of cyclic di-GMP giving half-maximal activation of cellulose synthesis in vitro is 0.35 MM, which would be within the biologically relevant range however. Environmental signals affecting bioilm formation through alterations in cyclic di-GMP levels Very little is known about the environmental signals or cues that are recognized by the sensory domains of GGDEF, EAL and HD-GYP domain proteins or which activate twocomponent or chemotaxis-like signal transduction pathways involving these proteins. Although bioinformatic analysis can give some clues through the nature of the input domains (see Table 5.1), only in a few cases are the cognate signals known. he best-studied example is PdeA1, the cyclic di-GMP phosphodiesterase from G. xylinus, which has a PAS-GAFGGDEF-EAL domain structure (Chang et al., 2001). he PAS domain of this protein contains a heme moiety that binds molecular oxygen with a Kd of approximately 10 MM. Removal of the heme results in a dramatic loss of enzymatic activity, whereas binding of oxygen to the heme reduces the activity approximately threefold. he consequences of oxygen sensing are thus a reduced ability to degrade cyclic di-GMP, which could conceivably lead to an elevation in the cellular levels, promoting cellulose production and pellicle formation. PdeA1 is highly homologous over its entire length to the Dos protein of Escherichia coli, which is also has a heme-binding PAS domain involved in oxygen sensing (Delgado-Nixon et al., 2000). A second example of a defined environmental signal whose recognition is linked to cyclic di-GMP turnover is the cell–cell signaling molecule DSF of Xanthomonas campestris (Barber et al., 1997: Slater et al., 2000; Crossman and Dow, 2004). DSF has been characterized as a cis-unsaturated fatty acid (Wang et al., 2004). Perception and transduction of the DSF signal is thought to involve a two-component system comprising the sensor histidine kinase RpfC and the CheY-HD-GYP domain regulator RpfG (Slater et al., 2000; Crossman and Dow, 2004; Ryan et al., 2006). It is proposed that recognition of DSF by RpfC leads to phosphorylation of RpfG and its activation in cyclic di-GMP hydrolysis, although this has not been directly demonstrated. Mutation of rpfC, rpfG or rpfF (which directs DSF synthesis), all lead to the formation of biofilms by X. campestris (Dow et al., 2003). he polyamine norspermidine, which is present in a wide range of prokaryotes and eukaryotes, has been shown to activate the formation of biofilms in Vibrio cholerae (Karatan et al., 2005). his activity depends upon NspS, a periplasmic binding protein and MbaA, a GGDEF-EAL domain protein, previously characterized as a repressor of V. cholerae

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biofilm formation (Bomchil et al., 2003; Karatan et al., 2005). he nspS and mbaA genes are arranged in an operon with a third gene that has little or no role under the conditions tested. It is proposed that NspS acts as a sensor for norspermidine and that interaction of a norspermidine-NspS complex with the periplasmic portion of MbaA reduces its ability to inhibit biofilm formation. his may occur through destabilization of an MbaA dimer or through conformational changes affecting MbaA activity (Karatan et al., 2005). Two laboratories have described effects of addition of antibiotics on biofilm formation in Pseudomonas aeruginosa that were influenced by the action of specific EAL domain proteins (Drenkard and Ausubel, 2002; Hoffman et al., 2005). When plated in the presence of a number of antibiotics including kanamycin, tobramycin and tetracycline, antibiotic-resistant small colony variants of P. aeruginosa PA14 arise at a frequency of 10–7 to 10–6 (Drenkard and Ausubel, 2002). A percentage of these variants have a rough colony appearance and were called RSCV (for rough small-colony variant). he RSCV cells show increased attachment to surfaces and increased biofilm formation compared to the wild type. Small colony variants are also found in clinical samples from cystic fibrosis patients. RSCV cells can revert to the wild-type phenotype in the absence of antibiotic. A CheYEAL domain two-component regulator, PvrR (for phenotype variant regulator), positively influences the rate of conversion. he mechanistic basis of these phenotypic variations and the role of cyclic di-GMP are however not known. A more recent report (Hoffman et al., 2005) shows that subinhibitory concentrations of tobramycin trigger biofilm formation in P. aeruginosa. his effect depends upon Arr (for aminoglycoside response regulator), a membrane-associated EAL domain protein with a periplasmic domain. he arr mutant phenotype is not due to altered frequency of RSCV variants. An arr mutant showed reduced membrane cyclic di-GMP phosphodiesterase activity, but no apparent enhancement in biofilm formation in the absence of tobramycin. A variant Arr protein with a mutation in the conserved EAL domain was not able to restore tobramycin-induced biofilm formation to an arr mutant of P. aeruginosa. Hoffman et al. (2005) propose that tobramycin, either directly or indirectly, enhances the phosphodiesterase activity of Arr leading to cyclic di-GMP degradation and increased biofilm formation, through a localized effect on a discrete pool of cyclic di-GMP (see also above). Since biofilms and phase variation contribute to the increased resistance of Pseudomonas aeruginosa to antibiotics, compounds that affect PvrR or Arr function could have a role in the treatment of infections associated with cystic fibrosis (Drenkard and Ausubel, 2002; Hoffman et al., 2005). Mechanisms related to those discussed above may have evolved to allow Pseudomonas aeruginosa to evade the action of antibiotics produced by other microorganisms co-inhabiting environments such as the soil and plant root surfaces. he above examples illustrate the ability of bacteria to adopt different modes of growth as an adaptation to different environmental conditions. Studies of the effects of carbon starvation in Pseudomonas putida have shown that transitions from a biofilm to planktonic state can occur quite rapidly, within minutes of the application of the stress (Gjermansen et al., 2005). A TM-GGDEF-EAL domain protein PP0165 is implicated in these transitions, although mechanistic details of the role of cyclic di-GMP in this rapid switch in lifestyle remain to be uncovered.

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Regulation by transcription and protein degradation In addition to modulation of the activity of the signaling proteins by ligand binding, cyclic di-GMP signaling is also subject to regulation at the level of transcription of the cognate genes and by signal protein degradation. A number of studies have shown that the expression of genes encoding GGDEF and/or EAL domain proteins is differentially regulated under different environmental conditions. As we have seen above, the two-component regulator CsgD of Salmonella regulates transcription of adrA, a gene that encodes a GGDEF domain protein that is required for cellulose synthesis in the multicellular rdar phenotype (Römling et al., 2000; Römling, 2005). CsgD is activated by as yet unidentified histidine kinase, which presumably responds to environmental cues. Whole-genome microarray technology has revealed a number of genes in P. aeruginosa whose expression responds to signals found in muco-purulent airway liquids collected from chronically infected cystic fibrosis patients (Wolfgang et al., 2004). PA2567, encoding a cyclic di-GMP phosphodiesterase, is downregulated by more than 5-fold. his repression requires P. aeruginosa quorum sensing system since these effects are not observed in an rhlRlasR double mutant (Wolfgang et al., 2004). he formation of mixed biofilms between the hyperthermophiles ermotoga maritima and Methanococcus jannaschii is accompanied by increased expression in T. maritima of genes encoding synthesis of an extracellular polysaccharide. In addition two genes encoding GGDEF domain proteins are upregulated whereas a third gene encoding a GGDEF domain protein is down regulated ( Johnson et al., 2004). he synthesis of extracellular polysaccharide is also regulated by a peptide cell–cell signal ( Johnson et al., 2005). In Vibrio cholerae, the quorum-sensing regulator AphA, which influences the expression of virulence genes, also strongly represses the expression of an operon encoding proteins involved in acetoin biosynthesis (Kovacikova et al., 2005). his molecule prevents intracellular acidification and is required for growth of V. cholerae on glucose. he acetoin biosynthesis operon also contains genes encoding a GGDEF domain protein and an EAL domain protein (Kovacikova et al., 2005). It has yet to be determined whether these proteins influence biofilm formation or motility or whether cyclic di-GMP influences expression of the acetoin operon. Several of these examples illustrate interplay between cyclic di-GMP signaling and extracellular cell–cell signaling (quorum-sensing), which also has a critical role in biofilm formation. his has led to the suggestion that the two signaling processes converge in the regulation of diverse bacterial behaviors (Camilli and Bassler, 2006). A direct connection between cell–cell signaling and cyclic di-GMP turnover has been proposed; perception and transduction of the DSF cell–cell signal in Xanthomonas campestris is thought to involve a two-component system involving the CheY-HD-GYP domain regulator RpfG (Slater et al., 2000; Crossman and Dow, 2004), which is active in hydrolysis of cyclic di-GMP (Ryan et al., 2006). Activation of a GGDEF/EAL domain protein by binding of a quorum-sensing molecule has not yet been reported however. Further regulation of cyclic di-GMP signaling occurs at the level of protein degradation. In Yersinia pestis, the Congo red binding Hms+ biofilm phenotype occurs at temperatures up to 34oC but not at higher temperatures. Biofilm formation depends upon the GGDEF domain protein HmsT, as well as other Hms proteins (Table 5.2, Perry et al., 2004). he

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cellular level of the HmsT protein is very low at 37oC, although transcription from the hmsT promoter or the level of hmsT mRNA is not significantly affected by growth temperature. However, HmsT at 37oC is sensitive to degradation by cellular proteases, thus contributing to the temperature regulation of the Hms+ phenotype (Perry et al., 2004). Regulation at the level of transcription and/or proteolytic degradation may be particularly important in determining the activity of “stand alone” EAL or GGDEF domain proteins, which lack sensory input domains. However it cannot be excluded that these EAL and GGDEF domains also bind small molecules that influence their enzymatic activity. How does cyclic di-GMP exert its action? Almost nothing is known about the mechanisms whereby cyclic di-GMP exerts its action on diverse cellular functions including biofilm formation. he best-studied effect of cyclic di-GMP is its allosteric activation of cellulose synthesis in G. xylinus, where cyclic di-GMP has been shown to bind to BcsB, the B-subunit of cellulose synthase and to an uncharacterized 200 kDa protein (see above). Recent bioinformatic studies have suggested that in contrast cyclic di-GMP binds to BcsA, the A-subunit of cellulose synthase and that PilZ, a domain at the C-terminus of BcsA, is part of the binding site (Amikam and Galperin, 2006). PilZ was originally described as a protein involved in assembly of functional pili in Pseudomonas aeruginosa (Alm et al., 1996). Several lines of evidence support the proposal that PilZ is also involved in the wider cellular activities of cyclic di-GMP. PilZ can be present as a “stand-alone” domain but can also be found associated with other domains including CheY, GGDEF, EAL and HD-GYP, suggesting a role in regulation and signaling. he phyletic distribution of the PilZ domain is generally similar to those of the GGDEF and EAL and some phenotypes of mutation of genes encoding PilZ domain proteins are consistent with a role in cyclic di-GMP regulation (Huang et al., 2003; Amikam and Galperin, 2006). hese bioinformatic predictions of the role of PilZ domain proteins are open to experimental verification. Key questions include what are cellular roles of PilZ proteins, do they bind cyclic di-GMP and is this binding required for their activity? Other possible targets of cyclic di-GMP action should not be overlooked. By analogy with cyclic AMP, cyclic di-GMP may affect transcription of genes involved in virulence and biofilm formation by binding to transcriptional regulators. Since in a number of cases cognate regulators of virulence genes have been identified, it may be possible to examine the effects of cyclic di-GMP by in vitro promoter binding assays. One caveat is that transcriptome profiling reveals that elevation of the level of cyclic di-GMP alters the expression of a substantial number of genes in Pseudomonas aeruginosa (Hickman et al., 2005) and Escherichia coli (Méndez-Ortiz, et al., 2006). Such large-scale changes may make it difficult to distinguish direct and indirect effects of cyclic di-GMP and consequently to identify candidate transcription factors for further study. How will an understanding of cyclic di-GMP signaling have an impact in biotechnology? An understanding of the role of cyclic-di-GMP signaling in the regulation of bacterial biofilm formation and virulence could underpin the design of new strategies for the control of disease in both animals and plants. In the biomedical context, directed interference

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to promote biofilm dispersal may improve the efficacy of antibiotic treatment to control persistent infections. his could potentially be achieved through blocking the perception of the environmental signal(s), modulation of the biochemical activity of the signaling domains or blocking the sites or targets of cyclic di-GMP action. An in-depth understanding of this sophisticated signaling system is needed to achieve the desired aims. For example knowledge of the biochemical activity of the domains involved in cyclic di-GMP turnover suggests that chemicals that inhibit GGDEF domain proteins or those that activate EAL domain proteins may have a role in triggering biofilm dispersal. However we now appreciate that there are particular signaling systems in which EAL or GG(D/E)EF proteins have opposite effects on biofilm formation from that expected from their activities against cyclic di-GMP. Furthermore potential effects on the activation of virulence factor synthesis cannot be overlooked. An understanding of how cyclic di-GMP exerts its effects on different cellular functions may allow blocking of biofilm formation without promotion of virulence factor synthesis. he definition of those signaling elements that are key to biofilm formation and/or virulence under certain conditions may identify specific targets for intervention. Two examples from Pseudomonas aeruginosa illustrate this point; Arr, which positively influences tobramycin-induced biofilm formation and antibiotic resistance and PvrR, which influences phase variation, biofilm formation and virulence in thermally injured mice (Drenkard and Ausubel, 2002; Hoffman et al., 2005; Kulesekara et al., 2006). Although most attention has been focused on animal and human pathogens, the ability of plant pathogenic bacteria to form and detach from biofilms may equally have considerable implications for the completion of their disease cycle. An understanding of these processes at the molecular level and their role in virulence opens a new perspective for the control of plant disease. Interference with the ability of pathogens to attach to and form biofilms during epiphytic growth phases on leaf surfaces, to attach and from aggregates on internal plant surfaces such as within xylem elements or to associate with insect vectors may all have a role in controlling disease. More research is also required to examine potential industrial applications for cyclic di-GMP “technologies.” For example will alteration of cyclic di-GMP levels enhance the production of microbial products such as polysaccharides, secondary metabolites and enzymes of commercial importance? Will the ability to control biofilm formation or dispersal by regulated ectopic expression of GGDEF or EAL domains have a role in microbial fermentations? Concluding remarks he appreciation of the role of cyclic di-GMP as an important and almost ubiquitous signaling molecule in bacteria has occurred relatively recently and has been strongly promoted by the increased interest in microbial biofilm formation and by microbial genome sequencing. he research in this area has built upon the seminal findings of the group of Moshe Benziman, both in the discovery of cyclic di-GMP in 1987 and in the cloning of genes encoding proteins involved in cyclic di-GMP turnover in 1998. Subsequently the biochemical roles of domains involved in turnover of cyclic di-GMP have been defined and proteins with GGDEF/EAL/HD-GYP domains have been implicated in a number

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of important aspects of bacterial behavior such as developmental transitions, adhesion and virulence, in addition to biofilm formation. Despite this significant progress, there are many questions that remain unanswered. Principal amongst these are the mechanism(s) by which cyclic di-GMP exerts its action on different cellular functions, about which almost nothing is known. We might also expect that investigation of the cell biology and spatial aspects of the various signaling systems will come to the fore, as researchers address the issue of specificity of individual signaling systems. here are a host of other questions. Are there networks of interacting systems? What are the cues that activate or inactivate the signaling proteins? Are further domains involved in cyclic di-GMP turnover? Can the cellular level of cyclic di-GMP be regulated by export from the cell? Can bacteria take up cyclic di-GMP? he ultimate goal of this research is a deeper understanding of a signaling system whose importance to the lifestyle of diverse bacteria is now emerging. References Aldridge, P., Paul, R., Goymer, P., Rainey, P., and Jenal, U. (2003). Role of the GGDEF regulator PleD in polar development of Caulobacter crescentus. Mol. Microbiol. 47, 1695–1708. Alm, R.A., Bodero, A.J., Free, P.D., and Mattick, J.S. (1996). Identification of a novel gene, pilZ, essential for type 4 fimbrial biogenesis in Pseudomonas aeruginosa. J. Bacteriol, 178, 46–53. Amikam, D., and Benziman, M. (1989). Cyclic diguanylic acid and cellulose synthesis in Agrobacterium tumefaciens. J. Bacteriol. 171, 6649–6655. Amikam, D., and Galperin, M.Y. (2006). PilZ is part of the bacterial c-di-GMP binding protein. Bioinformatics 22, 3–6. Ausmees, N., Jonsson, H., Höglund, S., Ljunggren, H., and Lindberg, M. (1999). Structural and putative regulatory genes involved in cellulose synthesis in Rhizobium leguminosarum bv. trifolii. Microbiology 145, 1253–1262. Ausmees, N., Mayer, R., Weinhouse, H., Volman, G., Amikam, D., Benziman, M., and Lindberg, M. (2001). Genetic data indicate that proteins containing the GGDEF domain possess diguanylate cyclase activity. FEMS Microbiol. Lett. 204, 163–167. Barber, C.E., Tang, J.L., Feng, J.X., Pan, M.Q., Wilson, T.J., Slater, H., Dow, J.M., Williams, P., and Daniels, M.J. (1997). A novel regulatory system required for pathogenicity of Xanthomonas campestris is mediated by a small diffusible signal molecule. Mol. Microbiol. 24, 555–566. Bobrov, A.G., Kirillina, O., and Perry, R.D. (2005). he phosphodiesterase activity of the HmsP EAL domain is required for negative regulation of biofilm formation in Yersinia pestis. FEMS Microbiol. Lett. 247, 123–130. Boles, B.R., and McCarter, L.L. (2002). Vibrio parahaemolyticus scrABC, a novel operon affecting swarming and capsular polysaccharide regulation. J. Bacteriol. 184, 5946–5954. Bomchil, N., Watnick, P., and Kolter, R. (2003). Identification and characterization of a Vibrio cholerae gene, mbaA, involved in maintenance of biofilm architecture. J. Bacteriol. 185, 1384–1390. Camilli, A., and Bassler, B.L. (2006). Bacterial small-molecule signaling pathways. Science 311, 1113– 1116. Chan, C., Paul, R., Samoray, D., Amiot, N.C., Giese, B., Jenal, U., and Schirmer, T. (2004). Structural basis of activity and allosteric control of diguanylate cyclase. Proc. Natl. Acad. Sci. USA 101, 17084– 17089. Chang, A.L., Tuckerman, J.R., Gonzalez, G., Mayer, R., Weinhouse, H., Volman, G. Amikam, D., Benziman, M., and Gilles-Gonzalez, M.A. (2001). Phosphodiesterase A1, a regulator of cellulose synthesis in Acetobacter xylinum, is a heme-based sensor. Biochemistry 40, 3420–3426. Christen, M., Christen, B., Folcher, M., Schauerte, A., and Jenal, U. (2005). Identification and characterization of a cyclic di-GMP-specific phosphodiesterase and its allosteric control by GTP. J. Biol. Chem. 280, 30829–30837. Cook, K.E., and Colvin, J.R. (1980). Evidence for a beneficial influence of cellulose production on growth of Acetobacter xylinum in liquid medium. Curr. Microbiol. 3, 203–205.

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Crossman, L., and Dow, J.M. (2004). Biofilm formation and dispersal in Xanthomonas campestris. Microbes Infect. 6, 623–629. D’Argenio, D.A., Calfee, M.W., Rainey, P.B., and Pesci, E.C. (2002). Autolysis and autoaggregation in Pseudomonas aeruginosa colony morphology mutants. J. Bacteriol. 184, 6481–6489. D’Argenio, D.A., and Miller, S.I. (2004). Cyclic di-GMP as a bacterial second messenger. Microbiology 150, 2497–2502. Delgado-Nixon, V.M., Gonzalez, G., and Gilles-Gonzalez, M.A. (2000). Dos, a heme-binding PAS protein from Escherichia coli, is a direct oxygen sensor. Biochemistry 39, 2685–2691. Dow, J.M., Crossman, L., Findlay, K., He, Y.Q., Feng, J.X., and Tang, J.L. (2003). Biofilm dispersal in Xanthomonas campestris is controlled by cell–cell signaling and is required for full virulence to plants. Proc. Natl. Acad. Sci. USA. 100, 10995–11000. Drenkard, E., and Ausubel, F.M. (2002). Pseudomonas biofilm formation and antibiotic resistance are linked to phenotypic variation. Nature 416, 740–743. Friedman, L., and Kolter, R. (2004). Genes involved in matrix formation in Pseudomonas aeruginosa PA14 biofilms. Mol. Microbiol. 51, 675–690. Galperin, M.Y., Natale, D.A., Aravind, L., and Koonin, E.V. (1999). A specialized version of the HD hydrolase domain implicated in signal transduction. J. Mol. Microbiol. Biotechnol. 1, 303–305. Galperin, M.Y., Nikolskaya, A.N., and Koonin, E.V. (2001). Novel domains of the prokaryotic two-component signal transduction systems. FEMS Microbiol. Lett. 203, 11–21. Galperin, M.Y. (2005). A census of membrane-bound and intracellular signal transduction proteins in bacteria: Bacterial IQ, extroverts and introverts. BMC Microbiology 5, 35 Garcia, B., Latasa, C., Solano, C., Portillo, F.G., Gamazo, C., and Lasa, I. (2004). Role of the GGDEF protein family in Salmonella cellulose biosynthesis and biofilm formation. Mol. Microbiol. 54, 264–277. Gjermansen, M., Ragas, P., Sternberg, C., Molin, S., and Tolker-Nielsen, T. (2005). Characterization of starvation-induced dispersion in Pseudomonas putida biofilms. Environ. Microbiol. 7, 894–906. Goymer P. (2002). Characterisation of the Pseudomonas fluorescens WspR locus. D.Phil thesis, University of Oxford, UK. Güvener, Z.T., and McCarter, L.L. (2003). Multiple regulators control capsular polysaccharide production in Vibrio parahaemolyticus. J. Bacteriol. 185, 5431–5441. Hecht, G.B., and Newton, A. (1995). Identification of a novel response regulator required for the swarmerto-stalked-cell transition in Caulobacter crescentus. J. Bacteriol. 177, 6223–6229. Hickman, J.W., Tifrea, D.F., and Harwood, C.S. (2005). A chemosensory system that regulates biofilm formation through modulation of cyclic diguanylate levels. Proc. Natl. Acad. Sci. USA. 102, 14422– 14427. Hisert, K.B., MacCoss, M., Shiloh, M.U., Darwin, K.H., Singh, S., Jones, R.A., Ehrt, S., Zhang, Z., Gaffney, B.L., Gandotra, S. et al. (2005). A glutamate-alanine-leucine (EAL) domain protein of Salmonella controls bacterial survival in mice, antioxidant defence and killing of macrophages: role of cyclic diGMP. Mol Microbiol. 56, 1234–1245. Hoffman, L.R., D’Argenio, D.A., MacCoss, M.J., Zhang, Z., Jones, R.A., and Miller, S.I. (2005). Aminoglycoside antibiotics induce bacterial biofilm formation. Nature 436, 1171–1175. Huang, B., Whitchurch, C.B., and Mattick, J.S. (2003). FimX, a multidomain protein connecting environmental signals to twitching motility in Pseudomonas aeruginosa. J. Bacteriol. 185, 7068–7076. Jenal, U. (2004). Cyclic di-guanosine-monophosphate comes of age: a novel secondary messenger involved in modulating cell surface structures in bacteria? Curr. Opin. Microbiol. 7, 185–191. Johnson, M.R., Montero, C.I., Conners, S.B., Shockley, K.R., Bridger, S.L., and Kelly R.M. (2005). Population density-dependent regulation of exopolysaccharide formation in the hyperthermophilic bacterium ermotoga maritima. Mol. Microbiol. 55, 664–674. Jones, H.A., Lillard, J.W., Jr., and Perry, R.D. (1999). HmsT, a protein essential for expression of the haemin storage (Hms+) phenotype of Yersinia pestis. Microbiology 145, 2117–2128. Karatan, E., Duncan, T.R., and Watnick, P.I. (2005). NspS, a predicted polyamine sensor, mediates activation of Vibrio cholerae biofilm formation by norspermidine. J. Bacteriol. 187, 7434–7443. Kirillina, O., Fetherston, J.D., Bobrov, A.G., Abney, J., and Perry, R.D. (2004). HmsP, a putative phosphodiesterase, and HmsT, a putative diguanylate cyclase, control Hms-dependent biofilm formation in Yersinia pestis. Mol. Microbiol. 54, 75–88.

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Kovacikova, G., Lin, W., and Skorupski, K. (2005). Dual regulation of genes involved in acetoin biosynthesis and motility/biofilm formation by the virulence activator AphA and the acetate-responsive LysR-type regulator AlsR in Vibrio cholerae. Mol. Microbiol. 57, 420–433. Kulasekara, H.D., Ventre, I., Kulasekara, B.R., Lazdunski, A., Filloux, A., and Lory, S. (2005). A novel two-component system controls the expression of Pseudomonas aeruginosa fimbrial cup genes. Mol. Microbiol. 55, 368–380. Kulesekara, H., Lee, V., Brencic, A., Liberati, N., Urbach, J., Miyata, S., Lee, D.G., Neely, A.N., Hyodo, M., Hayakawa, Y. et al. (2006). Analysis of Pseudomonas aeruginosa diguanylate cyclases and phosphodiesterases reveals a role for bis-(3a-5a)-cyclic-GMP in virulence. Proc. Natl. Acad. Sci. USA 103, 2839–2844. Mayer, R., Ross, P., Weinhouse, H., Amikam, D., Volman, G., Ohana, P., Calhoon, R.D., Wong, H.C., Emerick, A.W., and Benziman M. (1991). Polypeptide composition of bacterial cyclic diguanylic aciddependent cellulose synthase and the occurrence of immunologically crossreacting proteins in higher plants. Proc. Natl. Acad. Sci. USA. 88, 5472–5476. Méndez-Ortiz, M.M., Hyodo, M., Hayakawa, Y., and Membrillo-Hernández, J. (2006). Genome wide transcriptional profile of Escherichia coli in response to high levels of the second messenger c-di-GMP. J. Biol. Chem. 281, 8090–8099. Merkel, T.J., and Stibitz, S. (1995). Identification of a locus required for the regulation of bvg-repressed genes in Bordetella pertussis. J. Bacteriol. 177, 2727–2736. Merkel, T.J., Barros, C., and Stibitz, S. (1998a). Characterization of the bvgR locus of Bordetella pertussis. J. Bacteriol. 180, 1682–1690. Merkel, T.J., Stibitz, S., Keith, J.M., Leef, M., and Shahin, R. (1998b). Contribution of regulation by the bvg locus to respiratory infection of mice by Bordetella pertussis. Infect. Immun. 66, 4367–4373. Milton, D.L., Norqvist, A., and Wolf-Watz, H. (1995). Sequence of a novel virulence-mediating gene, virC, from Vibrio anguillarum. Gene 164, 95–100. Paul, R., Weiser, S., Amiot, N.C., Chan, C., Schirmer, T., Giese, B., and Jenal, U. (2004). Cell cycle-dependent dynamic localization of a bacterial response regulator with a novel di-guanylate cyclase output domain. Genes Dev. 18, 715–727. Pei, J., and Grishin, N.V. (2001). GGDEF domain is homologous to adenylyl cyclase. Proteins 42, 210–216. Perry, R.D., Bobrov, A.G., Kirillina, O., Jones, H.A., Pedersen, L., Abney, J., and Fetherston, J.D. (2004). Temperature regulation of the hemin storage (Hms+) phenotype of Yersinia pestis is posttranscriptional. J. Bacteriol. 186, 1638–1647. Rashid, M.H., Rajanna, C., Ali, A., and Karaolis, D.K. (2003). Identification of genes involved in the switch between the smooth and rugose phenotypes of Vibrio cholerae. FEMS Microbiol. Lett. 227, 113–119. Römling, U. (2002). Molecular biology of cellulose production in bacteria. Res. Microbiol. 153, 205–12. Römling, U. (2005). Characterization of the rdar morphotype, a multicellular behaviour in Enterobacteriaceae. Cell. Mol. Life Sci. 62, 1–13. Römling, U., Rohde, M., Olsen, A., Normark, S., and Reinköster, J. (2000). AgfD, the checkpoint of multicellular and aggregative behaviour in Salmonella typhimurium regulates at least two independent pathways. Mol. Microbiol. 36, 10–23. Römling, U., Gomelsky, M., and Galperin, M.Y. (2005). C-di-GMP: he dawning of a novel bacterial signalling system. Mol. Microbiol. 57, 629–639. Ross, P., Weinhouse, H., Aloni, Y., Michaeli, D., Weinberger-Ohana, P., Mayer, R., Braun, S., de Vroom, E., van der Marel, G.A., van Boom, J.H., and Benziman, M. (1987). Regulation of cellulose synthesis in Acetobacter xylinum by cyclic diguanylate. Nature 325, 279–281. Ryan, R.P., Fouhy, Y., Lucey, J.F., Crossman, L.C., Spiro, S., He, Y.W., Zhang L.H., Heeb, S., Cámara, M., Williams, P., and Dow, J.M. (2006). Cell–cell signaling in Xanthomonas campestris involves an HD-GYP domain protein that functions in cyclic di-GMP turnover. Proc. Natl. Acad. Sci. USA. 103, 6712–6717. Ryjenkov, D.A., Tarutina, M., Moskvin, O.M., and Gomelsky, M. (2005). Cyclic diguanylate is a ubiquitous signaling molecule in Bacteria: insights into biochemistry of the GGDEF protein domain. J. Bacteriol. 187, 1792–1798. Saxena, I.M., and Brown, R.M., Jr. (1995). Identification of a second cellulose synthase gene (acsAII) in Acetobacter xylinum. J. Bacteriol. 177, 5276–5283.

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Schmidt, A.J., Ryjenkov, D.A., and Gomelsky, M. (2005). Ubiquitous protein domain EAL encodes cyclic diguanylate-specific phosphodiesterase: enzymatically active and inactive EAL domains. J. Bacteriol. 187, 4774–4781. Simm, R., Morr, M., Kader, A., Nimtz, M., and Römling, U. (2004). GGDEF and EAL domains inversely regulate cyclic di-GMP levels and transition from sessility to motility. Mol. Microbiol. 53, 1123–1134. Simm, R., Fetherston, J.D., Kader, A., Römling, U., and Perry, R.D. (2005). Phenotypic convergence mediated by GGDEF-domain-containing proteins. J. Bacteriol. 187, 6816–23. Slater, H., Alvarez-Morales, A., Barber, C.E., Daniels, M.J., and Dow, J.M. (2000). A two-component system involving an HD-GYP domain protein links cell–cell signalling to pathogenicity gene expression in Xanthomonas campestris. Mol. Microbiol. 38, 986–1003. Solano, C., García, B., Valle, J., Berasain, C., Ghigo, J.M., Gamazo, C., and Lasa, I. (2002). Genetic analysis of Salmonella enteritidis biofilm formation: critical role of cellulose. Mol. Microbiol. 43, 793–808. Spiers, A.J., Bohannon, J., Gehrig, S.M., and Rainey, P.B. (2003). Biofilm formation at the air-liquid interface by the Pseudomonas fluorescens SBW25 wrinkly spreader requires an acetylated form of cellulose. Mol. Microbiol. 50, 15–27. Tal, R., Wong, H.C., Calhoon, R., Gelfand, D., Fear, A.L., Volman, G., Mayer, R., Ross, P., Amikam, D., Weinhouse, H., Cohen, A., Sapir, S., Ohana, P., and Benziman, M. (1998). hree cdg operons control cellular turnover of cyclic di-GMP in Acetobacter xylinum: genetic organization and occurrence of conserved domains in isoenzymes. J. Bacteriol. 180, 4416–4425. Taylor, B.L., and Zhulin, I.B. (1999). PAS domains: internal sensors of oxygen, redox potential, and light. Microbiol. Mol. Biol. Rev. 63, 479–506. Tischler, A.D., Lee, S.H., and Camilli, A. (2002). he Vibrio cholerae vieSAB locus encodes a pathway contributing to cholera toxin production. J. Bacteriol. 184, 4104–4113. Tischler, A.D., and Camilli, A. (2004). Cyclic diguanylate (c-di-GMP) regulates Vibrio cholerae biofilm formation. Mol. Microbiol. 53, 857–869. Tischler, A.D., and Camilli, A. (2005). Cyclic diguanylate regulates Vibrio cholerae virulence gene expression. Infect. Immun. 73, 5873–5882. Vallet, I., Olson, J.W., Lory, S., Lazdunski, A., and Filloux, A. (2001). he chaperone/usherpathways of Pseudomonas aeruginosa: identification of fimbrial gene clusters (cup) and their involvement in biofilm formation. Proc. Natl. Acad. Sci. USA 98, 6911–6916. Wang, L.H., He, Y., Gao, Y., Wu, J.E., Dong, Y.H., He, C., Wang, S.X., Weng, L.X., Xu, J.L., Tay, L., et al. (2004). A bacterial cell–cell communication signal with cross-kingdom structural analogues. Mol. Microbiol. 51, 903–912. Weinhouse, H., Sapir, S., Amikam, D., Shilo, Y., Volman, G., Ohana, P., and Benziman, M. (1997). C-diGMP-binding protein, a new factor regulating cellulose synthesis in Acetobacter xylinum. FEBS Lett. 416, 207–211. Williams, W.S., and Cannon, R.E. (1989). Alternative environmental roles for cellulose produced by Acetobacter xylinum. Appl. Environ. Microbiol. 55, 2448–2452. Wolfgang, M.C., Jyot, J., Goodman, A.L., Ramphal, R., and Lory, S. (2004). Pseudomonas aeruginosa regulates flagellin expression as part of a global response to airway fluid from cystic fibrosis patients. Proc. Natl. Acad. Sci. USA. 101, 6664–6668. Zhulin, I.B., Nikolskaya, A.N., and Galperin, M.Y. (2003). Common extracellular sensory domains in transmembrane receptors for diverse signal transduction pathways in bacteria and archaea. J. Bacteriol. 185, 285–294. Zogaj, X., Nimtz, M., Rohde, M., Bokranz, W., and Römling, U. (2001). he multicellular morphotypes of Salmonella typhimurium and Escherichia coli produce cellulose as the second component of the extracellular matrix. Mol. Microbiol. 39, 1452–1463.

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N-Acylhomoserine Lactones, Quorum Sensing, and Bioilm Development in Gram-negative Bacteria

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Steve Atkinson, Miguel Cámara, and Paul Williams

Abstract Many different Gram negative bacteria employ N-acylhomoserine lactones (AHLs) as diffusible signal molecules which enable bacterial populations to co-ordinate gene expression as a function of cell population density. Such co-ordinated community behavior, termed “quorum sensing” (QS) regulates diverse physiological processes including secondary metabolite production, motility, DNA transfer and pathogenicity. AHLs are produced during the biofilm mode of growth and AHL-dependent QS influences the development, integrity and architecture of biofilm communities as well as orchestrating the optimal timing and production of secondary metabolites to combat predators and host defense mechanisms. In most cases the identity of the QS-regulated target structural genes which contribute to biofilm development have not yet been identified. However, the increased susceptibility of biofilms formed by Pseudomonas aeruginosa QS mutants to conventional antibacterial agents and host defenses highlights the utility of AHL-dependent QS as a novel antimicrobial target. Introduction Bacteria have evolved multiple integrated sensory systems to facilitate adaptation to environmental challenges at both the individual cell and population levels. he perception and processing of chemical information forms a pivotal component of the regulatory mechanisms necessary for such population-dependent adaptive behavior. Many bacteria employ self-generated small diffusible signal molecules to control gene expression as a function of cell population density. In this process, termed “quorum sensing” (QS), the concentration of a signal molecule which accumulates in the extracellular environment reflects the cell number such that the perception of a threshold concentration of signal molecule determines when the population is “quorate” and ready to make a collective behavioral adaptation (Salmond et al., 1995; Williams et al., 2000; Swift et al., 2001; Cámara et al., 2002). QS signal molecule diffusion between spatially separated bacterial subpopulations may also convey information about their physiological state, their numbers, and the specific environmental conditions being encountered. he notion that a QS signal molecule can accumulate is based on the assumption that there will be a diffusion barrier over which more molecules will accumulate than are lost from a given micro-habitat (Winzer et al., 2002;

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Redfield, 2002). his has been suggested as a “compartment” or “diffusion” sensing system where QS signal molecule accumulation is both the measure for the degree of compartmentalization and the means to distribute this information among the community (Redfield, 2002; Williams et al., 2007). QS may therefore be considered as an alternative version of diffusion sensing (quorum diffusion sensing might be a more appropriate term), where the threshold concentration of the QS signal eliciting a response can only be achieved by a population rather than by a single cell. Consequently, the size of the quorum is not fixed but will depend on the relative rates of synthesis and loss of QS signal molecule. here are also situations where a single bacterium can switch from the non-quorate to quorate state. For example individual Staphylococcus aureus cells trapped intracellularly within an endosome in mammalian endothelial cells are unable to escape and replicate within the cytoplasm unless they are able to express their agr QS system (Qazi et al., 2001). In such situations, the QS system is clearly functioning in the “compartment” not “population” sensing mode. It is also important to consider QS as only one of multiple environmental parameters which a bacterial cell must integrate in order to make a behavioral decision (Withers et al., 2001). Consequently QS systems have been identified as integral components of global gene regulatory networks in diverse bacterial genera. he QS signal molecule classes so far identified are chemically diverse and include Nacylhomoserine lactones (AHLs), G-butyrolactones, 2-alkyl-4-quinolones, furanones, fatty acid derivatives and peptides. No “universal” QS language has been discovered and only the autoinducer-2/LuxS QS system appears to be shared by both Gram negative and Grampositive bacteria (Winzer et al., 2002; Winzer and Williams, 2003; Vendeville et al., 2005) (see also Chapters 7 and 8). However, many more chemically distinct QS signal molecules are likely to exist and it has been argued that the majority of extracellular bacterial metabolites including compounds with antibiotic activity are likely to function as signal molecules (Yim et al., 2006). Consequently, a number of criteria have been proposed to distinguish between QS signal molecules and other extracellular metabolites. Firstly, the biosynthesis of the QS signal should occur during a specific stage of growth, under certain physiological conditions, or in response to particular environmental conditions. Secondly, the QS signal must accumulate in the extracellular milieu, attain a critical threshold concentration and be recognized by a specific bacterial receptor. hirdly, the cellular response to the molecule should extend beyond the physiological changes required to metabolize or detoxify the molecule (Winzer et al., 2002). hese criteria, in conjunction with the concept of diffusible signal molecule-mediated density dependent QS, define the boundary from which the role of diffusible signal molecules in co-ordinating bacterial behavior can be conceptualized. In this chapter the contribution of N-acylhomoserine lactone (AHL)-dependent QS to the biofilm lifestyle of Gram-negative bacteria will be explored. AHL-dependent quorum sensing AHLs are employed as QS signal molecules by Gram negative proteobacteria belonging to A, B and G subdivisions. So far no AHL-producing Gram positive bacteria have been identified (Swift et al., 1998; Withers et al., 2001; Cámara et al., 2002; Chhabra et al., 2005). Structurally AHLs consist of a homoserine lactone ring covalently linked via an amide bond to an acyl side chain (ranging between 4 and 18 carbons) which may be saturated or

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unsaturated and with or without a hydroxy-, oxo- or no substituent on the carbon at the 3 position of the N-linked acyl chain (Figure 6.1). AHLs are usually synthesized by enzymes belonging to the LuxI family of AHL synthases, around 100 of which are currently present in the genome databases. Studies of the crystal structures of EsaI and LasI have revealed that this large protein family belongs to the GCN5-related N-acetyltransferase protein family (Watson et al., 2002; Gould et al., 2004). Most LuxI proteins share low homologies although ten invariant amino acid residues have been identified which are essential for activity and are localized within the N-terminal domain of the protein (Fuqua et al., 2001). hree members of a second family of AHL synthases, the LuxM family, has also been identified in the genus Vibrio (Hanzelka et al., 1999; Milton et al., 2001). Both LuxI and LuxM proteins employ the same substrates for AHL synthesis, namely S-adenosylmethionine (SAM) and the appropriately charged acyl-acyl carrier protein (acyl-ACP; (Hanzelka et al., 1999)). To date it has not been possible to predict the nature of the AHL(s) produced by a given LuxI or LuxM protein from bioinformatic analyses alone. However, the availability of several different biosensors have greatly facilitated the detection and preliminary identification of AHLs although unequivocal chemical identification requires mass spectrometry and/or NMR spectroscopy (Swift et al., 1999; Chhabra et al., 2005). From such experiments it has become clear that many LuxI homologues direct the synthesis of a variety of different AHLs although in most cases 1–2 different compounds predominate (Ortori et al., 2006). Once synthesized, AHLs accumulate extracellularly and diffuse into neighboring bacterial cells where they usually interact with members of the LuxR family of transcriptional regulators (Fuqua et al., 1996; Swift et al., 2001; Cámara et al., 2002). AHLs bind to, and activate LuxR homologous proteins and the resulting LuxR protein/AHL complex activates or represses the relevant target structural gene(s) (Swift et al., 2001; Zhang et al., 2002; Cámara et al., 2002). LuxR proteins bind to a region within the promoter/operator O

O O

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Figure 6.1 General structures of 3-oxo-AHL, N-(3-oxoacyl)homoserine lactone; 3-hydroxyAHL, N-(3-hydoxyacyl) homoserine lactone and AHL, N-acylhomoserine lactone, where R ranges from C1 to C15. The acyl side chains may also contain one or more double bonds.

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of the target genes which contains a region of dyad symmetry, a 20 base pair palindrome called the lux box (Fuqua et al., 2001; Zhang et al., 2002). Such regulatory regions are often found upstream of the transcriptional start site for genes coding for LuxI-type AHL synthases such that AHL synthesis is subject to auto-amplification and hence AHLs are often referred to as “autoinducers.” Furthermore, a LuxRI pair can be considered as a quorum sensing “module” several of which may be linked as for example in Pseudomonas aeruginosa where LasRI regulates the rhlRI genes (Latifi et al., 1996). he LuxR/AHL paradigm forms the basis of AHL-mediated multicellular behavior in Gram-negative bacteria and is responsible for controlling a variety of population-dependent phenotypes including bioluminescence, swarming, swimming and sliding motility, antibiotic biosynthesis, plasmid conjugal transfer and the production of virulence determinants in animal, fish and plant pathogens (for reviews see (Swift et al., 1999; Swift et al., 2001; Williams, 2002). It is also apparent that AHL-mediated QS systems are expressed during the biofilm mode of growth and contribute to the development and dispersal of surface-associated bacterial biofilm communities. AHL production during the bioilm mode of growth he possibility that bacterial cell-to-cell communication might contribute to the biofilm lifestyle was proposed in 1993 (Williams and Stewart, 1993). One of the earliest experimental indications that AHL-dependent QS contributed to the biofilm mode of growth came from studies of the recovery rates of the ammonia oxidizer, Nitrosomonas europaea (Batchelor et al., 1997). Planktonic cells starved of ammonia exhibited long lag phases prior to nitrite production following supplementation with ammonium. In contrast, biofilm populations of N. europaea colonizing sand or soil particles in continuous flow, fixed column reactors exhibited no lag phase prior to nitrite production even after long term ammonium starvation. Since supplementation of starved planktonic cells with both ammonium and N-(3-oxohexanoyl)homoserine lactone (3-oxo-C6-HSL) reduced the lag phase by a factor of five and as AHLs were detected in N. europaea culture supernatants, the authors suggested that the rapid recovery of high-density biofilm populations was due to the production and accumulation of AHLs to levels, not possible in low density cell suspensions, where they may be responsible for controlling metabolism or other properties of the biofilm community (Batchelor et al., 1997). Preliminary evidence for AHL production in natural biofilms was provided by McLean et al. (1997) who reported that AHLs could be detected in aquatic biofilms growing on submerged stones but not from rocks lacking a biofilm. Similarly, AHLs have been detected in microbial mats (Bachofen and Schenk, 1998). hese are coherent macroscopic accumulations of micro-organisms which have formed layered structures on solid surfaces and sediments and are found in habitats such as thermal springs, pond bottoms, tidal regions and in algal blooms. However in both studies, neither the identity of the biofilm organisms present nor their in vitro AHL profiles were presented. Stickler et al. (1998) screened bacterial isolates taken from biofilms growing on indwelling urethral catheters for AHL production using an AHL biosensor. Of the 19 isolates obtained, 14 P. aeruginosa strains were reported to be AHL positive and a further five isolates (Providencia stuartii, Proteus mirabilis, Morganella morganii, Escherichia coli, and Klebsiella pneumoniae) were either weakly positive or negative.

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Indwelling urethral catheters which had become colonized with bacterial biofilms were also removed from patients undergoing long-term bladder catheterization and cut into sections. Four of nine catheter sections gave positive biosensor reactions for AHLs while unused catheters were negative suggesting that AHLs were being produced in situ in the catheter associated biofilms (Stickler et al., 1998). P. aeruginosa biofilms were also shown to produce AHLs in the catheter in both an in vitro bladder model and ex vivo in catheters which had been freshly removed from the patient. hese data suggest that AHLs are present in biofilms and produced in vivo during a clinically significant infection. However, the use of AHL biosensors alone can give rise to false positive results given that AHL biosensors also can respond to unrelated molecules such as the cyclic dipeptides (Holden et al., 1999). Consequently this work needs to be interpreted with some caution and requires further chemical characterization of the compounds responsible for AHL biosensor activation. For P. aeruginosa, the major AHLs produced are N-(3-oxododecanoyl)homoserine lactone (3-oxo-C12-HSL) and N-butanoylhomoserine lactone (C4-HSL). hese AHLs are synthesized via LasI and RhlI and activate LasR and RhlR respectively (Pearson et al., 1994; Winson et al., 1995). To unequivocally establish that AHLs are produced during the growth of P. aeruginosa biofilms in flow cells, Charlton et al. (2000) devised a quantitative method for 3-oxo-AHLs using gas chromatography (GC) coupled with mass spectrometry. 3-oxo-AHLs such as 3-oxo-C12-HSL but not un-substituted AHLs such as C4-HSL can be assayed sensitively after derivatization with pentafluorobenzyloxime (PFBO) which converts the B-ketone of the 3-oxo-AHL to an oxime (Charlton et al., 2000). Using this approach, the concentration of 3-oxo-C12-HSL present in the P. aeruginosa biofilm was found to be in excess of 600 MM and around 14 MM in the biofilm effluent. P. aeruginosa was also found to produce significant amounts of N-(3-tetradecanoyl) homoserine lactone (3-oxo-C14-HSL), N-(3-oxodecanoyl) homoserine lactone (3-oxo-C10-HSL) and N-(3oxo-octanoyl) homoserine lactone (3-oxo-C8-HSL) (Charlton et al., 2000). Although the biological significance of these additional AHLs is not known, 3-oxo-C12-HSL levels in biofilms appear to be far higher than those previously estimated to be present in cell free supernatants prepared from planktonic cultures of P. aeruginosa (1 MM) (Pearson et al., 1995). he high biofilm concentration of 3-oxo-C12-HSL may reflect the partitioning and equilibration of the hydrophobic 3-oxo-C12-HSL molecule between the intra- and extracellular compartments or the restricted diffusion of 3-oxo-C12-HSL possibly as a consequence of interactions between the QS signal molecule and extracellular biofilm matrix components. Since AHLs are rapidly inactivated through lactonolysis at alkaline pHs (Yates et al., 2002), it is also possible that conditions within the biofilm matrix are less likely to promote ring opening. hus the concentration of 3-oxo-C12-HSL which accumulates is a balance between production and turnover within the biofilm. 3-oxo-C12-HSL production will clearly be dependent on substrate availability, the levels of LasI and LasR and also RsaL, which negatively regulates lasI expression (Rampioni et al., 2006). P. aeruginosa chronically colonizes the lungs of individuals with cystic fibrosis (CF) (Lyczak et al., 2002). Both AHLs, lasR and lasI transcripts have been detected directly in CF sputum obtained from patients colonized with P. aeruginosa (Middleton et al., 2002; Erickson et al., 2002). Using an indirect method to assay for AHL production in CF sputum, Singh et al. (2000) incubated 14C-methionine (which is incorporated into AHLs via SAM

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during AHL biosynthesis) to show that de novo AHL synthesis occurs in sputum incubated ex vivo for 4 hours. he method was subsequently used to determine the relative ratios of 3-oxo-C12-HSL and C4-HSL produced by laboratory and CF isolates of P. aeruginosa in broth, biofilms and CF sputum. From the profiles obtained, the authors concluded that the similarity in the ratios obtained from biofilms and CF sputa in contrast to broth (where 3-oxo-C12-HSL was produced at a rate between 3 and 10 times that of C4-HSL) supports the hypothesis that P. aeruginosa forms biofilms in CF sputum (Singh et al., 2000). AHL-dependent quorum sensing and bioilm development Biofilms are generally considered to develop in a stepwise manner from initial attachment to cell migration, aggregation and microcolony stages through to maturation and dispersal (O’Toole et al., 2000; Kjelleberg and Molin, 2002; Sauer et al., 2002; Klausen et al., 2003; Webb et al., 2003) (see also MacEachran and O’Toole, this volume; Pamp et al., this volume). Intuitively gene regulation in such a complex community would be predicted to be tightly regulated and to involve communication between the individuals within the population such that a concerted response can be elicited. he detection of AHL production within biofilms raises questions with respect to the contribution of QS to each stage of biofilm development and the identity of any AHL-regulated biofilm specific genes. hese questions have been addressed experimentally by examining the biofilm phenotypes of strains with mutations in genes which code for AHL synthesis (e.g. luxI homologues) or mediate the AHL-response (luxR homologues). AHL synthase mutants are especially useful in this context since the parental phenotype can be restored by the exogenous provision of the cognate AHL as well as by supplying an intact copy of the mutated gene. Here we review the evidence for the contribution of specific QS-dependent genes to adherence, aggregation, biofilm maturation and dispersal in AHL-producing Gram-negative bacteria. Table 6.1 summarizes the current literature with respect to the key organisms in which AHL-dependent QS is known to play a role in biofilm development. Adherence and aggregation Bacterial cell attachment to biotic or abiotic surfaces constitutes the initial phase of biofilm development (see also MacEachran and O’Toole, this volume). However there are few examples currently in the literature where AHL-dependent QS is essential for this process. he timing of QS-controlled exopolysaccharide (EPS) production does however play a major role in the surface adherence and colonization potential of Pantoea stewartii (see also Eberl and Fuqua, this volume). his plant pathogen causes vascular wilt and leaf blight in sweet corn and maize. he EPS, stewartan, represents a major virulence factor in P. stewartii, which accumulates in the plant xylem vessels post-infection, and by blocking the free flow of water leads to the characteristic wilting condition (Koutsoudis et al., 2006). he cps operon responsible for stewartan biosynthesis is negatively regulated via a QS system consisting of the LuxRI homologues EsaRI and 3-oxo-C6-HSL (von Bodman et al., 1998). A P. stewartii esaI mutant which is both AHL and EPS negative produces dense bacterial mats in vitro and tightly packed bacterial agglomerates in vivo when compared with the parent which exhibits low levels of adhesion. he addition of exogenous synthetic AHLs

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restores the parental phenotype in a concentration dependent manner. However, the esaRI double mutant was unable to adhere to surfaces but could form aggregates (flocs) in liquid culture (Koutsoudis et al., 2006). In the parent P. stewartii strain, during the initial attachment event, negative regulation of EPS enhances the probability of attachment taking place in an EPS-free environment. In contrast, the esaI mutant is locked into the adhesion phase of biofilm development while the double esaRI mutant is locked into the high cell density mode (Koutsoudis et al., 2006). In P. stewartii the QS system does not actually promote attachment per se but prevents the expression of EPS which would otherwise inhibit attachment at the early stages of biofilm development. As QS systems activate their targets when a population achieves a particular size, the cell numbers present during the initial attachment stages of biofilm development will presumably not be sufficient to elicit a response. In P. aeruginosa, for example, differences in attachment between QS mutants and the parental strains have only been observed in vitro in nutrient-rich growth media (Davies et al., 1998; De Kievit et al., 2001; Shih and Huang, 2002). It is possible therefore that inverse coordinate regulatory systems such as the one described for P. stewartii, which de-repress as the cell population reaches a threshold level may allow early stage attachment to proceed prior to biofilm maturation. Burkholderia cenocepacia is ubiquitous in the environment and is frequently isolated from the sputum of cystic fibrosis sufferers where, in common with P. aeruginosa, it causes chronic respiratory infections (Isles et al., 1984). he QS systems of B. cenocepacia, which are termed cepIR and cciIR respectively employ N-octanoyl homoserine lactone (C8-HSL) and N-hexanoyl homoserine lactone (C6-HSL) (Huber et al., 2001; 2002; Tomlin et al., 2005). Both QS systems regulate virulence factor expression and interestingly the ccIR locus is located within the B. cenocepacia pathogenicity island associated with epidemic strains (Huber et al., 2001; 2002; Tomlin et al., 2005). A detailed quantitative analysis of biofilm attachment and maturation indicated that the cepIR system was not involved in the regulation of attachment in strain B. cenocepacia H111 (Huber et al., 2001;2002). However, in B. cenocepacia strain K56-2, the cciR and cciR cepIR mutants but not the cciI or cciI cepI mutants were reported to show significant reductions in a simple attachment assay (Tomlin et al., 2005). In flow cell chambers however the K56-2 cepR and cepI mutants were attachment-impaired. hus in B. cenocepacia, the regulation of attachment by AHLdependent QS is both strain and growth environment dependent. Floc formation and aggregation in liquid cultures has been correlated with an increased likelihood of biofilm development (Parsek and Greenberg, 2005). he properties of flocs are similar to those of static biofilms with the component cells embedded within an extracellular matrix (Parsek and Greenberg, 2005). he enteropathogen Yersinia pseudotuberculosis possesses two pairs of LuxRI homologues (YpsRI and YtbRI) and produces multiple AHLs (Atkinson et al., 1999). Mutation of ypsR resulted in severe flocculation in liquid culture, upregulation of flagellin synthesis and a hypermotile phenotype on semi-solid agar plate assays (Atkinson et al., 1999). Interestingly, mutation of the AHL synthase gene, ypsI did not promote cellular aggregation although the same motility phenotype is apparent. Since ypsI expression and hence AHL production is not YpsR-dependent (Atkinson et al., 1999) it is possible that YpsR, in common with EsaR from E. stewartii, represses the target structural genes required for cellular aggregation. Consequently flocculation may be repressed at

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Serratia marcescens (liquefaciens)

Burkholderia cenocepacia

Pantoea stewartii

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N H H

O

O

O

O

O

Major AHL(s) produced

SwrI/R

CepR/I CciR/I

EsaR/I

QS loci

Maturation

Adherence Maturation

Adherence Aggregation

AHL regulated bioilm phenotype

Huber et al., 2001; 2002; Tomlin et al., 2005

von Bodman et al., 1998; Koutsoudis et al., 2006

References

bsmA (adhesin), bsmB Labbate et al., 2004

?

cps (EPS synthesis)

AHL regulated structural genes contributing to the bioilm phenotype

Table 6.1 Gram-negative bacteria in which AHL-dependent QS contributes to bioilm development

Aeromonas hydrophila

Pseudomonas putida

Pseudomonas aeruginosa

Rhodobacter sphaeroides

Yersinia pseudotuberculosis

O

N H H

N H H

N H H

O

O

O

C4-HSL

O

N H H

O

O

3-oxo-C12-HSL

3-oxo-C10-HSL

C4-HSL

O

3-oxo-C12-HSL

C14:1-HSL

C6-HSL

O

3-oxo-C6-HSL

O

O

O

O

O

O

O

O

O

O

O

N H H

OH

N H H

O

O

N H H

N H H

O

O

O

O

O

O

AhyR/I

PpuR/I

LasR/I RhlR/I

CerR/I

YpsR/I

Maturation

Maturation

Maturation Dispersal

Aggregation

Aggregation

?

psoA (Biosurfactant)

rhlA (Rhamnolipids) lecA, lecB (lectin production)

?

?

Lynch et al., 2002

Dubern et al., 2006

Davies et al., 1998; Heydorn et al., 2002; Hentzer et al., 2003; Davy et al., 2003; Bjarnsholt et al., 2005a; Bjarnsholt et al., 2005b; Diggle et al., 2006; Tielker et al., 2005

Puskas et al., 1997

Atkinson et al., 1999

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low cell densities but becomes activated by AHL-dependent de-repression of YpsR as the population becomes “quorate.” Y. pseudotuberculosis is very closely related to Yersinia pestis, the causative agent of bubonic and pneumonic plague which possesses a closely related AHL-dependent quorum sensing system (Atkinson et al., 2006). his pathogen is transmitted mainly via the bites of infected fleas and forms dense aggregates within the flea gut. Physical blockage of the flea proventriculus which connects the midgut to the esophagus by bacterial biofilm-like aggregates impede the passage of a blood meal into the mid-gut such that Y. pestis is conveyed to the bite site by regurgitation when blocked fleas next attempt to feed, so transmitting the pathogen. Given that Y. pseudotuberculosis ypsR mutants aggregate it is possible that AHL-dependent quorum sensing is involved in biofilm formation in the flea gut. However, Jarrett et al. (2004) have shown that a Y. pestis strain with mutations in both AHL synthases as well as in luxS was still able to produce a biofilm on a glass surface and infect and block fleas as well as the parent strain. However, this does not rule out a role for QS in flea blockage if the Y. pestis QS system negatively regulates aggregation in a manner analogous to that of YpsR and EsaR. he aggregation phenotype would only become apparent on mutating the corresponding luxR homologue. Flocculation is also QS-controlled in the free living photoheterotroph Rhodobacter sphaeroides which possesses a QS system consisting of the LuxRI homologues CerRI and 7,8-cis-N-(tetradecanoyl)homoserine lactone (C14:1-HSL). When grown in liquid culture R. sphaeroides cerI mutants flocculate but the aggregates can be dispersed by adding an exogenous source of C14:1-HSL. he cerI mutant also produces up to 30 times more exopolysaccharide than the parent and these data, when taken together, indicate that the QS system of R. sphaeroides is involved in repressing a biofilm-associated phenotype (Puskas et al., 1997). AHL-dependent QS therefore regulates cell aggregation in both Yersinia and Rhodobacter. Maturation Although insights into the contribution and role of AHL-dependent QS to biofilm maturation have primarily been derived from studies with P. aeruginosa, the impact of QS on biofilm maturation has also been investigated in Aeromonas, Burkholderia and Serratia. Pseudomonas he LasRI/RhlRI QS system of P. aeruginosa is the most extensively investigated and understood QS cascade in Gram-negative bacteria. Transcriptomic studies of planktonic cells have revealed that the LasRI- and RhlRI-regulated genes (over 500) are scattered throughout the chromosome supporting the view that the P. aeruginosa quorum sensing circuitry constitutes a true global regulatory system (Hentzer et al., 2003; Schuster et al., 2003; Wagner et al., 2003). P. aeruginosa readily forms biofilms on both biotic and abiotic surfaces which are problematic in both industrial and healthcare settings. Initial work on the impact of QS on biofilm formation in P. aeruginosa indicated that LasRI but not the RhlRI system was an integral part of the biofilm maturation process and governed microcolony formation (Davies et al., 1998). his was because lasI mutants formed flat undifferentiated biofilms which were much more susceptible to dispersal by detergents such as sodium dodecyl sulfate than the parent strain. hese interesting data suggested that

Quorum Sensing and Bioilms

AHL-dependent QS played an important role in determining the architecture of a biofilm. However, other laboratories reported that P. aeruginosa parent and QS mutant biofilms were either architecturally indistinguishable or showed only minor differences (Heydorn et al., 2002; Bjarnsholt et al., 2005a; Bjarnsholt et al., 2005b). hese discrepancies are probably the consequence of the different P. aeruginosa PAO1 strains, environmental conditions (e.g. carbon source) and biofilm fermenter systems employed. Interestingly, both Hentzer et al. (2003) and Bjarnsholt and Givskov (2007) suggest that the structural stability of P. aeruginosa biofilms may be compromised in QS mutants. Furthermore, it has been suggested that one possible explanation for the contradictory results on the role of QS in biofilm formation in P. aeruginosa may be the potential accumulation of secondary mutations in QS mutants (Beatson et al., 2002). If this is the case a biofilm population will be heterogeneous and might actually be composed of a series of distinct niches populated by clonally compartmentalized bacteria (Parsek and Greenberg, 2005; Bjarnsholt and Givskov, 2007). To determine whether, when and where the AHL synthase genes lasI and rhlI are expressed during P. aeruginosa biofilm growth, both gfp and lacZ reporter gene fusions have been employed (De Kievit et al., 2001; Sauer et al., 2002). During the course of an 8 day biofilm flow cell experiment, lasI expression was found to decrease progressively over time whereas rhlI expression did not change appreciably. For both genes, spatial analysis indicated that expression was maximal in cells located at the substratum but decreased with increasing biofilm height (De Kievit et al., 2001). In a complementary study which employed lasB and rhlA reporter fusions respectively as markers for las and rhl activity, Sauer et al. (2002) reported that the las system was expressed before rhl during the early irreversible attachment stage whereas the rhl system was not induced until the early stages of maturation when cell clusters became progressively layered. Interestingly using alcian blue as a stain for acidic EPS, lasI mutant cells, in contrast to the parent PAO1 cells, were observed to be more closely packed together within the biofilm and not separated by EPS (Sauer et al., 2002). Scanning confocal microscopy of P. aeruginosa biofilms formed in continuous flow chambers has revealed a complex architecture where the bacteria are embedded in an extracellular matrix containing open channels which facilitate the flow of fresh nutrients and bacterial metabolites including QS signals throughout the biofilm. Since QS has been linked to the formation of structured biofilms, it is likely that the products of certain QS-regulated genes are essential for this process. In this context, rhamnolipid biosurfactants appear to be required for the maintenance but not formation of microcolonies and open channels in P. aeruginosa biofilms. his is because mutation of the rhamnolipid biosynthesis gene rhlA results in a strain which retains the ability to form but not maintain channels surrounding P. aeruginosa biofilm macrocolonies (Davey et al., 2003). his finding was confirmed by the exogenous provision of rhamnolipids to the rhlA mutant which partially restored the biofilm architecture to that of the parent strain. Furthermore, rhlA::gfp expression coincided with microcolony formation and continued throughout biofilm development where cell density was high. Since rhlA is controlled by RhlR/C4-HSL, these findings establish a direct link between QS and biofilm functionality. Indeed the importance of the timing of rhlA expression was further highlighted by experiments in which inappropriate overproduction or supply of rhamnolipids disrupted biofilm development (Davey et al., 2003). hese

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findings were extended by Lequette and Greenberg (2005) who showed that expression of a rhlA-gfp reporter in P. aeruginosa biofilms was low during early biofilm development even in the presence of exogenously supplied C4-HSL but was induced once microcolonies of > 20 Mm had formed. hey also noted that rhlA was preferentially expressed in the stalks rather than caps of mature microcolonies. In common with P. aeruginosa, biosurfactant production in Pseudomonas fluorescens, Serratia liquefaciens and B. cenocepacia is regulated by AHL-dependent QS. In Pseudomonas putida PCL1445 the LuxRI pair, PpuRI control the production of two cyclic lipodepsipeptides termed putisolvins I and II respectively in conjunction with long chain AHLs such as 3-oxo-C10-HSL and 3-oxo-C12-HSL (Dubern et al., 2006). Mutation of either ppuR or ppuI abolishes putisolvin production and results in the formation of dense biofilms on wells in PVC microtiter plates. Putisolvin production in the ppuI mutant can be restored by the addition of the long chain AHLs which consequently no longer develops the characteristic dense biofilm on PVC (Dubern et al., 2006). Other AHL-regulated P. aeruginosa genes which are involved in biofilm development are lecA and lecB which code for lectins which preferentially bind hydrophobic galactosides and L-fucose respectively Diggle et al., 2006; Tielker et al., 2005). Both genes are regulated by RhlR/C4-HSL as well as by the alternative sigma factor RpoS (Winzer et al., 2000). By using a lecA-gfp translation fusion and immunoblot analysis of the biofilm extracellular matrix, Diggle et al. (2006) showed that lecA is expressed in biofilm grown P. aeruginosa cells. In static biofilm assays on both polystyrene and stainless steel, biofilm depth and surface coverage was reduced by mutation of lecA. Biofilm surface coverage on steel coupons was also inhibited by growth in the presence of either isopropylthiol-A-D-galactoside (IPTG) or p-nitrophenyl-A-D-galactoside (NPG). Furthermore, mature wild-type biofilms formed in the absence of these hydrophobic galactosides could be dispersed by the addition of IPTG. In contrast, addition of p-nitrophenyl-A-L-fucose (NPF) which has a high affinity for the P. aeruginosa LecB lectin, had no effect on biofilm formation or dispersal. hese data suggest that the observed effects on biofilm formation were due to the competitive inhibition of LecA-ligand binding. Similar results were also presented for P. aeruginosa biofilms grown under dynamic flow conditions on steel coupons, suggesting that LecA contributes to P. aeruginosa biofilm architecture under different environmental conditions. Mutation of lecB has also been reported to result in defective biofilm formation (Tielker et al., 2005). hus both lectins can contribute to biofilm formation although only LecA specific ligands appear to be capable of disrupting pre-formed biofilms. As yet, the nature of the carbohydrate ligands recognized by LecA and LecB and present in P. aeruginosa biofilms have not been identified. In P. aeruginosa, it has recently been established that extracellular, genomic DNA is a major component of the biofilm matrix (Whitchurch et al., 2002) (see also Pamp et al., this volume). he DNA released appears to be to be localized primarily in the stalks of microcolonies (Allesen-Holm et al., 2006). Biofilm formation in flow chambers was also inhibited by DNase I which was also shown to be capable of dissolving pre-formed biofilms in a biofilm age-dependent manner (Whitchurch et al., 2002). he mechanism by which this DNA is released may involve the lysis of a subpopulation of the culture although other mechanisms dependent on phage or membrane vesicles cannot be ruled out. DNA release

Quorum Sensing and Bioilms

in P. aeruginosa does however involve AHL-dependent quorum sensing. his is because the level of extracellular DNA released by a lasI rhlI double mutant was significantly reduced compared with the parent strain and could be restored by supplying C4-HSL and 3-oxoC12-HSL (Allesen-Holm et al., 2006). he differences noted between the biofilms formed by the parent and lasI rhlI double mutant with respect to the amount of extracellular DNA in the region of the biofilm closest to the substratum also correlated with the spatial expression of lasI and rhlI in P. aeruginosa biofilms as reported by De Kievit et al. (2001). Although the AHL-dependent target genes involved in mediating DNA release have yet to be identified, a pqsA mutant defective in the production of the 4-quinolone QS signal molecule, 2-heptyl-3-hydroxy-4-quinolone (PQS), in common with the lasI rhlI mutant, generates low amounts of extracellular DNA (Allesen-Holm et al., 2006). Furthermore in a static biofilm model, Diggle et al. (2003) have shown that the addition of PQS to a wild-type P. aeruginosa strain greatly enhances biofilm surface coverage. Since AHL-dependent QS and PQS signaling are intricately linked, the target genes involved may well be co-regulated by both QS systems. Aeromonas he fish pathogen Aeromonas hydrophila which readily forms biofilms in a variety of environmental niches employs an AHL-based QS system consisting of the LuxR homologue, AhyR and the LuxI homologue AhyI which directs the synthesis of C4-HSL (Swift et al., 1997; Lynch et al., 2002). Mutation of either ahyR or ahyI result in the loss of extracellular serine- and metalloproteases which can be restored in an ahyI mutant by the addition of C4-HSL. A. hydrophila readily forms complex three dimensional biofilms on stainless steel with microcolonies covering up to 50% of the available surface area (Lynch et al., 2002). In a continuous flow chamber, C4-HSL and the AhyI protein have both been detected in the developing biofilm while mutation of ahyI resulted in the formation of immature biofilms lacking microcolonies. he parenteral biofilm architecture could be partially restored by the provision of C4-HSL. Mutation of ahyR does not result in the loss of C4-HSL biosynthesis, since ahyR is not absolutely required for ahyI expression, and does not influence biofilm depth or microcolony formation. However, biofilm surface coverage in the ahyR mutant increased to around 80% of the available surface area (Lynch et al., 2002). While these data support a role for AHL-dependent QS in A. hydrophila biofilm maturation, the specific QS dependent genes involved have not yet been identified. Burkholderia From a transposon insertion library, Huber et al. (2001) obtained a number of B. cenocepacia H111 mutants defective in biofilm maturation on polystyrene surfaces one of which was also AHL-negative. Mapping of the insertion site revealed that the transposon had inserted into the cepR gene. Since CepR positively regulates cepI, this would also account for the loss of AHL production. Both cepI and cepR mutants exhibited biofilm maturation defects in that both were arrested at the microcolony stage and, in contrast to the parent strain, failed to colonize the entire surface. he parental biofilm phenotype was restored to the cepI mutant by providing exogenous C8-HSL firmly establishing a role for QS in B. cenocepacia biofilm maturation (Huber et al., 2001). Similar architectural defects in B.

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cenocepacia QS mutant biofilms have also been noted by Tomlin et al. (2005). Although the QS-regulated structural genes responsible for biofilm maturation in B. cenocepacia have not yet been characterized, three higher level regulators (yciR, suhB and yciL) of the cep QS system have been identified in strain H111 as a consequence of their biofilm phenotype and reduced C8-HSL synthesis (Huber et al., 2002). However, neither exogenous C8-HSL addition nor provision of cepR in trans restored the biofilm defects in any of these mutants. Consequently while the mechanism by which they influence cep-dependent QS remains to be elucidated it is clear that the cep quorum-sensing system is a major checkpoint for biofilm maturation in B. cenocepacia in strains H111 and K56-2. To determine whether AHL-dependent QS contributed generally to biofilm maturation in the Burkholderia cepacia complex, Wopperer et al. (2006) introduced an AHL-degrading enzyme (AiiA) via a plasmid into a large number of different strains. Although dramatic differences in the biofilm forming capacity of these strains on polystyrene was noted, it was apparent that the expression of aiiA reduced biofilm biomass in many but not all strains. Serratia Serratia liquefaciens MG1 (also termed Serratia marcescens MG1) has a well characterized AHL-driven QS system which governs surface colonization through swarming motility by regulating the expression of the biosurfactant, serrawettin (Eberl et al., 1996). his strain forms an unusual biofilm when grown in flow chambers which consists of cell aggregates, chains of cells and elongated filamentous cells. Disruption of the AHL synthase swrI resulted in the loss of C4-HSL synthesis and the formation of thin, immature biofilms lacking aggregates and filamentous cells. Restoration of wild-type biofilm architecture to the swrI mutant was achieved by providing C4-HSL exogenously (Labbate et al., 2004). Two C4-HSL-dependent genes termed bsmA and bsmB were identified from a screen of transposon mutants for poor biofilm producers. Both mutants failed to form the biofilm cell aggregates and cell chains characteristic of the parent strain unless genetically complemented. Both BsmA and BsmB appear to be involved in cellular aggregation since the bsmA mutant exhibited excess aggregation while the bsmB mutant failed to aggregate, data which implies that cellular aggregation is likely to be a highly regulated process in S. liquefaciens. Although the functions of BsmA and BsmB are not known, BsmA has been suggested to be an adhesin given the excess adhesion and aggregation observed when bsmA was expressed in multicopy (Labbate et al., 2004). Interestingly, Serratia biofilm morphology is also dependent on nutrient cues such that the contribution of QS to biofilm architecture can be over-ridden by the growth medium used to culture the organism (Rice et al., 2005). Dispersal As biofilms develop to maturity there will be dispersal of aggregates from the main body of the multicellular structure. Dispersal has been considered as a passive process which is a consequence of the immediate environmental conditions and has usually been considered to occur by erosion, sloughing, abrasion or predator grazing (Bryers, 1988). While passive dispersal will undoubtedly be a factor in the dissemination of biofilm aggregates, active dispersal driven by the individuals within the population is accepted as an alternative strategy which bacteria employ to control dissemination of single cells or clusters of cells still

Quorum Sensing and Bioilms

bound by the biofilm matrix. Different types of dispersal mechanism have been reported in Actinobacillus actinomycetemcomitans Staphylococcus epidermidis and P. aeruginosa (Kaplan et al., 2003; Purevdorj-Gage and Stoodley, 2004). An active dispersal mechanism could conceivably be triggered at a predetermined point such as the attainment of a critical mass. Such a mechanism could be QS-mediated. While this is an attractive notion, dispersal driven by AHL-mediated QS is poorly understood. Purevdorj-Gage et al. (2005) examined this process in P. aeruginosa and found that although there was no difference in biofilm development and coverage between the parent strain and a lasI rhlI double mutant, seeding dispersal was severely impaired in the mutant. Furthermore, this phenomenon could not be attributed to swimming motility since both the double mutant and parent remained motile on semi-solid agar plate assays. he authors suggested that QS was likely to be involved in the differentiation process possibly by sensing cell population density and nutrient depletion within the periphery of biofilm clusters (Purevdorj-Gage et al., 2005). As noted previously, biosurfactant production in a variety of Gram-negative bacteria is AHL-dependent and required for swarming motility since biosurfactants promote population migration by reducing surface tension (Daniels et al., 2004). In biofilms, coupling the production of biosurfactants with high cell population densities may promote biofilm dispersal. While rhamnolipid synthesis is involved in biofilm channel maintenance in P. aeruginosa, rhamnolipids do not appear to be required for seeding dispersal (PurevdorjGage et al., 2005). In P. putida, putisolvins inhibit biofilm formation and can promote the breakdown of existing biofilms (Kuiper et al., 2004). Consequently it has been suggested that high P. putida biofilm population densities stimulate the release of bacterial cells from the biofilm as a dispersal mechanism for escaping nutrient depletion and encouraging the colonization of other environmental niche(s) (Dubern et al., 2006). AHL-dependent QS is also clearly involved in the dispersal of S. marcescens biofilms (Rice et al. (2005). After 70 to 80 hours of growth in a flow cell, the mature filamentous biofilm formed by the wild-type strain sloughed off from the surface. Similar results were obtained with an S. marcescens swrI mutant supplied with C4-HSL. However, no sloughing was observed in the swrI mutant grown in the absence of C4-HSL. Since the sloughing profile of a biosurfactant-deficient S. marcescens strain was the same as the parent, this suggests that the QS-mediated dispersal is likely to involve a range of gene products (Rice et al., 2005). Transcriptomic and proteomic investigations of the bioilm phenotype Rapid developments in microarray technology and mass spectrometry have facilitated the genome wide analysis of bacterial adaptations to different growth environments at both the transcriptional and proteomic levels. However, the application of these technologies to biofilm grown bacteria have so far had to treat biofilms largely as homogeneous entities and consequently the profiles obtained only show the average across a biofilm grown to a specific developmental stage. Different cells within distinct areas of a biofilm will experience differential access to nutrients and oxygen and this in turn results in a variety of micro-environmental conditions which will dictate diversity in gene expression patterns (Werner et al., 2004) (see also Kjelleberg and Givskov, this volume; Webb, this volume).

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Genome-wide studies of biofilms have been viewed either by treating biofilms as a single entity in which gross phenotypic changes emerge as particular genes are mutated or alternatively from the perspective of a specific developmental stage, i.e. attachment, maturation or dispersal. In each situation, there have been conflicting views as to whether a particular biofilm architecture is governed by a designated set of genes which constitute the “biofilm regulon.” In a microarray study by (Whiteley et al., 2001) in which the transcriptomes of planktonic and continuous culture biofilm P. aeruginosa cells were compared, only 1% of the genes (73 about of 5570) were found to be differentially expressed (2-fold or greater). he most highly activated genes in the biofilm array were those of a temperate bacteriophage related to Pf1 while genes for pili and flagellar biosynthesis were strongly downregulated a finding which correlates with their role in the early but not later stages of biofilm maturation (Whiteley et al., 2001). No differences in the expression of the las or rhl QS systems was reported. In a comparable study by Hentzer et al. (2005) the gene expression profiles of biofilm grown P. aeruginosa cells were noted to most closely resemble those of stationary phase cells. Similar findings were also reported by Waite et al. (2005). hese studies generally support the notion that biofilm development does not depend on a unique genetic program, i.e. the “biofilm regulon,” but instead depends on a series of adaptive responses to the prevailing micro-environmental conditions. Hentzer et al. (2005) did however identify a subset of AHL-regulated genes in biofilm grown P. aeruginosa that were differentially expressed compared with planktonic cells. hese included, for example, the lectins, LecA (PA-1L) and LecB, the mutation of which results in biofilm-defective strains (Tielker et al., 2005; Diggle et al., 2006) the rhamnolipid biosynthetic gene, rhlC, genes coding for pyocyanin and hydrogen cyanide biosynthesis and pvdQ (Hentzer et al., 2005). While the physiological function of the latter is not known, it is required for synthesis of the siderophore pyoverdine and also has AHL-inactivating activity. PvdQ is an amidase/acylase which cleaves the amide bond of AHLs with fatty acid side chains ranging from 11 to 14 carbons (Sio et al., 2006) and may therefore represent a post-translational mechanism for modulating 3-oxo-C12-HSL levels within biofilms. Proteomic approaches have also provided useful data highlighting the multiple phenotypes displayed by P. aeruginosa during biofilm development (Sauer et al., 2002). he levels of proteins involved in motility, alginate production and quorum sensing were observed to vary significantly with over 50% of the proteome reported to exhibit a six-fold or greater change in expression level when planktonic cells were compared with mature biofilm cells (Sauer et al., 2002). When a lasI mutant was compared with the parental strain, las-dependent QS was observed to be important in late but not early biofilm stages (Sauer et al., 2002). Apart from P. aeruginosa, proteomics has only been used to investigate the role of QS in biofilm development in P. putida (Arevalo-Ferro et al., 2005) where it has emerged that a major set of QS-regulated proteins overlaps with those identified as differentially regulated in sessile compared with planktonic cells. Interestingly, the majority of surfaceinduced proteins were negatively regulated by QS in biofilm cells, i.e. were upregulated in the ppuI mutant compared with the parent strain (Arevalo-Ferro et al., 2005). As post-genomic and imaging technologies are further refined particularly with respect to investigating gene expression in single bacterial cells, the spatial and temporal impact of AHL-dependent QS on surface adaptation and the nature of the AHL-regulated structural

Quorum Sensing and Bioilms

genes involved in determining biofilm architecture in pseudomonads and other AHL-producers should become more apparent (see also Kjelleberg and Givskov, this volume). QS in multi-species bioilms Much of the work on AHL-dependent QS and biofilms has focused on single-species biofilms. However, given that natural biofilms are often polymicrobial and that many different Gram-negative bacteria produce the same or different AHLs, it is possible that AHL-dependent QS also contributes to the maturation of mixed species biofilms. Although LuxR homologues preferentially respond to a specific AHL, they are capable of responding to a range of structurally related AHLs albeit less sensitively (McClean et al., 1997; Winson et al., 1998). Furthermore, bacteria such as Escherichia coli, Klebsiella and Salmonella which do not themselves produce AHLs, nevertheless possess a LuxR homologue termed SdiA which responds to AHLs produced by other bacteria (Ahmer, 2004). In Salmonella enterica SdiA regulates a number of genes including several involved in adherence to host cells, extracellular matrix proteins and resistance to complement-mediated killing (Ahmer, 2004). In E. coli, SdiA regulates genes involved in glutamate metabolism including gadA which codes for a glutamate decarboxylase which plays an important role in acid resistance. Interestingly C6-HSL was reported to enhance the tolerance of E. coli to acid in an sdiA-dependent manner (Houdt et al., 2006). Consequently such AHL signal interception could also play a role in the maturation of biofilms in mixed communities containing AHL-producers and bacteria such as E. coli or S. enterica. When dialysis tubing containing spent culture supernatants with or without AHLs was placed in lakewater, McLean et al. (2005) noted that the nature of the adherent population changed with a notable reduction in the adherent population biodiversity. Exogenously supplied AHLs have also been shown to influence bacterial sludge community composition (Valle et al., 2004). In laboratory studies, AHL-mediated communication between P. aeruginosa and B. cenocepacia in mixed biofilms has been investigated and reported to be unidirectional in that B. cenocepacia responds to the P. aeruginosa AHLs but not vice versa (Riedel et al., 2001). Interestingly, differences in biofilm architecture with respect both to the spatial distribution of microcolonies and the formation of mixed microcolonies was observed when B. cenocepacia H111 (AHL positive) was grown with either a P. aeruginosa AHL producer or a non-producer (Riedel et al., 2001). QS has also been reported to influence interactions between P. aeruginosa and Agrobacterium tumefaciens in mixed cultures (An et al., 2006). Although P. aeruginosa, in both planktonic and biofilm modes of growth, rapidly outgrows A. tumefaciens, P. aeruginosa quorum sensing mutants showed reduced growth yields in planktonic co-cultures. In older biofilms, while the amount of A. tumefaciens biomass decreased in co-culture with the wild-type P. aeruginosa, it remained constant with P. aeruginosa lasR rhlR mutant (An et al., 2006). hese experiments provide a useful basis from which to begin dissecting the role of QS in promoting competitive interactions within microbial communities. AHLs, bioilms, and interactions with higher organisms Apart from their role as signal molecules in QS, certain AHLs and notably 3-oxo-C12HSL, have a broad range of biological activities which can influence the interactions

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between AHL-producing bacteria and other organisms. For example, 3-oxo-C12-HSL inhibits both growth and agr-mediated quorum sensing in S. aureus (Qazi et al., 2006), filamentation in Candida albicans (Hogan et al., 2004) (see also Hogan et al., this volume), has immune modulatory activity (Telford et al., 1998; Chhabra et al., 2003) and elicits both pro- and anti-inflammatory responses (Smith et al., 2002; Pritchard et al., 2005). 3-oxo-C12-HSL also influences smooth muscle contraction in blood vessels (Lawrence et al., 1999) and exerts a marked bradycardia in rats (Gardiner et al., 2001). hus, for P. aeruginosa, an opportunistic pathogen, 3-oxo-C12-HSL not only appears to function as a QS signal molecule controlling expression of key virulence determinants but also as a means to gain a competitive survival advantage in the presence of other organisms occupying the same ecological niche. Zoospore settlement here is also evidence that higher organisms can “tune-in” to AHL signaling between planktonic and biofilm bacterial communities. An intriguing example of this involves the intertidal green macroalga Ulva which can reproduce asexually via the release of vast numbers of motile, asexual zoospores which initiate a process of surface selection involving the sensing of a surface followed by temporary adhesion. If a surface is suboptimal, the zoospore will detach. One important factor which influences zoospore surface-selection is the presence of a bacterial biofilm and there is a positive correlation between zoospore settlement and bacterial biofilm density ( Joint et al., 2000). his observation led to the discovery that AHLs produced by bacterial biofilms are involved in the attraction of Ulva zoospores to surfaces. From investigations of the response of Ulva zoospores to (a) Vibrio anguillarum mutants defective in AHL production or expressing an AHL-inactivating enzyme, (b) E. coli strains expressing AHL synthases from recombinant plasmids and (c) synthetic AHLs, it emerged that zoospore attraction to a surface is, at least in part, mediated via zoosporemediated AHL detection ( Joint et al., 2002; Tait et al., 2005). Image analysis using GFPtagged V. anguillarum biofilms revealed that zoospores settle directly on bacterial cells and in particular on microcolonies which are sites of concentrated AHL production (Tait et al., 2005) (Figure 6.2). he mechanism of attraction of zoospores does not appear to involve a chemotactic orientation towards the AHLs but instead a chemokinesis in which spore swimming speed rapidly decreases in the presence of AHLs such that they accumulate at the AHL source. Zoospore swimming speed for example decreased more rapidly over wild-type V. anguillarum biofilms when compared with those of an isogenic AHL negative mutant. AHL detection by zoospores causes an influx of calcium and it has been suggested that the reduction in swimming speed occurs through calcium-dependent modulation of the flagellar beat pattern ( Joint et al., 2002). Ulva zoospores can sense a range of different AHLs (Tait et al., 2005) although the chemoresponse was reported to be most marked towards 3-oxo-C12-HSL (Wheeler et al., 2006), an interesting finding given the broad biological activity of this AHL molecule and its capacity to modulate mammalian host cell responses through calcium signaling (Shiner et al., 2006). he reason(s) why Ulva zoospores target bacterial biofilms as a preferred site for attachment remains unclear but bacterial biofilms are clearly an important factor in biofouling. Many studies have shown that microbial biofilms influence the settlement of

Quorum Sensing and Bioilms

A

B

Figure 6.2 Attraction of Ulva zoospores to AHL-producing Vibrio anguillarum bioilm microcolonies. (A) AHL production in bioilm microcolonies of V. anguillarum carrying the gfp-based AHL biosensor luxR-PluxI-RBSII::gfpmut3*-T0 (B) DAPI-stained bioilm showing zoospores settled on a V. anguillarum microcolony. Images are transmission images overlaid with the luorescent color image. Reproduced with permission from Tait et al. (2005). A color version of this igure is available at http://www.horizonpress.com/hsp/supplementary/bioilm/ ch6ig2.jpg.

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marine invertebrates. It has been assumed that grazing organisms can exploit bacterial biofilms as a food source but this is unlikely to be a factor in the attraction of an alga. Consequently, the presence of a bacterial biofilm may signal that the environment is benign for Ulva zoospores. However, given that the interaction is very specific, with zoospores settling directly on bacterial cells in the biofilm, rather than merely attaching in the vicinity, there are likely to be other reasons why zoospores prefer attaching to bacterial biofilms. For example, many green algae do not develop normal morphology when grown under axenic conditions (Provasoli and Pintner, 1980). Recently, Matsuo et al. (2003) have shown that differentiation in the green alga Monostroma oxyspermum depends on the presence of specific bacterial strains, i.e. normal morphology depends on particular bacteria and not on bacteria in general. Consequently the preference of Ulva zoospores for settlement on AHL-producing bacterial biofilms may facilitate a close association between the developing thallus and certain essential bacteria (Tait et al., 2005). Protozoan grazing he growth and survival of bacteria in many different natural environments are constrained to some extent by the activities of predatory protozoa. Adoption of the biofilm mode of growth and the ability to produce toxic secondary metabolites are important bacterial defense mechanisms against grazing by protozoans although the effectiveness of the resistance mechanism depends on the protozoan feeding type (Matz and Kjelleberg, 2005; Weitere et al., 2005) (see also Matz, this volume). Selective predation by protozoa has been suggested to drive the evolution of bacterial grazing resistance strategies including biofilm adaptations, cell-to-cell signaling and the emergence of pathogenic traits (Matz and Kjelleberg, 2005). In contrast to stationary phase bacterial cultures, predation is not likely to be such a problem for exponentially growing cultures since the high rates at which bacterial cell numbers are increasing should be sufficient to compensate for grazing losses. In situations where nutrients are limiting and bacterial growth is restricted, biofilm formation is a useful mechanism for combating predation. Whether bacteria are growing in planktonic cultures or as biofilms, the co-ordination of secondary metabolite production as a function of cell population density through QS is likely to constitute a useful strategy for ensuring that sufficient local concentrations of these toxic metabolites are attained. QS may therefore facilitate resistance to predation by controlling the architecture of biofilms and by regulating the production of anti-protozoal secondary metabolites. Consequently it can be predicted that mutants with defective AHL-dependent QS systems might be more susceptible to predation. So far, this has been shown to be the case for S. marcescens, Chromobacterium violaceum and P. aeruginosa (Matz and Kjelleberg, 2005). he differentiated biofilm phenotype of S. marcescens MG1 has been described earlier in this chapter. Under static growth conditions, MG1 forms biofilms with microcolony-like structures but in flow cells forms highly differentiated filamentous biofilms consisting of cell chains and clusters (Labbate et al., 2004). Experiments with a suspension feeder flagellate (Bodo saltans) and a surface feeder (Acanthamoaeba polyphaga) have revealed that while QS is not involved in the grazing resistance of S. marcescens microcolony-type biofilms, the QScontrolled biofilm specific differentiation into filaments and cell chains does confer effective

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resistance against protozoan grazing (Queck et al., 2006). While there was no apparent role for QS in the development of resistance against protozoan grazing during the early stages of biofilm development by S. marcescens, QS clearly places a key role in protection of the late stage differentiated biofilm. Comparable findings have also been obtained for P. aeruginosa where the surface feeding flagellate Rhynchomonas nasuta induced microcolony formation in both parent and QS mutant (lasR and lasR/rhlR) strains although the latter showed reduced resistance to grazing (Matz et al., 2004). he lasR/rhlR mutant however showed a distinct lack of toxicity against the flagellates during late biofilm development. Hence AHL-dependent QS clearly contributes to the resistance of Gram-negative bacteria to protozoan gazing particularly with respect the production of inhibitors which confer protection against a wide range of protozoan feeding types (Weitere et al., 2005). A detailed account of bacterial biofilm-protozoan interactions is provided by Matz, this volume. AHL-dependent QS in bioilms as an antibacterial target Bacterial biofilm-centered infections are notoriously difficult to treat and conventional antibiotic therapy often fails as a consequence of the tolerance of biofilm bacteria (Davies, 2003). When compared with planktonic cells, biofilm bacteria almost always exhibit a large (up to 1000 fold) increase in resistance to diverse antimicrobial agents (see also Kjelleberg and Givskov, this volume). While the mechanisms underlying tolerance are not well understood, they are clearly multifactorial and include slow growth and metabolic rates, restricted penetration and specific resistance mechanisms (Davies, 2003). Given that more than 80% of bacterial infections in humans have been proposed to involve biofilms (Davies, 2003), there is an urgent need to discover novel targets against which effective therapeutic agents for the treatment of biofilm-centered chronic infections can be developed. In this context, AHL-dependent QS is an attractive target. his is because biofilms formed by P. aeruginosa QS mutants exhibit enhanced susceptibility to biocides such as sodium dodecyl sulfate and hydrogen peroxide and to antibiotics such as tobramycin (Davies et al., 1998; Bjarnsholt et al., 2005a). Agents which block QS either by inhibiting AHL synthesis, by inactivating AHLs or preventing the response to AHLs are likely to render P. aeruginosa biofilms susceptible to conventional antibiotics and to aid clearance by host defense mechanisms (Williams, 2002; Rasmussen and Givskov, 2006). Proof of this principle both in vitro and in vivo in experimental animal infection models has been presented for natural products such as furanones, patulin and penicillic acid and garlic extracts (Hentzer et al., 2003; Bjarnsholt et al., 2005b; Rasmussen et al., 2005b; Rasmussen and Givskov, 2006). Using transcriptomic approaches, many of these natural products clearly downregulate the las and rhl regulons (Hentzer et al., 2003; Rasmussen et al., 2005a). Further development of QS blockade as an anti-biofilm strategy will clearly depend on drug discovery programs uncovering safe effective QS inhibitors and in identifying the QS target structural genes which are responsible for the antibiotic tolerance of P. aeruginosa biofilms. Whether the inhibition of QS will render the biofilms of other AHL producers more susceptible to antimicrobials remains to be established although B. cenocepacia biofilms formed by cepI and cepR mutants have been reported to be more susceptible to sodium dodecyl sulfate while a cepI cciI double mutant biofilm was more sensitive to ciprofloxacin that the parent strain (Tomlin et al., 2005).

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Concluding remarks Among Gram-negative bacteria known to co-ordinate gene expression via AHL-dependent QS, there is now convincing evidence that AHLs are produced during the biofilm mode of growth in both laboratory and natural environments. It is also apparent that AHL-dependent QS is neither essential nor is it a dominant gene regulatory pathway required for facilitating bacterial adaptation to the biofilm lifestyle. here is also no evidence that biofilm development depends on a genetically programmed series of events driven by QS or indeed any other master regulatory system. QS is clearly a useful strategy for co-ordinating bacterial behavior within a population and undoubtedly can influence the structural development, integrity and architecture of a biofilm community as well as orchestrating the optimal timing and production of toxic secondary metabolites by the biofilm community to combat predators and host defense mechanisms. With the exception of the rhamnolipids which play a role in biofilm channel maintenance, the identity of the QS-regulated structural genes which contribute to biofilm development in P. aeruginosa or indeed to any other AHL-producing Gram-negative bacterium has yet to be uncovered. However, it is strikingly apparent that the loss by mutation of AHL-dependent QS in P. aeruginosa renders the biofilm population more susceptible to antimicrobial agents and host defenses. Consequently, the inhibition of QS offers an exciting target for the development of novel antibacterial agents for the treatment of chronic biofilm-centered infections. Acknowledgments Research in the authors’ laboratories has been funded by the Biotechnology and Biological Sciences Research Council, UK, the Medical Research Council UK, the Natural Environmental Research Council, the Wellcome Trust and the European Union which are gratefully acknowledged. References Ahmer, B.M.M. (2004). Cell-to-cell signalling in Escherichia coli and Salmonella enterica. Mol. Microbiol. 52, 933–945. Allesen-Holm, M., Barken, K.B., Yang, L., Klausen, M., Webb, J.S., Kjelleberg, S., Molin, S., Givskov, M., and Tolker-Nielsen, T. (2006). A characterization of DNA release in Pseudomonas aeruginosa cultures and biofilms. Mol. Microbiol. 59, 1114–1128. An, D.D., Danhorn, T., Fuqua, C., and Parsek, M.R. (2006). Quorum sensing and motility mediate interactions between Pseudomonas aeruginosa and Agrobacterium tumefaciens in biofilm cocultures. Proc. Natl. Acad. Sci. USA 103, 3828–3833. Arevalo-Ferro, C., Reil, G., Gorg, A., Eberl, L., and Riedel, K. (2005). Biofilm formation of Pseudomonas putida IsoF: the role of quorum sensing as assessed by proteomics. Syst. Appl. Microbiol. 28, 87–114. Atkinson, S., hroup, J.P., Stewart, G.S.A.B., and Williams, P. (1999). A hierarchical quorum-sensing system in Yersinia pseudotuberculosis is involved in the regulation of motility and clumping. Mol. Microbiol. 33, 1267–1277. Atkinson, S., Sockett, R.E., Cámara, M., and Williams, P. (2006). Quorum sensing and the lifestyle of Yersinia. Curr. Issues Mol. Biol. 8. 1–10. Bachofen, R., and Schenk, A. (1998). Quorum sensing autoinducers: Do they play a role in natural microbial habitats? Microbiol. Res. 153, 61–63. Batchelor, S.E., Cooper, M., Chhabra, S.R., Glover, L.A., Stewart, G.S.A.B., Williams, P., and Prosser, J.I. (1997). Cell density-regulated recovery of starved biofilm populations of ammonia-oxidizing bacteria. Appl. Environ. Microbiol. 63, 2281–2286. Beatson, S.A., Whitchurch, C.B., Semmler, A.B.T., and Mattick, J.S. (2002). Quorum sensing is not required for twitching motility in Pseudomonas aeruginosa. J. Bacteriol. 184, 3598–3604.

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Bjarnsholt, T., Jensen, P.O., Burmolle, M., Hentzer, M., Haagensen, J.A.J., Hougen, H.P., Calum, H., Madsen, K.G., Moser, C., Molin, S., Hoiby, N., and Givskov, M. (2005a). Pseudomonas aeruginosa tolerance to tobramycin, hydrogen peroxide and polymorphonuclear leukocytes is quorum-sensing dependent. Microbiology 151, 373–383. Bjarnsholt, T., Jensen, P.O., Rasmussen, T.B., Christophersen, L., Calum, H., Hentzer, M., Hougen, H.P., Rygaard, J., Moser, C., Eberl, L., Hoiby, N., and Givskov, M. (2005b). Garlic blocks quorum sensing and promotes rapid clearing of pulmonary Pseudomonas aeruginosa infections. Microbiology 151, 3873–3880. Bjarnsholt, T., and Givskov, M. (2007). Quorum sensing blockade as a strategy for enhancing host defences against bacterial pathogens. Philosophical Transactions of the Royal Society of London, Series B-Biological Sciences. In Press. Bryers, J.D. (1988). Modelling biofilm accumulation. In: Physiological Models in Microbiology, M.J. Bazin and J.I. Prosser, eds. (Boca Raton: CRC Press), pp. 109–144. Cámara, M., Williams, P., and Hardman, A. (2002). Controlling infection by tuning in and turning down the volume of bacterial small-talk. Lancet Infect. Dis. 2, 667–676. Charlton, T.S., de Nys, R., Netting, A., Kumar, N., Hentzer, M., Givskov, M., and Kjelleberg, S. (2000). A novel and sensitive method for the quantification of N-3-oxoacyl homoserine lactones using gas chromatography-mass spectrometry: application to a model bacterial biofilm. Environ. Microbiol. 2, 530–541. Chhabra, S.R., Harty, C., Hooi, D.S.W., Daykin, M., Williams, P., Telford, G., Pritchard, D.I., and Bycroft, B.W. (2003). Synthetic analogues of the bacterial signal (quorum sensing) molecule N-(3oxododecanoyl)-L-homoserine lactone as immune modulators. J. Med. Chem. 46, 97–104. Chhabra, S.R., Philipp, B., Eberl, L., Givskov, M., Williams, P., and Cámara, M. (2005). Extracellular Communication in Bacteria (Berlin/Heidelberg: Springer). Daniels, R., Vanderleyden, J., and Michiels, J. (2004). Quorum sensing and swarming migration in bacteria. FEMS Microbiol. Rev. 28, 261–289. Davey, M.E., Caiazza, N.C., and O’Toole, G.A. (2003). Rhamnolipid surfactant production affects biofilm architecture in Pseudomonas aeruginosa PAO1. J. Bacteriol. 185, 1027–1036. Davies, D. (2003). Understanding biofilm resistance to antibacterial agents. Nature Reviews Drug Discovery 2, 114–122. Davies, D.G., Parsek, M.R., Pearson, J.P., Iglewski, B.H., Costerton, J.W., and Greenberg, E.P. (1998). he involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280, 295–298. De Kievit, T.R., Gillis, R., Marx, S., Brown, C., and Iglewski, B.H. (2001). Quorum-sensing genes in Pseudomonas aeruginosa biofilms: heir role and expression patterns. Appl. Environ. Microbiol. 67, 1865–1873. Diggle, S.P., Stacey, R.E., Dodd, C., Cámara, M., Williams, P., and Winzer, K. (2006). he galactophilic lectin, LecA, contributes to biofilm development in Pseudomonas aeruginosa. Environ. Microbiol. 8, 1095–1104. Diggle, S.P., Winzer, K., Chhabra, S.R., Worrall, K.W., Cámara, M., and Williams, P. (2003). he Pseudomonas aeruginosa quinolone signal overcomes the cell density-dependency of the quorum sensing hierarchy, regulates rhl-dependent genes at the onset of stationary phase and can be produced in the absence of LasR. Mol. Microbiol. 50, 29–43. Dubern, J.F., Lugtenberg, B.J.J., and Bloemberg, G.V. (2006). he ppuI-rsaL-ppuR quorum-sensing system regulates biofilm formation of Pseudomonas putida PCL1445 by controlling biosynthesis of the cyclic lipopeptides putisolvins I and II. J. Bacteriol. 188, 2898–2906. Eberl, L., Winson, M.K., Sternberg, C., Stewart, G.S.A.B., Christiansen, G., Chhabra, S.R., Bycroft, B., Williams, P., Molin, S., and Givskov, M. (1996). Involvement of N-acyl-L-homoserine lactone autoinducers in controlling the multicellular behaviour of Serratia liquefaciens. Mol. Microbiol. 20, 127–136. Erickson, D.L., Endersby, R., Kirkham, A., Stuber, K., Vollman, D.D., Rabin, H.R., Mitchell, I. and Storey, D.G. (2002). Pseudomonas aeruginosa quorum-sensing systems may control virulence factor expression in the lungs of patients with cystic fibrosis. Infect. Immun. 70, 1783-1790. Fuqua, C., Parsek, M.R., and Greenberg, E.P. (2001). Regulation of gene expression by cell-to-cell communication: Acyl-homoserine lactone quorum sensing. Annu. Rev. Genet. 35, 439–468. Fuqua, C., Winans, S.C., and Greenberg, E.P. (1996). Census and consensus in bacterial ecosystems: he LuxR-LuxI family of quorum-sensing transcriptional regulators. Annu. Rev. Microbiol. 50, 727–751.

|

117

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|

Atkinson et al.

Gardiner, S.M., Chhabra, S.R., Harty, C., Williams, P., Pritchard, D.I., Bycroft, B.W., and Bennett, T. (2001). Haemodynamic effects of the bacterial quorum sensing signal molecule, N-(3-oxododecanoyl)-L-homoserine lactone, in conscious, normal and endotoxaemic rats. Br. J. Pharmacol. 133, 1047–1054. Gould, T.A., Schweizer, H.P., and Churchill, M.E.A. (2004). Structure of the Pseudomonas aeruginosa acyl-homoserine lactone synthase LasI. Mol. Microbiol. 53, 1135–1146. Hanzelka, B.L., Parsek, M.R., Val, D.L., Dunlap, P.V., Cronan, J.E., and Greenberg, E.P. (1999). Acylhomoserine lactone synthase activity of the Vibrio scheri AinS protein. J. Bacteriol. 181, 5766– 5770. Hentzer, M., Eberl, L., and Givskov, M. (2005). Transcriptome analysis of Pseudomonas aeruginosa biofilm development: anaerobic respiration and iron limitation. Biofilms 2, 37–61. Hentzer, M., Wu, H., Andersen, J.B., Riedel, K., Rasmussen, T.B., Bagge, N., Kumar, N., Schembri, M.A., Song, Z.J., Kristoffersen, P., Manefield, M., Costerton, J.W., Molin, S., Eberl, L., Steinberg, P., Kjelleberg, S., Hoiby, N., and Givskov, M. (2003). Attenuation of Pseudomonas aeruginosa virulence by quorum sensing inhibitors. EMBO J. 22, 3803–3815. Heydorn, A., Ersboll, B., Kato, J., Hentzer, M., Parsek, M.R., Tolker-Neilsen, T., Givskov, M., and Molin, S. (2002). Statistical analysis of Pseudomonas aeruginosa biofilm development: impact of mutations in genes involved in twitching motility, cell-to-cell signalling and stationary-phase sigma factor expression. Appl. Environ. Microbiol. 68, 2008–2017. Hogan, D.A., Vik, A., and Kolter, R. (2004). A Pseudomonas aeruginosa quorum-sensing molecule influences Candida albicans morphology. Mol. Microbiol. 54, 1212–1223. Holden, M.T.G., Chhabra, S.R., de Nys, R., Stead, P., Bainton, N.J., Hill, P.J., Manefield, M., Kumar, N., Labatte, M., England, D., Rice, S., Givskov, M., Salmond, G.P.C., Stewart, G.S.A.B., Bycroft, B.W., Kjelleberg, S.A., and Williams, P. (1999). Quorum-sensing cross talk: isolation and chemical characterisation of cyclic dipeptides from Pseudomonas aeruginosa and other Gram-negative bacteria. Mol. Microbiol. 33, 1254–1266. Houdt, R., Aertsen, A., Moons, P., Vanoirbeek, K., and Michiels, C.W. (2006). N-acyl-L-homoserine lactone signal interception by Escherichia coli. FEMS Microbiol. Lett. 256, 83–89. Huber, B., Riedel, K., Hentzer, M., Heydorn, A., Gotschlich, A., Givskov, M., Molin, S., and Eberl, L. (2001). he cep quorum-sensing system of Burkholderia cepacia H111 controls biofilm formation and swarming motility. Microbiology 147, 2517–2528. Huber, B., Riedel, K., Kothe, M., Givskov, M., Molin, S., and Eberl, L. (2002). Genetic analysis of functions involved in the late stages of biofilm development in Burkholderia cepacia H111. Mol. Microbiol. 46, 411–426. Isles, A., Maclusky, I., Corey, M., Gold, R., Prober, C., Fleming, P., and Levison, H. (1984). Pseudomonas Cepacia infection in cystic-fibrosis—an emerging problem. J. Pediatr. 104, 206–210. Jarrett, C.O., Deak, E., Isherwood, K.E., Oyston, P.C., Fischer, E.R., Whitney, A.R., Kobayashi, S.D., DeLeo, F.R., and Hinnebusch, B.J. (2004). Transmission of Yersinia pestis from an infectious biofilm in the flea vector. J. Infect. Dis. 190, 783–792. Joint, I., Callow, M.E., Callow, J.A., and Clarke, K.R. (2000). he attachment of Enteromorpha zoospores to a bacterial biofilm assemblage. Biofouling 16, 151–158. Joint, I., Tait, K., Callow, M.E., Callow, J.A., Milton, D., Williams, P., and Cámara, M. (2002). Cell-to-cell communication across the prokaryote-eukaryote boundary. Science 298, 1207. Kaplan, J.B., Ragunath, C., Ramasubbu, N., and Fine, D.H. (2003). Detachment of Actinobacillus actinomycetemcomitans biofilm cells by an endogenous beta-hexosaminidase activity. J. Bacteriol. 185, 4693–4698. Kjelleberg, S., and Molin, S. (2002). Is there a role for quorum sensing signals in bacterial biofilms? Curr. Opin. Microbiol. 5, 254–258. Klausen, M., Aaes-Jorgensen, A., Molin, S., and Tolker-Nielsen, T. (2003). Involvement of bacterial migration in the development of complex multicellular structures in Pseudomonas aeruginosa biofilms. Mol. Microbiol. 50, 61–68. Koutsoudis, M.D., Tsaltas, D., Minogue, T.D., and von Bodman, S.B. (2006). Quorum-sensing regulation governs bacterial adhesion, biofilm development, and host colonization in Pantoea stewartii subspecies stewartii. Proc. Natl. Acad. Sci. USA 103, 5983–5988. Kuiper, I., Lagendijk, E.L., Pickford, R., Derrick, J.P., Lamers, G.E.M., homas-Oates, J.E., Lugtenberg, B.J.J., and Bloemberg, G.V. (2004). Characterization of two Pseudomonas putida lipopeptide biosur-

Quorum Sensing and Bioilms

factants, putisolvin I and II, which inhibit biofilm formation and break down existing biofilms. Mol. Microbiol. 51, 97–113. Labbate, M., Queck, S.Y., Koh, K.S., Rice, S.A., Givskov, M., and Kjelleberg, S. (2004). Quorum sensingcontrolled biofilm development in Serratia liquefaciens MG1. J. Bacteriol. 186, 692–698. Latifi, A., Foglino, M., Tanaka, K., Williams, P., and Lazdunski, A. (1996). A hierarchical quorum-sensing cascade in Pseudomonas aeruginosa links the transcriptional activators LasR and RhIR (VsmR) to expression of the stationary-phase sigma factor RpoS. Mol. Microbiol. 21, 1137–1146. Lawrence, R.N., Dunn, W.R., Bycroft, B., Cámara, M., Chhabra, S.R., Williams, P., and Wilson, V.G. (1999). he Pseudomonas aeruginosa quorum-sensing signal molecule, N-(3-oxododecanoyl)-L-homoserine lactone, inhibits porcine arterial smooth muscle contraction. Br. J. Pharmacol. 128, 845–848. Lequette, Y., and Greenberg, E.P. (2005). Timing and localization of rhamnolipid synthesis gene expression in Pseudomonas aeruginosa biofilms. J. Bacteriol. 187, 37–44. Lyczak, J.B., Cannon, C.L., and Pier, G.B. (2002). Lung infections associated with cystic fibrosis. Clin. Microbiol. Rev. 15, 194-+. Lynch, M.J., Swift, S., Kirke, D.F., Keevil, C.W., Dodd, C.E.R., and Williams, P. (2002). he regulation of biofilm development by quorum sensing in Aeromonas hydrophila. Environ. Microbiol. 4, 18–28. Matsuo, Y., Suzuki, M., Kasai, H., Shizuri, Y., and Harayama, S. (2003). Isolation and phylogenetic characterization of bacteria capable of inducing differentiation in the green alga Monostroma oxyspermum. Environ. Microbiol. 5, 25–35. Matz, C., Bergfeld, T., Rice, S.A., and Kjelleberg, S. (2004). Microcolonies, quorum sensing and cytotoxicity determine the survival of Pseudomonas aeruginosa biofilms exposed to protozoan grazing. Environ. Microbiol. 6, 218–226. Matz, C., and Kjelleberg, S. (2005). Off the hook—how bacteria survive protozoan grazing. Trends Microbiol. 13, 302–307. McClean, K.H., Winson, M.K., Fish, L., Taylor, A., Chhabra, S.R., Cámara, M., Daykin, M., Lamb, J.H., Swift, S., Bycroft, B.W., Stewart, G.S.A.B., and Williams, P. (1997). Quorum sensing and Chromobacterium violaceum: exploitation of violacein production and inhibition for the detection of N-acylhomoserine lactones. Microbiology 143, 3703–3711. McLean, R.J.C., Whiteley, M., Stickler, D.J., and Fuqua, W.C. (1997). Evidence of autoinducer activity in naturally occurring biofilms. FEMS Microbiol. Lett. 154, 259–263. McLean, R.J.C., Barnes, M.B., Windham, M.K., Merchant, M., Forstner, M.R.J., and Fuqua, C. (2005). Cell–cell influences on bacterial community development in aquatic biofilms. Appl. Environ. Microbiol. 71, 8987–8990. Middleton, B., Rodgers, H.C., Cámara, M., Knox, A.J., Williams, P., and Hardman, A. (2002). Direct detection of N-acylhomoserine lactones in cystic fibrosis sputum. FEMS Microbiol. Lett. 207, 1–7. Milton, D.L., Chalker, V.J., Kirke, D., Hardman, A., Cámara, M., and Williams, P. (2001). he luxM Homologue vanM from Vibrio anguillanrum directs the synthesis of N-(3-hydroxyhexanoyl)homoseri ne lactone and N-hexanoylhomoserine lactone. J. Bacteriol. 183, 3537–3547. O’Toole, G., Kaplan, H.B., and Kolter, R. (2000). Biofilm formation as microbial development. Annu. Rev. Microbiol. 54, 49–79. Ortori, C.A., Atkinson, S., Chhabra, S.R., Cámara, M., Williams P., and Barrett, D.A. (2006). Comprehensive profiling of N-acylhomoserine lactones produced by Yersinia pseudotuberculosis using liquid chromatography coupled to hybrid quadruple-linear ion trap mass spectrometry. Analytical and Bioanalytical Chemistry. In Press Parsek, M.R., and Greenberg, E.P. (2005). Sociomicrobiology: the connections between quorum sensing and biofilms. Trends Microbiol. 13, 27–33. Pearson, J.P., Gray, K.M., Passador, L., Tucker, K.D., Eberhard, A., Iglewski, B.H., and Greenberg, E.P. (1994). Structure of the autoinducer required for expression of Pseudomonas-aeruginosa virulence genes. Proc. Natl. Acad. Sci. USA 91, 197–201. Pearson, J.P., Passador, L., Iglewski, B.H., and Greenberg, E.P. (1995). A 2nd N-acylhomoserine lactone signal produced by Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 92, 1490–1494. Pritchard, D.I., Todd, I., Brown, A., Bycroft, B.W., Chhabra, S.R., Williams, P., and Wood, P. (2005). Alleviation of insulitis and moderation of diabetes in NOD mice following treatment with a synthetic Pseudomonas aeruginosa signal molecule, N-(3-oxododecanoyl)-L-homoserine lactone. Acta Diabetol. 42, 119–122.

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Atkinson et al.

Provasoli, L., and Pintner, I.J. (1980). Bacteria induced polymorphism in an axenic laboratory strain of Ulva-Lactuca (Chlorophyceae). J. Phycol. 16, 196–201. Purevdorj-Gage, B., Costerton, W.J., and Stoodley, P. (2005). Phenotypic differentiation and seeding dispersal in non-mucoid and mucoid Pseudomonas aeruginosa biofilms. Microbiology 151, 1569–1576. Purevdorj-Gage, B., and Stoodley, P. (2004). Hydrodynamic considerations of biofilm structure and behaviour. In: Microbial Biofilms, M.A. Ghannoum and G. O’Toole, eds. (Washington: American Society for Microbiology), pp. 160–173. Puskas, A., Greenberg, E.P., Kaplan, S., and Schaeffer, A.L. (1997). A quorum-sensing system in the freeliving photosynthetic bacterium Rhodobacter sphaeroides. J. Bacteriol. 179, 7530–7537. Qazi, S.N.A., Middleton, B., Muharram, S.H., Cockayne, A., Hill, P., O’Shea, P., Chhabra, S.R., Cámara, M., and Williams, P. (2006). N-acylhomoserine lactones antagonize virulence gene expression and quorum sensing in Staphylococcus aureus. Infect. Immun. 74, 910–919. Qazi, S.N.A., Rees, C.E.D., Mellits, K.H., and Hill, P.J. (2001). Development of gfp vectors for expression in Listeria monocytogenes and other low G+C Gram positive bacteria. Microb. Ecol. 41, 301–309. Queck, S.Y., Weitere, M., Moreno, A.M., Rice, S.A., and Kjelleberg, S. (2006). he role of quorum sensing mediated developmental traits in the resistance of Serratia marcescens biofilms against protozoan grazing. Environ. Microbiol. 8, 1017–1025. Rampioni, G., Bertani, I., Zennaro, E., Polticelli, F., Venturi, V., and Leoni, L. (2006). he quorum-sensing negative regulator RsaL of Pseudomonas aeruginosa binds to the lasI promoter. J. Bacteriol. 188, 815–819. Rasmussen, T.B., Bjarnsholt, T., Skindersoe, M.E., Hentzer, M., Kristoffersen, P., Kote, M., Nielsen, J., Eberl, L., and Givskov, M. (2005a). Screening for quorum-sensing inhibitors (QSI) by use of a novel genetic system, the QSI selector. J. Bacteriol. 187, 1799–1814. Rasmussen, T.B., and Givskov, M. (2006). Quorum sensing inhibitors: a bargain of effects. Microbiology 152, 895–904. Rasmussen, T.B., Skindersoe, M.E., Bjarnsholt, T., Phipps, R.K., Christensen, K.B., Jensen, P.O., Andersen, J.B., Koch, B., Larsen, T.O., Hentzer, M., Eberl, L., Hoiby, N., and Givskov, M. (2005b). Identity and effects of quorum-sensing inhibitors produced by Penicillium species. Microbiology 151, 1325–1340. Redfield, R.J. (2002). Is quorum sensing a side effect of diffusion sensing? Trends Microbiol. 10, 365– 370. Rice, S.A., Koh, K.S., Queck, S.Y., Labbate, M., Lam, K.W., and Kjelleberg, S. (2005). Biofilm formation and sloughing in serratia marcescens are controlled by quorum sensing and nutrient cues. J. Bacteriol. 187, 3477–3485. Riedel, K., Hentzer, M., Geisenberger, O., Huber, B., Steidle, A., Wu, H., Hoiby, N., Givskov, M., Molin, S., and Eberl, L. (2001). N-acylhomoserine-lactone-mediated communication between Pseudomonas aeruginosa and Burkholderia cepacia in mixed biofilms. Microbiology 147, 3249–3262. Salmond, G.P.C., Bycroft, B.W., Stewart, G.S.A.B., and Williams, P. (1995). he bacterial enigma—cracking the code of cell–cell communication. Mol. Microbiol. 16, 615–624. Sauer, K., Camper, A.K., Ehrlich, G.D., Costerton, J.W., and Davies, D.G. (2002). Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J. Bacteriol. 184, 1140–1154. Schuster, M., Lostroh, C.P., Ogi, T., and Greenberg, E.P. (2003). Identification, timing, and signal specificity of Pseudomonas aeruginosa quorum-controlled genes: a transcriptome analysis. J. Bacteriol. 185, 2066–2079. Shih, P.C., and Huang, C.T. (2002). Effects of quorum-sensing deficiency on Pseudomonas aeruginosa biofilm formation and antibiotic resistance. J. Antimicrob. Chemother. 49, 309–314. Shiner, E.K., Terentyev, D., Bryan, A., Martinez-Zaguilan, R., Gyorke, S., Williams, S.C., and Rambaugh, K.P. (2006). Pseudomonas aeruginosa autoinducer modulates host cell responses through calcium signalling. Cell. Microbiol. 8, 1601–1610. Singh, P.K., Schaefer, A.L., Parsek, M.R., Moninger, T.O., Welsh, M.J., and Greenberg, E.P. (2000). Quorum-sensing signals indicate that cystic fibrosis lungs are infected with bacterial biofilms. Nature 407, 762–764. Sio, C.F., Otten, L.G., Cool, R.H., Diggle, S.P., Braun, P.G., Bos, R., Daykin, M., Cámara, M., Williams, P., and Quax, W.J. (2006). Quorum quenching by an N-acyl-homoserine lactone acylase from Pseudomonas aeruginosa PAO1. Infect. Immun. 74, 1673–1682.

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Smith, R.S., Harris, S.G., Phipps, R., and Iglewski, B. (2002). he Pseudomonas aeruginosa quorum-sensing molecule N-(3-oxododecanoyl)homoserine lactone contributes to virulence and induces inflammation in vivo. J. Bacteriol. 184, 1132–1139. Stickler, D.J., Morris, N.S., McLean, R.J.C., and Fuqua, C. (1998). Biofilms on indwelling urethral catheters produce quorum sensing signal molecules in situ and in vitro. Appl. Environ. Microbiol. 64, 3486–3490. Swift, S., Downie, J.A., Whitehead, N.A., Barnard, A.M.L., Salmond, G.P.C., and Williams, P. (2001). Quorum sensing as a population-density-dependent determinant of bacterial physiology. Adv. Microb. Physiol. 45, 199–270. Swift, S., Karlyshev, A.V., Fish, L., Durant, E.L., Winson, M.K., Chhabra, S.R., Williams, P., MacIntyre, S., and Stewart, G.S.A.B. (1997). Quorum sensing in Aeromonas hydrophila and Aeromonas salmonicida: Identification of the LuxRI homologs AhyRI and AsaRI and their cognate N-acylhomoserine lactone signal molecules. J. Bacteriol. 179, 5271–5281. Swift, S., hroup, J.P., Bycroft, B., Williams P., and Stewart, G.S.A.B. (1998). Quorum sensing: bacterial cell–cell signalling from bioluminescence to pathogenicity. In: NATO ASI series Molecular Microbiology, S.J. Busby, C.M. homas, and N.L. Brown, eds., pp. 185–207. Swift, S., Williams P., and Stewart, G.S.A.B. (1999). N-acylhomoserine lactones and quorum sensing in proteobacteria. In: Cell–cell Signaling in Bacteria, G.M. Dunny and S.C. Winans, eds. (Washington DC: ASM Press), pp. 291–314. Tait, K., Joint, I., Daykin, M., Milton, D.L., Williams, P., and Cámara, M. (2005). Disruption of quorum sensing in seawater abolishes attraction of zoospores of the green alga Ulva to bacterial biofilms. Environ. Microbiol. 7, 229–240. Telford, G., Wheeler, D., Williams, P., Tomkins, P.T., Appleby, P., Sewell, H., Stewart, G.S.A.B., Bycroft, B.W., and Pritchard, D.I. (1998). he Pseudomonas aeruginosa quorum-sensing signal molecule N-(3oxododecanoyl)-L-homoserine lactone has immunomodulatory activity. Infect. Immun. 66, 36–42. Tielker, D., Hacker, S., Loris, R., Strathmann, M., Wingender, J., Wilhelm, S., Rosenau, F., and Jaeger, K.E. (2005). Pseudomonas aeruginosa lectin LecB is located in the outer membrane and is involved in biofilm formation. Microbiology 151, 1313–1323. Tomlin, K.L., Malott, R.J., Ramage, G., Storey, D.G., Sokol, P.A., and Ceri, H. (2005). Quorum-sensing mutations affect attachment and stability of Burkholderia cenocepacia biofilms. Appl. Environ. Microbiol. 71, 5208–5218. Valle, A., Bailey, M.J., Whiteley, A.S., and Manefield, M. (2004). N-acyl-L-homoserine lactones (AHLs) affect microbial community composition and function in activated sludge. Environ. Microbiol. 6, 424–433. Vendeville, A., Winzer, K., Heurlier, K., Tang, C.M., and Hardie, K.R. (2005). Making “sense” of metabolism: Autoinducer-2, LuxS and pathogenic bacteria. Nature Rev. Microbiol. 3, 383–396. von Bodman, S.B., Majerczak, D.R., and Coplin, D.L. (1998). A negative regulator mediates quorumsensing control of exopolysaccharide production in Pantoea stewartii subsp. stewartii. Proc. Natl. Acad. Sci. USA 95, 7687–7692. Wagner, V.E., Bushnell, D., Passador, L., Brooks, A.I., and Iglewski, B.H. (2003). Microarray analysis of Pseudomonas aeruginosa quorum-sensing regulons: Effects of growth phase and environment. J. Bacteriol. 185, 2080–2095. Waite, R.D., Papakonstantinopoulou, A., Littler, E., and Curtis, M.A. (2005). Transcriptome Analysis of Pseudomonas aeruginosa Growth: Comparison of Gene Expression in Planktonic Cultures and Developing and Mature Biofilms. J. Bacteriol. 187, 6571–6576. Watson, W.T., Minogue, T.D., Val, D.L., von Bodman, S.B., and Churchill, M.E.A. (2002). Structural basis and specificity of acyl-homoserine lactone signal production in bacterial quorum sensing. Mol. Cell 9, 685–694. Webb, J.S., hompson, L.S., James, S., Charlton, T., Tolker-Nielsen, T., Koch, B., Givskov, M., and Kjelleberg, S. (2003). Cell death in Pseudomonas aeruginosa biofilm development. J. Bacteriol. 185, 4585–4592. Weitere, M., Bergfeld, T., Rice, S.A., Matz, C., and Kjelleberg, S. (2005). Grazing resistance of Pseudomonas aeruginosa biofilms depends on type of protective mechanism, developmental stage and protozoan feeding mode. Environ. Microbiol. 7, 1593–1601.

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121

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|

Atkinson et al.

Werner, E., Roe, F., Bugnicourt, A., Franklin, M.J., Heydorn, A., Molin, S., Pitts, B., and Stewart, P.S. (2004). Stratified growth in Pseudomonas aeruginosa biofilms. Appl. Environ. Microbiol. 70, 6188– 6196. Wheeler, G.L., Tait, K., Taylor, A.R., Brownlee, C., and Joint, I. (2006). Acyl-homoserine lactones modulate the settlement rate of zoospores of the marine alga Ulva intestinalis via a novel chemokinetic mechanism. Plant Cell Environ. 29, 608–618. Whitchurch, C.B., Tolker-Nielsen, T., Ragas, P.C., and Mattick, J.S. (2002). Extracellular DNA required for bacterial biofilm formation. Science 295, 1487. Whiteley, M., Bangera, M.G., Bumgarner, R.E., Parsek, M.R., Teitzel, G.M., Lory, S., and Greenberg, E.P. (2001). Gene expression in Pseudomonas aeruginosa biofilms. Nature 413, 860–864. Williams, P. (2002). Quorum sensing: an emerging target for antimicrobial chemotherapy? Exp. Opin. her. Targets 6, 257–274. Williams, P., Cámara, M., Hardman, A., Swift, S., Milton, D., Hope, V.J., Winzer, K., Middleton, B., Pritchard, D.I., and Bycroft, B.W. (2000). Quorum sensing and the population-dependent control of virulence. Phil. Trans. R. Soc. London Series B-Biol. Sci. 355, 667–680. Williams, P., and Stewart, G.S.A.B. (1993). Cell density dependent control of gene expression in bacteriaimplications for biofilm development and control. British Biofilm Club Meeting. Williams, P., Winzer, K., Chan, W.C., and Cámara, M. (2007). Look who’s talking: communication and quorum sensing in the bacterial world. Phil. Trans. R. Soc. London Series B-Biol. Sci. In Press. Winson, M.K., Cámara, M., Latifi, A., Foglino, M., Chhabra, S.R., Daykin, M., Bally, M., Chapon, V., Salmond, G.P.C., Bycroft, B.W., Lazdunski, A., Stewart, G.S.A.B., and Williams, P. (1995). Multiple N-acyl-L-homoserine lactone signal molecules regulate production of virulence determinants and secondary metabolites in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 92, 9427–9431. Winson, M.K., Swift, S., Fish, L., hroup, J.P., Jorgensen, F., Chhabra, S.R., Bycroft, B.W., Williams, P., and Stewart, G.S.A.B. (1998). Construction and analysis of luxCDABE-based plasmid sensors for investigating N-acyl homoserine lactone-mediated quorum sensing. FEMS Microbiol. Lett. 163, 185–192. Winzer, K., Falconer, C., Garber, N.C., Diggle, S.P., Cámara, M., and Williams, P. (2000). he Pseudomonas aeruginosa lectins PA-IL and PA-IIL are controlled by quorum sensing and by RpoS. J. Bacteriol. 182, 6401–6411. Winzer, K., Hardie, K.R., and Williams, P. (2002). Bacterial cell-to-cell communication: Sorry, can’t talk now-gone to lunch! Curr. Opin. Microbiol. 5, 216–222. Winzer, K., and Williams, P. (2003). Escherichia coli gets the message. Nat. Med. 9, 1118–1119. Withers, H., Swift, S., and Williams, P. (2001). Quorum sensing as an integral component of gene regulatory networks in Gram-negative bacteria. Curr. Opin. Microbiol. 4, 186–193. Wopperer, J., Cardona, S.T., Huber, B., Jacobi, C.A., Valvano, M.A., and Eberl, L. (2006). A quorumquenching approach to investigate the conservation of quorum-sensing-regulated functions within the Burkholderia cepacia complex. Appl. Environ. Microbiol. 72, 1579–1587. Yates, E.A., Philipp, B., Buckley, C., Atkinson, S., Chhabra, S.R., Sockett, R.E., Goldner, M., Dessaux, Y., Cámara, M., Smith, H., and Williams, P. (2002). N-acylhomoserine Lactones undergo lactonolysis in a pH-, temperature-, and acyl chain length-dependent manner during growth of Yersinia pseudotuberculosis and Pseudomonas aeruginosa. Infect. Immun. 70, 5635–5646. Yim, G., Wang, H.H.M., and Davies, J. (2006). he truth about antibiotics. Int. J. Med. Microbiol. 296, 163–170. Zhang, R.G., Pappas, T., Brace, J.L., Miller, P.C., Oulmassov, T., Molyneauz, J.M., Anderson, J.C., Bashkin, J.K., Winans, S.C., and Joachimiak, A. (2002). Structure of a bacterial quorum-sensing transcription factor complexed with pheromone and DNA. Nature 417, 971–974.

Signaling in Escherichia coli Bioilms Thomas K. Wood and William E. Bentley

7

Abstract Cell signaling in Escherichia coli biofilms is more important than originally known, in that autoinducer two (AI-2), AI-1 (N-acylhomoserine lactones), indole, hydroxylated indoles, norepinephrine, and epinephrine are all functional signals in this organism. his gives the bacterium the ability to monitor not only the presence of cells of its own species (through AI-2 and indole) but also the activity of the human host (through norepinephrine and epinephrine) as well as that of the other commensal bacteria (through hydroxylated indoles). We have also found that E. coli monitors the presence of bacteria that produce acylhomoserine lactones as these signals influence formation of its biofilms through the LuxR homolog SdiA. Hence, E. coli monitors its surroundings through signals it synthesizes as well as those synthesized by both prokaryotes and eukaryotes. In this chapter, we present recent findings on the role of these different signals as well as provide a detailed account of the complex features of uptake and regulation of the AI-2 signal in E. coli. AI-2 signaling through the motility quorum sensing regulator, MqsR Cell signaling (quorum sensing) is established for biofilm formation in bacteria. For example, it controls the production and secretion of exopolysaccharides for Vibrio cholerae biofilms (Hammer and Bassler, 2003). his signaling may be complex as V. harveyi uses three quorum sensing signals including N-(3-hydroxybutanoyl) homoserine lactone (AI1), furanosyl borate diester or related compounds (AI-2) (Lombardia et al., 2006), and a signal, synthesized by CqsA, whose structure is unknown (Henke and Bassler, 2004). Other examples include the quorum signal N-(3-oxododecanoyl)-L-homoserine lactone which controls biofilm formation in P. aeruginosa (Davies et al., 1998), and the quorum signal N-butanoyl-L-homoserine lactone which controls biofilm formation in Serratia liquefaciens MG1 (Labbate et al., 2004). AI-2 is produced by the enzymes Pfs (nucleosidase) and LuxS (terminal synthase) from S-adenosylhomocysteine and is a species non-specific signal used by both Gram-negative and Gram-positive bacteria (Surette et al., 1999) which has been found in at least 55 strains (Bassler et al., 1997). We have focused our AI-2 based studies on E. coli biofilms since E. coli is the most thoroughly studied bacterium (Blattner et al., 1997), its genome is sequenced

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(Blattner et al., 1997), microarrays are available (Selinger et al., 2000), the function of many of its proteins have been elucidated (Riley and Labedan, 1996), and many isogenic mutants are available (Kang et al., 2004). Also, our group has experience in determining the genetic basis of E. coli biofilm formation and its inhibition with natural, plant-derived antagonists such as furanone and ursolic acid (Ren et al., 2001; Ren et al., 2005). For E. coli, three groups have used DNA microarrays to show that AI-2 controls 166 to 404 genes including those for chemotaxis, flagellar synthesis, motility, and virulence factors (Beloin et al., 2004; Ren et al., 2004a; Schembri et al., 2003). Remarkably, 80% of these same phenotypes were found to be regulated by the quorum sensing inhibitor furanone in an opposite manner (Ren et al., 2004b); addition of furanone reduces biofilm formation for E. coli (Ren et al., 2001; Ren et al., 2004b). his shows the importance of AI-2 as a quorum signal in E. coli. A recent report (Walters et al., 2006) has attributed the function of AI-2 as more growth-related in E. coli because its deletion affects homocysteine synthesis; however, the authors used a non-robust phenotype method (Biolog) to survey the impact of a luxS mutation so that their results were restricted to changes in these growth phenotypes, rather than being able to address the regulatory functions of AI-2. As the first clear example of the impact of AI-2 and biofilm formation and of the importance of AI-2 on phenotypes of a bacterium other than Vibrio, we demonstrated that in vitro synthesized AI-2 from purified Pfs and LuxS directly stimulates E. coli biofilm formation by 30-fold in the absence of a conjugation plasmid (González Barrios et al., 2006). Note that in the absence of AI-2, E. coli BW25113 $luxS made 50% less biofilm compared to the isogenic wild-type strain, which indicates again that AI-2 stimulates biofilm formation in E. coli since LuxS forms AI-2 (González Barrios et al., 2006). Biofilm could be restored by complementing luxS in trans using plasmid pCA24N luxS+ (46% of the wildtype biofilm was formed at 0 mM IPTG and 110% of the wild-type biofilm was formed at 0.25 mM IPTG in LB medium). To confirm that AI-2 is the cause of the increase in biofilm formation, we measured (González Barrios et al., 2006) biofilm stimulation with the isogenic MG1655 lsrK mutant because this mutation dramatically impairs the AI-2 uptake compared with other mutations in the lsr system (Wang et al., 2005a; Xavier and Bassler, 2005a). AI-2 phosphokinase, LsrK, is one of the essential components of the LuxS regulated AI-2 uptake operon, lsrADCBFG (Wang et al., 2005a). As expected, AI-2 was not able to induce biofilm formation of the lsrK mutant at 6.4 MM; hence, AI-2 induces biofilm formation through the LsrK transport pathway (Wang et al., 2005a). Note that the presence of a conjugation plasmid alters dramatically how E. coli forms a biofilm (Ghigo, 2001; Reisner et al., 2003). Showing that purified AI-2 controls biofilm formation is significant, as other groups have relied on the use of conditioned medium or luxS mutations (Balestrino et al., 2005; Cole et al., 2004; Labbate et al., 2004; Li et al., 2001; McNab et al., 2003), and this result demonstrates that AI-2 regulates more than the lsr operon (Walters et al., 2006). We then set about to determine how AI-2 controls biofilms by studying motility since this trait has been linked to biofilm formation in the absence of a conjugation plasmid (Pratt and Kolter, 1998) and since our microarrays (Ren et al., 2004a) indicated that motility genes were induced by AI-2 (determined using a luxS mutant). It was found that the motility of several E. coli K-12 strains (ATCC 25404, MG1655, DH5A, and BW25113)

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increased by about 30–80% upon addition of 0.8 to 3.2 MM AI-2 (González Barrios et al., 2006). To discern the genetic basis of this increase in motility upon AI-2 addition, we probed the ability of AI-2 to induce the promoters of motility genes qseB, flhD, fliA, fliC, and motA (González Barrios et al., 2006). Upon addition of 6.4 MM AI-2, the quorum-sensing flagella regulon qseB (Sperandio et al., 2002) was induced 8 fold (González Barrios et al., 2006). hese results corroborate the ones reported by Sperandio et al. (2002) who previously found qseB to be induced 17-fold compared with the luxS mutant through DNA microarray studies using E. coli O157:H7 and its isogenic luxS mutant. he induction of qseB here led to a 4.0 fold increase in transcription of flhD (master controller of the flagella regulon), 2.6 fold increase of fliA (sigma factor S28), 3.6-fold of fliC (flagellin), and 6-fold increase of motA (proton conductor for flagella movement) (González Barrios et al., 2006). Given that the purified AI-2 controlled motility (González Barrios et al., 2006), that b3022 is induced 8-fold in biofilms (Ren et al., 2004a) and is nearby qseB (b3025) which is clearly induced by the addition of purified AI-2, we hypothesized that B3022 may interact with AI-2 and control biofilm formation. B3022 is a conserved regulator protein (98 amino acids) since it has more than 50% identity with hypothetical proteins from Yersinia pseudotuberculosis, Y. pestis, Cupriavidus oxalaticus, Bordetella bronchiseptica, P. fluorescens, and Bordetella pertussis (Chain et al., 2004; Parkhill et al., 2003; Paulsen et al., 2005; Song et al., 2004; Toussaint et al., 2003). As part of the 300-gene, quorum-sensing regulon in E. coli (Beloin et al., 2004; Ren et al., 2004a; Schembri et al., 2003), qseBC (b3025, b3026) are organized in an operon in the E. coli chromosome with QseB playing a role as a response regulator and QseC, the sensor kinase (Sperandio et al., 2002). Flagella expression is temporally regulated, and the operons are divided into early, middle, and late genes. QseBC regulates transcription of the master regulon flhDC and therefore expression of the middle operon (e.g. fliA encoding sigma factor S28) and late operon (e.g. fliC encoding flagellin, motA encoding the proton exchange conductor for flagella movement) (Chilcott and Hughes, 2000). We found that deletion of b3022 abolished motility which was restored by expressing b3022 in trans (González Barrios et al., 2006). Deletion of b3022 also decreased biofilm formation significantly relative to the wild-type strain in three media (46–74%) in 96 well plates as well as decreased biomass (8-fold) and substratum coverage (19-fold) in continuous flow cells with minimal medium (growth rate not altered and biofilm restored by expressing b3022 in trans). Deleting b3022 changed the wild-type biofilm architecture from a thick (54-Mm), complex structure to one that contained only a few microcolonies (González Barrios et al., 2006). B3022 positively regulates expression of qseBC, flhD, fliA, and motA since deleting b3022 decreased their transcription by 61-, 25-, 2.4-, and 18fold, respectively (González Barrios et al., 2006). Transcriptome analysis also revealed that B3022 induces crl (26-fold) and flhCD (8 to 27-fold). Adding AI-2 (6.4 MM) increased biofilm formation of wild-type K12 MG1655 but not that of the isogenic b3022, qseBC, fliA, and motA mutants. Adding AI-2 also increased motA transcription for the wild-type strain but did not stimulate motA transcription for the b3022 and qseB mutants; therefore, the induction of motility is likely mediated by both MqsR and QseBC.

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Evidence that mqsR is first in the cascade was provided by measuring the transcription of qseB with the wild-type strain and the b3022 mutant upon addition of AI-2. If B3022 is first in the cascade and necessary for the transduction of the AI-2 signal, then the addition of AI-2 should only increase qseB transcription when b3022 is present (González Barrios et al., 2006). We found that adding AI-2 at 6.4 MM induced the expression of qseB 3.2 fold in the wild-type strain in M9C glu but did not induce the qseB in the b3022 mutants. As expected, the wild-type strain responded to AI-2 addition in a dose-dependent manner. To show further that MqsR is first in the cascade, the expression of qseB from pVS159 was also measured while inducing B3022 expression in trans in the mqsR mutant by adding IPTG to strains harboring pVLT31 b3022+. As expected if B3022 is required for signal transduction to QseB, expression of qseB was induced 4-fold in a dose-dependent manner in M9C glu (see Figure 7.6B). Together, these results indicate that AI-2 induces biofilm formation in E. coli through B3022 (either directly or indirectly), which then regulates QseBC and motility; hence, b3022 has been re-named motility quorum sensing regulator (mqsR), and B3022 is the master regulator of motility (Figure 7.1). AI-2 import through LsrACDB and export through the quorumsensing transport protein TqsA Since AI-2 is highly polar, we reasoned that it must be actively transported from the cell. It has been previously established that internalization of AI-2 requires an ABC transporter. Moreover, we have hypothesized that the formation of AI-2 from its precursor, 4,5-dihy-

+

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b3021

b3022

b3023

+ + AI-2

b3024

+ csrA

qseB

qseC

b3025

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b3027

+ flhDC fliA motA crl

+ Biofilm

Figure 7.1 Model for AI-2-mediated quorum-sensing regulation of motility and bioilms in E. coli K-12. AI-2 increases expression of MqsR (B3022) which increases expression of QseBC and CsrA which in turn increase expression of the motility regulon that includes FlhDC, FliA, MotA, and Crl, which results in stimulation of bioilm formation. Plus signs indicate positive regulation (shown by dashed lines).

Signaling in Escherichia coli Bioilms

droxy-2,3-pentanedione (DPD) may require biological assistance. However, the means by which AI-2 is exported has not been identified. AI-2 induces transcription of the lsrACDBFGE operon of Salmonella typhimurium in which the first four genes encode an ATP binding cassette-type import protein with high homology to the ribose transporter (Taga et al., 2001; Taga et al., 2003). In E. coli, the proteins for AI-2 transport are encoded by lsrACDBFG (Wang et al., 2005a; Wang et al., 2005b). AI-2 is phosphorylated inside the cell by LsrK, and the phosphorylated AI-2 induces inactivation of the lsr operon repressor LsrR (Taga et al., 2003; Wang et al., 2005a); hence, AI-2 uptake is enhanced. Surprisingly, expression of lsrR, lsrK, tam, and yneE, which flank the lsrACDBFG operon, were significantly induced by luxS (2.2, 5.5, 2.8 and 3.5-fold respectively (Wang et al., 2005b)). We investigated their regulation in detail and found that deletion of lsrR significantly increased lsrR expression, indicating that the lsrR transcription was autorepressed. Additionally, the intergenic region between the lsrR and lsr operon, which are divergently transcribed, revealed a CRP binding site which has a typical 6-bp spacer between two conserved motifs. Gel mobility shifts were used to demonstrate CRP binding to the intergenic region containing this site. Moreover, we discovered that lsrR and lsrK are transcribed into one mRNA, forming an lsrRK operon, which is repressed by LsrR and activated by CRP. Interestingly, the tam gene was also found to be a component of the lsr operon. his gene encodes an S-adenosyl-L-methionine-dependent methyltransferase, which catalyzes the methyl esterification of trans-aconitate. he trans-aconitate appears to be formed spontaneously from the citric acid cycle intermediate cis-aconitate. he benefit of methylation of the trans-aconitate to the E. coli cells is not clear. he addition of sugars that are part of the phospho-transferase system increase AI-2 extracellular activity to its maximum level in the mid to late exponential phase (Surette and Bassler, 1998) through a decrease of adenosine 3a,5a-cyclic monophosphate (cAMP) concentration (Wang et al., 2005a). Once glucose is depleted, cAMP concentrations increase, the lsr operon is activated by cAMP, and AI-2 is internalized by cells (Surette and Bassler, 1998). While it is tempting to speculate that the enormous increase in extracellular AI-2 in the presence of PTS sugar, glucose, is due to the tight restriction of its uptake, we have performed stochastic model analysis of the synthesis/uptake mechanism in E. coli and predicted that the rate of DPD to AI-2 conversion requires either (i) additional biological steps or (ii) fueling from another reaction pathway (non LuxS-mediated). While the details of this analysis are found elsewhere (Li et al., 2006), we have shown that the addition of adenosine, the product of eukaryotic SAH hydrolase activity, to extracts of luxS and pfs mutants leads to AI-2 activity. It is also important to note that dihydroxyacetone phosphate (DHAP) represses the lsr operon (Xavier and Bassler, 2005a). Hence, mutations leading to changes in DHAP synthesis affect AI-2 concentrations. For example, both the gatC and agaY mutations increase intracellular AI-2 concentrations and biofilms (Domka et al., 2007). While studying another one of the genes identified by us as induced in biofilms through DNA microarrays (Ren et al., 2004a), we found that the uncharacterized protein YdgG is involved in export of AI-2 (Herzberg et al., 2006). he evidence for this discovery is that deleting ydgG decreased extracellular and increased intracellular concentrations of AI-2 by 13-fold and 16-fold, respectively (Herzberg et al., 2006). Consistent with this hypothesis,

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deleting ydgG resulted in a 7000-fold increase in biofilm thickness and 574-fold increase in biomass in flow cells (Herzberg et al., 2006). Also consistent with the hypothesis, deleting ydgG increased cell motility by increasing transcription of flagella genes (genes induced by AI-2). By expressing ydgG in trans, the wild-type phenotypes for extracellular AI-2 activity, motility, and biofilm were restored. YdgG is predicted to be a membrane-spanning protein that is conserved in many bacteria (Figure 7.2), and it influences resistance against several antimicrobials including crystal violet and streptomycin (this phenotype could also be complemented). Deleting ydgG also caused 31% of the bacterial chromosome to be differentially expressed in biofilms as expected since AI-2 affects hundreds of genes (Beloin et al., 2004; Ren et al., 2004a; Schembri et al., 2003). YdgG was found to negatively modulate expression of flagella- and motility-related genes as well as other known products essential for biofilm formation including operons for type 1 fimbriae, the autotransporter protein Ag43, curli production, colanic acid production, and production of polysaccharide adhesin. Eighty genes not previously related to biofilm formation were also identified including those that encode transport proteins (yihN and yihP), polysialic acid production (gutM and gutQ), CP4-57 prophage functions (y R and alpA), methionine biosynthesis (metR), biotin and thiamine biosynthesis (bioF and thiDFH), anaerobic metabolism (focB, hyfACDR, ttdA, and fumB), and genes with unknown function (ybfG, yceO, yjhQ, and yjbE); ten of these genes were verified through mutation to decrease biofilm formation by 40% or more (y R, bioF, yccW, yjbE, yceO, ttdA, fumB, yjiP, gutQ, and yihR). Hence, the gene that encodes YdgG was renamed tqsA (for transport of the quorum sensing signal AI-2 (Herzberg et al., 2006)).

Figure 7.2 Schematic of YdgG combining data predicted by different bioinformatics programs.

Signaling in Escherichia coli Bioilms

AI-3 signaling Enterohemorrhagic E. coli 0157:H7 and several other bacteria (e.g. Shigella sp.) also have another signal known as AI-3 whose synthesis depends on LuxS (Sperandio et al., 2003). he composition of AI-3 is not known but it appears to require tyrosine for its synthesis (Walters et al., 2006). Furthermore, it remains to be shown that AI-3 is a quorum sensing signal in that the cellular response has not been shown to extend beyond the metabolism required to generate it (Winzer et al., 2002). he importance of AI-3 lies in its ability to activate transcription of pathogenicity genes found in the locus of enterocyte effacement (Sperandio et al., 2003). Both AI-3 (4 MM) (Sperandio et al., 2003) and AI-2 (6.4 MM) (González Barrios et al., 2006) have been shown to control motility directly by inducing transcription of qseBC. N-Acylhomoserine lactone signaling through SdiA he physiological role of SdiA has been unclear in E. coli (Lindsay and Ahmer, 2005). SdiA is a LuxR homologue that is a quorum-sensing-regulated transcription factor in E. coli (Rahmati et al., 2002); in other bacteria, LuxR systems control density-dependent gene regulation through acyl homoserine lactones (AI-1) but E. coli does not have an AI-1 synthase. In E. coli O157-H7, SdiA has been shown to regulate virulence factors (Kanamaru et al., 2000), and SdiA has been shown (by overexpressing SdiA from a plasmid) to inhibit chemotaxis and motility genes in E. coli K-12, to repress tnaA (encodes tryptophanase), as well as to induce indole export via AcrEF (Wei et al., 2001). Recently, it has been determined that purified SdiA responds to three different AI-1 signals (Yao et al., 2006) and that SdiA controls acid resistance via a synthetic AI-1 (van Houdt et al., 2006). Since AI-1 controls biofilms in other strains (Davies et al., 1998), and since we found indole controls biofilms through SdiA (below) (Lee et al., 2006), we investigated if AI-1 signals can control biofilm formation in E. coli (Lee et al., 2006). Adding three naturally occurring AI-1 signals (N-butyryl-DL-homoserine lactone produced by Pseudomonas aeruginosa (Pesci et al., 1999), N-hexanoyl-DL-homoserine lactone produced by P. syringae (Elasri et al., 2001), and N-octanoyl-DL-homoserine lactone produced by P. fluorescens (Elasri et al., 2001)) inhibited K-12 biofilm formation in LB medium in a dose-dependent manner without inhibiting growth by 25%, 27%, and 18%, respectively; however, the isogenic sdiA mutant does not respond to the AI-1 signals (see Figure 7.3 for N-butyryl-DL-homoserine lactone). Hence, E. coli responds to AI-1 signals (signals that are produced by neighboring bacteria but not by itself ) by altering its own biofilm formation and it does so through SdiA (Lee et al., 2006). Indole signaling through SdiA Indole is an extracellular signal in E. coli as it has been shown to regulate expression of astD (encodes arginine succinyltransferase), tnaB (encodes a low-affinity tryptophan permease), and gabT (encodes glutamate:succinate semialdehyde aminotransferase) in the stationary phase for suspension cells (Wang et al., 2001). Indole has also been shown to control multidrug exporters in E. coli (Hirakawa et al., 2005) as well as to regulate the pathogenicity island of pathogenic E. coli (Anyanful et al., 2005) (note tryptophanase activity has also been linked to killing of nematodes by E. coli but indole is not directly responsible for this

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Figure 7.3 Effect of N-butyryl-DL-homoserine lactone on bioilm formation of E. coli K-12 BW25113 and the sdiA mutant in LB medium at 30oC. Biomass was measured at 540 nm after 24 hours. The experiments were repeated ive times (one representative data set shown), and one standard deviation is shown.

effect (Anyanful et al., 2005)). Using DNA microarrays, we discovered that genes for the synthesis of indole (tnaAL) were induced by a stationary phase signal (Ren et al., 2004c) and that the gene encoding tryptophanase, tnaA, was repressed 13-fold in 6-day-old E. coli biofilms in complex medium (Ren et al., 2004a). hese results implied that indole plays a role in biofilm formation since biofilm cells most closely resemble stationary-phase cells (Beloin and Ghigo, 2005; Lazazzera, 2005). Using two E. coli mutants yliH and yceP, which are involved in regulating biofilm formation via both AI-2 and indole, we found that indole probably inhibits biofilm formation since these two mutations lead to biofilms with lower intracellular indole concentrations which leads to dramatic increases in biofilm formation and since the addition of extracellular indole reduced biofilm formation for these mutants (Domka et al., 2006). In contrast, others have reported that indole induces biofilm formation in E. coli as the tnaA deletion decreased biofilm formation and the addition of indole restored it (Di Martino et al., 2003). To determine the genetic basis of indole regulation of biofilms, we utilized a series of tryptophan mutants (Figure 7.4, tnaA encodes tryptophanase synthase, trpE encodes anthranilate synthase component I, tnaL (tnaC) encodes the tryptophanase leader peptide, and trpL encodes the Trp operon leader peptide (Neidhardt, 1996)) as well as conducted three sets of microarray experiments using RNA extracted from biofilm cells grown on glass wool (Lee et al., 2006): (i) direct addition of 600 MM indole to K-12 yceP in LB glu since this strain has elevated biofilm formation due to low intracellular indole and the biofilm responds to added indole (Domka et al., 2006), (ii) K-12 trpE vs. wild type in LB glu since the trpE cells had significantly reduced indole levels (Figure 7.5) and had elevated biofilm in LB glu at 24 hours, and (iii) K-12 tnaA vs. wild type in LB since the tnaA cells also had little detectable indole in this medium.

Signaling in Escherichia coli Bioilms

Chorismate

TrpL

TrpE, TrpD, TrpC

1-(o-carboxyphenilamino)-1’-deoxyribulose-5’-phosphate

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Indole-3-glycerol-phosphate

-

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Indole H2S

Acetate

Acetyl-CoA

L-serine

L-cysteine CysK

+

TnaA + H2 O

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CoenzymeA

CysE

TrpB

H2O

+

Pyruvate + NH3 +H2S

L-tryptophan

TnaL

+

TnaA + H2 O

Protein synthesis

Indole + Pyruvate + NH3

Figure 7.4 Major components of the tryptophan pathway. Dashed lines indicate regulation.

Notably SdiA was one of the most-induced genes (2.9-fold) upon addition of 600 MM indole (Lee et al., 2006). Since SdiA inhibits chemotaxis and motility (Wei et al., 2001), it was expected that a sdiA deletion should lead to enhanced motility and biofilm formation. Corroborating our microarray results, the motility of the isogenic sdiA strain was increased 2-fold and biofilm formation was increased 6.4-fold in LB glu at 37oC at 24 hours and increased 3.5-fold in LB at 30oC at 24 hours. In addition, for short time experiments (8 hours), the sdiA mutation caused a 51-fold increase in biofilm formation at 30oC in LB; however, there was no change in biofilm formation upon deleting sdiA at 37oC with LB (Lee et al., 2006). As expected, the addition of 1000 MM indole to the sdiA mutant in LB glu did not appreciably decrease its elevated biofilm levels. Hence, indole induces expression of SdiA which results in SdiA repressing biofilm formation by decreasing motility which leads to decreased biofilm formation. hese are some of the first results with SdiA showing a phenotypic change in E. coli K12 (Walters and Sperandio, 2006). Similarly, the trpE mutation, which diminishes intracellular indole, led to both increased biofilm and motility which indicates again that indole is a biofilm inhibitor which controls biofilms by reducing motility via SdiA (Lee et al., 2006). In addition, two research groups have found that SdiA induces the multi-drug efflux pump AcrAB of E. coli (Rahmati et al., 2002; Wei et al., 2001), and AcrAB has been hypothesized to control the efflux of quorum signals (Rahmati et al., 2002). Given that we demonstrated that TqsA of E. coli controls the efflux of the quorum signal AI-2 (Herzberg et al., 2006), and others demonstrated that MexAB-OprM of P. aeruginosa controls the efflux of the quorum signal N-(3-oxododecanoyl)-L-homoserine lactone (Evans et al., 1998;

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Figure 7.5 Intracellular and extracellular indole concentration in LB for the E. coli K-12 strains BW25113, BW25113 $trpE, BW25113 $tnaL, BW25113 $tnaA, and BW25113 $trpL. Each experiment was performed in duplicate, and one standard deviation is shown.

Pearson et al., 1999), it appears that indole via its control of SdiA, may also control the efflux of quorum signals as well as control antibiotic resistance (Hirakawa et al., 2005). Indole addition also repressed the glutamate decarboxylase acid-resistance genes gadABCEX 4-fold (Lee et al., 2006). he genes encoding chaperones to prevent aggregation of periplasmic proteins under extremely acidic conditions (Masuda and Church, 2003), hdeABD, were also repressed 3- to 5-fold as was a new locus related to acid resistance, ymgABC (Lee et al., 2006). To show clearly that indole is directly related to acid resistance, 2 mM indole was added to the wild type at pH 2.5, and we found that its survival was decreased 350 to 650-fold. In addition, the trpE mutant (which produces 10 times less indole), was 53 times less sensitive to pH 2.5 than the wild type. Hence, indole decreases acid resistance as shown by these two independent experiments (Lee et al., 2006). In addition, isogenic mutants showed these genes are directly related to biofilm formation (6.4-fold increase for sdiA, 6-fold for gadA, 7.3-fold for gadE, 4.3-fold for hdeA, 1.7-fold for hdeB, and 2-fold for hdeD) (Lee et al., 2006). Because indole accumulates extracellulary during a specific phase of growth (stationary) (Baca-Delancey et al., 1999), is recognized by a specific receptor (Mtr) (Wang et al., 2001), and has been shown to control genes beyond those used to synthesize it or detoxify it (e.g. sdiA, gadABCEX, hdeABD, ymgABC), it is likely that it is a true quorum sensing signal (Winzer et al., 2002). Toluene o-monooxygenase (TOM) of the soil bacterium Burkholderia cepacia G4 converts indole into isoindigo (Rui et al., 2005); hence, we hypothesized that if indole represses E. coli biofilm formation, then E. coli would be present in higher numbers in a dual-species biofilm in which TOM was expressed. TOM was integrated into the chromosome of the P. fluorescens strain to diminish the metabolic burden of this locus (Yee et al., 1998) and

Signaling in Escherichia coli Bioilms

because this strain does not produce indole. When the Pseudomonas was tagged with red fluorescent protein and E. coli with a green fluorescent protein so that both bacteria could be visualized, constitutive expression of TOM led to a 12-fold increase in E. coli biofilm after five days in the flow cell (Figure 7.6). By expressing TOM to remove indole, both substratum coverage and mean thickness increased 10-fold. hus, including an organism that actively expressed TOM increased the amount of E. coli biofilm by decreasing the amount of indole by 22-fold (Lee et al., 2006). hese results show that indole is an interspecies signal and indicate that it may be manipulated to control biofilms in complex communities (Lee et al., 2006). Indole is also a signal for pseudomonads since indole at 500 MM increased biofilm formation 1.6-fold and 1000 MM increased biofilm formation 2.5-fold; hence, indole, although not synthesized by P. aeruginosa (extracellular concentration was 0 MM), indole is a signal that stimulates biofilm in this strain. In addition, removing the 1000 MM added indole by expressing TOM in P. fluorescens 2–79 results in a 5.7-fold reduction in its biofilm again showing that indole stimulates biofilm formation in this pseudomonad. Hence, indole is an

Figure 7.6 Bioilm formation in LB medium after 5 days in low cells for (A) dual species of E. coli K-12 XL1-Blue/pCM18 (stained with GFP) and P. luorescens 2-79TOM/pHKT3 expressing toluene O-monooxygenase (TOM) (stained with RFP), and (B) dual species of E. coli K-12 XL1Blue/pCM18 (stained with GFP) and P. luorescens 2-79/pHKT3 (stained with RFP). Scale bar is 10 Mm. Both pseudomonads contained RFP via the broad-host-range plasmid pHKT3 (Tomlin et al., 2004) and E. coli K-12 XL1-Blue was used with GFP from pCM18 (Hansen et al., 2001); in this way, both bacteria were tagged with a luorophore. TOM was active in P. luorescens 279TOM/pHKT3 (RFP) (0.24 nmol/minute mg protein) based on a naphthalene to naphthol assay and high-pressure liquid chromatography (HPLC) (Tao et al., 2005) (constitutively expressed TOM converts indole to isoindigo (Yee et al., 1998)). See also Plate 7.6.

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interspecies signal that may be manipulated by oxygenases of another bacterium (one that does not necessarily synthesize it) to control biofilm formation. he type of signal manipulation reported above suggests that competition and control for signals are intense in biofilms. Competition for signals for planktonic cells has been demonstrated previously for AI-2 (Xavier and Bassler, 2005b) and for AI-1 (Zhang and Dong, 2004). his competition extends beyond prokaryotes as eukaryotes are well known for manipulating the quorum sensing signals of bacteria, too. For example, algae block bacterial biofilm formation by controlling both AI-1 and AI-2 signaling via furanones (Ren et al., 2001; Ren et al., 2004b), and mammals (including humans) blocking AI-1 signaling via lactonase in sera (Yang et al., 2005). Furthermore, it appears that the mechanism by which prokaryotes (such as the pseudomonads in this study) manipulate the biofilm signal indole is through the relaxed substrate range of many oxygenases found in bacteria that bring about indole hydroxylation (Rui et al., 2005); i.e. we propose that some of the oxygenases bacteria use for catabolism (Fishman et al., 2005) have also evolved to regulate concentrations of the inter-species signal indole by removing it via precipitation: competitors that wish to remove indole simply oxidize it in one step to indigo which is insoluble and hence leaves the system (Lee et al., 2006). 5-Hydroxyindole and isatin signaling Since competition in biofilms for signals is intense (there are as many as 500 to 1000 different bacteria in the gastrointestinal tract) and given that the human host itself makes indolelike hormones, it is expected that the large quantity of indole produced by E. coli would be manipulated by surrounding intestinal bacteria and that the intestinal bacteria would utilize the indole-like compounds epinephrine and norepinephrine produced by the human host. We reasoned that given the high indole concentrations for E. coli in rich medium (Figure 7.5) (Lee et al., 2006) and given the ability of oxygenases to convert indole into insoluble indigoids (Rui et al., 2005), other bacteria would take advantage of this interspecies signal indole and manipulate it. Hence we tested a series of hydroxyindoles for their ability to alter biofilm formation of E. coli O157:H7. We found that the doubly hydroxylated isatin increases the bottom biofilm formation in a 96-well plate by 4-fold whereas 5-hydroxyindole reduces biofilm formation by 6-fold (84%) (unpublished). Hence, hydroxylated indoles affect E. coli in different ways which opens the possibility that this bacterium can discern the presence of other bacteria by detecting the way its signal is manipulated. Indole, norepinephrine, and epinephrine signaling in EHEC We also propose that the DNA microarray analyses and results with isogenic mutants provide insight into how the bacteria of the gastrointestinal (GI) tract may help to restrict access to pathogens. Since indole represses the genes for acid resistance (gad, hde, and ymg acid-resistance operons) in E. coli K-12 (Lee et al., 2006), increases in extracellular indole in the GI tract may result in the exclusion of the pathogen E. coli O157:H7 from the GI tract environment by affecting its ability to resist the low-pH environment of the stomach. In addition, since indole represses genes that encode virulence regulators such as gadX (gadX was repressed 2-fold by indole (Lee et al., 2006)), indole may reduce virulence in the duodenum since GadX activates virulence genes there (Moreira et al., 2006). Alternatively,

Signaling in Escherichia coli Bioilms

the repression of motility genes by indole could also lead to increased washout of pathogen from the GI tract and, thereby, decrease their colonization. We speculate that the extracellular levels of indole increase only after the biofilm has reached a certain critical thickness so that indole impacts acid resistance and motility only in the colonizing pathogenic bacteria. he pathogen E. coli O157:H7 may use indole as a signal since SdiA represses expression of the virulence factors EspD and intimin (Kanamaru et al., 2000). In addition, indole from bacteria is absorbed into the body (Gillam et al., 2000) so cells of the gastrointestinal tract may also manipulate indole levels to control bacteria. It is becoming clear that prokaryotes and eukaryotes signal not only themselves but also one another; for example, there appears to be crosstalk between E. coli O157:H7 (EHEC) and cells of the gastrointestinal tract through the hormones epinephrine and norepinephrine (catecholamines) (Kaper and Sperandio, 2005). Using microarrays, we have found that the interspecies signal indole represses key virulence genes of EHEC as well as that the human hormones epinephrine and norepinephrine stimulate virulence (unpublished). Other hormones are also present in the gastrointestinal tract including melatonin (Lee and Pang, 1993) and serotonin (Meyer and Brinck, 1997); both are neural hormones which maintain homeostasis and both reduce chlamydial infection (Rahman et al., 2005). In addition, plants use indole 3-acetic acid as their main hormone (for cell growth, division, tissue differentiation, and response to light and gravity), and bacteria are known to interrupt this eukaryotic signaling by using indole 3-acetic acid as a source of carbon, nitrogen, and energy (Leveau and Lindow, 2005). All five hormones have indole-like chemical structures (Figure 7.7); hence, although it is highly speculative, it is intriguing to ponder whether indole that was produced by bacteria was incorporated into the metabolism of eukaryotic hosts (plants and animals) and is the archetypal hormone.

H N

H N

H N OH

NH2 O

Indole

HO

Serotonin

Indole-3-acetic acid

OH H N

HO

H N CH3

O H3 C

O

N H

Melatonin

CH3

HO

Epinephrine

Figure 7.7 Chemical structures of indole, melatonin, serotonin, epinephrine, and indole-3acetic acid. Indole motifs are in bold.

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Acknowledgments Financial support was provided by the NIH (EB003872-01A1), the National Science Foundation (BES-0124401), and the US Environmental Protection Agency. References Anyanful, A., Dolan-Livengood, J.M., Lewis, T., Sheth, S., DeZalia, M.N., Sherman, M.A., Kalman, L.V., Benian, G.M., and Kalman, D. (2005). Paralysis and killing of Caenorhabditis elegans enteropathogenic Escherichia coli requires the bacterial tryptophanase gene. Mol. Microbiol. 57, 988–1007. Baca-Delancey, R.R., South, M.M.T., Ding, X., and Rather, P.N. (1999). Escherichia coli genes regulated by cell-to-cell signaling. Proc. Natl. Acad. Sci 96, 4610–4614. Balestrino, D., Haagensen, J.A.J., Rich, C., and Forestier, C. (2005). Characterization of type 2 quorum sensing in Klebsiella pneumoniae and relationship with biofilm formation. J. Bacteriol. 187, 2870– 2880. Bassler, B.L., Greenberg, E.P., and Stevens, A.M. (1997). Cross-species induction of luminescence in the quorum-sensing bacterium Vibrio harveyi. J. Bacteriol. 179, 4043–4045. Beloin, C., Valle, J., Latour-Lambert, P., Faure, P., Kzreminski, M., Balestrino, D., Haagensen, J.A.J., Molin, S., Prensier, G., Arbeille, B., and Ghigo, J.-M. (2004). Global impact of mature biofilm lifestyle on Escherichia coli K-12 gene expression. Mol. Microbiol. 51, 659–674. Beloin, C., and Ghigo, J.-M. (2005). Finding gene-expression patterns in bacterial biofilms. Trends Microbiol. 13, 16–19. Blattner, F.R., III, G.P., Bloch, C.A., Perna, N.T., Burland, V., Riley, M., Collado-Vides, J., Glasner, J.D., Rode, C.K., Mayhew, G.F., Gregor, J., Davis, N.W., Kirkpatrick, H.A., Goeden, M.A., Rose, D.J., Mau, B., and Shao, Y. (1997). he complete genome sequence of Escherichia coli K-12. Science 277, 1453–1462. Chain, P.S.G., Carniel, E., Larimer, F.W., Lamerdin, J., Stoutland, P.O., Regala, W.M., Georgescu, A.M., Vergez, L.M., Land, M.L., Motin, V.L., Brubaker, R.R., Fowler, J., Hinnebusch, J., Marceau, M., Medigue, C., Simonet, M., Chenal-Francisque, V., Souza, B., Dacheux, D., Elliott, J.M., Derbise, A., Hauser, L.J., and Garcia, E. (2004). Insights into the evolution of Yersinia pestis through whole-genome comparison with Yersinia pseudotuberculosis. Proc. Natl. Acad. Sci. USA 101, 13826–13831. Chilcott, G.S., and Hughes, K.T. (2000). Coupling of flagellar gene expression to flagellar assembly in Salmonella enterica Serovar typhimurium and Escherichia coli. Microbiol. Mol. Biol. Rev. 64, 694–708. Cole, S.P., Hardwood, J., Lee, R., She, R., and Guiney, D.G. (2004). Characterization of monospecies biofilm formation by Helicobacter pylori. J. Bacteriol. 186, 3124–3132. Davies, D.G., Parsek, M.R., Pearson, J.P., Iglewski, B.H., Costerton, J.W., and Greenberg, E.P. (1998). he involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280, 295–298. Di Martino, P., Fursy, R., Bret, L., Sundararaju, B., and Phillips, R.S. (2003). Indole can act as an extracellular signal to regulate biofilm formation of Escherichia coli and other indole-producing bacteria. Can. J. Microbiol. 49, 443–449. Domka, J., Lee, J., and Wood, T.K. (2006). YliH (BssR) and YceP (BssS) regulate Escherichia coli K-12 biofilm formation by influencing cell signaling. Appl. Environ. Microbiol. 72, 2449–2459. Domka, J., Lee, J., Bansai, T., and Wood, T.K. (2007). Temporal gene-expression in Escherichia coli K-12 biofilms. Environ. Microbiol. 9, 322–346. Elasri, M., Delorme, S., Lemanceau, P., Stewart, G., Laue, B., Glickmann, E., Oger, P.M., and Dssaux, Y. (2001). Acyl-homoserine lactone production is more common among plant-associated Pseudomonas spp. than among soilborne Pseudomonas spp. Appl. Environ. Microbiol. 67, 1198–1209. Evans, K., Passador, L., Srikumar, R., Tsang, E., Nezezon, J., and Poole, K. (1998). Influence of the MexAB-OprM multidrug efflux system on quorum sensing in Pseudomonas aeruginosa. J. Bacteriol. 180, 5443–5447. Fishman, A., Tao, Y., Rui, L., and Wood, T.K. (2005). Controlling the regiospecific oxidation of aromatics via active site engineering of toluene para-monooxygenase of Ralstonia pickettii PKO1. J. Biol. Chem. 280, 506–514. Ghigo, J.-M. (2001). Natural Conjugative plasmids induce bacterial biofilm development. Nature 412, 442–445.

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Gillam, E., Notley, L., Cai, H., De Voss, J., and Guengerich, F. (2000). Oxidation of indole by cytochrome P450. Enzymes 39, 13817–13824. González Barrios, A.F., Zuo, R., Y. Hashimoto, Yang, L., Bentley, W.E., and Wood, T.K. (2006). Autoinducer 2 controls biofilm formation in Escherichia coli through a novel motility quorum sensing regulator (MqsR, B3022). J. Bacteriol. 188, 305–306. Hammer, B.K., and Bassler, B.L. (2003). Quorum sensing controls biofilm formation in Vibrio cholerae. Mol. Microbiol. 50, 101–104. Hansen, M.C., Palmer, R.J., Jr., Udsen, C., White, D.C., and Molin, S. (2001). Assessment of GFP fluorescence in cells of Streptococcus gordonii under conditions of low pH and low oxygen concentration. Microbiology 147, 1383–1391. Henke, J.M., and Bassler, B.L. (2004). hree parallel quorum-sensing systems regulate gene expression in Vibrio harveyi. J. Bacteriol. 186, 6902–6914. Herzberg, M., Kaye, I.K., Peti, W., and Wood, T.K. (2006). YdgG (TqsA) controls biofilm formation in Escherichia coli K-12 by enhancing autoinducer 2 transport. J. Bacteriol. 188, 587–598. Hirakawa, H., Inazumi, Y., Masaki, T., Hirata, T., and Yamaguchi, A. (2005). Indole induces the expression of multidrug exporter genes in Escherichia coli. Mol. Microbiol. 55, 1113–1126. Kanamaru, K., Kanamaru, K., Tasuno, I., Tobe, T., and Sasakawa, C. (2000). SdiA, an Escherichia coli homologue of quorum-sensing regulators, controls the expression of virulence factors in enterohaemorrhagic Escherichia coli O157:H7. Mol. Microbiol. 38, 805–816. Kang, Y., Durfee, T., Glasner, J.D., Qiu, Y., Frisch, D., Wintemberg, K.M., and Blatnner, F.R. (2004). Systematic mutagenesis of the Escherichia coli genome. J. Bacteriol. 186, 4921–4930. Kaper, J.B., and Sperandio, V. (2005). Bacterial cell-to-cell signaling in the gastrointestinal tract. Infect. Immun. 73, 3197–3209. Labbate, M., Queck, S.Y., Koh, K.S., Rice, S.A., Givskov, M., and Kjelleberg, S. (2004). Quorum-sensingcontrolled biofilm development in Serratia liquefaciens MG1. J. Bacteriol. 186, 692–698. Lazazzera, B.A. (2005). Lessons from DNA microarray analysis: the gene expression profile of biofilms. Curr. Opin. Microbiol. 8, 222–227. Lee, J., Jayaraman, A., and Wood, T.K. (2006). Indole and acyl-homoserine lactones are inter-species Escherichia coli biofilm signals mediated by SdiA. BMC Microbiol. in revision. Lee, P.P.N., and Pang, S.F. (1993). Melatonin and its receptors in the gastrointestinal tract. Biol. Signals 2, 181–193. Leveau, J.H.J., and Lindow, S.E. (2005). Utilization of the plant hormone indole-3-acetic acid for growth by Pseudomonas putida strain 1290. Appl. Environ. Microbiol. 71, 2365–2371. Li, J., Wang., L., Hashimoto, Y., Tsao, C.-Y., Wood, T.K., Valdes, J.J., Zafiriou, E., and Bentley, W.E. (2006). A stochastic model of E. coli AI-2 quorum signal circuit reveals alternative synthesis pathways. Mol. Systems Biol. 2, 67. Li, Y.-H., Lau, P.C.Y., Lee, J.H., Ellen, R.P., and Cvitkovitch, D.G. (2001). Natural genetic transformation of Streptococcus mutans growing in biofilms. J. Bacteriol.183, 897–908. Lindsay, A., and Ahmer, B.M.M. (2005). Effect of sdiA on biosensors of N-acylhomoserine lactones. Appl. Environ. Microbiol. 187, 5054–5068. Lombardia, E., Rovetto, A.J., Arabolaza, A.L., and Grau, R.R. (2006). A LuxS-dependent cell-to-cell language regulates social behavior and development in Bacillus subtilis. J. Bacteriol. 188, 4442–4452. Masuda, N., and Church, G.M. (2003). Regulatory network of acid resistance genes in Escherichia coli. Mol. Microbiol. 48, 699–712. McNab, R., Ford, S.K., El-Sabaeny, A., Barbieri, B., Cook, G.S., and Lamont, R.J. (2003). LuxS-based signaling is Streptococcus gordonii:autoinducer 2 controls carbohydrate metabolism and biofilm formation with Porphyromonas gingivalis. J. Bacteriol. 185, 274–284. Meyer, T., and Brinck, U. (1997). Differential distribution of serotonin and tryptophan hydroxylase in the human gastrointestinal tract. Digestion 60, 63–68. Moreira, C.G., Palmer, K., Whiteley, M., Sircili, M.P., Trabulsi, L.R., Castro, A.F.P., and Sperandio, V. (2006). Bundle-forming Pili and EspA are involved in biofilm formation by enteropathogenic Escherichia coli. J. Bacteriol. 188, 3952–3961. Neidhardt, F.C. (ed) (1996). Escherichia coli and Salmonella: Cellular and Molecular Biology. Washington, D.C.: ASM Press. Parkhill, J., Sebaihia, M., Preston, A., Murphy, L.D., homson, N., Harris, D.E., Holden, M.T., Churcher, C.M., Bentley, S.D., Mungall, K.L., Cerdeno-Tarraga, A.M., Temple, L., James, K., Harris, B., Quail,

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M.A., Achtman, M., Atkin, R., Baker, S., Basham, D., N. Bason, Cherevach, I., Chillingworth, T., Collins, M., Cronin, A., Davis, P., Doggett, J., Feltwell, T., Goble, A., Hamlin, N., Hauser, H., Holroyd, S., Jagels, K., Leather, S., S. Moule, Norberczak, H., O’Neil, S., Ormond, D., Price, C., Rabbinowitsch, E., Rutter, S., Sanders, M., Saunders, D., Seeger, K., Sharp, S., Simmonds, M., Skelton, J., Squares, R., Squares, S., Stevens, K., Unwin, L., Whitehead, S., Barrell, B.G., and Maskell, D.J. (2003). Comparative analysis of the genome sequences of Bordetella pertussis, Bordetella parapertussis and Bordetella bronchiseptica. Nat. Genet. 35, 32–40. Paulsen, I.T., Press, C.M., Ravel, J., Kobayashi, D.Y., Myers, G.S.A., Mavrodi, D.V., DeBoy, R.T., Seshadri, R., Ren, Q., Madupu, R., Dodson, R.J., Durkin, A.S., Brinkac, L.M., Daugherty, S.C., Sullivan, S.A., Rosovitz, M.J., Gwinn, M.L., Zhou, L., Schneider, D.J., Cartinhour, S.W., Nelson, W.C., Weidman, J., Watkins, K., Tran1, K., Khouri1, H., Pierson, E.A., III, L.S.P., homashow, L.S., and Loper, J.E. (2005). Complete genome sequence of the plant commensal Pseudomonas fluorescens Pf-5. Nat. Biotech. 23, 873–878. Pearson, J.P., Van Delden, C., and Iglewski, B.H. (1999). Active efflux and diffusion are involved in transport of Pseudomonas aeruginosa cell-to-cell signals. J. Bacteriol. 181, 1203–1210. Pesci, E.C., Milbank, J.B.J., Pearson, J.P., McKnight, S., Kende, A.S., Greenberg, E.P., and Iglewski, B.H. (1999). Quinolone signaling in the cell-to-cell communication system of Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 96, 11229–11234. Pratt, L.A., and Kolter, R. (1998). Genetic analysis of Escherichia coli Biofilm formation: roles of flagella, motility, chemotaxis and type I pili. Mol. Microbiol. 30, 285–293. Rahman, M.A., Azuma, Y., Fukunaga, H., Murakami, T., Sugi, K., Fukushi, H., Miura, K., Suzuki, H., and Shirai, M. (2005). Serotonin and melatonin, neurohormones for homeostasis, as novel inhibitors of infections by the intracellular parasite chlamydia. J. Antimicrob. Chemother. 56, 861–868. Rahmati, S., Yang, S., Davidson, A.L., and Zechiedrich, E.L. (2002). Control of the AcrAB multidrug efflux pump by quorum-sensing regulator SdiA. Mol. Microbiol. 43, 677–685. Reisner, A., Haagensen, J.A.J., Schembri, M.A., Zechner, E.L., and Molin, S. (2003). Development and maturation of Escherichia coli K-12 biofilms. Mol. Microbiol. 48, 933–946. Ren, D., Sims, J.J., and Wood, T.K. (2001). Inhibition of biofilm formation and swarming of Escherichia coli by (5Z)-4-bromo-5-(bromomethylene)-3-butyl-2 (5H)-furanone. Environ. Microbiol. 3, 731–736. Ren, D., Bedzyk, L.A., homas, S.M., Ye, R.W., and Wood, T.K. (2004a). Gene expression in Escherichia coli biofilms. Appl. Microbiol. Biotech. 64, 515–524. Ren, D., Bedzyk, L.A., homas, S.M., Ye, R.W., and Wood, T.K. (2004b). Differential gene expression shows natural brominated furanones interfere with the autoinducer-2 bacterial signaling system of Escherichia coli. Biotech. Bioengineer. 88, 630–642. Ren, D., Bedzyk, L.A., Ye, R.W., homas, S.M., and Wood, T.K. (2004c). Stationary phase quorum-sensing signals affect autoinducer-2 and gene expression in Escherichia coli. Appl. Environ. Microbiol.70, 2038–2043. Ren, D., Zuo, R., Barrios, A.F.G., Bedzyk, L.A., Eldridge, G.R., Pasmore, M.E., and Wood, T.K. (2005). Differential gene expression to investigate Escherichia coli biofilm inhibition by plant extract Ursolic Acid. Appl. Environ. Microbiol. 71, 4022–4034. Riley, M., and Labedan, B. (eds) (1996). Escherichia coli Genes Products: Physiological Functions and Common Ancestries. Washington, D.C: ASM Press. Rui, L., Reardon, K.F., and Wood, T.K. (2005). Protein engineering of toluene ortho-monooxygenase of Burkholderia cepacia G4 for regiospecific hydroxylation of indole to form various indigoid compounds. Appl. Microbiol. Biotech. 66, 422–429. Schembri, M.A., Kjærgaard, K., and Klemm, P. (2003). Global Gene Expression in Escherichia coli Biofilms. Mol. Microbiol. 48, 253–267. Selinger, D.W., Cheung, K.J., Mei, R., Johansson, E.M., Richmond, C.S., Blattner, F.R., Lockhart, D.J., and Church, G.M. (2000). RNA Expression Analysis Using a 30 Base Pair Resolution Escherichia coli Genome Array. Nature Biotech. 18, 1262–1268. Song, Y., Tong, Z., Wang, J., Wang, L., Guo, Z., Han, Y., Zhang, J., Pei, D., Zhou, D., Qin, H., Pang, X., Han, Y., Zhai, J., Li, M., Cui, B., Z. Qi, L. Jin, R. Dai, Chen, F., Li, S., Ye, C., Du, Z., Lin, W., Wang, J., Yu, J., Yang, H., Wang, J., Huang, P., and Yang, R. (2004). Complete genome sequence of Yersinia pestis strain 91001, an isolate avirulent to humans. DNA Res. 11, 179–197.

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Sperandio, V., Torres, A.G., and Kaper, J.B. (2002). Quorum sensing Escherichia coli regulators B and C (QseBC): a novel two-component regulatory system involved in the regulation of flagella and motility by quorum sensing in E. coli. Mol. Microbiol. 43, 809–821. Sperandio, V., Torres, A.G., Jarvis, B., Nataro, J.P., and Kaper, J.B. (2003). Bacteria–host communication: the language of hormones. Proc. Natl. Acad. Sci. USA 100, 8951–8956. Surette, M.G., and Bassler, B.L. (1998). Quorum sensing in Escherichia coli and Salmonella typhimurium. Proc. Natl. Acad. Sci. USA 95, 7046–7050. Surette, M.G., Miller, M.B., and Bassler, B.L. (1999). Quorum sensing in Escherichia coli, Salmonella typhimurium, and Vibrio harveyi: a new family of genes responsible for autoinducer production. Proc. Natl. Acad. Sci. USA 96, 1639–1644. Taga, M.E., Semmelhack, J.L., and Bassler, B.L. (2001). he LuxS-dependent autoinducer AI-2 controls the expression of an ABC transporter that functions in AI-2 uptake in Salmonella typhimurium. Mol. Microbiol. 42, 777–793. Taga, M.E., Miller, S.T., and Bassler, B.L. (2003). Lsr-Mediated Transport and Processing of AI-2 in Salmonella typhimurium. Mol. Microbiol. 50, 1411–1427. Tao, Y., Bentley, W.E., and Wood, T.K. (2005). Phenol and 2-naphthol production by toluene 4-monooxygenases using an aqueous/dioctyl phthalate system. Appl. Microbiol. Biotech. 68, 614–621. Tomlin, K.L., Clark, S.R.D., and Ceri, H. (2004). Green and red fluorescent protein vectors for use in biofilm studies of the intrinsically resistant Burkholderia cepacia complex. J. Microbiol. Methods 57, 95–106. Toussaint, A., Merlin, C., S. Monchy, Benotmane, M.A., Leplae, R., Mergeay, M., and Springael, D. (2003). he biphenyl- and 4-chlorobiphenyl-catabolic transposon Tn4371, a member of a new family of genomic islands related to IncP and Ti plasmids. Appl. Environ. Microbiol. 69, 4837–4845. van Houdt, R., Aertsen, A., Moons, P., Vanoirbeek, K., and Michiels, C.W. (2006). N-Acyl-L-homoserine lactone signal interception by Escherichia coli. FEMS Microbiol. Letters 256, 83–89. Walters, M., Sircili, M.P., and Sperandio, V. (2006). AI-3 Synthesis is not dependent on luxS in Escherichia coli. J. Bacteriol. 188, 5668–5681. Walters, M., and Sperandio, V. (2006). Quorum sensing in Escherichia coli and Salmonella. Int. J. Med. Microbiol. 296, 125–131. Wang, D., Ding, X., and Rather, P.N. (2001). Indole can act as an extracellular signal in Escherichia coli. J. Bacteriol.183, 4210–4216. Wang, L., Hashimoto, Y., Tsao, C.-Y., Valdes, J.J., and Bentley, W.E. (2005a). Cyclic AMP (cAMP) and cAMP receptor protein influence both synthesis and uptake of extracellular autoinducer 2 in Escherichia coli. J. Bacteriol.187, 2066–2076. Wang, L., Li, J., March, J.C., Valdes, J.J., and Bentley, W.E. (2005b). LuxS-dependent gene regulation in Escherichia coli K12 revealed by genomic expression profiling. J. Bacteriol. 187, 8350–8360. Wei, Y., Lee, J.-M., Smulski, D.R., and LaRossa, R.A. (2001). Global impact of sdiA amplification revealed by comprehensive gene expression profiling of Escherichia coli. J. Bacteriol.183, 2265–2272. Winzer, K., Hardie, K.R., and Williams, P. (2002). Bacterial cell-to-cell communications: sorry, can’t talk now-gone to lunch! Curr. Opin. Microbiol. 5, 216–222. Xavier, K.B., and Bassler, B.L. (2005a). Regulation of uptake and processing of the quorum-sensing autoinducer AI-2 in Escherichia coli. J. Bacteriol. 187, 238–248. Xavier, K.B., and Bassler, B.L. (2005b). Interference with AI-2-mediated bacterial cell–cell communication. Nature 437, 750–753. Yang, F., Wang, L.-H., Wang, J., Dong, Y.-H., Hu, J.Y., and Zhang, L.-H. (2005). Quorum quenching enzyme activity is widely conserved in the sera of mammalian species. FEBS Lett. 579, 3713–3717. Yao, Y., Martinez-Yamout, M.A., Dickerson, T.J., Brogan, A.P., Wright, P.E., and Dyson, H.J. (2006). Structure of the Escherichia coli quorum sensing protein SdiA: activation of the folding switch by acyl homoserine lactones. J. Mol. Biol. 355, 262–273. Yee, D.C., Maynard, J.A., and Wood, T.K. (1998). Rhizoremediation of trichloroethylene by a recombinant, root-colonizing Pseudomonas fluorescens strain expressing toluene ortho-monooxygenase constitutively. Appl. Environ. Microbiol. 64, 112–118. Zhang, L.-H., and Dong, Y.-H. (2004). Quorum sensing and signal interference: diverse implications. Mol. Microbiol. 53, 1563–1571.

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Abstract he study of peptide signaling is yielding both fascinating and valuable information regarding bacterial biofilm development and helping to elucidate the disease processes caused by several human pathogens. Peptide signaling potentially impacts all stages of the biofilm “life-cycle” for many bacterial species, from attachment to maturation and detachment. While the particular roles of the signaling systems in biofilm formation varies among species, the implications of several phenomena, from natural transformation of streptococci to quorum sensing variant generation in staphylococci, may only be fully appreciated in the context of biofilms and cell-to-cell signaling. Understanding the mechanisms by which the peptide signaling systems exert their effects on biofilms may yield therapeutic strategies with limited, but important, uses. Introduction he role of peptide signaling (also known as quorum sensing) in bacterial biofilm development continues to attract significant attention, in no small part due to the potential of disrupting biofilm formation, preventing expression of pathogenic factors, and understanding certain disease processes. However, as quickly becomes evident through review of relevant literature, the contribution of peptide signaling to pathogenesis and biofilm behavior can vary significantly not only species to species but even within species. he pleotropic regulatory effects exerted by these signaling systems, their interaction with other regulatory elements, and the effects of varied growth conditions can make for somewhat conflicting results from study to study. Still, important insights are being made into the physiological and signaling processes involved in biofilm development, including both the commonalities and differences between species. While bacteria have historically been studied in planktonic cell cultures, it is clear that biofilm-associated organisms are different by just about any measure. hey exhibit markedly different gene and protein expression profiles as compared to planktonic cells and are remarkably resistant to treatment with antimicrobials. Furthermore, the frequently heterogeneous nature of the biofilm environment (gradients in carbon sources, oxygen, metabolic by-products, etc.) makes for a correspondingly heterogeneous population of cells unlike those in fairly uniform planktonic cultures. Yet, elements of the traditional batch culture appear to translate meaningfully to various stages of biofilm formation, perhaps best

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illustrated by the staphylococci (Figure 8.1). Expression of staphylococcal cell surface-associated adhesins occurs during early growth at low cell densities in planktonic cell culture when peptide signal concentrations are also low. hese adhesins, such as the fibrinogenbinding and fibronectin-binding proteins, can mediate attachment to biological surfaces through ligand–receptor interactions. Large cell-surface proteins, including the adhesins, can also enhance attachment to abiotic substrates through non-specific physiochemical means, such as hydrophobic interactions. Correspondingly, these colonizing factors would be important in vivo when initial cell densities are relatively low and nutrient supply nonlimiting. In contrast, once staphylococcal planktonic cell cultures reach high densities and nutrient supply is limited, signal concentrations are high and the cultures tend to produce extracellular enzymes (e.g. proteases, lipases and hemolysins), immunostimulatory factors (e.g. superantigens) and even some molecules with surfactant properties (e.g. staphylococcal D-toxin). In vivo, such cell density and limited nutrient supply could be found in biofilms or abscesses, and these extracellular factors may contribute to cell detachment and dissemination though cleavage of cell-to-cell or cell-to-host bonds and degradation of surrounding host tissue. he increased vascular permeation that accompanies expression of extracellular virulence factors may also enhance the spread of organisms from one site to another. Once

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Spread to new sites Figure 8.1 Model of staphylococcal virulence gene expression in vitro and in vivo. Expression of cell surface-associated adhesins enhances colonization and proliferation of staphylococci at the site of infection. Growth of the staphylococcal colony leads to a mature bioilm. Expression of extracellular enzymes, toxins and surfactants facilitates escape of staphylococci from the localized infection and subsequent colonization of secondary sites or hosts.

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released from the colony, cells experience low signal concentrations and revert to expression of surface adhesins that mediate attachment, enabling colonization of secondary sites. his model can only be presented in a simplistic fashion in Figure 8.1 as the specific bacterial agents involved in this process, the timing of their induction, and their activity will vary from species to species, and even within species, based on the disease process. Yet, as this chapter will show, evidence of interplay between peptide signaling, cell density, and bacterial products involved in colonization and dissemination is widespread enough among certain bacteria to deserve further investigation into the regulatory systems that control the biofilm “life-cycle” of cell attachment, biofilm growth and maturation, cell detachment, and finally, cell attachment at secondary sites. hough peptide-based signaling systems have been identified in some Gram-negative bacteria, little is known regarding their role, if any, in biofilm formation. hus, this chapter will focus on the biology of peptide signaling and biofilm formation by Gram-positive bacteria. Among these, the signaling systems have been best characterized in three genera, Staphylococcus, Streptococcus, and Enterococcus. Unlike the acyl-homoserine lactone signals found in many Gram-negative bacteria (Atkinson et al., this volume), the peptide signals that mediate quorum sensing in these Gram-positive bacteria cannot freely diffuse through the cell membrane. Instead, these signaling systems are generally characterized by twocomponent regulatory systems that sense and respond to secreted signals. Usually, distinct genetic loci exist that encode the two-component system, the peptide signal precursor, and proteins likely involved in the processing and/or secretion of the signal. Peptide signaling and Staphylococcus bioilms Staphylococci are remarkably adept at causing a variety of human and animal diseases. hese range from relatively benign skin infections, such as impetigo, to much more serious ones, including toxic shock syndrome. Many staphylococcal infections appear to take the form of biofilms, including endocarditis, osteomyelitis, and even some skin infections. In fact, biofilm formation is considered to be the primary “virulence factor” of certain coagulase-negative staphylococci, including S. epidermidis, a common cause of implanted medical device-related infections. The Agr quorum-sensing system Since the identification of the accessory gene regulator (Agr) quorum sensing system in Staphylococcus aureus and subsequently in other staphylococcal species it has been assigned a central role in the regulation of staphylococcal virulence (Kong et al., 2006; Novick, 2003; Novick, 2006; Yarwood, 2006). As such, it has attracted substantial attention as a potential target for controlling staphylococcal disease. While virulence gene regulation by Agr appears to be considerably more complex in vivo than initially understood from in vitro studies, expression of Agr, or even lack thereof, remains an important determinant in staphylococcal disease development. agr mutants have been shown to be attenuated for virulence in some animal models of infection, including a murine arthritis model, an osteomyelitis model, and a skin abscess model (Novick, 2003). It has also been shown that expression of Agr, and Agr-regulated exotoxins, facilitates escape of S. aureus internalized by epithelial cells (Shompole et al., 2003). Now evidence is accumulating that the Agr sys-

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tem plays a significant role in biofilm formation, development and behavior, with important implications for human disease. his chapter will focus primarily on the Agr system of S. aureus, in which Agr, and virulence in general, has been best studied, and to a lesser extent, S. epidermidis. Agr homologs have been identified in many additional staphylococcal species, but little or no investigation has been conducted into regulation of biofilm formation by Agr in these staphylococci. he agr locus consists of two divergent operons (Figure 8.2) (Novick, 2003). he P2 operon (agrACDB) encodes the proteins necessary for signal synthesis, processing, secretion, and recognition, while the transcript of the P3 operon, RNAIII, mediates the regulatory effects of Agr expression. he autoinducing peptide (AIP) signal is formed by cleavage and processing of the AgrD protein. A characteristic thiolactone ring is formed between a generally conserved central cysteine and the peptide’s C-terminal carboxyl group, and this cyclical structure is generally required for the activity of AIP. Both the cleavage of AgrD and the secretion of AIP are thought to be mediated by the membrane protein AgrB, though other proteins may be involved as well. In comparison of AIP sequence from multiple Staphylococcus species, only the central cysteine and the five-membered thiolactone ring are generally conserved (Novick, 2003; Otto, 2001). he length of the N-terminal

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? AIP=

Ag rB

Figure 8.2 Model of the Staphylococcus aureus Agr system (see text for additional description). The agrD product is processed in part by AgrB into AIP and secreted into the extracellular environment. Recognition of AIP by AgrC triggers transfer of a phosphate group between AgrC and AgrA. AgrA, together with the transcriptional regulator SarA, acts to increase transcription of both the P2 and P3 operons. The transcript of the P3 operon, RNAIII, is the effector molecule of the agr locus and encodes D-toxin (hld) as well.

Peptide Signaling in Bacterial Bioilms

tail varies from two and four amino acids, which results in pheromones with seven to nine residues in all. Lengthening or shortening the tail of an S. epidermidis AIP by a single amino acid residue results in a lack of biological activity in this bacterium, suggesting that AIP length is important for biological activity. Four distinct Agr groups, based on the identity of the AIP produced, have been described in S. aureus (Novick, 2003). AIP produced by one group generally inhibits signaling by staphylococci from a different Agr group by competitive binding to the AgrC receptor. In addition, one AIP produced by S. epidermidis inhibits signaling by three of the four S. aureus Agr groups. Conversely, S. aureus Agr group IV is the only one capable of inhibiting S. epidermidis signaling. In planktonic cultures, the amount of AIP in the medium generally increases in correlation with increasing cell density. Upon reaching sufficient AIP concentration (this has been reported to be during the mid-log phase, though the timing may vary from strain to strain) signaling via the AgrA-AgrC system leads to increased transcription of both the P2 and P3 operons. AgrA and AgrC form the response regulator and histidine kinase receptor, respectively, of a two-component regulatory system that responds to the secreted AIP. AgrA is thought to be constitutively phosphorylated, thus its activation may require dephosphorylation. Binding of AgrA to the agr promoters has not been demonstrated, and other regulatory proteins such as staphylococcal accessory regulator A (SarA) likely are a critical component of Agr autoinduction and response to the AIP. he transcript of the P3 operon, RNAIII, is considered to be the effector of the agr locus in mediating the repression or induction of quorum-controlled genes. Levels and timing of RNAIII transcription vary from strain to strain, and can be correlated with the relative levels of Agr-regulated secreted or surface factors (Li et al., 1997). D-toxin (hld) is also translated from the RNAIII molecule, though disruption of D-toxin translation does not appear to impair the regulatory capabilities of RNAIII. hough the RNAIII nucleotide sequence is not well conserved, the secondary structure is, including several stem-loop motifs. When RNAIII from S. epidermidis, S. simulans, and S. warneri were expressed in S. aureus, they completely repressed expression of the protein A (spa) gene, similar to native S. aureus RNAIII, and they stimulated expression of A-toxin (hla) and serine protease, suggesting conservation of some important regulatory function among species (Tegmark et al., 1998). Much remains unknown of how RNAIII exerts its regulatory effects, though RNAIII is capable of regulation both at the transcriptional and translational levels (Novick, 2003). For instance, the transcription termination loop of RNAIII is necessary for repression of spa transcription, whereas the 3a end of RNAIII is complementary to the translation initiation site of spa mRNA and reportedly blocks its translation. In regulation of hla translation, the 5a region of RNAIII is complementary to the hla mRNA leader sequence. he hla mRNA leader folds into an untranslatable form unless prevented from doing so by RNAIII. Benito et al. (2000) proposed that several structurally different populations of RNAIII might coexist in vivo, and that RNAIII undergoes conformational changes necessary for specific functions. In fact, many of the regulatory effects exerted by RNAIII are likely to be indirect. An analysis of Agr regulation of sed expression suggests that the late log growth phase

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increase in sed transcription occurs via the Agr-mediated reduction in Rot (repressor of toxin) activity rather than a direct effect of Agr (Tseng et al., 2004). his was supported by the observation that the sed promoter is not regulated by the Agr system in a rot mutant background. Agr does not appear to affect rot transcription. However, it has been shown to downregulate Rot activity by an as yet undefined mechanism. One explanation is that Rot might be an RNAIII-binding protein, and activation of agr results in titration of Rot from its gene targets (McNamara et al., 2000). Agr-regulated genes In general, Agr expression in planktonic cultures of S. aureus leads to the increased expression of secreted virulence factors and the decreased expression of several surface-associated adhesins and virulence factors. For most strains this occurs during the transition from latelog growth to early stationary phase, also known as the postexponential phase. A model for how this might affect biofilm development in vivo was presented in the introduction to this chapter (see Figure 8.1). With the exception of capsular polysaccharides, secreted factors upregulated by the Agr system with particular significance for virulence can generally be divided into two classes, exoenzymes and exotoxins (Novick, 2006). Exoenzymes, such as lipase and the proteases, contribute to host tissue degradation, perhaps to create a source of nutrients or to facilitate escape from the localized infection. Also, proteases degrade host proteins important for the immune response, such as the neutrophil defensins, platelet microbicidal proteins, and antibodies, thus providing some protection from host immunity. Proteases may also contribute to the degradation of cell-to-cell or cell-to-host bonds formed by staphylococcal cell surface adhesins. Regulation of lipase production itself may be due to the upregulation by Agr of proteolytic activity, enhancing conversion of the pro-form of the lipase to mature lipase, rather than regulation at the transcriptional level. Fatty-acid modifying enzyme (FAME) is also strongly activated by Agr. FAME can inactivate bactericidal lipids often found in staphylococcal abscesses through esterification of the lipids into alcohols. hese lipids are frequently released from glycerides in the abscess, perhaps by the action of staphylococcal lipase. Accordingly, most S. aureus strains that produce lipase also produce FAME. Exotoxins upregulated by Agr include the hemolysins (A-, B-, D-, and G-toxin), which have general lytic activity against a broad range of host cells, and the pyrogenic toxin superantigens (SAgs), which have broad immunostimulatory activity. A-and D-toxins are both thought to be pore-forming agents strongly regulated by Agr. A-toxin is positively regulated at both the transcriptional and translational levels by RNAIII, whereas D-toxin is directly encoded via RNAIII. A-toxin is a highly potent toxin, killing erythrocytes, mononuclear immune cells, epithelial and endothelial cells. D-toxin is a small, surfactive protein and is active against many types of membranes. B-toxin is likely a sphingomyelinase, causing hydrolysis of sphingomyelin in the membrane outer leaflet patches of erythrocytes and eventual collapse of the lipid bilayer. G-toxin is a member of a family of bi-component toxins of S. aureus in which their pore-forming activity is mediated by two synergistically acting proteins, and is strongly hemolytic but much less leukotoxic. SAgs generally exert their effects through non-antigen specific binding of professional antigen-presenting cells and T-cells. Typical antigens might stimulate one out of 10 000

Peptide Signaling in Bacterial Bioilms

T-cells that specifically recognize that particular antigen; superantigens may stimulate and cause polyclonal proliferation of 20% or more of all circulating T-cells. his results in the release of high levels of cytokines, leading to the symptoms of toxic shock such as vascular dilation, loss of blood pressure, and subsequent organ damage and failure. It is possible that the increased permeability of organs and tissues due to this immune response could also facilitate the spread of staphylococci from one site to another by fluid flow. In general, most of the SAgs are activated by the Agr system, though there are some exceptions. Staphylococcal enterotoxin A (SEA), for instance, is produced throughout growth in an Agr-independent manner. It is not clear what the significance of this Agrindependent expression and the relative contribution of Agr-independent toxins versus Agr-activated toxins in a particular infection type might be. Many of the SAgs are also enterotoxins, exhibiting emetic activity which is separable within the protein from their superantigenic activity, and are responsible for the dominance of S. aureus as a leading cause of food poisoning. here are numerous factors important for colonization and virulence that are downregulated by Agr as well. hese include several surface-associated adhesins such as protein A, fibronectin-binding proteins (fnbA, fnbB), vitronectin-binding protein and coagulase (Novick, 2006). Many of these proteins can also be found in substantial quantities in the growth medium, suggesting that their release from the cell may have importance as well. his release may be due in part to proteolytic activity, particularly at high cell densities. Protein A was the first staphylococcal surface protein to be characterized and is noted for its ability to bind the Fc region of mammalian IgG. By binding IgG, Protein A may interfere with phagocytosis of opsonized bacteria. Protein A can also mediate staphylococcal adherence to von Willebrand factor, a host extracellular matrix protein, suggesting that protein A influences several aspects of the colonization and infectious processes. Two structurally similar proteins, FnbpA and FnbpB, have been shown to mediate S. aureus binding to fibronectin. Fibronectin is a ubiquitous protein found in the extracellular matrix of most tissues, as well as in soluble form in many body fluids, and is necessary for the adhesion of almost all cell types. Fibronectin is one of the host proteins that rapidly coat foreign objects, such as an intravascular catheter, thus facilitating adherence of staphylococci to this de facto biological surface. he fibronectin-binding proteins may also play a role in invasion of host cells by binding soluble fibronectin which is then recognized by integrins on the host cell. his results in phagocytosis of the host protein-coated bacteria. Vitronectin is an adhesive glycoprotein found in circulation at several extracellular matrix sites, particular during tissue or vascular remodeling. Similar to fibronectin- or fibrinogen-binding proteins, vitronectin-binding likely facilitates colonization by staphylococci of host tissues or host-protein coated implanted devices. Coagulase production is the primary criterion used to distinguish S. aureus from other staphylococcal species in a clinical microbiology setting. Coagulase binds soluble fibrinogen and also binds human prothrombin to form a complex which converts soluble fibrinogen to insoluble fibrin. Coagulase is cell-wall associated, though does not have a cell-wall anchoring sequence. he role of coagulase in staphylococcal pathogenesis is not well understood. It could be that fibrin clotting around infection foci protects the bacteria from elements of host immunity.

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A transcriptome analysis of Agr function in S. aureus identified 104 genes induced and 34 genes repressed in an Agr-dependent manner (Dunman et al., 2001). his study supports in general the idea that extracellular virulence factors are activated by Agr and surface adhesins repressed. However, the majority of genes identified as being Agr-regulated were in fact not known virulence factors, but were instead involved in such cellular processes as amino acid metabolism and nutrient transport. Considering this evidence, as well as the identification of Agr homologs in other, less virulent staphylococci, one can speculate that the Agr system may not have evolved originally to facilitate virulence, but perhaps for the coordination of more basic biological functions. Some of these biological functions could well include colonization and dissemination at appropriate cell densities—functions essential to biofilm development. Several aspects of the in vitro model of the Agr regulatory circuit were confirmed in an experimental endocarditis model, although with an intriguing exception (Xiong et al., 2002). As might be expected, maximal RNAII activation in vegetations occurred early, followed by increasing RNAIII expression. his correlated with increased bacterial densities within the vegetations (as compared to lower densities in kidney and spleen tissues), supporting the idea that RNAIII activation in vivo is time and cell-density dependent, and perhaps also tissue-specific. Surprisingly, RNAIII activation was also observed in vegetations formed using Agr signaling mutants (though to a lesser extent than the wild type), suggesting that an RNAII-independent mechanism of RNAIII activation may exist in vivo. Also, there was no correlation between RNAII promoter activity and vegetation densities. he plasma protein fibrinogen is an important component of the acute inflammatory response. It helps to promote neutrophil migration and adhesion, induction of cytokine synthesis, coating of foreign bodies, walling off of infection sites, and initiation of wound healing. As discussed earlier, S. aureus possesses several cell-associated and secreted factors that directly interact with fibrinogen or its soluble precursor, fibrin. In a murine abscess model, transient depletion of the animal of fibrinogen significantly reduced the bacterial burden and overall morbidity and mortality in the animals (Rothfork et al., 2003). his was not observed in infection by an agr mutant. Fibrinogen depletion also inhibited in vivo activation of RNAIII transcription, as well as expression of the quorum-activated virulence factors A-toxin and capsule. he data suggest that fibrinogen-mediated clumping is sufficient to concentrate the autoinducer and promote quorum sensing. he same effects could also mediated by fibronectin. his study provides an important mechanistic link between the innate immune response and pathogenesis of S. aureus, as well as insight into regulation of agr expression in vivo. Peptide signaling in Staphylococcus epidermidis In general, the Agr system in S. epidermidis appears to be highly similar to that in S. aureus (Van Wamel et al., 1998; Vuong et al., 2000a). Agr expression is growth-phase dependent, and with a few exceptions, upregulates exoprotein production while downregulating several surface-associated proteins. In particular, both lipase and protease activity are greatly downregulated in a S. epidermidis agr mutant. Overall, homology of the agr loci between S. aureus and S. epidermidis is 68%. he D-toxin presumably encoded by the S. epidermidis RNAIII molecule differs in three amino acids from that produced by S. aureus, and is upregulated in

Peptide Signaling in Bacterial Bioilms

post-exponential phase, as is RNAIII. D-toxin activity was found in 21 of 23 S. epidermidis strains tested. Agr was also shown to be indirectly involved in production of the lantibiotic epidermin by S. epidermidis via regulation of EpiP, a protease involved in the formation of mature epidermin (Kies et al., 2003). Agr and staphylococcal bioilms here are at least three important stages in staphylococcal biofilm development and behavior, similar to those for many other bacterial species. he first is the initial attachment of cells to a biotic or abiotic surface, usually mediated by surface adhesins. he second, or maturation, stage involves the accumulation of cells into multi-layered clusters enclosed in an at least partly self-produced matrix, or glycocalyx. he third stage of biofilm development involves detachment of cells from the biofilm which may facilitate the colonization of distant sites from the original infection site. he factors contributing to detachment are both external and internal to the biofilm. Physical factors such as shear and physical disruption of the biofilm induce large-scale detachment, while emerging evidence suggests that biofilm-associated bacteria may also actively promote their own detachment (see also Webb, this volume). here are mechanisms whereby Agr expression might impact each of these stages of biofilm development, based on a limited number of in vitro studies. Initial attachment here appear to be two general mechanisms by which staphylococci attach to a surface as illustrated by colonization of an intravascular catheter. During insertion of the catheter, attachment to the naked polymer surface occurs through non-specific, physiochemical interactions, such as hydrophobic interactions. Subsequent to implantation, the catheter surface becomes coated with components of the host matrix, such as fibrinogen, fibronectin, and collagen. his facilitates more specific interactions between the staphylococci and what is now a biological surface mediated by specific receptors on the staphylococci, such as the fibrinogen- and fibronectin-binding proteins. Several of these specific staphylococcal receptors are negatively regulated by Agr. In some staphylococcal species large proteins that might mediate non-specific, hydrophobic interactions with the uncoated polymer surface are also regulated by Agr (e.g. the autolysin AtlE in S. epidermidis). Maturation Little evidence exists for or against a contribution of the Agr system to the maturation of biofilms. he matrix is thought to usually consist of the polysaccharide intercellular adhesion (PIA), and the expression of PIA (encoded by the ica locus) is not regulated by Agr (Vuong et al., 2003). In the host milieu, however, it is not entirely clear whether PIA is in fact required to form a biofilm-like community. hrough intracellular binding mediated by host cell matrix components [e.g. fibrinogen (Rothfork et al., 2003)], a biofilm-like structure could be achieved together with the important characteristics of a biofilm (nutritional gradients, protection from host immune factors and predation, etc.) without the presence of PIA. hus, continued expression of surface adhesins through downregulation of Agr might enhance accumulation in the host of staphylococcal cells in a biofilm.

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Detachment Expression of D-toxin, a protein with surfactant properties and encoded by the agr locus is though to contribute to detachment of cells from a biofilm (Vuong et al., 2000b). hus, in combination with the downregulation of surface adhesins, Agr may well play an important role in facilitating release of staphylococcal cells from the biofilm. Indeed, we have observed enhanced detachment of Agr-expressing cells from a biofilm (Yarwood et al., 2004), but have not been able to confirm the contribution of Agr to this phenomenon. Large-scale detachment events would also be expected to influence mature biofilm structure, at least temporarily, thus potentially influencing the maturation stage of biofilm development as well. At first glance, studies of the role of the Agr system in staphylococcal biofilm formation and behavior appear somewhat inconsistent in their conclusions. A survey of S. aureus strains found a strong correlation between lack of Agr activity (as measured by D-toxin production) and ability to adhere to polystyrene (Vuong et al., 2000b). his was attributed, at least in part, to the surfactant properties of D-toxin, as addition of increasing concentrations of D-toxin decreased attachment of S. aureus to polystyrene. In two studies with somewhat conflicting results agr mutants were first found to demonstrate increased adherence to immobilized fibrinogen, increased induction of platelet aggregation, and had little impact on adherence to immobilized fibronectin, von Willebrand factors, bovine corneal extracellular matrix and endothelial cells (Shenkman et al., 2001). he difference in adherence properties developed primarily under flow conditions, suggesting different adhesion mechanisms under static and flow conditions. In the second study, it was concluded that RNAIII downregulated S. aureus adherence to fibrinogen under static conditions while upregulating S. aureus adherence to fibronectin and endothelial cells under both static and flow conditions (Shenkman et al., 2002). In addition, the contribution of activated platelets to S. aureus adherence to endothelial cells was downregulated by RNAIII, likely due to decreased adherence to fibrinogen, a plasma protein thought to bridge S. aureus, platelets, and endothelial cells. Finally, pleotropic effects were shown of both the agr and sar operons on expression of surface molecules responsible for binding to substrata (Pratten et al., 2001). To address whether the variable results found in the literature were the result of different strains or different growth conditions, biofilms of an isogenic pair (wild type versus agr mutant) were grown under several conditions (Yarwood et al., 2004). In this study, the contribution of Agr to biofilm development was found to be dependent on growth conditions. In some cases, Agr expression decreased bacterial attachment and biofilm formation. Under other conditions, it enhanced biofilm formation or, in the case of flow-cell biofilms, appeared to have little effect on biofilm structure at discrete time points even when clearly expressed. However, in time-course studies Agr expression did often precede cell detachment. Given these results and those of other studies, it is clear that the Agr contribution to biofilm formation is heavily dependent on growth conditions (medium, shear, temperature, etc.), surface character (e.g. biological or abiotic), and strains used. Going forward, the most useful results will likely come from either in vitro studies that closely mimic the in vivo environment, or from animal models of staphylococcal biofilm infection. In the first study of its kind to address directly the biofilm-forming capabilities of agr mutants in vivo, Vuong et al. (2004) found that a S. epidermidis agr mutant shows increased

Peptide Signaling in Bacterial Bioilms

binding to epithelial cells and a higher colonization rate in a rabbit model of an indwelling medical device-related infection. hey also confirmed that deletion of agr or inhibition of Agr activity leads to thicker biofilms in vitro. hese results were consistent with a study conducted earlier by the same laboratory group in which a S. epidermidis agr mutant showed increased primary attachment and biofilm formation, as well as expression of the cell surface-associated autolysin AtlE (Vuong et al., 2003). (Repetitive sequences in AtlE are thought to interact hydrophobically with abiotic surfaces.) Like S. aureus, production of PIA by the S. epidermidis agr mutant was similar to the wild type. As expected, the agr mutant lacked D-hemolysin production. Addition of increasing concentrations of D-toxin resulted in decreased attachment of S. epidermidis cells to polystyrene, where 10 Mg/ml D-toxin was sufficient to reduce biofilm formation of the agr mutant strain to the same levels found using the agr wild-type strain. Interestingly, there may also be some role for Agr expression in the resistance of staphylococcal biofilms to antibiotic exposure. Under conditions where an agr mutant formed a smaller biofilm than its wild-type parent, the mutant was also more sensitive to rifampicin treatment, but not oxacillin (Yarwood et al., 2004). he basis for this variation in sensitivity is unknown, though there is precedent for the regulation of other antibiotic resistance mechanisms by Agr. Regulation of NorA, a multi-drug efflux pump involved in resistance to quinolones, by the DNA-binding protein NorR was found to require an intact Agr system (Truong-Bolduc et al., 2003). Agr variants Agr variants (cells either in which expression of Agr is significantly higher or lower as compared to the parental strain) have been frequently isolated from cultures in vitro, suggesting that staphylococci maintain some capacity to alter their Agr phenotype or maintain Agr-negative subpopulations. Somerville et al. (2002) found that repeated passage of S. aureus in vitro resulted in the loss of Agr function in a large percentage of the population, along with corresponding hemolytic and aconitase activity. he authors hypothesized that frequent mutations of agr create a mixed population of bacteria, with some cells expressing colonization factors, while others would tend to express secreted exotoxins. Under a particular environment with specific ecological and/or immunological selection, the Agr variant best able to adapt would emerge. Agr mutants are frequently found among clinical isolates. One study (Vuong et al., 2004) showed that the percentage of strains with defective quorum-sensing systems was significantly higher among isolates from patients with infections of joint prostheses than among isolates from the skin of healthy controls (36% versus 5%, respectively). Another study found that 26% of S. aureus isolates failed to produce D-toxin, indicating that they were deficient in quorum-sensing-mediated regulation (Vuong et al., 2000b). When staphylococci were isolated from the lungs of cystic fibrosis patients, not only did the strains generally express low levels of RNAIII, but several isolates were also found to be Agr-negative (Goerke et al., 2000). Fowler et al. (2004) showed that the percentage of S. aureus isolates recovered from patients with persistent bacteremia with defective D-toxin production (a consistent indicator of Agr activity) was higher than in isolates from patients with resolving bacteremia

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(71% versus 39%, respectively). he authors postulated that lack of Agr expression might contribute to persistent bacteremia through the increased expression of the S. aureus surface adhesion gene, fnbA, in these mutants. he fibronectin-binding protein encoded by fnbA has been shown to enhance S. aureus adhesion to, invasion of, and persistence within endothelial cells. Intracellular invasion may contribute to resistance to antibiotics, as vancomycin penetrates poorly into endothelial cells. hus, lack of Agr expression may facilitate a protected intracellular reservoir for S. aureus. Indeed, an agr mutant is incapable of escape from the endosome (Shompole et al., 2003) or inducing apoptosis (Wesson et al., 1998), suggesting a prominent role for Agr in invasion of and persistence in host cells. One area of particular concern in staphylococcal pathogenesis is the emergence of staphylococci with intermediate resistance to glycopeptide antibiotics (GISA). Interestingly, GISA are frequently isolated from biomedical device-related infection, which are also likely to be biofilm-associated, and these same GISA have been shown to be predominantly Agrnegative (Sakoulas et al., 2002). he same study also suggested that loss of Agr function might in fact contribute to the development of vancomycin tolerance, an intriguing idea yet to be confirmed. Behavior of mixed populations of hyper-hemolytic, hemolytic, and non-hemolytic variants was examined in a murine abscess model of infection (Schwan et al., 2003). he percentage of non-hemolytic variants, likely representing Agr-negative bacteria, recovered from the wound increased over time, whereas the number of hyper-hemolytic variants (Agr overexpressors) decreased dramatically over the same time period. A wound infection model demonstrated the same trend, though to a lesser degree. In contrast, hemolytic variants seemed to be favored in isolates recovered from murine livers and spleens in a model of systemic infection. hus, Agr activity likely facilitates survival and pathogenesis in some host environments, but not others. Additional studies will be of great importance in monitoring Agr phenotypes of clinical isolates from various infection types (preferably multiple isolates from each patient) and determining any correlation to disease progression and outcome. It can be hypothesized that Agr-negative variants are better suited to biofilm formation and long-term, chronic infection as they tend to (1) express the surface adhesins that mediate cell-to-cell and cell-to-surface interactions, while downregulating factors that may facilitate detachment, such as D-toxin, and (2) express more immuno-evasive factors, such as protein A, than immuno-stimulatory ones (such as the superantigens). In addressing the first idea, our laboratory has found that Agr-negative variants become the predominant form in biofilms grown in a serum-based medium (Yarwood, 2004; Yarwood and Greenberg, 2006). It is not yet clear whether this is due to a selective pressure against Agr-positive cells, increased generation of Agr variants in the biofilm, active detachment of cells expressing Agr, or some combination of all three. Also, the results indicate that the Agr-positive population is not completely lost from the biofilm ( J.M. Yarwood, unpublished data), suggesting a mechanism to retain the capability to express invasive factors at an appropriate stage of infection. Indeed, we have detected the frequent detachment of cells expressing Agr from the biofilm (Yarwood et al., 2004). his may have important clinical implications, as detaching cells expressing Agr are also likely to be expressing extracellular virulence factors important in causing acute infection. he frequency at which variants arise also appears

Peptide Signaling in Bacterial Bioilms

to vary from strain to strain, and with cell density ( J.M. Yarwood, unpublished data), and much work remains to be done to understand the mechanisms of variant generation and the importance of these functional Agr variants in the disease process. One potential model of S. aureus Agr evolution in the context of a chronically infected host was presented previously (Yarwood, 2006) and is summarized here. Upon establishment of infection, mutations (often point mutations) accumulate in the agr loci of S. aureus cells. hese mutations result in the conversion of a significant part of the population to a quorum-sensing negative phenotype. he Agr-negative phenotype confers some protection to the staphylococcal population as a whole due to increased expression of immuno-evasive factors and facilitates attachment and accumulation through increased expression of host protein-binding factors. his protected environment is conducive to the continued growth of staphylococci and additional accumulation of mutations in the agr locus. On very rare occasions, appropriate mutations are acquired by the Agr-negative variant to return functionality to the agr locus, such as alteration of the AgrC receptor to recognize an AIP variant. hese new Agr specificity group cells detach from the biofilm through expression of invasive virulence factors or production of D-toxin and establish infection elsewhere in the host or, alternatively, colonize a secondary host. In some cases, appropriate ecological pressures are present to allow emergence of this new Agr specificity group from among the established groups. he combined rarity of these events—accumulation of several, eventually positive mutations, and selection for any emergent Agr specificity group would only give rise to a major new S. aureus Agr group very infrequently. his would be consistent with the identification of only four distinct S. aureus Agr groups thus far, despite frequent mutation of the agr locus. he driving force behind this cycle is, in part, the advantage conferred by maintaining a mixed population of cells, where Agr-negative variants prevent recognition by immune surveillance, and cells expressing Agr provide additional nutrient sources through host tissue degradation or facilitate escape from the localized infection at appropriate times. hus, the emergence of distinct Agr groups may be a byproduct of this mode of Agr evolution in which the generation of variants is itself important. However, it is noteworthy that this Agr-negative phenotype is often generated through non-reversible mutation of the agr locus, rather than a reversible, conditional switching of Agr expression on and off. his is consistent with some evolutionary or pathological advantage for generation of distinct Agr specificity groups. RIP/RAP With some degree of controversy, a second quorum-sensing system has been described in S. aureus that is proposed to regulate Agr activity (see references Balaban et al., 1998; Dell’Acqua et al., 2004; Novick, 2003 and other studies by N. Balaban and colleagues). his system consists of the auto-inducer RNAIII activating protein (RAP) and its target molecule TRAP. RAP is described as an ortholog of the ribosomal protein L2 that is synthesized early in growth. Reportedly, when RAP reaches a threshold concentration, it induces the histidine phosphorylation of the membrane protein TRAP. his event leads to the upregulation of agr transcription through an undescribed mechanism. Once AIP is made, it has been reported to lead to the downregulation of TRAP phosphorylation. Immunization against RAP was shown to mitigate pathology in a murine cutaneous S.

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aureus infection model. A protein produced by S. xylosus and resembling in the N-terminal sequence of RAP, RNAIII inhibiting peptide (RIP), has been reported act as an agonist of TRAP, inhibiting its phosphorylation and, consequently, agr expression. Treatment with synthetic RIP inhibits several types of S. aureus and S. epidermidis infections, including those that are biofilm-related or caused by multiple drug-resistance staphylococci. Apparently, these therapeutic effects can be observed when RIP is applied locally or systemically. he RAP/TRAP/RIP story thus far is somewhat unsatisfying. Little has been described regarding the properties of RIP, and even its amino acid sequence and structure remain questionable. It is also conceivable that when used at high concentrations, RIP has sufficient amphipathic, or surfactant, properties to prevent bacterial attachment, not unlike many other proteins, including staphylococcal D-toxin. Most of the in vivo experiments have administered RIP before or coincident with bacterial challenge and it is clear that, in these cases, RIP inhibits bacterial adherence to surfaces. However, there is no evidence that RIP has any effect against established biofilm, and little data is available as to the overall effect of RIP on bacterial physiology or virulence. It is also not clear whether a RIP-impregnated intravascular catheter would continue to inhibit staphylococcal adhesion against the more or less continual “challenge” that likely occurs in vivo—either from the epithelium or transient bacteremias. Implanted devices are soon coated with host matrix proteins, a fact that limits the efficacy of many device surface treatments as the now biotic surface provides several protein receptor specific targets for staphylococci to bind to. Furthermore, very little is known as to the regulatory targets of this system. Besides being reported to inhibit Agr activation, no other gene targets have been conclusively identified. his is particularly unsatisfying, as inhibition of Agr activity would be expected to increase bacterial adhesion, yet, in fact, the RIP-treated cells are less likely to adhere, and suggests that RIP may in fact simply be acting as a surfactant-like molecule. Finally, other laboratory groups have been unable to detect the RNAIII-activating activity in supernatants of agr-null strains, despite the presumed presence of RAP, contributing to the controversy as to the true nature of this molecule (Novick, 2003; Novick et al., 2000). Peptide signaling and Streptococcus bioilms Streptococci are the causative agents of numerous diseases, from indigenous microflora that cause dental caries to exogenous pathogens that are the etiologic agents of both relatively benign infections, such as impetigo, and potentially fatal diseases, including necrotizing fasciitis. First described in S. pneumoniae in its relationship to natural transformation, peptide signaling has also been shown to be involved in streptococcal virulence, biofilm formation, acid tolerance and bacteriocin production (Cvitkovitch et al., 2003; Suntharalingam and Cvitkovitch, 2005). Highly homologous competence-stimulating peptide (CSP) quorum sensing systems have been identified in several streptococcal species, including strains in the mitis (including S. pneumoniae), anginosus (including S. intermedius), and mutans (including S. mutans) groups. Streptococcal quorum sensing, particularly in its relationship to competence induction and transformation, has been most extensively studied in S. pneumoniae (Figure 8.3). he CSP precursor is encoded by comC. Two secretory proteins, ComA, an ATP-binding cassette transporter, and ComB, described as an accessory protein to ComA, are involved in

Peptide Signaling in Bacterial Bioilms

ComE ComD

P comX

P ComE

comA

comC

comD

comB

comE

ComB ComA

CSP =

Figure 8.3 Model of the Streptococcus pneumoniae competence-stimulating peptide (CSP) system (see text for additional description). The comC product is processed by the ComA/ ComB complex and secreted as CSP into the extracellular environment. Recognition of CSP by ComD triggers transfer of a phosphate group between ComD and ComE. ComE acts to increase transcription of both the operons encoding the CSP system itself (comAB and comCDE) as well as comX, encoding the alternative sigma factor ComX. ComX activates the transcription of late competence genes, including those involved in DNA uptake and integration into the host cell genome.

the processing and export of a 17-amino acid CSP. Interaction of the CSP with its histidine kinase receptor, ComD, initiates a series of temporally distinct transcription profiles via the response regulator ComE. Among the earliest operons to be induced are those encoding the cell-to-cell signaling system itself, comAB and comCDE, and an alternative sigma factor, ComX. ComX is integral to later stages of competence development, inducing genes involved in DNA uptake and integration. Many of the S. pneumoniae operons regulated by ComX have a com-box consensus sequence (5a-TACGAATA-3a) in their promoter regions recognized by the sigma factor. he presence of com-boxes in S. mutans late-competence genes suggest ComX behaves similarly in this organism. ComX also regulates a mechanism whereby a small fraction of the bacterial population initiates lysis and release of donor DNA. In vitro, accumulation of the CSP in the growth medium triggers a short (20–40 minutes) window of competence throughout the culture. In vivo, the ability to take up DNA through natural transformation may enable the streptococci to acquire novel genes, such as antibiotic-resistance cassettes and virulence factors, that provide a selective advantage over

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neighboring bacteria competing for the same resources. Also of note, recent transcriptional analyses during S. pneumoniae competence development showed that only 23 of 124 CSPinducible genes were required for transformation, at least in laboratory conditions. It remains to be seen what role many of the remaining gene products might have in cell-density dependent responses, such as nutrient adaptation, biofilm formation, or stress responses. he mechanics and effects of the CSP quorum sensing systems vary somewhat among the streptococci. While some streptococcal species (particularly the anginosus group) encode and respond to identical CSPs, CSPs are often species, and sometimes strain, specific (streptococci responding to the same CSP signal are identified as a “pherotype”). In many species the genes encoding the CSP, histidine kinase receptor and the cognate response regulator are organized as a single operon. In S. mutans, however, comC is divergently transcribed from the comDE operon. Competence can be completely abolished in S. pneumoniae by inactivation of the ComDE two-component system; in S. mutans, inactivation of this system only reduces competence. In S. pneumoniae, competence development affects most of the cell culture; in S. mutans, competence development affects only a small number of cells. While many species of streptococci have been found to form biofilms, the relationship between biofilm formation, peptide signaling, and pathogenesis has been best studied in the oral streptococci (Cvitkovitch et al., 2003; Suntharalingam and Cvitkovitch, 2005). he oral cavity is one of the most microbiologically diverse environments on earth, with as many as 500 species residing in the human mouth. Dental plaque is also one of the more complex representations of bacterial biofilms, with multiple species colonizing the tooth surface in multiple stages. Furthermore, the oral cavity is a particularly stressful environment for bacteria, with wide swings in nutrient availability, low pH, high osmolarity, and presence of host-produced enzymes and antimicrobials. hus, biofilm formation likely provides oral bacteria a protected environment, as well as facilitating genetic exchange. he first indication that a CSP quorum sensing system was involved in streptococcal biofilm formation came when transposon mutants of comD in the oral bacterium S. gordonii were shown to be defective in biofilm formation (Loo et al., 2000). A subsequent study in S. mutans found that inactivation of any component of the ComCDE pathway in results in abnormal biofilms (Li et al., 2002). Biofilms of the comC mutant lack the wild-type architecture (which can be restored by addition of synthetic CSP), whereas biofilms of the comD and comE mutants have reduced biomass. In S. intermedius, biofilm formation is enhanced in the presence of CSP without affecting the organism’s growth rate (Petersen et al., 2004). While little is known as to the mechanisms by which the CSP systems contribute to streptococcal biofilm development, initial evidence indicates that CSP influences the early stages of biofilm formation rather than later maturation steps, at least for S. mutans and S. intermedius. S. mutans comD or comE mutants adhere less to surfaces, while S. intermedius CSP appears to enhance the early buildup of cells in a biofilm. Of particular interest is the observation that S. mutans and S. intermedius biofilm cells are much more efficient in incorporating foreign DNA than corresponding planktonic cells (Li et al., 2001b). Furthermore, the CSP quorum sensing system appears to be transcriptionally upregulated in biofilm-associated S. gordonii and S. mutans. It seems likely that the high cell density of the biofilm likely enhances transformation efficiency through both

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increased CSP cell-to-cell signaling and the presence of relatively high concentrations of extracellular genetic material. It may well be that the heterogeneous biofilm environment also provides gradients of CSP signal and other growth conditions that sustain localized clusters of cells with a competence window significantly longer than the 20–40 minutes typically seen in planktonic cultures (Suntharalingam and Cvitkovitch, 2005). Indeed, observations via confocal microscopy of a comX-gfp reporter in a S. mutans biofilm demonstrated spatial heterogeneity; it appeared that cells in denser areas of the biofilm had increased comX activity and likely were genetically competent (Aspiras et al., 2004). Peptide signaling and the acid-tolerance response One of the hallmarks of tooth decay (dental caries) associated with colonization by S. mutans is the production of acid from fermentable dietary carbohydrates that leads to demineralization of the tooth surface. In the dental biofilm, S. mutans encounters pH shifts from above 7 to nearly 3 during ingestion of these carbohydrates. hus, S. mutans’ tolerance to low pH is critical to its survival and pathogenicity. he acid tolerance response (ATR) by S. mutans requires de novo synthesis of proteins apparently required for adaptation to an acidic environment and is pH-inducible—exposure of S. mutans to mildly acidic pH (5–6) results in enhanced survival at lower pH values (3.0–3.5). In many bacteria, the ATR is also growth phase- and time-dependent, leading to speculation that it might be quorum-regulated in S. mutans. Indeed, mutations in the comC, comD, or comE genes result in a diminished log-phase ATR in S. mutans, whereas addition of synthetic CSP to a comC mutant restores the ATR (Li et al., 2001a). Correspondingly, cell density enhances the ATR—planktonic S. mutans taken from a high cell density culture or from a biofilm are more resistant to a killing pH (3.5) than planktonic cells taken from a lower cell density culture. hus, both low pH induction and cell-to-cell communication (including, but not limited to, CSP) appear important for optimal development of acid adaptation. Peptide signaling and bacteriocin production Bacteriocins are antimicrobial peptides produced by many bacteria, including streptococci, presumably to enhance their ecological fitness by controlling competing populations of bacteria. In planktonic culture, S. pneumoniae releases chromosomal DNA after addition of CSP, a process that is ComDE-dependent (Moscoso and Claverys, 2004). Concentrations of released DNA are highest in stationary phase, coinciding with maximum bacteriocin production. hus, it has been hypothesized that bacteriocins might aid in release of DNA from surrounding organisms through permeabilization of their cytoplasmic membranes. his liberated DNA might contribute to the biofilm extracellular matrix, serve as a nutrient source, and enhance overall DNA uptake and recombination. Some evidence is emerging to support these hypotheses. Expression of nlmAB, genes which encode the two-peptide non-lantibiotic bacteriocin mutacin IV in S. mutans, is highest at high cell density and is abolished with disruption of the comDE genes (van der Ploeg, 2005). Kreth et al. (2005) also found that CSP induced coordinated expression of competence and mutacin IV in S. mutans. Furthermore, in mixed cultures, plasmid transfer from S. mutans to S. gordonii, which is sensitive to mutacin IV, was CSP and mutacin-dependent. he authors proposed that the coordinated expression of bacteriocins and competence may

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exist to effectively acquire DNA from other species living in the same ecological niche. his is consistent with the extensive genomic diversity among S. mutans strains, which may well have resulted in part from horizontal gene transfer. Peptide signaling and Enterococcus bioilms he Gram-positive commensal bacterium Enterococcus faecalis, a normal member of the human intestinal microflora, has emerged as a leading cause of nosocomial infections. Diseases caused by this organism range from endocarditis to urinary tract and dental infections. A majority of clinical isolates form biofilms in vitro (Mohamed et al., 2004), though biofilm formation per se has yet to be demonstrated as essential for enterococcal virulence. E. faecalis biofilms have been observed on dental root canals (Distel et al., 2002), urethral catheters (Tunney and Gorman, 2002), and heart valves (Donlan and Costerton, 2002). It is also worth nothing that, while an E. faecalis strain may not form a biofilm in vitro in the classical sense (i.e. surrounded by a self-produced polymeric matrix), the vegetations that the organism forms in vivo may functionally and physiologically mimic a classic biofilm. hese vegetations are thought to be an aggregation, in part, of fibrin and platelets and contain very high cell densities (McCormick et al., 2001; McCormick et al., 2002). he vegetation provides an immunologically protected environment for the bacteria similar to what would be expected of an entirely self-produced biofilm (McCormick et al., 2002). In a survey of the 17 two-component regulatory systems identified in the genome of E. faecalis, only one system was identified as affecting biofilm formation (Hancock and Perego, 2004a; Hancock and Perego, 2004b). his system, known as Enterococcus faecalis regulator (encoded by the fsr locus; Figure 8.4), is similar in many respects to the staphylococcal Agr system. fsrA, fsrB, and fsrC are homologous to the staphylococcal agrA, agrB, and agrC genes, respectively. Furthermore, he 3a-end of fsrB is homologous to agrD, the gene responsible for encoding the staphylococcal auto-inducing peptide. Indeed, a 11-residue cyclic peptide was identified in culture supernatants of E. faecalis that was able to induce early gelatinase production (Nakayama et al., 2001). his pheromone corresponds to a Cterminal portion of FsrB. he Fsr system is autoregulated (expression of fsrB and fsrC is cell density dependent), and regulates two downstream genes, gelE (encoding gelatinase, a secreted thermolysin-like M4 protease) and sprE (encoding a secreted serine protease) in a cell-density dependent manner (Qin et al., 2001). he reported percentage of E. faecalis isolates carrying the fsr locus varies widely depending on patient group from 24% to 100% ( Jones and Deshpande, 2003; Pillai et al., 2002; Roberts et al., 2004). Some studies have found that isolates from patients with endocarditis (Pillai et al., 2002) and from ICU patients with urinary tract infections ( Jones and Deshpande, 2003) are enriched for the fsr locus. While one study contradicted these results by finding similar levels of the fsr locus among isolates from diseased and healthy patients (Roberts et al., 2004), it did not address whether presence of the fsr locus or gelatinase affects the severity of disease, as they do in animal models of infection (Engelbert et al., 2004; Sifri et al., 2002; Singh et al., 1998). he Fsr system appears to regulate E. faecalis biofilm formation, at least in part, through control of gelE expression (Hancock and Perego, 2003; Hancock and Perego, 2004a). Disruption of both the fsr and gelE genes attenuates E. faecalis biofilm development. Expression of the gelatinase gene in trans restores a biofilm positive phenotype,

Peptide Signaling in Bacterial Bioilms

FsrA

P FsrC FsrA PA

fsrA

PB

fsrB

P fsrC

PE

gelE

sprE

? GBAP=

Figure 8.4 Model of the Enterococcus faecalis Fsr system (see text for additional description). The GBAP pheromone is liberated from the C-terminal portion of FsrB and secreted by an as yet undeined mechanism into the extracellular environment. Recognition of GBAP by FsrC leads to the phosphorylation of FsrA, which acts to upregulate the fsrBC operon as well as the adjacent gelE and sprE genes. “P” indicates distinct promoter regions.

even in the absence of a functional Fsr system. Observations by Pillai et al. (2004) also indicated that Fsr exerted its effects on biofilm control via the downstream protease genes (they did not isolate GelE activity from that of SprE). Interestingly, the same study found that glucose-mediated augmentation of biofilm occurred in wild-type E. faecalis, but not in either a fsrA mutant or a protease mutant. his indicated that catabolite control of biofilm formation occurs via the Fsr system. Catabolites, such as glucose, have dramatic effects on biofilm production by several bacterial species, yet the mechanisms by which these effects are exerted are poorly understood. he mechanism by which gelatinase might exert its effect on enterococcal biofilm development is still unclear, nor is it well established what role gelatinase plays in virulence overall (Carniol and Gilmore, 2004). Mohamed et al. (2004) found that disruption of the fsr locus resulted in less E. faecalis biofilm formation, and proposed that the effect was due to decreased primary attachment. Since gelatinase, a secreted protease, tends to cleave its substrates at hydrophobic residues, it is possible that gelatinase activity increases the hydrophobicity of the cell surface, thus promoting non-specific interaction between many surfaces and the cell (Carniol and Gilmore, 2004; Hancock and Perego, 2004a). It has also been proposed that expression of gelatinase, induced in late stages of growth by cell-density dependent mechanisms, enables the dissemination of the organism in vivo from vegetations (Waters et al., 2003). Gelatinase appears capable of degrading polymerized fibrin, which likely coats the vegetation, as well as a broad range of other substrates (Carniol and

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Gilmore, 2004). Gelatinase could aid in bacterial dissemination by cleaving bacterial attachment proteins and host tissue proteins. he Fsr-controlled serine protease SprE may also be involved in this step, even though it doesn’t appear to be involved in biofilm formation (Hancock and Perego, 2004a). his would be consistent with the model presented by Rasmussen and Bjorck (2002) who argued that initial stages of streptococcal infection and colonization are characterized by low levels of protease activity, whereas higher levels of protease activity during later stages of infection, when bacterial density is high, facilitate detachment and spreading of the bacteria. In the end, many questions remain to be answered regarding the mechanisms by which enterococcal biofilms form, and how peptide signaling contributes to that process in vivo. While many endocarditis-derived isolates do not produce gelatinase, a majority of endocarditis-derived isolates do form biofilms in vitro, and recent studies have failed to find a correlation between gelatinase activity and biofilm formation in E. faecalis (Mohamed and Murray, 2005; Rosa et al., 2006). It may well be that Fsr exerts its effects on biofilm formation both through gelatinase production as well as other mechanisms, particularly in vivo, depending on environmental conditions. Indeed, fsr locus mutants were found to be more attenuated in a rabbit endophthalmitis than the protease mutants, suggesting additional pleotropic effects by Fsr disruption (Engelbert et al., 2004). Disruption of gelE does attenuate virulence in several animal models of infection (Carniol and Gilmore, 2004). Inhibition of peptide signaling as a therapeutic tool Given the role that peptide signaling plays in biofilm control and in virulence factor regulation in several Gram-positive species, the control of peptide signaling by artificial means remains an area of significant interest for inhibiting pathogenesis. However, due to the varied nature of the signals, the complex responses to those signals, and the potential tradeoffs for manipulating the quorum response, the practical use of signaling inhibitors remains a challenging proposition. For instance, inhibition of quorum sensing has been proposed as one mechanism for controlling staphylococcal infections ( Ji et al., 1997). In a skin abscess model co-administration of the synthetic Agr group II AIP together with the bacterial inoculation significantly attenuated an infection caused by an Agr group I strain (Mayville et al., 1999). However, use of cross-inhibiting pheromones mimics agr mutations in both S. aureus and S. epidermidis and enhances biofilm formation (Vuong et al., 2003; Vuong et al., 2004; Vuong et al., 2000b). his result calls for extreme caution in the use of signaling inhibitors, as it is conceivable that such treatments, while mitigating the acute phase of infections, might facilitate chronic, biofilm-associated infections, at least by staphylococci. Use of quorum sensing inhibitors has also been proposed to as a method for preventing CSP-mediated biofilm formation in streptococci (Suntharalingam and Cvitkovitch, 2005). Since early evidence suggests quorum sensing appears to positively regulate attachment by streptococci, downregulating CSP activity might inhibit biofilm formation without the additional risk of upregulating other virulence factors, particularly in the oral streptococci. Indeed, synthetic CSPs have been used to study and manipulate streptococcal quorum sensing already. Interestingly, addition of CSP beyond levels required for induction of competence inhibits the growth of S. mutans, and at higher levels, even lead to cell death (Qi et al., 2005).

Peptide Signaling in Bacterial Bioilms

Whether these peptide signaling systems can be effectively manipulated for therapeutic purposes is an open question. Regardless of the answer, the role of peptide signaling in biofilm formation remains an intriguing area of study, and promises to greatly enhance our understanding of the physiology and pathogenesis of many important human pathogens. References Aspiras, M.B., Ellen, R.P., and Cvitkovitch, D.G. (2004). ComX activity of Streptococcus mutans growing in biofilms. FEMS Microbiol. Lett. 238, 167–174. Balaban, N., Goldkorn, T., Nhan, R.T., Dang, L.B., Scott, S., Ridgley, R.M., Rasooly, A., Wright, S.C., Larrick, J.W., Rasooly, R., and Carlson, J.R. (1998). Autoinducer of virulence as a target for vaccine and therapy against Staphylococcus aureus. Science 280, 438–440. Benito, Y., Kolb, F.A., Romby, P., Lina, G., Etienne, J., and Vandenesch, F. (2000). Probing the structure of RNAIII, the Staphylococcus aureus agr regulatory RNA, and identification of the RNA domain involved in repression of protein A expression. RNA 6, 668–679. Carniol, K., and Gilmore, M.S. (2004). Signal transduction, quorum-sensing, and extracellular protease activity in Enterococcus faecalis biofilm formation. J. Bacteriol. 186, 8161–8163. Cvitkovitch, D.G., Li, Y.H., and Ellen, R.P. (2003). Quorum sensing and biofilm formation in streptococcal infections. J. Clin. Invest. 112, 1626–1632. Dell’Acqua, G., Giacometti, A., Cirioni, O., Ghiselli, R., Saba, V., Scalise, G., Gov, Y., and Balaban, N. (2004). Suppression of drug-resistant staphylococcal infections by the quorum-sensing inhibitor RNAIII-inhibiting peptide. J. Infect. Dis. 190, 318–320. Distel, J.W., Hatton, J.F., and Gillespie, M.J. (2002). Biofilm formation in medicated root canals. J. Endod. 28, 689–693. Donlan, R.M., and Costerton, J.W. (2002). Biofilms: survival mechanisms of clinically relevant microorganisms. Clin. Microbiol. Rev. 15, 167–193. Dunman, P.M., Murphy, E., Haney, S., Palacios, D., Tucker-Kellogg, G., Wu, S., Brown, E.L., Zagursky, R.J., Shlaes, D., and Projan, S.J. (2001). Transcription profiling-based identification of Staphylococcus aureus genes regulated by the agr and/or sarA loci. J. Bacteriol. 183, 7341–7353. Engelbert, M., Mylonakis, E., Ausubel, F.M., Calderwood, S.B., and Gilmore, M.S. (2004). Contribution of gelatinase, serine protease, and fsr to the pathogenesis of Enterococcus faecalis endophthalmitis. Infect. Immun. 72, 3628–3633. Fowler, V.G., Jr., Sakoulas, G., McIntyre, L.M., Meka, V.G., Arbeit, R.D., Cabell, C.H., Stryjewski, M.E., Eliopoulos, G.M., Reller, L.B., Corey, G.R., et al. (2004). Persistent bacteremia due to methicillinresistant Staphylococcus aureus infection is associated with agr dysfunction and low-level in vitro resistance to thrombin-induced platelet microbicidal protein. J. Infect. Dis. 190, 1140–1149. Goerke, C., Campana, S., Bayer, M.G., Doring, G., Botzenhart, K., and Wolz, C. (2000). Direct quantitative transcript analysis of the agr regulon of Staphylococcus aureus during human infection in comparison to the expression profile in vitro. Infect. Immun. 68, 1304–1311. Hancock, L., and Perego, M. (2003). he fsr signal transduction system of Enterococcus faecalis controls biofilm development through the production of gelatinase. Paper presented at: Functional Genomics of Gram-Positive Microorganisms, 12th International Conference on Bacilli (Baveno, Italy). Hancock, L.E., and Perego, M. (2004a). he Enterococcus faecalis fsr two-component system controls biofilm development through production of gelatinase. J. Bacteriol. 186, 5629–5639. Hancock, L.E., and Perego, M. (2004b). Systematic inactivation and phenotypic characterization of twocomponent signal transduction systems of Enterococcus faecalis V583. J. Bacteriol. 186, 7951–7958. Ji, G., Beavis, R., and Novick, R.P. (1997). Bacterial interference caused by autoinducing peptide variants. Science 276, 2027–2030. Jones, R.N., and Deshpande, L.M. (2003). Distribution of fsr among Enterococcus faecalis isolates from the SENTRY antimicrobial surveillance program. J. Clin. Microbiol. 41, 4004–4005. Kies, S., Vuong, C., Hille, M., Peschel, A., Meyer, C., Gotz, F., and Otto, M. (2003). Control of antimicrobial peptide synthesis by the agr quorum sensing system in Staphylococcus epidermidis: activity of the lantibiotic epidermin is regulated at the level of precursor peptide processing. Peptides 24, 329–338. Kong, K.-F., Vuong, C., and Otto, M. (2006). Staphylococcus quorum sensing in biofilm formation and infection. Int. J. Med. Microbiol. 296, 133–139.

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Kreth, J., Merritt, J., Shi, W., and Qi, F. (2005). Co-ordinated bacteriocin production and competence development: a possible mechanism for taking up DNA from neighbouring species. Mol. Microbiol. 57, 392–404. Li, S., Arvidson, S., and Mollby, R. (1997). Variation in the agr-dependent expression of alpha-toxin and protein A among clinical isolates of Staphylococcus aureus from patients with septicaemia. FEMS Microbiol. Lett. 152, 155–161. Li, Y.H., Hanna, M.N., Svensater, G., Ellen, R.P., and Cvitkovitch, D.G. (2001a). Cell density modulates acid adaptation in Streptococcus mutans: implications for survival in biofilms. J. Bacteriol. 183, 6875–6884. Li, Y.H., Lau, P.C., Lee, J.H., Ellen, R.P., and Cvitkovitch, D.G. (2001b). Natural genetic transformation of Streptococcus mutans growing in biofilms. J. Bacteriol. 183, 897–908. Li, Y.H., Tang, N., Aspiras, M.B., Lau, P.C.Y., Lee, J.H., Ellen, R.P., and Cvitkovitch, D.G. (2002). A quorum-sensing signaling system essential for genetic competence in Streptococcus mutans is involved in biofilm formation. J. Bacteriol. 184, 2699–2708. Loo, C.Y., Corliss, D.A., and Ganeshkumar, N. (2000). Streptococcus gordonii biofilm formation: identification of genes that code for biofilm phenotypes. J. Bacteriol. 182, 1374–1382. Mayville, P., Ji, G., Beavis, R., Yang, H., Goger, M., Novick, R.P., and Muir, T.W. (1999). Structure-activity analysis of synthetic autoinducing thiolactone peptides from Staphylococcus aureus responsible for virulence. Proc. Natl. Acad. Sci. USA 96, 1218–1223. McCormick, J.K., Hirt, H., Waters, C.M., Tripp, T.J., Dunny, G.M., and Schlievert, P.M. (2001). Antibodies to a surface-exposed, N-terminal domain of aggregation substance are not protective in the rabbit model of Enterococcus faecalis infective endocarditis. Infect. Immun. 69, 3305–3314. McCormick, J.K., Tripp, T.J., Dunny, G.M., and Schlievert, P.M. (2002). Formation of vegetations during infective endocarditis excludes binding of bacterial-specific host antibodies to Enterococcus faecalis. J. Infect. Dis. 185, 994–997. McNamara, P.J., Milligan-Monroe, K.C., Khalili, S., and Proctor, R.A. (2000). Identification, cloning, and initial characterization of rot, a locus encoding a regulator of virulence factor expression in Staphylococcus aureus. J. Bacteriol. 182, 3197–3203. Mohamed, J.A., Huang, W., Nallapareddy, S.R., Teng, F., and Murray, B.E. (2004). Influence of origin of isolates, especially endocarditis isolates, and various genes on biofilm formation by Enterococcus faecalis. Infect. Immun. 72, 3658–3663. Mohamed, J.A., and Murray, B.E. (2005). Lack of correlation of gelatinase production and biofilm formation in a large collection of Enterococcus faecalis isolates. J. Clin. Microbiol. 43, 5405–5407. Moscoso, M., and Claverys, J.P. (2004). Release of DNA into the medium by competent Streptococcus pneumoniae: kinetics, mechanism and stability of the liberated DNA. Mol. Microbiol. 54, 783–794. Nakayama, J., Cao, Y., Horii, T., Sakuda, S., Akkermans, A.D., de Vos, W.M., and Nagasawa, H. (2001). Gelatinase biosynthesis-activating pheromone: a peptide lactone that mediates a quorum sensing in Enterococcus faecalis. Mol. Microbiol. 41, 145–154. Novick, R.P. (2003). Autoinduction and signal transduction in the regulation of staphylococcal virulence. Mol. Microbiol. 48, 1429–1449. Novick, R.P. (2006). Staphylococcal pathogenesis and pathogenicity factors: genetics and regulation. In: Gram-positive Pathogens, V.A. Fischetti, R.P. Novick, J.J. Ferretti, D.A. Portnoy, and J. I. Rood, eds. (Washington, D.C., ASM Press), pp. 496–516. Novick, R.P., Ross, H.F., Figueiredo, A.M.S., Abramochkin, G., Muir, T., Balaban, N., Singh, B., Goldkorn, T., Rasooly, A., Torres, J.V., and Uziel, O. (2000). Activation and inhibition of the staphylococcal AGR system. Science 287, 391a. Otto, M. (2001). Staphylococcus aureus and Staphylococcus epidermidis peptide pheromones produced by the accessory gene regulator system. Peptides 22, 1603–1608. Petersen, F.C., Pecharki, D., and Scheie, A.A. (2004). Biofilm mode of growth of Streptococcus intermedius favored by a competence-stimulating signaling peptide. J. Bacteriol. 186, 6327–6331. Pillai, S.K., Sakoulas, G., Eliopoulos, G.M., Moellering, R.C., Jr., Murray, B.E., and Inouye, R.T. (2004). Effects of glucose on fsr-mediated biofilm formation in Enterococcus faecalis. J. Infect. Dis. 190, 967–970. Pillai, S.K., Sakoulas, G., Gold, H.S., Wennersten, C., Eliopoulos, G.M., Moellering, R.C., Jr., and Inouye, R.T. (2002). Prevalence of the fsr locus in Enterococcus faecalis infections. J. Clin. Microbiol. 40, 2651–2652.

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Pratten, J., Foster, S.J., Chan, P.F., Wilson, M., and Nair, S.P. (2001). Staphylococcus aureus accessory regulators: expression within biofilms and effect on adhesion. Microbes Infect. 3, 633–637. Qi, F., Kreth, J., Levesque, C.M., Kay, O., Mair, R.W., Shi, W., Cvitkovitch, D.G., and Goodman, S.D. (2005). Peptide pheromone induced cell death of Streptococcus mutans. FEMS Microbiol. Lett. 251, 321–326. Qin, X., Singh, K.V., Weinstock, G.M., and Murray, B.E. (2001). Characterization of fsr, a regulator controlling expression of gelatinase and serine protease in Enterococcus faecalis OG1RF. J. Bacteriol. 183, 3372–3382. Rasmussen, M., and Bjorck, L. (2002). Proteolysis and its regulation at the surface of Streptococcus pyogenes. Mol. Microbiol. 43, 537–544. Roberts, J.C., Singh, K.V., Okhuysen, P.C., and Murray, B.E. (2004). Molecular epidemiology of the fsr locus and of gelatinase production among different subsets of Enterococcus faecalis isolates. J. Clin. Microbiol. 42, 2317–2320. Rosa, R., Creti, R., Venditti, M., D’Amelio, R., Arciola, C.R., Montanaro, L., and Baldassarri, L. (2006). Relationship between biofilm formation, the enterococcal surface protein (Esp) and gelatinase in clinical isolates of Enterococcus faecalis and Enterococcus faecium. FEMS Microbiol. Lett. 256, 145–150. Rothfork, J.M., Dessus-Babus, S., Van Wamel, W.J., Cheung, A.L., and Gresham, H.D. (2003). Fibrinogen depletion attenuates Staphylococcus aureus infection by preventing density-dependent virulence gene up-regulation. J. Immunol. 171, 5389–5395. Sakoulas, G., Eliopoulos, G.M., Moellering, R.C., Jr., Wennersten, C., Venkataraman, L., Novick, R.P., and Gold, H.S. (2002). Accessory gene regulator (agr) locus in geographically diverse Staphylococcus aureus isolates with reduced susceptibility to vancomycin. Antimicrob. Agents Chemother. 46, 1492–1502. Schwan, W.R., Langhorne, M.H., Ritchie, H.D., and Stover, C.K. (2003). Loss of hemolysin expression in Staphylococcus aureus agr mutants correlates with selective survival during mixed infections in murine abscesses and wounds. FEMS Immunol. Med. Microbiol. 38, 23–28. Shenkman, B., Rubinstein, E., Cheung, A.L., Brill, G.E., Dardik, R., Tamarin, I., Savion, N., and Varon, D. (2001). Adherence properties of Staphylococcus aureus under static and flow conditions: roles of agr and sar loci, platelets, and plasma ligands. Infect. Immun. 69, 4473–4478. Shenkman, B., Varon, D., Tamarin, I., Dardik, R., Peisachov, M., Savion, N., and Rubinstein, E. (2002). Role of agr (RNAIII) in Staphylococcus aureus adherence to fibrinogen, fibronectin, platelets and endothelial cells under static and flow conditions. J. Med. Microbiol. 51, 747–754. Shompole, S., Henon, K.T., Liou, L.E., Dziewanowska, K., Bohach, G.A., and Bayles, K.W. (2003). Biphasic intracellular expression of Staphylococcus aureus virulence factors and evidence for Agr-mediated diffusion sensing. Mol. Microbiol. 49, 919–927. Sifri, C.D., Mylonakis, E., Singh, K.V., Qin, X., Garsin, D.A., Murray, B.E., Ausubel, F.M., and Calderwood, S.B. (2002). Virulence effect of Enterococcus faecalis protease genes and the quorum-sensing locus fsr in Caenorhabditis elegans and mice. Infect. Immun. 70, 5647–5650. Singh, K.V., Qin, X., Weinstock, G.M., and Murray, B.E. (1998). Generation and testing of mutants of Enterococcus faecalis in a mouse peritonitis model. J. Infect. Dis. 178, 1416–1420. Somerville, G.A., Beres, S.B., Fitzgerald, J.R., DeLeo, F.R., Cole, R.L., Hoff, J.S., and Musser, J.M. (2002). In vitro serial passage of Staphylococcus aureus: changes in physiology, virulence factor production, and agr nucleotide sequence. J. Bacteriol. 184, 1430–1437. Suntharalingam, P., and Cvitkovitch, D.G. (2005). Quorum sensing in streptococcal biofilm formation. Trends Microbiol. 13, 3–6. Tegmark, K., Morfeldt, E., and Arvidson, S. (1998). Regulation of agr-dependent virulence genes in Staphylococcus aureus by RNAIII from coagulase-negative staphylococci. J. Bacteriol. 180, 3181– 3186. Truong-Bolduc, Q.C., Zhang, X., and Hooper, D.C. (2003). Characterization of NorR protein, a multifunctional regulator of norA expression in Staphylococcus aureus. J. Bacteriol. 185, 3127–3138. Tseng, C.W., Zhang, S., and Stewart, G.C. (2004). Accessory gene regulator control of staphylococcal enterotoxin D gene expression. J. Bacteriol. 186, 1793–1801. Tunney, M.M., and Gorman, S.P. (2002). Evaluation of a poly (vinyl pyrollidone)-coated biomaterial for urological use. Biomaterials 23, 4601–4608. van der Ploeg, J.R. (2005). Regulation of bacteriocin production in Streptococcus mutans by the quorumsensing system required for development of genetic competence. J. Bacteriol. 187, 3980–3989.

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Van Wamel, W.J., van Rossum, G., Verhoef, J., Vandenbroucke-Grauls, C.M., and Fluit, A.C. (1998). Cloning and characterization of an accessory gene regulator (agr)-like locus from Staphylococcus epidermidis. FEMS Microbiol. Lett. 163, 1–9. Vuong, C., Gerke, C., Somerville, G.A., Fischer, E.R., and Otto, M. (2003). Quorum-sensing control of biofilm factors in Staphylococcus epidermidis. J. Infect. Dis. 188, 706–718. Vuong, C., Gotz, F., and Otto, M. (2000a). Construction and characterization of an agr deletion mutant of Staphylococcus epidermidis. Infect. Immun. 68, 1048–1053. Vuong, C., Kocianova, S., Yao, Y., Carmody, A.B., and Otto, M. (2004). Increased colonization of indwelling medical devices by quorum-sensing mutants of Staphylococcus epidermidis in vivo. J. Infect. Dis. 190. Vuong, C., Saenz, H.L., Gotz, F., and Otto, M. (2000b). Impact of the agr quorum-sensing system on adherence to polystyrene in Staphylococcus aureus. J. Infect. Dis. 182, 1688–1693. Waters, C.M., Antiporta, M.H., Murray, B.E., and Dunny, G.M. (2003). Role of the Enterococcus faecalis GelE protease in determination of cellular chain length, supernatant pheromone levels, and degradation of fibrin and misfolded surface proteins. J. Bacteriol. 185, 3613–3623. Wesson, C.A., Liou, L.E., Todd, K.M., Bohach, G.A., Trumble, W.R., and Bayles, K.W. (1998). Staphylococcus aureus Agr and Sar global regulators influence internalization and induction of apoptosis. Infect. Immun. 66, 5238–5243. Xiong, Y.Q., Van Wamel, W., Nast, C.C., Yeaman, M.R., Cheung, A.L., and Bayer, A.S. (2002). Activation and transcriptional interaction between agr RNAII and RNAIII in Staphylococcus aureus in vitro and in an experimental endocarditis model. J. Infect. Dis. 186, 668–677. Yarwood, J.M. (2004). Quorum sensing in staphylococcal biofilms. Paper presented at: 11th International Symposium on Staphylococci and Staphylococcal Infections (Charleston, SC). Yarwood, J.M. (2006). Quorum-sensing-dependent regulation of staphylococcal virulence and biofilm development. In: Bacterial Cell-to-Cell Communication: Role in Virulence and Pathogenesis, D.R. Demuth, and R. J. Lamont, eds. (Cambridge, Cambridge University Press), pp. 199–231. Yarwood, J.M., Bartels, D.J., Volper, E.M., and Greenberg, E.P. (2004). Quorum sensing in Staphylococcus aureus biofilms. J. Bacteriol. 186, 1838–1850. Yarwood, J.M., and Greenberg, E.P. (2006). Generation of accessory gene regulator variants in Staphylococcus aureus biofilms. Paper presented at: American Society for Microbiology General Meeting (Orlando FL).

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Jeremy S. Webb

Abstract Biofilm formation is now commonly associated with concepts of development, differentiation, and dispersal of microorganisms, and often more broadly with multicellular biological systems. his underlying theme of multicellularity among sessile microorganisms has undoubtedly attracted significant fundamental research interest to the field. his chapter will summarize and discuss aspects of cellular differentiation in biofilms, including microcolony-based dispersal, autolysis of subpopulations of biofilm cells, and the recent finding that nitric oxide—a ubiquitous signal for cellular differentiation—can induce dispersal in Pseudomonas aeruginosa biofilms. Introduction—bioilms as primitive multicellular systems? Recent years have witnessed a dramatic expansion of research into bacterial biofilms, with rapid advances in molecular technologies and microscopy enabling detailed studies of the biofilm mode of life across a range of systems and organisms. his surge of interest is fueled in part by the ubiquitous impact of biofilms, and the need to manipulate, enhance or prevent biofilm formation in diverse environments. In addition, the discovery of cell–cell signaling or quorum sensing systems in bacteria was central in guiding researchers to study bacterial multicellularity and community behavior of bacteria, rather than the previous emphasis on single cell biological processes. Initially, two kinds of multicellular prokaryotic systems were envisaged. One system, as explored for example in differentiation and sporulation in Myxococcus xanthus, involves signaling-mediated development of specialized cells with the ensuing sharing of labor by the different types of cells in the population. he other system refers to the “mob” response displayed by a population of non-differentiating cells, in which the population adopts a new response, such as bioluminescence or virulence factor production, accommodated by the same and simultaneous behavior of all cells. However, since the discovery of a role of quorum sensing in organized surface motility in 1996 (Eberl et al., 1996) and later biofilm development in 1998 (Davies et al., 1998), a third concept has emerged; that of multicellularity and differentiation in biofilms. Biofilms have subsequently been found to display features reminiscent of multicellular systems, including co-ordinated dispersal events, and it now appears that complex development is a hallmark also of the community behavior of classically non-differentiating bacteria (Eberl et al., 1999; Eberl et al., 1996; Klausen et al., 2003; Sauer et al., 2002; Webb et al., 2003b). his chapter will

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discuss these and other recent findings in connection with the development and multicellular differentiation of biofilms. Overview of bioilm dispersal All organisms must disperse to and colonize new environments, and the ecological importance of the dispersal phase is well recognized. Colonization of new surfaces is a fundamental constraint on the life histories of sessile organisms; for example, most sessile eukaryotic organisms in aquatic environments have evolved dispersive propagules that represent a distinct stage in their life histories (e.g. invertebrate larvae, algal spores, etc). However, it is now also clear that (a) the principal mode of life of bacteria within the environment is as surface-dwelling biofilms, and that (b) biofilms exhibit dispersal stages on surfaces in many ways analogous to sessile eukaryotes. Biofilms are complex aggregates of cells on surfaces from which dispersal cells are at times released, e.g. in the language used for eukaryotes, there is a benthic phase in the “life cycle” from which distinct, dispersive “propagules” are released. While many bacterial cells can disperse from biofilms by passive processes, such as erosion or sloughing of cells from the biofilm caused by fluid shear (Stoodley et al., 2001a; Stoodley et al., 2001b) bacterial biofilms can also periodically undergo active dispersal events where sessile, matrix-encased biofilm cells convert en-masse to free-swimming, planktonic bacteria. Until recently, the mechanisms by which active bacterial dispersal from biofilms occurs remained almost completely unexplored, and little was known about the functions or regulatory pathways involved in release of bacteria from biofilms. Strategies to manipulate biofilm dispersal would find broad application in the control of microorganisms and this may in part be responsible for a recent surge of interest in biofilm dispersal. Mechanisms by which biofilms regulate dispersal are only beginning to be explored and will be an important area of research for the future. Processes now known to play a role in biofilm dispersal include enzyme-mediated breakdown of the biofilm matrix (Boyd and Chakrabarty, 1994; Kaplan et al., 2003a; Kaplan et al., 2003b; Lee et al., 1996), and the production of surfactants which loosen cells from the biofilm (Davey et al., 2003). Dispersal processes can also be under the control of quorum sensing systems (Rice et al., 2005), intracellular di-cyclic GMP levels (see also Chapter 5), changes in nutrient availability (Gjermansen et al., 2005; Hunt et al., 2004; Sauer et al., 2004) and the production of free radical species ((Barraud et al., 2006; Webb et al., 2003b), and see below). Biofilm dispersal is thus a dynamic process involving multiple genetic determinants and regulatory processes. Escape from within: development and dispersal of microcolonies Microcolony development With some notable exceptions (e.g. Hentzer et al., 2001; Heydorn et al., 2002), the vast majority of laboratory confocal microscopy biofilm investigations report multicell structures (microcolonies) that are often elegantly differentiated from the bulk biofilm. It has previously been proposed that three-dimensional biofilm structures are a consequence of complex nutrient gradients during growth, and hence that microcolony formation can be

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modeled mathematically (van Loosdrecht et al., 2002). However, recent years have also revealed numerous genetically encoded regulatory and structural determinants of biofilm development. Examples include the role of conjugative plasmids (Ghigo, 2001; Reisner et al., 2003) and antigen 43 (Danese et al., 2000; Reisner et al., 2003) in enhancing microcolony formation in E. coli biofilms, suggesting that these cell surface structures may act as cellular adhesins in stabilizing microcolony structures. Several studies have reported an involvement of cell–cell signaling in microcolony formation e.g. (Davies et al., 1998; Huber et al., 2001; Lynch et al., 2002), and pointed to the outward structural similarities between microcolony development and coordinated aggregation and fruiting body formation in social bacteria such as Myxococcus spp. In a clear example of the existence of different cell types within a population, Klausen et al. (Klausen et al., 2003) report a type IV pilus-mediated migration of a subpopulation of P. aeruginosa cells to form mushroom-like caps on the surface of microcolonies (Figure 9.1D). his process also shares similarities with the development of fruiting bodies in M. xanthus. Dispersal from microcolonies A paradigm of the biofilm mode of bacterial life is of a developmental sequence that culminates in the dispersal of physiologically differentiated free-living cells that can colonize new locations. Behaviors reminiscent of multicellular organization have been observed during dispersal processes occurring in mature microcolonies. Several recent studies have reported pronounced activity localized to the center of mature biofilm structures, leading to the dispersal of cells from inside the structure, and leaving behind large transparent cavities, or hollow “shells” made up of non-motile cells (Figure 9.1). his process of dispersal from the interior of microcolonies has been termed “seeding dispersal” in order to differentiate it from the process of erosion, which is the passive removal of cells from the biofilm by fluid shear. Seeding dispersal in Pseudomonas aeruginosa did not occur in a $lasI/rhlI quorum-sensing mutant strain, suggesting that this process is dependant on cell–cell signaling (Purevdorj-Gage et al., 2005). Such processes are likely also to involve enzymes that degrade the extracellular polysaccharide matrix (for example A

Motile cells

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Figure 9.1 Bacterial dispersal from multicellular bioilm structures (microcolonies), also termed “seeding dispersal.” The left pane shows a schematic “side-on” view of a mature bioilm undergoing dispersal; the right pane shows a “top-down” view of hollow microcolonies on a glass surface formed during dispersal of a Pseudomonas aeruginosa bioilm (bar = 50 µm). Panel A was kindly provided by Susse Kirkelund Hansen, Biocentrum-DTU, Danish Technical University.

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polysaccharide lyases) which are known to be of importance for their role in biofilm dissolution in several organisms (Kaplan et al., 2003b; Ott et al., 2001). Because these enzymes are not thought to be transported across the cell membrane, their release to the extracellular polysaccharide matrix may rely primarily upon lysis of cells within the biofilm (Sutherland, 1999). Cell lysis and dispersal One process that is often observed in association with microcolony seeding dispersal is the death and lysis of subpopulations of cells within the biofilm. Patterns of cell lysis within biofilms has now been observed as a normal feature of development across a broad range of biofilm forming bacteria. In biofilms, cell death commonly occurs with spatial organization inside mature microcolony structures, and kills only a proportion of cells within the biofilm (see Figure 9.3). It is proposed that autolysis impacts on dispersal processes in biofilms by destabilizing and disrupting the biofilm architecture, and that surviving cells in the biofilm benefit from the death of their siblings (Webb et al., 2003a, and see “Impacts of cell lysis in biofilms,” below), which facilitates conversion of surviving cells to the motile dispersal phenotype. he two organisms for which cell lysis during biofilm development are best understood are P. aeruginosa, and the ubiquitous marine bacterium, Pseudoaltermonas tunicata. In both of these organisms, specific genetic determinants control cell death during biofilm development. In P. tunicata, an autolytic protein, AlpP, is required for cell death during biofilm development (Mai-Prochnow et al., 2004; Mai-Prochnow et al., 2006) and remarkably, cell death was not observed at any stage in the development of a P. tunicata alpP mutant biofilm. In P. aeruginosa, cell death was linked to the expression of a Pf1-like filamentous prophage of Pseudomonas aeruginosa, and consequently genes that affected expression of receptors for the phage (RpoN-mediated regulation of type IV pili and flagellae) controlled cell death during development (Figure 9.1D) (Webb et al., 2003b). However, the molecular mechanism of killing in P. aeruginosa biofilms remains to be fully elucidated because these symbiotic filamentous phages are generally thought not to harm host cells. Intriguingly, the Pf1-like prophage of P. aeruginosa encodes homologues of proteins from two different E. coli toxin-antitoxin elements ((hompson et al., 2003) and M Lau, JS Webb and S Kjelleberg, unpublished data) which may play a role in cell killing within biofilms. In both P. aeruginosa and P. tunicata, mutants that did not show cell death did not develop hollow shell-like structures and dispersal of cells from internalized portions of the microcolony was not observed. Impacts of cell lysis in bioilms—enhanced metabolic activity and phenotypic variation among dispersal cells Recent studies have provided further evidence that cell lysis during biofilm development can indeed play an important role in biofilm dispersal and ecology. In the marine bacterium P. tunicata, cell death in wild-type biofilms led to a major reproducible dispersal event after 192 hours of biofilm development (Mai-Prochnow et al., 2006; Figure 9.2). A sudden increase in viable dispersal cells occurred only in the P. tunicata wild-type, but not in the alpP mutant. Using flow cytometry and the fluorescent dye DiBAC, it was also shown

Differentiation and Dispersal in Bioilms

Figure 9.2 A P. tunicata alpP mutant which does not undergo cell lysis is defective in bioilm dispersal. Bioilm dispersal shown in efluent viable counts (CFU). P. tunicata wild-type (black squares) shows a signiicant dispersal event at 192 hours. No major increase in dispersal can be detected for the ΔalpP mutant (grey triangles). Error bars represent the standard deviations for three independent experiments. reproduced from (Mai-Prochnow et al., 2006).

that P. tunicata wild-type cells that disperse from biofilms have enhanced metabolic activity compared to cells obtained from alpP mutant biofilms. Furthermore, using the BacLight Live/dead DNA stain, viable cells within the region of lysis appeared larger and with enhanced fluorescence intensity, presumably due to higher levels of DNA caused by greater nutrient uptake, suggesting that lysis material from dead cells may also contribute to the nutrient support of surviving cells. Studies of P. tunicata biofilms development therefore suggest that autocidal events mediated by an antibacterial protein can confer ecological advantages by enhancing dispersal and contributing to the production of a metabolically active subpopulation of dispersal cells. he use of dead cells as a nutrient source for surviving bacteria in this way has also been proposed to occur during differentiation processes in other organisms, including sporulation of Bacillus subtilis (Gonzalez-Pastor et al., 2003), mycelium formation of Streptomyces sp. (Mendez et al., 1985; Miguelez et al., 1999) and biofilm formation of Staphylococcus aureus (Resch et al., 2005). Cell lysis in biofilms may also be important for the colonization of new surfaces. Studies on P. tunicata have reported that there was considerable phenotypic variation among cells dispersing from wild-type biofilms, but not from the alpP mutant (without lysis). Wildtype cells that dispersed from biofilms showed significant increases in variation in growth, motility and biofilm formation which may all be important for successful colonization of new surfaces. Generally, it is recognized that a high diversity within a community protects against unfavorable conditions by increasing the range of conditions in which a community as a whole can thrive (Boles et al., 2005; McCann, 2000). In P. tunicata variation in colonization-relevant traits appears to be relatively stable in the dispersal cells as three culturing steps in culture media did not allow for reversion of the phenotypes. he highest variation was detected in the wild-type immediately after cell death had occurred, with some variants showing high growth rates and rapid biofilm while others showed slow growth rates and were mostly deficient in biofilm formation. Furthermore, some variants derived from wild-type biofilms showed increased motility. Each of these phenotypes may influence the

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ability of P. tunicata cells to colonize surfaces under different environmental conditions. For example, a higher growth and biofilm formation rate may enhance colonization under high nutrient conditions and a phenotype with increased motility could enhance settlement at more distant surfaces, which could lead to a wider distribution of the organism. he need to disperse to new habitats is an important constraint on all organisms with sessile stages in their lifecycle. Analogously, the generation of variability among dispersal propagules has been studied widely for many sessile colonizing eukaryotes (Marshall et al., 2003; Marshall and Keough, 2003; Moran and Emlet, 2001). It is well established for these organisms that variation in a range of phenotypes within propagule populations is a strategy used to ensure successful colonization at new surfaces in different habitats. For example differences in swimming ability due to variation in the size and nutritional status of larvae of the bryozoans Bugula neritina and Watersipora subtorquata and the ascidian Diplosoma listerianum lead to variation in settlement distances (Marshall and Keough, 2003). Overall, the studies described above suggest that developmental cell death and its consequences confer ecological advantages to groups of bacteria in P. tunicata biofilms and more generally the fitness of the species by generating a diverse but stable dispersal population. Because patterns of cell death during biofilm development are a feature of many bacteria, exploring the role of self induced lysis and generation of phenotypically different dispersal cells can lead to a better understanding of the ecology of the sessile lifestyle of non-differentiating bacteria as well as the development of potential control mechanisms of bacterial biofilms. Nitric oxide—a signal for differentiation and dispersal in bioilms? Nitric oxide (NO) is intimately associated with processes of central importance to biofilm development. Biofilms predominantly exhibit gene expression profiles consistent with anaerobic and iron limited growth (Hassett et al., 2002; Hentzer et al., 2005), and NO plays an important role in both of these growth modes. In P. aeruginosa biofilms, steep oxygen gradients can occur leading to anaerobic regions within the biofilm. Anaerobic respiratory metabolism in P. aeruginosa uses nitrate (NO3–), nitrite (NO2–) or nitrous oxide (N2O) as terminal electron acceptors and NO is generated in this process through the activity of the enzyme nitrite reductase. If NO is not reduced by NO reductase to N2O, it may compromise the viability of the biofilm (Yoon et al., 2002). NO is also closely linked with iron acquisition, often a limiting factor for microorganisms in biofilms. In diverse bacterial species, NO inhibits the DNA binding by the ferric uptake regulator (Fur), leading to upregulation of genes required for iron acquisition (D’Autreaux et al., 2002; Ochsner et al., 2002). While bacterial responses to NO have been studied extensively in planktonic bacterial physiology in the context of adaptive and protective mechanisms e.g. (Firoved et al., 2004; Mills et al., 2005; Poole, 2005; Tucker et al., 2006), there is a paucity of information as to its role or cellular targets in the context of multicellular biofilm development and differentiation processes. Recently it was found that nitric oxide (NO) is able to induce biofilm dispersal at concentrations that are non-toxic to Pseudomonas aeruginosa (Barraud et al., 2006). Using the NO donor sodium nitroprusside (SNP), concentrations as low as 10 nM were found

Differentiation and Dispersal in Bioilms

to cause a dispersal of biofilm cells from glass surfaces. Addition of SNP to established P. aeruginosa biofilms on glass slides caused up to 80% reduction in the amount of biomass on the glass surface. It was demonstrated that anaerobic denitrification occurs inside P. aeruginosa biofilms and that levels of peroxynitrite (a stable and highly damaging reaction product of NO) were enhanced in multicell structures that had undergone dispersal. A $nirS mutant strain of P. aeruginosa, lacking the only enzyme capable of generating metabolic NO, did not undergo dispersal in the study. he findings suggest a model for NOmediated differentiation during microcolony dispersal as described in Figure 9.3. A number of other studies within the literature also point towards a role for NO in the transition from the biofilm mode of growth to the planktonic, free-living form in P. aeruginosa. First, microarray studies have revealed that genes involved in adherence are downregulated in P. aeruginosa upon exposure to NO (Firoved et al., 2004). his suggests a mechanism by which NO-exposed bacteria detach from the biofilm leading to reduced biofilm biomass and increased number of planktonic organisms. Second, the transition from sessile to motile P. aeruginosa is known to be regulated by GGDEF and EAL protein domains that are involved in the turnover of c-di-GMP (Simm et al., 2004). Several signal transduction pathways are known to regulate the activity of these GGDEF and EAL domains, including sensing of oxygen, pH, temperature and other environmental stimuli (Galperin et al., 2001; Romling et al., 2005; Simm et al., 2004). Intriguingly, Aravind and colleagues (Iyer et al., 2003) have also found that NO sensing proteins, called heme nitric oxide binding (HNOB) domains, are frequently associated with GGDEF and EAL domains in diverse bacteria, suggesting a link between NO-sensing and c-di-GMP turnover. Further studies in our laboratory will establish whether NO-mediated biofilm dispersal in

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Figure 9.3 Model for NO-mediated dispersal and lysis. Anaerobic denitriication in the interior of bioilm microcolonies generates NO, as well as the cell-toxic radical ONOO– by the reaction between NO and superoxide (A). The reactive NO and ONOO– species lead to complex differentiation inside microcolonies. A confocal laser microscope image of a mature microcolony stained with the BacLight Live/Dead stain is shown in (B). It is hypothesized that NO induces a motile and planktonic phenotype in one subpopulation of bioilm cells (illed arrow, rod-shaped live cells (green in color version of this image)). In contrast, ONOO– causes cellular damage, bacteriophage induction (Webb et al., 2003b), and lysis of another subpopulation of cells within the bioilm (unilled arrow, irregular-shaped dead cells (red in color version of this image)). Ultimately, the combination of lysis and dispersal leads to the formation of “hollow colonies” typically observed during processes of seeding dispersal. Bar = 50 µm. The color version of this igure is available at http://www.horizonpress.com/hsp/supplementary/bioilm/ch9ig3.jpg.

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P. aeruginosa involves the GGDEF and EAL domains and variation in c-di-GMP levels and candidate gene products have already been identified for prioritization. Recent analyses of microbial genomes have suggested that homologous NO-sensing receptor domains are common to both prokaryotic and eukaryotic regulatory proteins (Aravind et al., 2003; Iyer et al., 2003). In eukaryotes, NO signaling is known to play an important role in the regulation of diverse processes, including apoptosis, cell proliferation and differentiation. Intriguing similarities exist between the signaling role of NO in eukaryotes, and its control of biofilm cell differentiation, death and dispersal. Biofilms are thought to exhibit developmental analogies with multicellular eukaryotes (Branda and Kolter, 2004; Webb et al., 2003a), and therefore it may be relevant to examine these bacterial biofilm populations for the origins of key regulatory processes observed in more complex organisms. Work on NO-mediated control of biofilm development in P. aeruginosa may point to a conserved role for NO signaling in the regulation of differentiation and developmental events across prokaryotic and eukaryotic physiology. References Aravind, L., Anantharaman, V., and Iyer, L.M. (2003). Evolutionary connections between bacterial and eukaryotic signaling systems: a genomic perspective. Curr. Opin. Microbiol. 6, 490–497. Barraud, N., Hassett, D.J., Hwang, S., Kjelleberg, S., and Webb, J.S. (2006). Nitric-oxide mediated biofilm dispersal in Pseudomonas aeruginosa. J. Bacteriol. 188, 7344–7353. Boles, B.R., hoendel, M., and Singh, P.K. (2005). Self-generated diversity produces “insurance effects” in biofilm communities (vol 101, pg 16630, 2004). Proc. Natl. Acad. Sci. USA 102, 955–955. Boyd, A., and Chakrabarty, A.M. (1994). Role of alginate lyase in cell detachment of Pseudomonas aeruginosa. Appl. Environ. Microbiol. 60, 2355–2359. Branda, S.S., and Kolter, R. (2004). Multicellularity in biofilms. In: Microbial Biofilms, M. Ghannoum, and G. O’Toole, eds. (Washington, D.C., ASM Press), pp. 20–29. D’Autreaux, B., Touati, D., Bersch, B., Latour, J.M., and Michaud-Soret, I. (2002). Direct inhibition by nitric oxide of the transcriptional ferric uptake regulation protein via nitrosylation of the iron. Proc. Natl. Acad. Sci. USA 99, 16619–16624. Danese, P.N., Pratt, L.A., Dove, S.L., and Kolter, R. (2000). he outer membrane protein, antigen 43, mediates cell-to-cell interactions within Escherichia coli biofilms. Mol. Microbiol. 37, 424–432. Davey, M.E., Caiazza, N.C., and O’Toole, G.A. (2003). Rhamnolipid surfactant production affects biofilm architecture in Pseudomonas aeruginosa PA01. J. Bacteriol. 185, 1027–1036. Davies, D.G., Parsek, M.R., Pearson, J.P., Iglewski, B.H., Costerton, J.W., and Greenberg, E.P. (1998). he involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280, 295–298. Eberl, L., Molin, S., and Givskov, M. (1999). Surface motility of serratia liquefaciens MG1. J. Bacteriol. 181, 1703–1712. Eberl, L., Winson, M.K., Sternberg, C., Stewart, G.S., Christiansen, G., Chhabra, S.R., Bycroft, B., Williams, P., Molin, S., and Givskov, M. (1996). Involvement of N-acyl-L-hormoserine lactone autoinducers in controlling the multicellular behaviour of Serratia liquefaciens. Mol. Microbiol. 20, 127–136. Firoved, A.M., Wood, S.R., Ornatowski, W., Deretic, V., and Timmins, G.S. (2004). Microarray analysis and functional characterization of the nitrosative stress response in nonmucoid and mucoid Pseudomonas aeruginosa. J. Bacteriol. 186, 4046–4050. Galperin, M.Y., Nikolskaya, A.N., and Koonin, E.V. (2001). Novel domains of the prokaryotic two-component signal transduction systems. FEMS Microbiol. Lett. 203, 11–21. Ghigo, J.M. (2001). Natural conjugative plasmids induce bacterial biofilm development. Nature 412, 442–445. Gjermansen, M., Ragas, P., Sternberg, C., Molin, S., and Tolker-Nielsen, T. (2005). Characterization of starvation-induced dispersion in biofilms. Environ. Microbiol. 7, 894–906. Gonzalez-Pastor, J.E., Hobbs, E.C., and Losick, R. (2003). Cannibalism by sporulating bacteria. Science 301, 510–513.

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Hassett, D.J., Cuppoletti, J., Trapnell, B., Lymar, S.V., Rowe, J.J., Yoon, S.S., Hilliard, G.M., Parvatiyar, K., Kamani, M.C., Wozniak, D.J., et al. (2002). Anaerobic metabolism and quorum sensing by Pseudomonas aeruginosa biofilms in chronically infected cystic fibrosis airways: rethinking antibiotic treatment strategies and drug targets. Adv. Drug Deliv. Rev. 54, 1425–1443. Hentzer, M., Eberl, L., and Givskov, M. (2005). Transcriptome analysis of Pseudomonas aeruginosa biofilm development. Biofilms 2, 37–61. Hentzer, M., Teitzel, G.M., Balzer, G.J., Heydorn, A., Molin, S., Givskov, M., and Parsek, M.R. (2001). Alginate overproduction affects Pseudomonas aeruginosa biofilm structure and function. J. Bacteriol. 183, 5395–5401. Heydorn, A., Ersboll, B., Kato, J., Hentzer, M., Parsek, M.R., Tolker-Nielsen, T., Givskov, M., and Molin, S. (2002). Statistical analysis of Pseudomonas aeruginosa biofilm development: impact of mutations in genes involved in twitching motility, cell-to-cell signaling, and stationary-phase sigma factor expression. Appl. Environ. Microbiol. 68, 2008–2017. Huber, B., Riedel, K., Hentzer, M., Heydorn, A., Gotschlich, A., Givskov, M., Molin, S., and Eberl, L. (2001). he cep quorum-sensing system of Burkholderia cepacia H111 controls biofilm formation and swarming motility. Microbiology 147, 2517–2528. Hunt, S.M., Werner, E.M., Huang, B., Hamilton, M.A., and Stewart, P.S. (2004). Hypothesis for the role of nutrient starvation in biofilm detachment. Appl. Environ. Microbiol. 70, 7418–7425. Iyer, L.M., Anantharaman, V., and Aravind, L. (2003). Ancient conserved domains shared by animal soluble guanylyl cyclases and bacterial signaling proteins. BMC Genomics 4, 5. Kaplan, J.B., Meyenhofer, M.F., and Fine, D.H. (2003a). Biofilm growth and detachment of Actinobacillus actinomycetemcomitans. J. Bacteriol. 185, 1399–1404. Kaplan, J.B., Ragunath, C., Ramasubbu, N., and Fine, D.H. (2003b). Detachment of Actinobacillus actinomycetemcomitans biofilm cells by an endogenous beta-hexosaminidase activity. J. Bacteriol. 185, 4693–4698. Klausen, M., Aaes-Jorgensen, A., Molin, S., and Tolker-Nielsen, T. (2003). Involvement of bacterial migration in the development of complex multicellular structures in Pseudomonas aeruginosa biofilms. Mol. Microbiol. 50, 61–68. Lee, S.F., Li, Y.H., and Bowden, G.H. (1996). Detachment of Streptococcus mutans biofilm cells by an endogenous enzymatic activity. Infect. Immun. 64, 1035–1038. Lynch, M.J., Swift, S., Kirke, D.F., Keevil, C.W., Dodd, C.E., and Williams, P. (2002). he regulation of biofilm development by quorum sensing in Aeromonas hydrophila. Environ. Microbiol. 4, 18–28. Mai-Prochnow, A., Evans, F., Dalisay-Saludes, D., Stelzer, S., Egan, S., James, S., Webb, J.S., and Kjelleberg, S. (2004). Biofilm Development and Cell Death in the Marine Bacterium Pseudoalteromonas tunicata. Appl. Environ. Microbiol. 70, 3232–3238. Mai-Prochnow, A., Webb, J.S., Ferrari, B.C., and Kjelleberg, K. (2006). Ecological advantages of autolysis during the development and dispersal of Pseudoalteromonas tunicata biofilms. Appl. Environ. Microbiol. 72, 5414–5420 Marshall, D.J., Bolton, T.F., and Keough, M.J. (2003). Offspring size affects the post-metamorphic performance of a colonial marine invertebrate. Ecology 84, 3131–3137. Marshall, D.J., and Keough, M.J. (2003). Variation in the dispersal potential of non-feeding invertebrate larvae: the desperate larva hypothesis and larval size. MEPS 255, 145–153. McCann, K.S. (2000). he diversity-stability debate. Nature 405, 228–233. Mendez, C., Brana, A.F., Manzanal, M.B., and Hardisson, C. (1985). Role of substrate mycelium in colony development in Streptomyces. Can. J. Microbiol. 31, 446–450. Miguelez, E.M., Hardisson, C., and Manzanal, M.B. (1999). Hyphal death during colony development in Streptomyces antibioticus: morphological evidence for the existence of a process of cell deletion in a multicellular prokaryote. J. Cell Biol. 145, 515–525. Mills, P.C., Richardson, D.J., Hinton, J.C., and Spiro, S. (2005). Detoxification of nitric oxide by the flavorubredoxin of Salmonella enterica serovar Typhimurium. Biochem. Soc. Trans. 33, 198–199. Moran, A.L., and Emlet, R.B. (2001). Offspring size and performance in variable environments: field studies on a marine snail. Ecology 82, 1597–1612. Ochsner, U.A., Wilderman, P.J., Vasil, A.I., and Vasil, M.L. (2002). GeneChip expression analysis of the iron starvation response in Pseudomonas aeruginosa: identification of novel pyoverdine biosynthesis genes. Mol. Microbiol. 45, 1277–1287.

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Ott CM, Day DF, Koenig DW, Pierson DL. (2001). he release of alginate lyase from growing Pseudomonas syringae pathovar phaseolicola. Curr. Microbiol. 2001, 42:78–81. Poole, R.K. (2005). Nitric oxide and nitrosative stress tolerance in bacteria. Biochem. Soc. Trans. 33, 176–180. Purevdorj-Gage, B., Costerton, W.J., and Stoodley, P. (2005). Phenotypic differentiation and seeding dispersal in non-mucoid and mucoid Pseudomonas aeruginosa biofilms. Microbiology 151, 1569–1576. Reisner, A., Haagensen, J.A., Schembri, M.A., Zechner, E.L., and Molin, S. (2003). Development and maturation of Escherichia coli K-12 biofilms. Mol. Microbiol. 48, 933–946. Resch, A., Fehrenbacher, B., Eisele, K., Schaller, M., and Gotz, F. (2005). Phage release from biofilm and planktonic Staphylococcus aureus cells. FEMS Microbiol. Lett. 252, 89–96. Rice, S.A., Koh, K.S., Queck, S.Y., Labbate, M., Lam, K.W., and Kjelleberg, S. (2005). Biofilm formation and sloughing in Serratia marcescens are controlled by quorum sensing and nutrient cues. J. Bacteriol. 187, 3477–3485. Romling, U., Gomelsky, M., and Galperin, M.Y. (2005). C-di-GMP: the dawning of a novel bacterial signalling system. Mol. Microbiol. 57, 629–639. Sauer, K., Camper, A.K., Ehrlich, G.D., Costerton, J.W., and Davies, D.G. (2002). Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J. Bacteriol. 184, 1140–1154. Sauer, K., Cullen, M.C., Rickard, A.H., Zeef, L.A., Davies, D.G., and Gilbert, P. (2004). Characterization of nutrient-induced dispersion in Pseudomonas aeruginosa PAO1 biofilm. J. Bacteriol. 186, 7312–7326. Simm, R., Morr, M., Kader, A., Nimtz, M., and Romling, U. (2004). GGDEF and EAL domains inversely regulate cyclic di-GMP levels and transition from sessility to motility. Mol. Microbiol. 53, 1123–1134. Stoodley, P., Hall-Stoodley, L., and Lappin-Scott, H.M. (2001a). Detachment, surface migration, and other dynamic behavior in bacterial biofilms revealed by digital time-lapse imaging. Methods Enzymol. 337, 306–319. Stoodley, P., Wilson, S., Hall-Stoodley, L., Boyle, J.D., Lappin-Scott, H.M., and Costerton, J.W. (2001b). Growth and detachment of cell clusters from mature mixed-species biofilms. Appl. Environ. Microbiol. 67, 5608–5613. Sutherland, I.W. (1999). Biofilm exopolysaccharides. In: Microbial Extracellular Polymeric Substances: Characterization, Structure and Function. Edited by Wingender J, Neu TR, Hans-Curt F. Berlin: Springer; pp 73–92. hompson, L.S., Webb, J.S., Rice, S.A., and Kjelleberg, S. (2003). he alternative sigma factor RpoN regulates the quorum sensing gene rhlI in Pseudomonas aeruginosa. FEMS Microbiol. Lett. 220, 187–195. Tucker, N.P., D’Autreaux, B., Spiro, S., and Dixon, R. (2006). Mechanism of transcriptional regulation by the Escherichia coli nitric oxide sensor NorR. Biochem. Soc. Trans. 34, 191–194. van Loosdrecht, M.C., Heijnen, J.J., Eberl, H., Kreft, J., and Picioreanu, C. (2002). Mathematical modelling of biofilm structures. Antonie Van Leeuwenhoek 81, 245–256. Webb, J.S., Givskov, M., and Kjelleberg, S. (2003a). Bacterial biofilms: prokaryotic adventures in multicellularity. Curr. Opin. Microbiol. 6, 578–585. Webb, J.S., hompson, L.S., James, S., Charlton, T., Tolker-Nielsen, T., Koch, B., Givskov, M., and Kjelleberg, S. (2003b). Cell death in Pseudomonas aeruginosa biofilm development. J. Bacteriol. 185, 4585–4592. Yoon, S.S., Hennigan, R.F., Hilliard, G.M., Ochsner, U.A., Parvatiyar, K., Kamani, M.C., Allen, H.L., DeKievit, T.R., Gardner, P.R., Schwab, U., et al. (2002). Pseudomonas aeruginosa anaerobic respiration in biofilms: relationships to cystic fibrosis pathogenesis. Dev. Cell 3, 593–603.

Human Oral Multi-species Bioilms: Bacterial Communities in Health and Disease

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Paul E. Kolenbrander, Nicholas S. Jakubovics, Natalia I. Chalmers, and Robert J. Palmer, Jr.

Abstract Possibly the first biofilm samples ever examined from a microbiological perspective were obtained from the oral cavity: Antonie van Leeuwenhoek’s tooth scrapings. Since that time, oral microbiologists have made major contributions to microbial taxonomy, physiology, and ecology. he oral cavity distinguishes itself from other environments by having over 700 phylotypes (taxonomic units), nearly half of which have culturable representatives. Aerobic, facultatively anaerobic, and obligately anaerobic physiologies are present. Members of the microbial kingdoms Archaea, Bacteria and Fungi are present. What generates and maintains this diversity? Why are these communities attractive targets for study? How does community analysis using modern molecular methods differ from that using classical bacteriological approaches? We strive to answer these questions in the following contribution and, as far as possible, we rely on knowledge obtained from studies of plaque in situ. The oral cavity as an environment for bioilm growth Sites for biofilm development in the human oral cavity can be segregated into tooth-associated sites and soft-tissue sites. Tooth surfaces are one of the few non-shedding surfaces in the human body and thus present unique opportunities for biofilm development. Each tooth varies somewhat from its neighbors with respect to flow rate of saliva across its surface, and with respect to wear through contact with the tongue or cheek. In contrast to lingual or buccal tooth surfaces, interproximal (between teeth) tooth surfaces are shielded from wear. Teeth thus present colonization sites not only along a gradient of nutrient quantity/quality that develops according to proximity to the different salivary glands and to gingival sulci (the source of gingival crevicular fluid) (Hannig, 1999), but also along a gradient of shear stress that develops according to salivary flow rate and abrasion (Dawes, et al., 1989). Softtissue surfaces likewise present sites along similar gradients, however these surfaces are continuously shed and thus must be constantly recolonized. Turnover time of oral epithelia ranges from roughly 6 days (tongue, cheek) to as much as 12 days (gingiva) (Itoiz and Carranza, 2002). Desquamated epithelial cells have been noted as components of tooth-surface biofilms (Nyvad, 1993) (Figure 10.1). Clearly, colonization of the desquamating epithelial cells is a different process compared to colonization of the non-shedding tooth surface.

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Figure 10.1 An epithelial cell detected on the enamel surface is colonized with multi-species bacterial bioilm communities in 8-hour supragingival dental plaque. Communities are documented with FISH probes (eubacterial probe EUB338, blue; Streptococcus spp. probe STR405, red) in conjunction with general nucleic acid stain (acridine orange, green). The nucleus of the epithelial cell is stained with acridine orange (green). Streptococcus spp. cells (purple, colocalization of red + blue) are closely associated with non-Streptococcus spp. cells (blue) on the epithelial surface. (Bar 5 Mm; inset—same region at lower magniication). Reprinted from (Kolenbrander, et al., 2006). See also Plate 10.1.

Although saliva bathes both surfaces, biofilms on these two kinds of surfaces are certainly distinct. Saliva is the fluid that transports nutrients of dietary origin, in the form of partially dissolved carbohydrates and peptides, to oral biofilms. In addition, the same proteins and glycoproteins that make up the salivary secretions themselves can be substrates for bacterial growth (Homer, et al., 1996; Palmer, et al., 2001). he majority of salivary secretions originate from the three different major glands: parotid, submandibular, and sublingual. Each gland secretes saliva of a different composition: for example, the parotid gland is the source of amylase, whereas the sublingual and submandibular glands secrete the bulk of the mucins (Scannapieco, 1994). hus, while the secretions of these glands combine within the oral cavity to create the mixture known as whole saliva, proximity of a particular bacterial colonization site to the ducts of the glands influences the composition and flow rate of the saliva across the site. he major component of whole saliva is the group of glycoproteins known as mucins. Mucins are also the major component in other secretions such as cervical mucous and sweat, but saliva differs dramatically from other secretions in the type and amount of other components. For example, amylase is the most prevalent protein component of saliva yet is found in only small amounts in other body secretions. Several minor

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salivary components, such as lactoferrin, histatins and lysozyme, have antimicrobial properties thought to hinder invasion of the oral environment by bacteria not specifically adapted to this niche (Scannapieco, 1994). Salivary components such as proline-rich proteins are adsorbed onto tooth surfaces to form the acquired enamel pellicle, a protein film containing molecules specifically recognized by bacterial adhesins during the initial colonization of teeth (Scannapieco, 1994). In fact, because pellicle formation is so rapid (within minutes), oral biofilm bacteria are not in direct contact with the tooth surface. Tooth surfaces in and near the gingival sulci are contacted by a second non-salivary secretion called gingival crevicular fluid (GCF). Like saliva, this secretion is a complex mixture, but GCF is closer in composition to serum than to saliva. GCF is roughly 30-fold higher in protein concentration than is saliva, and GCF is the major source of immunoglobulin G in the oral cavity (Cimasoni, 1983). Flow rates of GCF are very low (0.5–2.4 ml per day across the entire oral cavity) compared to saliva (0.5–2.0 ml per minute), and GCF is the primary fluid within the sulcus. hus, the hard tissue environments of subgingival (bathed in GCF) and supragingival (bathed in saliva) sites are different from each other and their topology, in turn, is different from the soft tissue surface structure. he constant flow of saliva and GCF make it imperative for oral bacteria to adhere to a surface to prevent being swallowed. he differences in molecular composition of the acquired pellicle on the supragingival surface, GCF-bathed subgingival surface, and soft tissue surface mediate specific bacterial adherence preferences called tissue tropism. Tissue tropism Early studies he first connection between the ability of oral bacteria to adhere and tissue tropism was reported in 1970 (Van Houte, et al., 1970). Using human volunteers, they showed that Streptococcus salivarius preferentially bound to the tongue whereas other streptococci presumed to be Streptococcus sanguis adhered preferentially to the enamel surface. Results from another early human study using S. salivarius and Streptococcus mutans isolated from human volunteers showed that S. mutans bound preferentially to enamel and, again, S. salivarius bound to epithelial surfaces (Gibbons and van Houte, 1971). In the latter experiment, streptomycin-resistant mutants of S. salivarius and S. mutans were first isolated and then re-introduced into a volunteer’s mouth. It had been known that up to 100 times more S. salivarius could be cultured from the tongue than from enamel, but these data from reintroduction of streptomycin-resistant strains proved the hypothesis that the specificity of adherence and colonization was a major ecological determinant in vivo. he retention time of only a few minutes for saliva in the oral cavity dictates that oral bacteria must adhere before they can grow. he fact that certain surfaces were selected by specific oral bacteria was a concept first known with oral bacteria and later applied to other medical pathogens such as the group A streptococci (Gibbons, 1996). Comprehensive studies of many sites in healthy subjects he distribution of bacterial species on several oral surfaces in healthy subjects has been conducted in depth by using molecular techniques (Aas, et al., 2005; Mager, et al., 2003).

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he first study (Mager, et al., 2003) employed a checkerboard DNA–DNA hybridization technique (Socransky, et al., 1994) where the genomic DNA of 40 commonly cultured oral species were used as probes to detect the presence of the species in the sample of interest. Samples from eight oral epithelial surfaces, saliva, supragingival plaque, and subgingival plaque were lyzed; the released DNA was applied in respective parallel lanes and UV-bound to a nylon membrane. he probe DNA was added to parallel lanes that were perpendicular to the sample lanes (hence checkerboard). By comparison to publications 20 years earlier, this report was a tour de force. Two clusters of sites from the 225 systemically healthy adult subjects were identified: cluster 1 comprised saliva, lateral and dorsal tongue surfaces, and cluster 2 comprised the ventral tongue surface, mouth floor, as well as the buccal, hard palate, vestibule lip and attached gingival surfaces. he mean proportions of Streptococcus mitis, Streptococcus oralis and Selenomonas noxia were significantly higher in cluster 2 than cluster 1. On the other hand, Veillonella parvula, Prevotella melaninogenica, Eikenella corrodens, Neisseria mucosa, Actinomyces odontolyticus, Fusobacterium periodonticum, Fusobacterium nucleatum ss. vincentii and Porphyromonas gingivalis were in significantly higher proportions in cluster 1. In fact, when considering single locations in the mouth, the proportions of 34 of the 40 species differed markedly. Notably, all 34 species were found on all surfaces including samples from teeth, which were distinguished by their increased proportions of Actinomyces naeslundii, Actinomyces israelii, and Actinomyces gerencseriae. However, Streptococcus sanguinis, Streptococcus anginosus, Streptococcus intermedius, and Streptococcus gordonii colonized tooth surfaces in comparable proportions to their proportions in saliva and on soft tissue surfaces; whereas, S. oralis, S. mitis, and S. constellatus colonized soft tissues and were present in saliva in higher proportions than the samples from teeth. his flood of data indicates that bacterial species colonize specific surfaces, and the data suggest that the soft tissues may serve as a reservoir for the bacteria that colonize hard surfaces. Secondly, these data indicate that the species found on teeth, either supragingivally or subgingivally are different from the species found on soft tissue surfaces. hus, biofilms formed on enamel are polymicrobial and are acquired from the same sources that bathe other oral surfaces. Yet, importantly, the proportions of the species acquired from the inocula and colonizing hard surfaces are distinct from the proportions of the species in the inocula and the proportions colonizing the soft tissue surfaces, indicating that the mixed-species biofilm communities on hard tissue are not formed randomly through non-specific accretion of cells from saliva or other inoculum sources. Furthermore, in support of this idea is the fact that similar compositions of initial plaque communities re-occur each day between oral hygiene regimens (Diaz, et al., 2006). he second study (Aas, et al., 2005) used culture-independent molecular techniques. Samples from nine sites from five healthy subjects were analyzed. he seven soft tissue sites were dorsum of the tongue, lateral sides of the tongue, buccal fold, hard palate, soft palate, labial gingival, and tonsils plus the two supragingival and subgingival dental plaques from teeth. Bacteria in the sample were lyzed, and the 16S rRNA genes were amplified by PCR and subsequently cloned and sequenced. A total of 2589 clones yielded 141 predominant species. As found in previous studies, several species were site-specific, while other species were subject specific and detected at most sites sampled. Actinomyces spp., S. sanguinis and S. gordonii were found preferentially on teeth, whereas S. salivarius was more frequently

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found on the tongue dorsum. Streptococcus australis and Gemella sanguinis did not colonize hard tissue. S. mitis was found in all subjects and at all sites sampled. Besides Streptococcus, the commonly detected genera in all subjects and at most sites were Gemella, Granulicatella and Veillonella. Although, collectively, more than 700 bacterial phylotypes have been found in the oral cavity in this study and previous studies, the number of phylotypes per person as reported in this study ranged from 34 to 72 (Aas, et al., 2005). Notably, more than 50% of the phylotypes have not yet been cultivated, however, that characteristic does not make them inherently less significant than the cultured species. Species such as Porphyromonas gingivalis, Tannerella forsythia, and Treponema denticola, which are associated with periodontal disease, were not seen. Likewise, species such as Streptococcus mutans, Lactobacillus spp., and Bi dobacterium spp., which are associated with dental caries, were not seen. Clearly, there is a distinctive bacterial species community in disease conditions that is not found easily in biofilms present in healthy oral cavities. Bioilms in health and disease Succession of species is tightly integrated with cell–cell adherence To relate in vitro analyses to in vivo conditions is the goal of basic science research in its attempt to connect with clinical studies. Occasionally, basic science research and clinical research proceed concurrently, and results from those two research approaches can be compared and integrated. To understand the totally intertwined relationship between succession of species and adherence requires a two-pronged analysis. Studies of coaggregation in vitro by using more than 1000 isolates from dental plaque, mostly cultured from subgingival samples, indicated that all oral bacteria adhere to other species (Kolenbrander, 1988). At the same time that those studies were revealing the coaggregation potential in vitro, bacteria were being cultured and identified from clinical sites (Loesche, et al., 1985; Moore, et al., 1987; Moore, et al., 1985; Moore, et al., 1983; Moore, et al., 1982; Socransky, et al., 1977). Collectively, these data indicated that a succession of bacteria developed in two ways: (i) on teeth with respect to time after the teeth had been professionally cleaned, and (ii) in the progression of disease from health through gingivitis to periodontal disease. A connection among these sets of data was proposed in 1993 (Kolenbrander and London, 1993). he connection was centered on the fact that coaggregation partnerships occurred selectively between the species of bacteria found together during the species succession associated with time after tooth cleaning and with progression of disease status. his proposal has been validated in various ways since then. For example, an extensive clinical analysis was reported in 1998 (Socransky, et al., 1998), which combined two aspects of bacterial communities in vivo. he results demonstrated that the bacterial species in the oral cavity undergo a succession from health through gingivitis to periodontal disease, and the study showed that certain groups of species are found at healthy sites, that other groups of species are found at periodontally diseased sites, and, based on statistically significant associations between species, that these species of bacteria can be organized as clusters. hese clusters of species were designated with different colors; the yellow complex was found at healthy sites and consisted solely of streptococci, the principal initial colonizers

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of freshly cleaned teeth; the purple complex, also associated with healthy sites contained Veillonella parvula and Actinomyces odontolyticus, two species found frequently within a few hours after teeth are cleaned; the green cluster contained three species of Capnocytophaga, Eikenella corrodens, Campylobacter concisus, and serotype a of Actinobacillus actinomycetemcomitans; whereas, A. naeslundii was an outlier, indicating that it associated with all clusters, including the orange cluster that contained fusobacteria and the red cluster that contained Porphyromonas gingivalis, Tannerella forsythia, and Treponema denticola (Socransky, et al., 1998). hese results were obtained with the checkerboard DNA–DNA hybridization technique. An earlier large scale investigation of cultured isolates from clinical samples taken from healthy and diseased sites yielded essentially the same outcome in that healthy sites contained predominantly streptococci and actinomyces, and a succession of different species accompanied the progression of disease status at the clinical sites (Moore and Moore, 1994). Interestingly, the checkerboard analysis showed conclusively that, in vivo, fusobacteria (orange complex) preceded the periodontal pathogens in the red complex (Socransky, et al., 1998). he culturing method also indicated that fusobacteria were the dominant Gram-negative species throughout the progression of disease status (Moore and Moore, 1994). he connection between these two large-scale studies of the species associated with dental plaque and the large-scale study of the coaggregation partnerships exhibited by freshly isolated species is that initial colonizers of teeth coaggregate with each other and with fusobacteria; whereas the late colonizers associated with disease coaggregate with fusobacteria but rarely with each other or with initial colonizers (Kolenbrander and London, 1993). hus, the fusobacteria precede the late colonizers and act as a coaggregation bridge that mediates the adherence of the red complex constituents. he role of coaggregation in vivo has been investigated and shown to be involved in the initial colonization of streptococci and actinomyces on enamel (Palmer, et al., 2003). hus, multi-species biofilms in vivo appear to involve the cell–cell specific interactions of coaggregation that assist in the succession of bacterial species associated with the temporal appearance of species each day between oral hygiene treatments and with the temporal appearance of species associated with the progression of disease that occurs over periods of several years to decades. Microbial diversity in human oral lora and succession of species Starting with a professionally cleaned enamel surface, the temporal succession of species forming supra-gingival plaque is easily documented. Because the oral cavity is a flowing environment with saliva bathing all surfaces, except the interproximal contact, it is likely that saliva is a primary conduit for movement of bacteria. his may be especially true for early colonizers, which are non-motile. If so, then the composition of bacterial species in saliva will be a sum of the variety of species on the oral surfaces. By using the checkerboard DNA–DNA hybridization assay, the bacterial species found in a subject’s saliva were compared with the early colonizers at 0, 2, 4, and 6 hours for each of 15 healthy subjects (Li, et al., 2004). Actinomyces spp. were the most common species at 0 and 2 hours and their numbers did not increase at the later time points. On the other hand, S. mitis and S. oralis increased to become the predominant species at 4 and 6 hours. hese species as well as members of the green and orange complexes (Socransky, et al., 1998) were present in

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saliva but poorly represented in dental plaque biofilms at these early times, indicating that the succession of members of these species into biofilm communities had not yet occurred. he three presumed periodontal pathogens (red complex), while detectable in saliva in low numbers, were present in extremely low levels or undetectable on oral surfaces throughout the experiment, again indicating the absence of this succession in these healthy subjects. Instead, these data describe a distinct bacterial distribution occurring in succession from saliva to biofilms that could occur through coaggregation (Kolenbrander and London, 1993; Kolenbrander, et al., 2006). he most abundant species found in these early dental plaques are members of the Actinomyces and purple and yellow complexes, which are considered beneficial bacteria (Socransky, et al., 1998). he first comprehensive analysis of the microbial diversity present in human subgingival plaque was conducted by PCR-amplification of 16S rRNA genes, subsequent cloning and sequencing (Paster, et al., 2001). About 60% of the 2522 clones analyzed were in 132 known species, but 215 were novel phylotypes. Previous reports of PCR-amplified samples and microbial diversity were narrower in scope (Choi, et al., 1994; Dymock, et al., 1996; Kroes, et al., 1999; Sakamoto, et al., 2000), but they were all in general agreement that many of the clones are known species but more were novel phylotypes. he comprehensive study involved healthy subjects as well as subjects with refractory periodontitis, adult periodontitis, human immunodeficiency virus periodontitis, and acute necrotizing ulcerative gingivitis (Paster, et al., 2001). he known putative periodontal pathogens P. gingivalis, T. forsythia, and T. denticola were identified in multiple subjects but usually as a minor component of subgingival plaque. In addition to these three species, 27 other species were identified as putative pathogens in subgingival plaque; these were found in at least four diseased subjects and not in any healthy subjects. hey included treponemes, fusobacteria, and Atopobium rimae among the most frequently isolated clones. Two phyla, TM7 and Deferribacteres, which do not have a cultured oral representative, were found. Together, these molecular studies clarified the breadth of cultured and not-yet-cultured species in the human oral cavity, and they showed that results of molecular methods were congruent with results by culturing bacteria with regard to the identity of species found. Compared with culture methods, the results from these molecular methods confirmed and expanded the distinction between species found in healthy sites versus those found in disease sites as well as confirming the succession of species associated with progression from health to disease. Ecological plaque hypothesis he progression of periodontal disease or caries over a period of several years is consistent with a dynamic relationship between plaque bacteria and the host during times of health and disease (not limited to oral diseases). Marsh (2003) proposed a novel hypothesis called the “ecological plaque hypothesis” to describe this relationship between bacteria and host. A key feature of the ecological plaque hypothesis is that interfering with the selection pressures responsible for pathogen enrichment can limit disease progression rate or possibly prevent disease. Directly targeting pathogens with antimicrobial or anti-adhesive strategies is not the only way to reduce pathogens. Changing the environment from one that promotes potential pathogens to one that promotes commensal bacteria will reduce the likelihood of disease progression. hus, the selection and promotion of pathogenic bacteria

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is directly coupled to changes in the environment. he ecological plaque hypothesis states that any species with relevant traits can contribute to the disease process. Oral diseases need not have a specific etiology; multiple species can be associated with disease. he data collected on the broad variety of species associated with disease and the environmental changes accompanying the progression of disease support the ecological plaque hypothesis, which is embodied in the concept that caries and periodontal diseases arise as a result of environmental perturbation to the ecological site (Marsh, 2003). Polymicrobial nature of periodontal disease As suggested by the ecological plaque hypothesis, several species of bacteria acting together might be the etiologic agents of periodontal disease. he primary etiologic agents might be any combination of species that together can cause disease. At least three species, Treponema denticola, Tannerella forsythia, and Porphyromonas gingivalis, are usually found together and have been placed in the red complex associated with severe periodontal disease (Socransky, et al., 1998). hose results were based on application of DNA probes from cultured organisms. However, using culture-independent, molecular identification methods, the three species were rarely detected (Kumar, et al., 2005). Interestingly, when detected, T. forsythia was associated with periodontal disease. he most numerous genera associated with periodontitis were Peptostreptococcus and Filifacter but others, such as Megasphaera, Desulfobulbus, Campylobacter, Selenomonas, Deferribacteres, Dialister, Catonella, Tannerella, Streptococcus, Atopobium, and Eubacterium were elevated in disease. Many of these genera were also found elevated in earlier studies by using culture methods (Moore and Moore, 1994), indicating that while both culture-dependent and culture-independent molecular methods have inherent respective biases, they can yield congruent results. Collectively, all of these studies tell us that the bacterial communities of mixed species assemble in a non-random, temporal succession that is dictated by tissue tropism, coaggregation, host receptors, and co-evolution with the human host. A second aspect of considering the ecological plaque hypothesis is to use bacterialsuccession information to predict early stages of periodontitis (Tanner, et al., 2006). Subgingival and tongue samples were taken from 141 healthy and early periodontitis adults. Species diversity was measured by two methods: oligonucleotide DNA probes used in reverse-capture checkerboard assay (Becker, et al., 2002; Paster, et al., 1998; Socransky, et al., 1994) and a direct PCR analysis for detection of P. gingivalis and T. forsythia. As was reported by many preceding studies, S. salivarius is the predominant colonizer of the tongue. By reverse-capture checkerboard assay, most species appeared to exhibit tissue tropism: most subgingival species were more frequently detected in subgingival samples, and most tongue-associated species were more frequently detected in tongue samples. Although this site specificity was observed, all species were detected in both sites, as is consistent with the ecological plaque hypothesis. To obtain higher sensitivity in the prediction of early periodontitis, direct PCR analysis of P. gingivalis and T. forsythia was conducted on 124 paired tongue and subgingival samples of 23 healthy, 59 early periodontitis 1, and 42 early periodontitis 2 individuals (Tanner, et al., 2006). he results revealed that higher frequency of detection of P. gingivalis from both sample sites and T. forsythia from subgingival samples was associated with early periodontitis. his comprehensive study indicates

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the potential for predicting early disease status of polymicrobial-caused diseases. However, different combinations of several species might form communities that collectively possess the relevant traits to initiate and cause disease. Polymicrobial nature of caries Unlike diseases that are caused by one species, oral diseases such as caries and periodontal disease appear to be associated with a change in the overall ecology of the site. his community-based idea has been gathering support through the analyses by molecular identification methods (Becker, et al., 2002; Corby, et al., 2005). Early childhood caries (ECC) was studied in children aged 2 to 8 years and the bacterial species found were compared to the species found at equivalent sites in healthy children. S. mutans is considered a causative agent of ECC, and indeed it was frequently enriched in numbers at diseased sites, but so were Actinomyces gerencseriae, Bi dobacterium sp. Clone CX010, Veillonella dispar or V. parvula (cannot be distinguished by 16S rRNA gene analysis), S. salivarius, S. constellatus, S. parasanguinis and Lactobacillus fermentum in order of decreasing cell numbers (Becker, et al., 2002). In contrast, S. sanguinis was associated with intact enamel and health. hese data strongly suggest that multiple species are associated with ECC. Collection and analysis of undisturbed plaque: spatial relationships between bacteria in vivo An important component of multispecies biofilm community analysis is the spatial relationship between the community members. Metabolic interdependence among oral bacteria has been investigated in vitro in well-mixed systems (Bradshaw, et al., 1994). he consequences of these interactions would be more pronounced in a diffusion-limited environment such as a plaque biofilm. herefore, the proximity of one physiology (phenotype) to another may play a key role in the ability of both phenotypes to flourish within the biofilm, and the maintenance of spatial relationships during biofilm sampling is paramount to detection and analysis of these interactions. Typically, oral biofilm material is obtained by placing a substratum in the oral cavity, allowing the biofilm to develop naturally on the substratum, then retrieving the substratum that now bears the intact biofilm. Substrata vary, but for tooth surface biofilms, the substratum-of-choice is enamel: human enamel or, when large amounts of enamel are required, bovine enamel. In the case of subgingival biofilms, a smaller and less obtrusive substratum is called for. he following paragraphs describe studies on intact supra- and subgingival biofilms developed on retrievable substrata. Early studies with retrievable enamel piece model A stent that holds enamel pieces is commonly used for supragingival oral biofilm investigations. Nyvad and coworkers, in an early example of time-resolved oral biofilm analysis, took a classical bacteriological approach to taxonomy coupled with electron microscopic observation of spatial relationships (Nyvad, 1993; Nyvad and Fejerskov, 1987a; Nyvad and Fejerskov, 1987b; Nyvad and Kilian, 1987). heir bacteriological data were obtained by sonication of the enamel piece in a small volume of cysteine-containing nutrient broth, followed by dilution-plating onto tryptone/yeast-extract/blood medium and anaerobic incubation for 4 days at 37oC. Colonies were characterized based on morphology and Gram stain-

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ing into five major groups: streptococci, actinomyces, G+ cocci (other than streptococci), G– cocci and “unidentified.” he streptococci were further subdivided based on physiologic characteristics into S. sanguis (now S. sanguinis), S. oralis, S. mitis (arg+), S. mitis (arg–), S. salivarius, and “other”; the actinomyces were further divided based on physiologic characteristics into A. naeslundii, A. viscosus, and “other.” While certain of their taxonomic criteria have been eliminated or otherwise changed as a result of molecular studies, this study pioneered a time-resolved complete-community approach to supragingival plaque and yielded conclusions that are to this day cornerstones in the concepts of plaque development. he study demonstrated that plaque biomass (as total colony counts) developed at essentially the same rate from individual to individual, and that a plateau was reached around 12 hours after stent insertion (Nyvad and Kilian, 1987). heir experimental system is probably best viewed as an analogue to smooth surface plaque: the plaque that develops away from the gingival margin on broad tooth faces. Smooth surface plaque is particularly subject to wear by the tongue and cheek, therefore it is thinner than the plaque that develops at other sites (such as interproximal locations). he results of Nyvad and colleagues place a 12-hour maturation time on this plaque and provided evidence that the maximum plaque thickness at this time was approximately 10 micrometers (Nyvad and Fejerskov, 1987b). Further, the 12-hour maturation time coincided with appearance of broad continuous sheets of bacteria (a confluent biofilm) on the enamel (Nyvad and Fejerskov, 1987a). Streptococci accounted for 60–90% of all bacteria isolated during the first 12 hours. he next most prominent group was the actinomyces, which represented 2–12% of the population, and similar amounts of G– cocci were found. Over the next 12 hours, streptococci decreased somewhat and actinomyces increased. he other major groups likewise increased slightly. Within the streptococci, S. sanguis increased in proportion whereas S. salivarius and arg+ S. mitis decreased. hese bacteriological results set the paradigm for early supragingival plaque development: dominance by streptococci and actinomyces for the first 12 hours, followed over the next 12 hours by changes within the streptococcal population together with a slight increase in non-streptococcal organisms. Electron microscopy confirmed from a morphological standpoint this change in species composition (Nyvad and Fejerskov, 1987a; Nyvad and Fejerskov, 1987b). While certain morphologies can be difficult to distinguish unambiguously with scanning electron microscopy (e.g. streptococci vs. actinomyces), electron microscopy clearly showed that cells of different wall types (G+ vs. G–) abutted one another in the plaque (Nyvad and Fejerskov, 1987b) thereby providing clear evidence for direct interaction between species in the biofilm. Coaggregation role in vivo A similar stent system was used by Palmer and colleagues to demonstrate the occurrence of interspecies recognition (coaggregation) in situ (Palmer, et al., 2003). hey used antibodies that target adhesins or their complementary receptors on bacteria for which coaggregation reactions have been well described in vitro. After extensive testing with cultured bacteria to verify the specificity of the antibodies, they applied the antibodies, together with nucleic acid stains as markers for antibody-unreactive bacteria, to investigate the spatial relationships in undisturbed plaque. Using immunofluorescence and laser confocal microscopy, they demonstrated that many of the cells reactive with the adhesin antibody were juxtaposed with

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cells reactive with the receptor antibody. Further, they showed that cell–cell recognition was important even in the very earliest stages of plaque development (at 4 hours). his study highlighted not only the multi-species nature of plaque biofilms but also the importance of cell–cell recognition in initial plaque development. he data fully supported and extended the conclusions of Nyvad et al. (Nyvad, 1993; Nyvad and Fejerskov, 1987a; Nyvad and Fejerskov, 1987b; Nyvad and Kilian, 1987) with respect to plaque development, while also proving that the coaggregation interactions studied for decades in vitro (Kolenbrander, et al., 2002) are translatable to bacterium–bacterium interactions in natural biofilms (Palmer, et al., 2003). Molecular phylogeny of initial dental plaque he same stent system was used by Diaz and colleagues to explore time-resolved development of initial plaque from a molecular taxonomic standpoint, while simultaneously examining spatial relationships between bacteria identified using ribosome-directed fluorescence in situ hybridization (FISH) (Diaz, et al., 2006). his approach paralleled that taken by Nyvad et al. (Nyvad, 1993; Nyvad and Fejerskov, 1987a; Nyvad and Fejerskov, 1987b; Nyvad and Kilian, 1987), but applied molecular taxonomic standards rather than classical bacteriological techniques. he molecular community analysis was performed by universal-primer PCR amplification of 16S rRNA genes from extracted community DNA. his aspect of the study showed that, at four and at eight hours of biofilm development, Streptococcus spp. comprised 60–80% of the phylotypes present; within these streptococci, S. mitis and S. oralis (cannot be distinguished by 16S rRNA gene analysis) were the dominant organisms and comprised between 25% and 100% of the streptococcal population. Twenty other genera were found in these communities; the most frequently recovered of these were Veillonella, Actinomyces, Neisseria, and Prevotella. Sixty-seven of the 513 sequences analyzed were classified as yet-to-be-cultured phylotypes. he results obtained by Diaz et al. (Diaz, et al., 2006) correlate well with those obtained by Nyvad et al. (Nyvad, 1993; Nyvad and Fejerskov, 1987a; Nyvad and Fejerskov, 1987b; Nyvad and Kilian, 1987). In addition, Diaz et al. (Diaz, et al., 2006) investigated community composition within each of the three subjects in their study and showed variation not only in phylotypes present but also in overall community composition between 4 and 8 hours. hus, while the predominant bacteria in initial plaque are similar between individuals, and while the overall time-line of development between individuals is also similar, it is possible to detect statistically significant differences between individuals in non-streptococcal components of the plaque community. hese differences may presage shifts in the oral flora towards diseaseassociated (caries, gingivitis, periodontitis) communities. he study also examined spatial relationships between all bacteria, streptococci, and Prevotella spp. using FISH (Figure 10.2). A “universal” eubacterial probe (EUB338), was combined with either a genus-level streptococcal probe (STR405) or a genus-level Prevotella probe (PRV392) to show that the vast majority of bacteria on the enamel pieces were streptococci (co-reactive with EUB and STR), and that streptococci as well as prevotellae were seen in association with other (solely EUB-reactive) bacteria. hese data first establish FISH as useful in detecting minor (prevotellae) components of the plaque community and, second, reinforce the interactive nature of phylotypes within the community.

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Figure 10.2 Confocal micrographs of typical multi-generic clusters of cells found on enamel chips at 4 hours (A and B) and 8 hours (C and D) of plaque development. Cells were simultaneously labeled with eubacterial probe EUB338 (red) and either the Streptococcusspeciic probe STR405 (green; A and B) or the Prevotella-speciic probe PRV392 (green; C and D). (A and B) Unidentiied bacterial cells (EUB338 reactive; red) juxtaposed with Streptococcus cells (EUB338 and STR405 reactive; red + green = yellow). (C and D) Unidentiied bacterial cells (EUB338 reactive; red) in association with Prevotella cells (EUB338 and PRV392 reactive; red + green = yellow). Scale bar for all images, 5 Mm. Reprinted from (Diaz, et al., 2006). See also Plate 10.2.

Rapid changes in Veillonella spp. population in vivo In another series of experiments using this stent system, Palmer and collaborators examined temporal changes in the Veillonella strains within an individual (Palmer, et al., 2006). Different populations of veillonellae were identified based on time of isolation: the 4-hour population and the 8-hour population. Using 16S ribosomal RNA gene analysis and ERIC-PCR analysis (a strain-level fingerprinting technique), the veillonellae were shown to cluster based on time of isolation. An antibody was developed which identified a subset of the 8-hour population, and a complementary antibody was developed which recognized veillonellae regardless of time of isolation. When these antibodies were used in immunofluorescence investigations of undisturbed plaque, the 4-hour-old biofilm was shown to harbor exclusively veillonellae that reacted with the broadly reactive antibody, whereas 8hour-old biofilms were shown to contain mostly veillonellae reactive with both antibodies together with a lower number of cells reactive solely with the narrowly reactive (8-hours) antibody (Figure 10.3). hus, the veillonellae present at 4 hours into plaque development differ from those present at 8 hours of biofilm formation. Also, the molecular-identification data combined with the phenotypic data showed that the veillonellae as a whole are mi-

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Figure 10.3 Rapid succession within the Veillonella population of early plaque. Two antibodies were employed simultaneously in these immunoluorescence experiments: one antibody, produced against a Veillonella strain isolated from 4-hour-old plaque, is broadly reactive and also recognizes many strains acquired from 8-hour-old plaque, whereas the second antibody, produced against a subgingival Veillonella isolate, recognizes a narrow range of strains isolated primarily from 8-hour-old plaque. Four-hour old plaque (left panel) contains veillonellae reactive solely with the broadly reactive antibody (green); cells reactive with the narrowly reactive antibody (red) are not seen in this plaque. In contrast, 8-hour-old plaque (right panel) contains veillonellae reactive solely with the narrowly reactive antibody (red) as well as veillonellae co-reactive with the broadly reactive antibody plus the narrowly reactive antibody (green + red = yellow). Thus, a shift in the phenotype (as measured by antibody reactivity) of the veillonellae population within plaque occurs between 4 and 8 hours of bioilm development. Antibody-reactive cells are typically associated with antibody-unreactive cells (blue = Syto 59 nucleic acid stain) showing the multi-species nature of plaque communities. Scale bar = 20 Mm. Reprinted from (Palmer, et al., 2006). See also Plate 10.3.

croheterogeneous, i.e. that differences not detectable at the ribosomal RNA gene sequence level are widespread in this group of organisms and that these differences may be relevant to fitness of particular veillonellae at different times in biofilm development. Lastly, in immunofluorescence experiments on intact plaque that combined a veillonella-directed antibody with an antibody that recognizes a subset of streptococci (those bearing a particular coaggregation-associated cell-surface polysaccharide), it could be shown that veillonellae were in immediate proximity to streptococci. his association has a physiological basis in the absolute requirement of veillonellae for short-chain organic acids, such as those produced by metabolism of streptococci. Retrievable subgingival plaque model Retrieval of intact biofilm from the periodontal pocket is difficult because the types of carriers and substrata are limited to those that cause the patient minimal discomfort in a sensitive area of the oral cavity. he devices must be small, thin, and lacking sharp edges. One device currently in use is a plastic rod around which a PTFE membrane is wrapped (Wecke, et al., 2000). he rod and membrane are inserted into the sulcus; a short length of membrane extends out of the sulcus and is cemented onto the supragingival tooth surface to stabilize the device in the pocket. he devices have been inserted into deep (ca. 8 mm) pockets of periodontally diseased individuals and carried for up to six days. After removal,

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the membrane is unfurled, the adherent biofilm is fixed and the membrane is embedded in resin for sectioning. Sections can then be processed for electron microscopy or for light microscopy. his approach can also accommodate ultra-thin dentin pieces (attached to the PTFE membrane), or gold foil instead of PTFE. Intact biofilms on the substrata can be analyzed with FISH, and single-cell resolution is achievable within the sections if confocal microscopy is used. Wecke et al. (2000) were able to demonstrate a variety of organisms in direct association within these biofilms. One feature of this approach is that, if the orientation of the membrane is maintained during sample workup, a record of the biofilm architecture from top to bottom of the pocket is obtained. In other studies, intact biofilms through removal of the tooth and associated tissues have been obtained. Such samples have been processed and analyzed using immunohistochemical (Noiri, et al., 2001; Noiri, et al., 1997), immunoelectron microscopical (Noiri and Ebisu, 2000), and FISH techniques (Sunde, et al., 2003). Mapping of organismal distribution throughout the entire periodontal pocket is possible (Noiri, et al., 2001). Species colonize preferentially at certain sites on the tooth and associated tissues resulting in multi-species communities composed of the same species but in different proportions, which is consistent with the ecological plaque hypothesis. Analyzing in vivo developed multi-species bioilms in vitro he retrievable-enamel-piece-model is an effective tool for analysis of accumulation rates in vivo and growth rates of the adherent bacteria. Enamel pieces (bovine origin) were placed on six teeth of each of 18 healthy subjects, and an enamel piece was retrieved after 2, 4, 8 and 24 hours (Bloomquist, et al., 1996). Measurements during these first 24 hours of plaque formation were conducted after gently sonicating the retrieved enamel piece, plating the released bacteria on culture media to enumerate different species of bacteria, and using radiolabeled nucleoside incorporation to determine DNA synthesis in the sonically released bacteria. Determinations of the rate and saturation of bacterial adherence to enamel for the initial 2 to 4 hours indicated that accumulation doubled every 12 to 24 minutes indicating an unexpectedly rapid rate of bacterial biofilm formation. Most (81%) of these initial colonizers were streptococci. Actinomyces and veillonellae composed the other species investigated. After the initial 4 hours, doubling of adherence occurred every 60 minutes or less until a population density of 2.5 s 105 to 6.3 s 105 cells per mm2 was obtained. his density was considered saturating. At and below this density, the bacteria, sonically removed from the saliva-coated enamel, incorporated radiolabeled thymidine at low levels per viable cell. A small increase in radionuclide incorporation per cell was seen as densities increased up to 2.0 s 105 cells per mm2. However, a marked increase in the incorporation per cell was seen at the density of 2.5 s 106 to 4.0 s 106 cells per mm2. At higher densities, 4.5 s 106 to 8.5 s 106 cells per mm2, the rate of incorporation dropped to levels seen at the lowest cell density. he increase in density of streptococci and of actinomyces as a function of the increase in total bacterial density was linear. In sharp contrast, the increase in veillonellae as a function of the increase in total bacterial density was bimodal. A rapid increase in density of veillonellae occurred at approximately 5 s 105 total cells per mm2, the saturation density. An interpretation of these observations is that the community metabolism changes at this density to favor the growth of veillonellae. he radionuclide incorporation period was only for 30 minutes; radiolabeled adenine gave similar results to radiolabeled thymidine. Also,

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radiolabeling adherent cells on the enamel piece before sonically releasing the cells yielded similar results to those obtained by labeling cells after sonication. hese analyses report the vital activity of in vivo biofilm bacteria in mixed-species natural populations. In addition, these results show the rapid binding of bacteria to form a biofilm on enamel and they show the importance of density of bacteria in the very early stages of development of multi-species biofilm communities. he apparent cell-density-related marked increase in DNA synthesis suggests that a signal/s was transmitted among the community. he signal/s was never identified, but, because it appeared to be associated with cell density, autoinducer-1 (AI-1; (Fuqua, et al., 1994; Atkinson et al., this volume)) or autoinducer-2 (AI-2; (Schauder, et al., 2001; Wood and Bentley, this volume)) or autoinducer-3 (AI-3; (Walters, et al., 2006)) might be involved. No evidence for AI-1 has been found in oral bacteria (Frias, et al., 2001; Whittaker, et al., 1996), and the presence of AI-3 in oral bacteria has yet to be examined. However, AI-2 has been reported in many oral bacteria (Blehert, et al., 2003; Burgess, et al., 2002; Chung, et al., 2001; Federle and Bassler, 2003; Fong, et al., 2001; Frias, et al., 2001; McNab, et al., 2003). his molecule has been proposed as a universal inter-species signal (Schauder, et al., 2001). Investigations exploring the role of AI-2 have been conducted in vitro and are reviewed by Wood and Bentley (this volume). We showed that AI-2 affected the architecture of S. gordonii mono-culture biofilms (Blehert, et al., 2003) and that mutualism in vitro between A. naeslundii and S. oralis is mediated by AI-2 (Rickard, et al., 2006). It is unlikely that these two species are the only ones that communicate by the interspecies signal AI-2, and therefore it is wise to initiate investigations of other logical communicating partnerships and communities. One way to initiate the search for mutualistic interactions and the role of AI-2 in celldensity regulated gene expression is to use the retrievable enamel piece model, as was done by Liljemark and collaborators (Bloomquist, et al., 1996) and discussed above. We propose that signals such as AI-2 are exchanged in vivo in dental plaque. Furthermore, we propose that coaggregation contributes to AI-2 mediated signaling between species, which in turn contributes to succession of species between oral hygiene procedures and succession of species accompanying progression of disease from health through gingivitis to periodontal disease. Summary he human oral cavity contains numerous ecologically distinct surfaces. Some are bathed in saliva and others in gingival crevicular fluid. Some are composed of non-shedding hard tissue such as enamel (tooth crown) or cementum (root surface) and other surfaces are desquamating soft tissues; some of the latter are keratinized and others are non-keratinized. All surfaces have access to all of the oral bacteria through the salivary and gingival crevicular fluid flow. However, the multispecies biofilms that develop on each surface are unique in the proportions of species that colonize. Many of the species are taxonomically identical, but that may be too simple a statement. Closer examination, as was done with the initial colonizing veillonellae (Palmer, et al., 2006) revealed that the veillonellae, while taxonomically identical are in fact distinct in other properties. Coaggregation has long been known to exist among oral bacteria (Gibbons and Nygaard, 1970), and investigations of more than

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1000 fresh isolates of more than 30 species revealed that all isolates have coaggregation partners (Cisar, et al., 1979; Kolenbrander, 1988; Kolenbrander, et al., 2002; Kolenbrander, et al., 1985; Kolenbrander, et al., 1989; Kolenbrander, et al., 1990; Kolenbrander and London, 1993; Kolenbrander, et al., 2006). he role for coaggregation in vivo in developing multispecies biofilms on enamel (dental plaque) was demonstrated by using highly specific antibodies against complementary coaggregation mediators (receptor polysaccharide on the streptococci and type 2 fimbriae on the actinomyces) and showing the tight juxtaposition of these labels (Palmer, et al., 2003). Culture independent molecular techniques for identification of oral species have given us a new look at the microbial diversity on oral surfaces. Results from these techniques have confirmed and extended the results obtained by culturing. Together, the approaches to in vivo biofilm research in the oral cavity have revealed that colonization of the diverse surfaces is non-random, community composition is repetitive on surfaces that have been cleaned, and multi-species biofilm development involves cell–cell adherence, Acknowledgment his research was supported in part by the Intramural Research Program of the National Institutes of Health, National Institute of Dental and Craniofacial Research. References Aas, J.A., B.J. Paster, L.N. Stokes, I. Olsen, and F.E. Dewhirst. (2005). Defining the normal bacterial flora of the oral cavity. J. Clin. Microbiol. 43, 5721–5732. Becker, M.R., B.J. Paster, E.J. Leys, M.L. Moeschberger, S.G. Kenyon, J.L. Galvin, S.K. Boches, F.E. Dewhirst, and A.L. Griffen. (2002). Molecular analysis of bacterial species associated with childhood caries. J. Clin. Microbiol. 40, 1001–1009. Blehert, D.S., R.J. Palmer, Jr., J.B. Xavier, J.S. Almeida, and P.E. Kolenbrander. (2003). Autoinducer 2 production by Streptococcus gordonii DL1 and the biofilm phenotype of a luxS mutant are influenced by nutritional conditions. J. Bacteriol. 185, 4851–4860. Bloomquist, C.G., B.E. Reilly, and W.F. Liljemark. (1996). Adherence, accumulation, and cell division of a natural adherent bacterial population. J. Bacteriol. 178, 1172–1177. Bradshaw, D.J., K.A. Homer, P.D. Marsh, and D. Beighton. (1994). Metabolic cooperation in oral microbial communities during growth on mucin. Microbiology 140, 3407–3412. Burgess, N.A., D.F. Kirke, P. Williams, K. Winzer, K.R. Hardie, N.L. Meyers, J. Aduse-Opoku, M.A. Curtis, and M. Cámara. (2002). LuxS-dependent quorum sensing in Porphyromonas gingivalis modulates protease and haemagglutinin activities but is not essential for virulence. Microbiology 148, 763–772. Choi, B.K., B.J. Paster, F.E. Dewhirst, and U.B. Gobel. (1994). Diversity of cultivable and uncultivable oral spirochetes from a patient with severe destructive periodontitis. Infect. Immun. 62, 1889–95. Chung, W.O., Y. Park, R.J. Lamont, R. McNab, B. Barbieri, and D.R. Demuth. (2001). Signaling system in Porphyromonas gingivalis based on a LuxS protein. J. Bacteriol. 183, 3903–3909. Cimasoni, G. 1983. Crevicular Fluid Updated, 2nd edn. Karger, Basel. Cisar, J.O., P.E. Kolenbrander, and F.C. McIntire. (1979). Specificity of coaggregation reactions between human oral streptococci and strains of Actinomyces viscosus or Actinomyces naeslundii. Infect. Immun. 24, 742–752. Corby, P.M., J. Lyons-Weiler, W.A. Bretz, T.C. Hart, J.A. Aas, T. Boumenna, J. Goss, A.L. Corby, H.M. Junior, R.J. Weyant, and B.J. Paster. (2005). Microbial risk indicators of early childhood caries. J. Clin. Microbiol. 43, 5753–5759. Dawes, C., S. Watanabe, P. Biglow-Lecomte, and G. H. Dibdin. (1989). Estimation of the velocity of the salivary film at some different locations in the mouth. J. Dent. Res. 68, 1479–1482.

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Diaz, P.I., N.I. Chalmers, A.H. Rickard, C. Kong, C.L. Milburn, R.J. Palmer, Jr., and P.E. Kolenbrander. (2006). Molecular characterization of subject-specific oral microflora during initial colonization of enamel. Appl. Environ. Microbiol. 72, 2837–2848. Dymock, D., A.J. Weightman, C. Scully, and W.G. Wade. (1996). Molecular analysis of microflora associated with dentoalveolar abscesses. J. Clin. Microbiol. 34, 537–42. Federle, M.J., and B.L. Bassler. (2003). Interspecies communication in bacteria. J. Clin. Invest. 112, 1291–1299. Fong, K.P., W.O. Chung, R.J. Lamont, and D.R. Demuth. (2001). Intra- and interspecies regulation of gene expression by Actinobacillus actinomycetemcomitans LuxS. Infect. Immun. 69, 7625–7634. Frias, J., E. Olle, and M. Alsina. (2001). Periodontal pathogens produce quorum sensing signal molecules. Infect. Immun. 69, 3431–3434. Fuqua, W.C., Winans, S.C., and Greenberg, E.P. (1994). Quorum sensing in bacteria, the LuxR-LuxI family of cell density-responsive transcriptional regulators. J. Bacteriol. 176, 269–275. Gibbons, R.J. (1996). Role of adhesion in microbial colonization of host tissues, a contribution of oral microbiology. J. Dent. Res. 75, 866–870. Gibbons, R.J., and M. Nygaard. (1970). Interbacterial aggregation of plaque bacteria. Arch. Oral Biol. 15, 1397–1400. Gibbons, R.J., and J. van Houte. (1971). Selective bacterial adherence to oral epithelial surfaces and its role as an ecological determinant. Infect. Immun. 3, 567–573. Hannig, M. (1999). Ultrastructural investigation of pellicle morphogenesis at two different intraoral sites during a 24-h period. Clin. Oral Invest. 3, 88–95. Homer, K.A., S. Kelley, J. Hawkes, D. Beighton, and M.C. Grootveld. (1996). Metabolism of glycoproteinderived sialic acid and N-acetylglucosamine by Streptococcus oralis. Microbiology 142, 1221–1230. Itoiz, M.E., and F.A. Carranza. (2002). he gingiva. In: Newman, M.G., H.H. Takei, and F.A. Carranza (ed.), Clinical Periodontology, 9th ed, pp. 16–35. W.B. Saunders Company, Philadelphia. Kolenbrander, P.E. (1988). Intergeneric coaggregation among human oral bacteria and ecology of dental plaque. Annu. Rev. Microbiol. 42, 627–656. Kolenbrander, P.E., R.N., andersen, D.S. Blehert, P.G. Egland, J.S. Foster, and R.J. Palmer, Jr. (2002). Communication among oral bacteria. Microbiol. Mol. Biol. Rev. 66, 486–505. Kolenbrander, P.E., R.N., andersen, and L.V. Holdeman. (1985). Coaggregation of oral Bacteroides species with other bacteria, central role in coaggregation bridges and competitions. Infect. Immun. 48, 741–746. Kolenbrander, P.E., R.N., andersen, and L.V. Moore. (1989). Coaggregation of Fusobacterium nucleatum, Selenomonas flueggei, Selenomonas infelix, Selenomonas noxia, and Selenomonas sputigena with strains from 11 genera of oral bacteria. Infect. Immun. 57, 3194–3203. Kolenbrander, P.E., R.N., andersen, and L.V. Moore. (1990). Intrageneric coaggregation among strains of human oral bacteria, potential role in primary colonization of the tooth surface. Appl. Environ. Microbiol. 56, 3890–3894. Kolenbrander, P.E., and J. London. (1993). Adhere today, here tomorrow, oral bacterial adherence. J. Bacteriol. 175, 3247–3252. Kolenbrander, P.E., R.J. Palmer, Jr., A.H. Rickard, N.S. Jakubovics, N.I. Chalmers, and P.I. Diaz. (2006). Bacterial interactions and successions during plaque development. Periodontology 2000 42, 1–33. Kroes, I., P.W. Lepp, and D.A. Relman. (1999). Bacterial diversity within the human subgingival crevice. Proc. Natl. Acad. Sci. USA 96, 14547–14552. Kumar, P.S., A.L. Griffen, M.L. Moeschberger, and E.J. Leys. (2005). Identification of candidate periodontal pathogens and beneficial species by quantitative 16S clonal analysis. J. Clin. Microbiol. 43, 3944–3955. Li, J., E.J. Helmerhorst, C.W. Leone, R.F. Troxler, T. Yaskell, A.D. Haffajee, S.S. Socransky, and F.G. Oppenheim. (2004). Identification of early microbial colonizers in human dental biofilm. J. Appl. Microbiol. 97, 1311–1318. Loesche, W.J., S.A. Syed, E. Schmidt, and E.C. Morrison. (1985). Bacterial profiles of subgingival plaques in periodontitis. J. Periodontol. 56, 447–456. Mager, D.L., L.A. Ximenez-Fyvie, A.D. Haffajee, and S.S. Socransky. (2003). Distribution of selected bacterial species on intraoral surfaces. J. Clin. Periodontol. 30, 644–654. Marsh, P.D. (2003). Are dental diseases examples of ecological catastrophes? Microbiology 149, 279– 294.

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McNab, R., S.K. Ford, A. El-Sabaeny, B. Barbieri, G.S. Cook, and R.J. Lamont. (2003). LuxS-based signaling in Streptococcus gordonii, autoinducer 2 controls carbohydrate metabolism and biofilm formation with Porphyromonas gingivalis. J. Bacteriol. 185, 274–284. Moore, L.V., W.E. Moore, E.P. Cato, R.M. Smibert, J.A. Burmeister, A.M. Best, and R.R. Ranney. (1987). Bacteriology of human gingivitis. J. Dent. Res. 66, 989–995. Moore, W.E., L.V. Holdeman, E.P. Cato, R.M. Smibert, J.A. Burmeister, K.G. Palcanis, and R.R. Ranney. (1985). Comparative bacteriology of juvenile periodontitis. Infect. Immun. 48, 507–519. Moore, W.E., L.V. Holdeman, E.P. Cato, R.M. Smibert, J.A. Burmeister, and R.R. Ranney. (1983). Bacteriology of moderate (chronic) periodontitis in mature adult humans. Infect. Immun. 42, 510–515. Moore, W.E., L.V. Holdeman, R.M. Smibert, I.J. Good, J.A. Burmeister, K.G. Palcanis, and R.R. Ranney. (1982). Bacteriology of experimental gingivitis in young adult humans. Infect. Immun. 38, 651–667. Moore, W.E.C., and L.V.H. Moore. (1994). he bacteria of periodontal diseases. Periodontology 5, 66–77. Noiri, Y., and S. Ebisu. (2000). Identification of periodontal disease-associated bacteria in the “plaque-free zone.” J. Periodontol. 71, 1319–1326. Noiri, Y., L. Li, and S. Ebisu. (2001). he localization of periodontal-disease-associated bacteria in human periodontal pockets. J. Dent. Res. 80, 1930–1934. Noiri, Y., K. Ozaki, H. Nakae, T. Matsuo, and S. Ebisu. (1997). An immunohistochemical study on the localization of Porphyromonas gingivalis, Campylobacter rectus and Actinomyces viscosus in human periodontal pockets. J. Periodontal Res. 32, 598–607. Nyvad, B. (1993). Microbial colonization of human tooth surfaces. Apmis 101, 7–45. Nyvad, B., and O. Fejerskov. (1987a). Scanning electron microscopy of early microbial colonization of human enamel and root surfaces in vivo. Scand. J. Dent. Res. 95, 287–296. Nyvad, B., and O. Fejerskov. (1987b). Transmission electron microscopy of early microbial colonization of human enamel and root surfaces in vivo. Scand. J. Dent. Res. 95, 297–307. Nyvad, B., and M. Kilian. (1987). Microbiology of the early colonization of human enamel and root surfaces in vivo. Scand. J. Dent. Res. 95, 369–380. Palmer, R.J., Jr., P.I. Diaz, and P.E. Kolenbrander. (2006). Rapid succession within the Veillonella population of a developing human oral biofilm in situ. J. Bacteriol. 188, 4117–4124. Palmer, R.J., Jr., S.M. Gordon, J.O. Cisar, and P.E. Kolenbrander. (2003). Coaggregation-mediated interactions of streptococci and actinomyces detected in initial human dental plaque. J. Bacteriol. 185, 3400–3409. Palmer, R.J., Jr., K. Kazmerzak, M.C. Hansen, and P.E. Kolenbrander. (2001). Mutualism versus independence, strategies of mixed-species oral biofilms in vitro using saliva as the sole nutrient source. J. Bacteriol. 69, 5794–5804. Paster, B.J., I.M. Bartoszyk, and F.E. Dewhirst. (1998). Identification of oral streptococci using PCRbased, reverse-capture, checkerboard hybridization. Meth. Cell Sci. 20, 223–231. Paster, B.J., S.K. Boches, J.L. Galvin, R.E. Ericson, C.N. Lau, V.A. Levanos, A. Sahasrabudhe, and F.E. Dewhirst. (2001). Bacterial diversity in human subgingival plaque. J. Bacteriol. 183, 3770–3783. Rickard, A.H., R.J. Palmer, Jr., D.S. Blehert, S.R. Campagna, M.F. Semmelhack, P.G. Egland, B.L. Bassler, and P.E. Kolenbrander. (2006). Autoinducer 2, a concentration-dependent signal for mutualistic bacterial biofilm growth. Mol. Microbiol. 60, 1446–1456. Sakamoto, M., M. Umeda, I. Ishikawa, and Y. Benno. (2000). Comparison of the oral bacterial flora in saliva from a healthy subject and two periodontitis patients by sequence analysis of 16S rDNA libraries. Microbiol. Immunol. 44, 643–52. Scannapieco, F.A. (1994). Saliva-bacterium interactions in oral microbial ecology. Crit. Rev. Oral Biol. Med. 5, 203–248. Schauder, S., K. Shokat, M.G. Surette, and B.L. Bassler. (2001). he LuxS family of bacterial autoinducers, biosynthesis of a novel quorum-sensing signal molecule. Mol. Microbiol. 41, 463–476. Socransky, S.S., A.D. Haffajee, M.A. Cugini, C. Smith, and R.L. Kent, Jr. (1998). Microbial complexes in subgingival plaque. J. Clin. Periodontol. 25, 134–144. Socransky, S.S., A.D. Manganiello, D. Propas, V. Oram, and J. van Houte. (1977). Bacteriological studies of developing supragingival dental plaque. J. Periodont. Res. 12, 90–106. Socransky, S.S., C. Smith, L. Martin, B.J. Paster, F.E. Dewhirst, and A.E. Levin. (1994). Checkerboard DNA–DNA hybridization. Biotechniques 17, 788–792.

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Sunde, P.T., I. Olsen, U.B. Gobel, D. heegarten, S. Winter, G.J. Debelian, L. Tronstad, and A. Moter. (2003). Fluorescence in situ hybridization (FISH) for direct visualization of bacteria in periapical lesions of asymptomatic root-filled teeth. Microbiology 149, 1095–1102. Tanner, A.C.R., B.J. Paster, S.C. Lu, E. Kanasi, R. Kent, Jr., T. Van Dyke, and S.T. Sonis. (2006). Subgingival and tongue microbiota during early periodontitis. J. Dent. Res. 85, 318–323. Van Houte, J., R.J. Gibbons, and S.B. Banghart. (1970). Adherence as a determinant of the presence of Streptococcus salivarius and Streptococcus sanguis on the human tooth surface. Arch. Oral Biol. 15, 1025–1034. Walters, M., M.P. Sircili, and V. Sperandio. (2006). AI-3 synthesis is not dependent on luxS in Escherichia coli. J. Bacteriol. 188, 5668–5681. Wecke, J., T. Kersten, K. Madela, A. Moter, U.B. Göbel, A. Friedmann, and J.P. Bernimoulin. (2000). A novel technique for monitoring the development of bacterial biofilms in human periodontal pockets. FEMS Microbiol. Lett. 191, 95–101. Whittaker, C.J., C.M. Klier, and P.E. Kolenbrander. (1996). Mechanisms of adhesion by oral bacteria. Annu. Rev. Microbiol. 50, 513–552.

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Bioilms as Refuge against Predation Carsten Matz

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Abstract Bacterial growth and survival in the environment as well as in association with human hosts are constrained by the action of phagocytic eukaryotic cells. Phagocytic predation on bacteria by host immune cells shares a number of cellular mechanisms with free-living protozoa. In and outside the human host, bacteria growing in biofilms appear to be less vulnerable to phagocytic predators than planktonic cells. Widespread resistance against predators is mediated by the interplay of biofilm-specific traits such as substratum adherence, exopolymer production, cellular cooperation, inhibitor secretion, and phenotypic variation. Selective predation is suggested to promote bacterial life in the biofilm niche and to govern structure-function relationships. here is increasing evidence that some of the pathogenicity traits may have their origin specifically in successful antipredator adaptations. Parallel selective pressures in and outside the human host may result in cross-adaptations of bacterial pathogens. Introduction Biofilm research today mostly deals with the question how to control problematic biofilms in medical and industrial settings. Particularly, researchers from different backgrounds try to understand biofilms as the preferred life style in persistent human infections and as pathogen reservoirs in the environment. hese studies are tightly intertwined with the search for intervention points for prevention and therapy. From a medical perspective, the key characteristic of bacterial biofilms is their tolerance to host defenses and antimicrobials. While antibiotic resistance poses a serious threat to human health in the 21st century, it needs to be stressed that the primary cause for persistent biofilm infections is the fact that intrinsic control mechanisms by the immune system do not unfold fully or efficiently. Similarly, the observation that bacterial biomass and activity in many natural ecosystems preferentially accumulate in association with surfaces raises the question whether the natural loss factors might be compromised by the biofilm mode-of-life. A key mortality factor in the control of bacterial populations is the uptake and killing of bacteria by phagocytic eukaryotic cells—no matter if we look at the microbial food web of natural ecosystems or at the host immune defense against bacterial pathogens. he antagonistic interaction describing the consumption of prey individuals of another species is generally defined as predation. From an ecological perspective, the control of pathogen

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growth at the stage of the non-specific, constitutive host response—as commonly represented by amoeboid phagocytes—is analogous to that of a microbial predator-prey system (Levin and Antia, 2001). Given the importance of predatory control mechanisms in and outside the human host, this chapter attempts to synthesize current knowledge to present a conceptual framework of the role of predator-prey interactions on biofilm formation and persistence. Predatory agents in environment and disease Before we can dwell on the question how bacterial biofilms cope with phagocytic predation, a brief discourse seems in place on who the predatory agents are that bacteria are confronted with. Interestingly, the major predatory pathway in and outside human hosts is mediated by the same cellular process: phagocytosis. Phagocytosis constitutes not only the primary line of host innate and adaptive defense against incoming microbial pathogens but is also employed by phagotrophic microeukaryotes, the protozoa, which are the primary consumers of bacteria in most soil, freshwater and marine ecosystems. Free-living protozoa in environmental bioilms he presence of protozoa in close association with environmental bacterial biofilms is well accepted. Surface-associated protozoan habitats range from sediments, rocks, water pipes and filters to dental unit waterlines and the oral cavity. Not only do protozoan numbers associated with biofilm communities often exceed those found in surrounding plankton, the number of protozoan taxa observed in natural biofilms communities also reveals great protozoan diversity (Arndt et al., 2003). Protozoa exhibit a variety of mechanisms to capture and engulf bacterial prey leading to a considerable diversification of protozoan morphologies, commonly grouped into flagellates, ciliates, and amoebae. Biofilm-associated protozoa comprise all three free-living groups, all of which have been shown to efficiently graze on surface-associated bacteria. Flagellates and ciliates contain many feeding types that are specialized on suspended bacteria and some that preferably feed on surface-bound bacterial prey; amoebae almost exclusively feed on biofilm bacteria. For instance, the kinetoplastid flagellate Rhynchomonas nasuta has been reported to feed on attached Pseudomonas spp. at rates between 13 and 120 bacteria per flagellate and hour (Boenigk and Arndt, 2000; Matz et al., 2004a). Ingestion rates by the hypotrich ciliate Euplotes sp. feeding on adherent Vibrio natriegens and Pseudomonas fluorescens were reported to be 120 and 882 bacteria per ciliate and hour, respectively (Lawrence and Snyder, 1998). Amoebae species such as Hartmanella cantabrigiensis, Platyamoeba placida, Saccamoeba limax, Vahlkamp a avara were shown to ingest attached Escherichia coli at rates of 15 to 440 bacteria per amoeba and hour (Heaton et al., 2001). A thorough review on protozoan grazing rates in freshwater biofilms has recently been published (Parry, 2004). Host immune cells and bioilm infection he immune response to microbial pathogens relies on both innate and adaptive components. A major mechanism for the destruction of bacteria that have invaded the host is killing by professional phagocytes (macrophages and neutrophil leukocytes). Resident macrophages and neutrophils constitute the primary line of innate defense against most

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bacterial pathogens by providing a means for their removal and destruction. Macrophages are found in all body tissues, where they serve as sentinels in wait for pathogens; the invaders shed a variety of chemotactic agents that alert the resident macrophages to the infection. Neutrophils or polymorphonuclear leukocytes (PMNs) are the first cells recruited from the bloodstream to sites of infection and are an essential component of the acute inflammatory response. One of the most classical examples of biofilms in human disease is the chronic lung infection of cystic fibrosis (CF) patients by P. aeruginosa. he inflammatory response to chronic P. aeruginosa lung infections is mainly characterized by the constant influx of PMNs. Analysis of bronchoalveolar lavage fluid has shown that the number of PMNs recovered from the lungs of patients with CF is 1000 times higher than that recovered from non-infected lungs (Brown and Kelly, 1994). Although PMNs have been described to efficiently feed on adherent bacteria (Lee et al., 1983; Hayashi et al., 1986), their significant phagocytic and secretory arsenal of toxic oxygen species, degrading enzymes, defensins, and lipid inflammatory mediators appears ineffective in combating P. aeruginosa infections of the lung. In fact, PMNs contribute with these mediators to the deterioration of lung tissue that is characteristic of inflammatory processes in CF lungs (Koch and Hoiby, 1993). Moreover, PMNs constitute the major leukocytes present in the blood and acutely inflamed tissue and would be expected to respond to growth of biofilms on surfaces implanted in the vasculature or other tissues. Hence, it would be important to understand how the cellular host defense interacts with biofilms under controlled conditions. For the sake of simplicity, this review concentrates mostly on components of the innate immune response; clearly humoral and adaptive immune response need to be included in future considerations. Shared predatory mechanisms between protozoa and immune cells Phagocytosis is an ancient eukaryotic feature; the first eukaryotic cell was likely to be phagotrophic as phagocytosis is thought to be essential for the uptake of the A-proteobacterial symbiont, from where the mitochondria evolved (Cavalier-Smith, 2002). Hence, bacteria and other prokaryotes should have experienced phagocytic predation since the dawn of eukaryotic life. Phagocytosis can be divided into a series of defined steps (Table 11.1): receptors on the cell membrane recognize and bind bacterial prey; intracellular signals are generated that induce actin polymerization at the site of contact; actin-rich membrane extensions reach out around the particle; the membranes fuse behind the particle, pulling it in toward the center of the cell; and the phagosome matures via a series of membrane fusion and fission events to become a phagolysosome. he phagolysosome is an acidic, hydrolytic compartment in which the bacterium is killed and digested. Locomotion and the ability to sense and respond to shallow gradients of extracellular bacterial signals are remarkably similar in amoebae such as Dictyostelium discoideum and phagocytic immune cells (Devreotes and Zigmond, 1988). Inflammatory signals however appear to be only important for phagocyte chemotaxis. A fundamental step in phagocytosis is the receptor-mediated recognition of an extracellular particle. he recognition mechanisms can be pattern based (non-opsonic phagocytosis) or humoral (opsonic phagocytosis), the latter of which does not apply to amoebae. While scavenger receptors which recognize bacterial surface components (e.g. lipopolysaccharide, lipoteichoic acid) have not yet been described for protozoa, surface lectins such as the mannose receptor are present in both

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Table 11.1 Summary of factors commonly involved in phagocyte attack by immune cells and free-living protozoa Step in predator-prey interaction

Factors involved Macrophages/leukocytes (PMNs)

Amoebae and other protozoa

Chemotaxis

LPS, formylated peptides, PAMPs, chemokines, IL-8

LPS, formylated peptides, lipoteichoic acid, cAMP, nutrients

Phagocytosis

Non-opsonic, opsonic

Non-opsonic

Prey recognition

Non-opsonic: Lectins (e.g. mannose-binding receptor), scavenger receptors, Toll-like receptors Opsonic: Fc-receptors, complement receptors, integrins

Non-opsonic: Lectins (e.g. mannose-binding receptor), scavenger receptors?, Toll-like receptors?

Oxygen-dependent killing

Reactive oxygen/nitrogen species

Reactive oxygen/nitrogen species

Oxygen-independent killing

Antimicrobial peptides/proteins

Antimicrobial peptides/proteins

Signaling

Cytokines

Pheromones?

amoebae and immune phagocytes. However, signals and motifs in prey recognition may differ significantly between protozoa and immune phagocytes as the feeding motivation is somewhat different (recognition of food particles versus recognition of “microbial non-self ” particles). In the phagolysosome the bacteria are killed after exposure to enzymes, antimicrobial peptides and reactive oxygen species (ROS). he arsenal of cytotoxic agents has been traditionally divided into oxygen-dependent and oxygen-independent mechanisms. he latter mechanism employs antimicrobial peptides and proteins such as defensins, lysozyme, elastase and cathepsin. Antimicrobial host defense peptides are widely distributed in animals and plants and are among the most ancient host defense factors (Hoffmann et al., 1999). Most of these peptides have cationic properties that allow interactions with the bacterial cytoplasmic membrane, which usually comprises negatively charged phospholipids. Accordingly, this class of peptides has been termed cationic antimicrobial peptides (CAMPs). Oxygen-dependent killing relies on the generation of oxygen radicals by NADPH oxidase, during the so-called oxidative burst, resulting in the accumulation of reactive oxygen and nitrogen intermediates. he production of both oxygen radicals and membrane-permeabilizing peptides has been described for environmental amoebae (Davies et al., 1991; Herbst et al., 2002). Taken together, the stresses and selective pressures imposed on bacteria by protozoa, specifically by amoebae, and mammalian phagocytes may show significant overlap. To avoid phagocytosis biofilm bacteria need to interfere with at least one of the steps, either with recognition, engulfment, antimicrobial agents or phagocytic activity.

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Are bioilms inherently protected from predation? he early biofilm concept proclaimed that the gel-like state of the biofilm matrix limits the access of antibacterial agents, such as antibodies and phagocytic eukaryotic cells, and inferred that biofilm bacteria are substantially protected from amoebae and immune cells, similar to the resistance against antibiotics (Costerton et al., 1987). Studies on the antiphagocytic effect of encapsulated streptococci and staphylococci and mucoid P. aeruginosa confirmed this concept and led to the common perception of biofilms being generally protected against predation. he finding of Yersinia pestis forming a biofilm that inhibits feeding by the nematode Caenorhabditis elegans has further added to the view that biofilms may be a bacterial defense against predation (Darby et al., 2002). he observation that biofilm bacteria predominate numerically and metabolically in many aquatic ecosystems may also have led to the current conception that the biofilm mode-of-growth is a universal antipredator strategy. his view, however, is challenged by the observation that protozoa not only occur widely in close association with natural biofilm communities, but that they are also able to disrupt biofilm structure and cause biofilm sloughing (Pedersen, 1982; Jackson and Jones, 1991; Weitere et al., 2005). Further support for a certain vulnerability of biofilms to predation comes from the simple fact that within the protozoa several feeding types have evolved that specifically feed on biofilms, such as amoebae, and exhibit a great species diversity. In addition, persistent biofilm infections most often occur in immunocompromised patients, which may suggest that a well-functioning immune defense would be capable to counteract biofilm formation and to avert biofilm manifestation. So, instead of painting a black-and-white picture and regarding biofilms as inherently protective, a more differential view seems to be needed to address the role of biofilms as a refuge against predation. It is important to note that resistance against predation should always be defined in relation to the other bacteria in the prey community. For example, a recent study on Vibrio cholerae demonstrated that biofilm cells resist flagellate grazing while their planktonic counterparts become eliminated (Matz et al., 2005). Findings of lower feeding rates on biofilms compared to suspended bacteria would therefore justify the term “refuge,” as the chances of survival are higher for biofilm bacteria. To understand why biofilms are less efficiently grazed, we need to take a closer look at biofilm-specific traits that are not characteristic for planktonic bacteria. Bioilm-speciic mechanisms of protection When contrasting planktonic and biofilm bacteria, four hallmarks can be noted that are characteristic for the biofilm mode-of-life: Adherence to a substratum; self-encasement into an extracellular matrix; life at high cell densities; and differentiation resulting in population heterogeneity. In the search for the most successful defensive strategy for biofilms, each of the four features is under suspicion to effectively compromise phagocytic attack (see Figure 11.1). Which impact these biofilm-specific traits have on bacterial survival during predation and on the predator’s fitness will be discussed below.

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A – Adherence

B – Matrix

C – Cell density

D – Diversity

• Cell surface physicochemistry • Pili, fimbriae, flagella

• EPS quantity • EPS composition

• Quorum sensing • Exoproducts

• Adaptive mutation • Phase variation

Figure 11.1 Bioilm-speciic traits and their role in antipredator resistance.

Adherence effects he first question is whether adherence to a substratum alone is sufficient to provide effective protection for bacteria facing phagocytic attack (see Figure 11.1A). Bacteria can adhere to almost any surface in any environment by means of cell surface structures, such as fimbriae, pili, and extracellular polymers, or simply by physicochemical interaction forces (see also MacEachran and O’Toole, this volume). he physicochemical basis of adhesion phenomena is the balance of electrostatic and van der Waals’ forces as well as hydrophobic surface interactions that result in either attraction or repulsion between particles. Hydrophobic interactions are strongly attractive and promote adhesion of microorganisms to abiotic surfaces and epithelial cells (Marshall, 1986). Interestingly, high bacterial surface hydrophobicity and low cell surface charge greatly increase the contact probability and the ingestion efficiency of bacterial cells during nonopsonic and opsonic phagocytosis (van Oss, 1978). For instance, binding of P. aeruginosa to phagocytes is inhibited by hydrophobic compounds such as R-nitrophenol, and strains that are hydrophobic and piliated are taken up more readily than hydrophilic, non-piliated

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strains (Speert et al., 1986). While studies on protozoa remain scarce, there are a few findings on planktonic protozoa suggesting similar physicochemical effects: feeding rates of interception-feeding nanoflagellates were lower with decreasing bacterial hydrophobicity (Monger et al., 1999) and increasing bacterial surface charge (Hammer et al., 1999; Matz et al., 2002a). In many cases, bacterial attachment to surfaces is mediated by cellular structures such as pili and flagella. At the same time these structures serve as ligands in phagocyte recognition. For example, pili and flagella were found to increase the susceptibility of P. aeruginosa to non-opsonic phagocytosis (Kelly et al., 1989; Mahenthiralingam and Speert, 1995). Similarly, E. coli possessing mannose-sensitive fimbriae adhered better to the amoeba Acanthamoeba castellanii and human PMNs than non-fimbriated cells (Lock et al., 1987). Apparently, the same bacterial cell surface traits that promote adherence to a substratum also increase contact probabilities with eukaryotic cell membranes and thus the probability of bacterial engulfment by phagocytic cells. Instead of finding a refuge from predation, surface-colonizing bacteria may initially face a higher predatory risk than their planktonic counterparts. Predator-prey studies of higher sessile organisms, such as seaweeds, suggest that sessile prey communities face intense grazing pressure due to the lack of avoidance or escape options (Duffy and Hay, 1990). herefore, high predatory risk and selective grazing pressure on adherent bacterial populations can be expected to drive the necessity to develop protective traits during biofilm formation. Matrix effects During surface colonization cell-to-cell contacts become established between bacteria on the substratum, which assist the cells in further development of the biofilm. he close proximity allows cells to interact and cooperate and thus may open up the door to more complex antipredator adaptations in bacterial biofilms compared to planktonic bacteria. In the course of biofilm development the resident bacteria produce extracellular polymeric substances (EPS), which help create and stabilize the biofilm by gluing the cells together (see also Pamp et al., this volume). Historically, it is this EPS matrix that has been made responsible for the protective nature of bacterial biofilms (see Figure 11.1B). Although the composition of the EPS matrix is expected to be highly diverse within and between bacterial species, three principal properties of the matrix can be identified to contribute to the resistance against phagocytic predation. One of the most obvious mechanisms is that the EPS matrix can form a physical barrier against the attacker. By gluing individual bacterial cells together, EPS allows biofilm bacteria to form defensive units, such as microcolonies, which reach a size beyond the feasible prey size spectrum of size-selective predators. Instead of handling bacterial cells one by one, phagocytic cells are confronted with many times more bacteria. Evidence for such a physical defense mechanisms comes from studies on P. aeruginosa and V. cholerae biofilms co-cultivated with flagellate grazers (Matz et al., 2004a; Matz et al., 2005). he flagellate predators used in theses studies handle single bacterial cells, so that two cells glued together lead to longer handling times and reduced feeding efficiencies, which ultimately favor colonial growth in bacteria. As a consequence, an alginate-overproducing strain of P. aeruginosa formed larger microcolonies in response to grazing (Matz et al., 2004a). Size-selective

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predation by flagellates stimulates the formation of bacterial cell clusters, so-called microcolonies, in a previously single-celled bacterial population (Matz et al., 2004a). Whether microcolony formation is induced mechanically or chemically remains unanswered at this point in time. Challenging P. aeruginosa biofilms with different protozoan feeding types (flagellate vs. ciliate vs. amoeba), however, provided evidence that microcolonies and alginate production alone do not suffice to allow biofilm persistence in the presence of amoebae and biofilm-browsing ciliates (Weitere et al., 2005). Alginate has previously been reported to inhibit phagocytosis, thereby decreasing susceptibility of planktonic P. aeruginosa to human leukocytes and macrophages. While there is evidence that P. aeruginosa mutant biofilms lacking the exopolymer alginate become susceptible to leukocyte killing (Leid et al., 2005), experimental data to support the notion of EPS serving as a “physical barrier” against host immune cells remains scarce. Observations of inactive, “paralyzed” or “frustrated” leukocytes settling on biofilms of P. aeruginosa without disrupting or clearing the biofilm (e.g. Jesaitis et al., 2003) do not exclude the possibility that phagocyte inactivation may be mediated chemically by biofilm-secreted effectors. Recent in vitro studies demonstrate that PMNs can penetrate S. aureus biofilms but are unable to engulf the surrounding bacteria which suggests that PMNs become inactive in this process (Leid et al., 2002). In fact, there is mounting evidence now of biofilms interfering chemically with phagocytic activity and cellular processes (see below). Based on the incomplete data available, one may conclude that the EPS matrix is able to reduce the ability of phagocytic predators to instantly penetrate and phagocytose biofilms. However, it may not offer total protection but forces the attackers to apply extracellular killing mechanisms. PMNs react to intruding foreign organisms either by phagocytosis or secretion; in both cases the PMNs launch a cocktail of antimicrobial agents, in particular free oxygen radicals. he EPS matrix keeps PMNs at a distance and thus provokes an extracellular attack which may not only be imprecise and costly but also less effective as the killing mechanisms in the vacuole. Another functional property of the EPS matrix is to act as a diffusional barrier and interfere with the powerful antibacterial cocktail (toxic ROS and antibacterial peptides/ proteins) released by activated neutrophils. EPS has repeatedly been described to interfere with the oxygen-dependent killing mechanisms of phagocytes by scavenging ROS produced by these cells. In a recent study it was shown that EPS isolated from a mucoid Burkholderia cenocepacia strain could inhibit neutrophil chemotaxis and scavenge neutrophil-derived ROS (Bylund et al., 2006). Similar findings have been presented for alginate from P. aeruginosa (Learn et al., 1987; Simpson et al., 1989). It is interesting to note that despite structural biochemical differences, the EPS material from P. aeruginosa and B. cenocepacia share many properties that could profoundly interfere with the effector functions of immune cells. Similarly, the EPS matrix can also be considered to function as a chemically reactive barrier to antimicrobial peptides (AMPs) charged during oxygen-independent phagocytic attack. Both P. aeruginosa and S. aureus tolerate considerable amounts of cationic AMPs (CAMPs). It has recently been demonstrated that alginate produced by P. aeruginosa promotes the aggregation and sequestration of CAMPs by inducing conformational changes in the peptide structure (Chan et al., 2004). his mechanism is suggested to result in the removal of the AMP from the cytoplasmic membrane, the prime target of AMP action. In

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addition, P. aeruginosa CF isolates produce a variant lipopolysaccharide (LPS) that renders the bacterial cells less susceptible to CAMPs (Ernst et al., 1999). Voung et al. (2004) found that the major matrix component in staphylococcal biofilms, the polysaccharide intercellular adhesin (PIA), protects against AMPs of human skin and of the neutrophil-specific granules LL-37 and B-defensin. It is suggested that the underlying mechanism is based on the cationic character of PIA which causes electrostatic repulsion of the commonly cationic AMPs. A similar mechanism has recently been described for poly-G-DL-glutamic acid (PGA) secreted by S. epidermidis (Kocianova et al., 2005). Taken together, resistance to AMPs in biofilms appears to be based, at least in part, on the interaction with specific extracellular biofilm polymers. A fundamental step in phagocytosis is the receptor-mediated recognition of a prey particle. One potential mechanism in biofilm protection which has received very little attention so far is the recognition barrier, which inhibits the binding of phagocyte-receptors, antibodies or complement on their surface. Biofilm bacteria may be less conspicuous to the immune system because antigens and ligands used by phagocytes may be hidden (see section on adherence effects above). It is conceivable that components of the biofilm matrix may function analogously to the protective function of capsular polymeric substances (CPS). Most clinically important mucosal pathogens make carbohydrate capsules that surround the organisms. Although the carbohydrate composition and biosynthetic pathways vary among organisms, it is well established that such structures protect the organism from complement lysis, antibody deposition, and ultimately from phagocytosis (Celli and Finlay, 2002). he pathogens that use this strategy include Streptococcus pneumoniae, E. coli K1, Klebsiella pneumoniae, Neisseria meningitides and S. aureus. Bacterial S-layers and some types of LPS also can inhibit antibody binding and phagocytosis, presumably by similar mechanisms (Navarre and Schneewind, 1999; van Putten and Robertson, 1995). Another strategy to avoid the immune system surveillance may be to bind host proteins on their surfaces, which disguises the bacterial surface antigens. Many pathogens are capable of binding several host proteins, such as the basement membrane components fibronectin and collagen. It will be interesting to examine the role of the biofilm matrix in averting recognition by phagocytic receptors other components of the host immune response. General knowledge on receptor-mediated phagocytosis in protozoa is much more limited. For amoebae such as Acanthamoeba castellanii and Dictyostelium discoideum, the involvement of mannose-sensitive receptors in phagocytosis have been described (see Table 11.1). Wildschutte et al. (2004) reported that Salmonella enterica serovars are grazed by intestinal amoebae at different rates depending on O-antigen variability at the bacterial cell surface. Interception-feeding flagellates were also found to differentially feed on synthetic prey items with specific biochemical coatings (proteins, polysaccharides) (Matz et al., 2002a). Although the specific mechanisms remain unclear, these studies suggest that receptor-mediated prey recognition should be included in future studies on biofilm-predator interactions in the environment. Cell density effects Besides interfering directly with phagocytic attack, the EPS matrix may also indirectly enhance antipredator fitness of biofilm cells as it promotes the formation of localized high

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density consortia. he localization of cells in close proximity allows bacterial populations to communicate and cooperate via quorum sensing (QS) (Parsek and Greenberg, 2005). By monitoring bacterial cell density and diffusion, QS is thought to synchronize bacterial behavior and to adjust the population response to changing environmental conditions (see Figure 11.1C), particularly during the transition from logarithmic to stationary growth. A central feature of QS regulated gene expression in pathogenic bacteria is the secretion of virulence factors that interfere with eukaryotic cell activity (Winzer and Williams, 2001). QS controlled exofactors are not strictly biofilm-specific but their production is clearly promoted by the localized high cell densities found in sessile matrix-encased communities (see also Atkinson et al., this volume). he semi-diffusible nature of the EPS matrix may further help to generate a hostile chemo-environment for phagocytic attackers. Potential chemical defense mechanisms may include the inhibition of phagocyte chemotaxis, inactivation of phagocytes, and phagocyte lysis via apoptosis or necrosis. Bjarnsholt et al. (2005) recently demonstrated that the limited penetration and elimination of P. aeruginosa wild-type biofilms by PMNs is dependent on a functional lasR/rhlR QS system. One mechanism involved in host defense evasion of P. aeruginosa is the production of extracellular factors such as proteases, toxins and phospholipases (Kharazmi, 1991; Smith and Iglewski, 2003). Two proteases, alkaline protease and elastase have been shown to inhibit chemotaxis, oxidative burst, phagocytosis and other microbicidal activities of phagocytes (Kharazmi et al., 1984a; Kharazmi et al., 1984b). In addition, the pigment pyocyanin induces neutrophil apoptosis and impairs neutrophil-mediated host defenses in vivo (Allen et al., 2005), and the rhamnolipid biosurfactant inhibits macrophage phagocytosis in vitro and in vivo (McClure and Schiller, 1996). QS controlled inhibitors secreted by P. aeruginosa further include cyanide (Pessi and Haas, 2000) with its general activity against most eukaryotic cells as it inhibits mitochondrial cytochrome oxidase c. Besides regulating the expression of exoenzymes and toxins, QS signal molecules can also directly affect phagocytic activity and inflammatory response. he 3-oxo-dodecanoylhomoserine lactone (3-oxo-C12-HSL) signal of P. aeruginosa has repeatedly been shown to act as immunomodulator (Telford et al., 1998; Smith et al., 2002; Ritchie et al., 2003). Tateda et al. (2003) reported that 3-oxo-C12-HSL, but not C4-HSL, in concentrations of 12–50 MM accelerate apoptosis in macrophages and neutrophils. Lower concentrations of P. aeruginosa QS signals were found to block the activation of PMNs and their oxidative burst, which may explain the observation that QS mutants cause a faster activation of the host defense system in vivo (Bjarnsholt et al., 2005). In the interaction between bacteria and protozoan predators, it has been shown in P aeruginosa, V. cholerae and Chromobacterium violaceum that QS mutants have a significantly reduced antipredator fitness compared to their isogenic wild-type strain (Matz et al., 2004a; Matz et al., 2004b; Matz et al., 2005). Mature biofilms of P. aeruginosa, for example, exhibit acute toxicity to flagellate and amoebal predators (Matz et al., 2004a; Weitere et al., 2005), which is achieved by the upregulation of exofactors mediated by the las/rhl QS system. Studies on the social amoeba D. discoideum also suggest the involvement of the las/rhl QS system in the killing of protozoan predators by P. aeruginosa (Pukatzki et al., 2002; Cosson et al., 2002). Similarly, V. cholerae biofilms were found to induce cell lysis of flagellate predators in a QS dependent fashion via the transcriptional regulator HapR (Matz et

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al., 2005). One factor to be responsible for protozoan cell death has been identified as the previously uncharacterized protease PrtV (Vaitkevicius et al., 2006). Interestingly, the same study shows that PrtV interferes with the production of the interleukin IL-8 from human intestinal epithelial cells. Diversity effects Bacteria living in a biofilm at high cell densities generate a complex three-dimensional structure with physicochemical gradients and numerous microenvironments. In biofilms, new niches arise through the actions of the organisms themselves. According to the niche exclusion principle, different types may be favored in each niche. he self-generated heterogeneity thus produces a variety of phenotypes within individual biofilms (Sauer et al., 2002) (see Figure 11.1D). he connection between environmental heterogeneity and diversity has a long history in ecology and population genetics. Rainey and Travisano (1998) documented the emergence of diversity in genetically identical founding populations of P. fluorescens when propagated in spatially heterogeneous environments. Diversity, in the form of niche-specific genotypes, emerged rapidly in spatially structured microcosms, but not in spatially homogeneous microcosms. Another study recently provided evidence that P. aeruginosa undergoes rapid genetic diversification during growth in biofilm communities (Boles et al., 2004). he genetic changes arise via a RecA-dependent mechanism, which likely involves recombination functions and affects multiple biofilm traits. Furthermore, experimental studies of colicin-producing, colicin-resistant, and colicin-sensitive populations of E. coli show that in an environment allowing for localized interactions diversity can be maintained (Kerr et al., 2002). Whereas these experiments show that diversity is rapidly produced in biofilms and/or structured environments, it remains elusive how it is generated. Besides the potentially strong selective pressure in structured environments, biofilm-specific conditions may also promote genetic innovation by higher mutation rates as found in the structured environments of agar surface colonies of E coli (Taddei et al., 1997; Bjedov et al., 2003) and increased rates of horizontal gene transfer due to the close spatial arrangement of bacteria in biofilms (Molin and Tolker-Nielsen, 2003). he concerted action of these mechanisms would render biofilms a true “hot spot” for phenotypic innovation. A good example for the innovative potential of biofilm populations may be the finding of a phenotypically diverse subpopulation of dispersal cells generated by biofilms of the marine bacterium Pseudoalteromonas tunicata (Mai-Prochnow et al., 2006). Ecological theory predicts that diversity renders a community more resistant to environmental challenge than bacteria displaying a single phenotype (McCann, 2000). To date, the only indication that functional diversity actually increases the ability of bacterial populations to resist predation was recently presented in a study on V. cholerae phase variants (Matz et al., 2005). In a variety of bacterial species, stress conditions are known to lead to the generation of phase variants. hese include the generation of a rugose variant of V. cholerae and a small colony variant of P. aeruginosa by random on-off switching of phenotypic expression. For example, the rugose variant of V. cholerae is found to occur at high frequencies under starvation conditions and exhibits enhanced biofilm formation and resistance to chlorine (Yildiz and Schoolnik, 1999). Grazing studies on the smooth and rugose variants of V. cholerae revealed that protozoan predation selects for the biofilm-en-

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hancing rugose type, which is adapted to the surface-associated niche by the production of exopolymers (Matz et al., 2005). he marked differences in grazing mortality between the two variants specify the fitness conflicts of the “explorer strategy” of the smooth phenotype versus the “persister strategy” of rugose biofilm cells. Similar observations were also made on mucoid and non-mucoid variants of P. fluorescens (Matz et al., 2002b). Predation represents a strong selective pressure on the prey population to evolve means of escaping. Predators as well as parasites are thought to play a significant role in the diversification of their prey or hosts. Several experiments with bacteria and their phage parasites, for example, have documented the spontaneous evolution of phage-resistant bacteria from a population of sensitive types (Bohannan and Lenski, 2000). Investigations of P. aeruginosa flow-chamber biofilms have indicated that the oxidative stress caused by the host immune response could be one of the factors that generates and/or selects the mucoid mutants. It also appears that the mutations in mucA induced by oxygen stress develop with higher frequencies in biofilm-associated cells than in suspended cells (Mathee et al., 1999; Ciofu et al., 2005). While studies on the direct impact of predation on biofilm diversity remain to be performed, it is well documented in planktonic multi-species communities that protozoan grazing causes shifts in the composition of the bacterial community and leads to the prevalence of grazing-resistant bacterial species ( Jürgens and Matz, 2002). Notably, strong grazing activity has led, in most of these studies, to the formation and accumulation of suspended biofilms, so-called flocs or aggregates. Tentative evidence for the potential impact of predation on biofilm diversification comes from a recent study where the functional diversity found in P. aeruginosa biofilms increased the biofilm’s ability to withstand an applied oxidative stress (Boles et al., 2004). Inducible responses to predation he formation of distinct microcolonies by Pseudomonas spp. and the cell elongation observed in Flectobacillus spp. in response to flagellate grazing have long been discussed as inducible by predator-specific signals (Hahn et al., 1999; Hahn et al., 2000). However, such chemically induced plasticity have not yet been convincingly demonstrated for bacterial grazing responses. Predator-induced plasticity has been described for many organisms from fish to protists and comprises alterations in morphology, chemistry, behavior and life history (Tollrian and Harvell, 1999). One inherent problem in the analysis of such a mechanism in microbial systems is that bacterial predators continuously change the chemical composition of the surrounding medium by their excretions which in turn influence bacterial physiology, growth and thus a number of phenotypic traits (see “physiological benefits” below). Without the identification of specific biologically active molecules it may not be feasible to differentiate between substrate-mediated and signal-mediated effects on predation-relevant bacterial phenotypes. First insights into inducible cellular responses can be gained from transcriptomic analyses of bacteria growing in co-culture with phagocytic predators. Microarray analysis of S. aureus strains that were resistant to killing by human PMNs and caused greater host cell lysis identified genes comprising a global response to PMN phagocytosis (Voyich et al., 2005). Genes involved in capsule synthesis, oxidative stress, and virulence were upregulated following ingestion of the pathogen. Preliminary data from the laboratory of Mike Givskov

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show that upon exposure to PMNs P. aeruginosa biofilms can mount either an aggressive or a defensive response (Alhede et al., unpublished data). In the aggressive response, many genes encoding secreted virulence factors, such as lasB, phz, rhlAB and prpL were found to be upregulated. he defensive response was characterized by genes involved in general stress responses, such as lexA, katB, ohr and recN. Palma et al. (2004) demonstrated by means of transcriptomic analysis that the early response of P. aeruginosa to hydrogen peroxide consists of an up regulation of protective mechanisms and a down regulation of primary metabolism. Physiological beneits of predation Although predator-prey interactions appear severely asymmetric and one-directional (positive for the predator, negative for the prey), it may be worthwhile considering potential benefits for bacteria coexisting with predators in biofilms. Protozoa are known to play an important role in the re-mineralization of nitrogen and phosphorous within planktonic and soil microbial communities (Clarholm, 1985; Caron and Goldman, 1990). herefore, bacterial populations limited in nitrogen or phosphorous may actually benefit from predation-mediated nutrient recycling. Interestingly, chemostat studies with nitrifying bacteria revealed that predation by a flagellate increased the metabolic activity per bacterial cell (Verhagen and Laanbroek, 1992). Nutrient-limited bacteria may not only benefit directly from substrates excreted by the predators; at the same time predators may also decrease the numbers of bacterial competitors and thus increase the supply of substrates per cell resulting in higher bacterial growth rates and metabolic activity. In one way or another, nutrient-limited biofilms may thus benefit from allowing (limited) predation to increase population maintenance and persistence. Moreover, predatory eukaryotes themselves offer an attractive nutrient source for biofilm bacteria to be exploited in direct vicinity. Certain pathogenic bacteria, e.g. Legionella, are known to avoid the digestive mechanism of protozoa by blocking the digestive processes and multiplying within the food vacuole, on the expense of the host (Greub and Raoult, 2004). As outlined in a previous section, a number of biofilm-forming bacteria are known to kill immune cells and/or protozoa by secreting exoproducts, particularly at the onset of stationary phase. he lysis of predatory cells does not only prevent bacteria from being grazed upon but may provide them with a welcome nutrient boost. Rapid necrosis of human neutrophils in the presence of P. aeruginosa was recently found to significantly enhance biofilm formation (Walker et al., 2005). he colonization of cellular debris comprising Factin and DNA resulted in a 3.5 fold increase in P. aeruginosa cell numbers in biofilms. Hence, the biofilm-phagocyte relationship does not appear to be one-sided but rather allow for far more complex interactions ranging from tolerance and defense to aggression and pathogenicity. Evolutionary beneits of predation Predation represents a strong selective force, so that adaptations that increase the antipredator fitness of bacteria should be evolutionarily favored. he formation of inedible microcolonies during protozoan grazing is a commonly observed phenomenon in natural bacterioplankton communities ( Jürgens and Matz, 2002). Similarly, the distinguishing fea-

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ture of biofilm infections is the presence of aggregated, matrix-enclosed microcolonies and the QS-mediated secretion of virulence factors. On the micro-evolutionary scale, Boraas et al. (1998) demonstrated that a size-selective protozoan predator selects for a self-replicating multicellular growth form in the unicellular green alga Chlorella vulgaris within less than 100 generations. Taken together, the relative fitness advantage of biofilms over planktonic bacteria in the presence of eukaryotic predators should make a crucial contribution to the evolution of multicellular traits and cooperative behavior found in bacterial biofilms (see also Webb, this volume). As outlined in the previous section, protozoan cells may offer an attractive nutrient source for bacteria once the predatory threat is overcome. What started as a struggle for life, may climax in the gradual transition of some bacterial species from a grazing resister to an aggressor and facultative pathogen. It is widely believed, for example, that the survival and successful replication of bacteria inside the protozoan niche gave rise to a number of facultative and obligate intracellular pathogens, such as Listeria, Rickettsia, Mycobacterium, Legionella, and Chlamydia. he fact that (i) these pathogens exhibit intracellular survival within both amoebae and human macrophages by using related mechanisms and that (ii) amoebae and macrophages share similar phagocytic mechanisms (see Table 11.1) support the notion that resistance to amoebae is an important prerequisite and a driving force in the evolution of some bacteria as pathogens (“training ground hypothesis”; Molmeret et al., 2005). Similar to intracellular survival strategies, protective mechanisms acting prior to the internalization by protozoa might be seen as important features for pathogenic bacteria to persist in the environment or to evade eukaryotic immune systems. Specifically, the antipredator mechanisms found for biofilms of P. aeruginosa (Matz et al., 2004a) and V. cholerae (Matz et al., 2005) suggest a causal link in some pathogenicity aspects between biofilm defense mechanisms against protozoa and professional phagocytes. he recent findings of predation-mediated variation at the rfb virulence locus of Salmonella enterica illustrate the potential impact of protozoan grazing on the origin of bacterial pathogenicity (Wildschutte et al., 2004). Hence, some of the virulence factors studied in the context of human disease are likely to have an ecological function within natural microbial communities and even have their origin specifically in successful antipredator adaptations. Concluding remarks and future directions Historically, biofilms have been viewed as the bacterial refuge against numerous abiotic and biotic stresses that bacteria encounter in natural and host environments. his causality was derived from the study of mono-culture biofilms of selected bacteria species, such as P. aeruginosa, which have dramatically increased our knowledge in recent years on the cellular and molecular mechanisms underlying attachment, migration, recombination, autoregulation, cell–cell signaling, dispersal, and antibiotic resistance. Compared to the rapid progress being made in these areas, our fundamental knowledge of species-species interactions in “real-world” biofilm communities still lags considerably behind. he findings summarized in this chapter support the notion that biofilms provide protection against phagocytic attack in environment and disease. However, our understanding of predator-prey interactions remains fragmentary, in part because the interactions are anticipated to be quite complex; phagocytosis can be viewed as either an obstacle or an opportunity for biofilms.

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However, as more focus is directed toward understanding these interactions, interesting patterns and principles may emerge. Biofilm adaptations to predation may be key to how some pathogens persist and diversify in the environment, reach the minimum infectious dose and undermine the first stages of the immune response. here are basic questions that remain to be answered, such as, whether bacteria know that all or parts of their population are under phagocytic attack and whether the attack induces a specific defense program in biofilms. Another emerging area of interest is how predation drives the evolution of virulence factors, the maintenance of their variability, and the emergence of new pathogens. Research in these areas would certainly benefit if some of the ecological thinking characteristic for microbial food web studies was combined with the molecular approaches used by infection microbiologists. Acknowledgments I thank Staffan Kjelleberg, Michael Givskov and Kenneth Timmis for their support and patience. his work was supported by the German Research Foundation (DFG) grant Ma 2491/3-1. References Allen, L., Dockrell, D.H., Pattery, T., Lee, D.G., Cornelis, P., Hellewell, P.G., and Whyte, M.K.B. (2005). Pyocyanin production by Pseudomonas aeruginosa induces neutrophil apoptosis and impairs neutrophil-mediated host defenses in vivo. J. Immunol. 174, 3643–3649. Arndt, H., Schmidt-Denter, K., Auer, B., and Weitere, M. (2003). Protozoans and biofilms. In: Fossil and Recent Biofilms, W E. Krumbein, D.M. Paterson, and G.A. Zavarzin, eds. (Dordrecht, Kluwer Academic Publ.), pp. 173–189. Bjarnsholt, T., Ostrup Jensen, P., Burmolle, M., Hentzer, M., Haagensen, J.A., Hougen, H.P., Calum, H., Madsen, K.G., Moser, C., Molin, S., et al. (2005). Pseudomonas aeruginosa tolerance to tobramycin, hydrogen peroxide and polymorphonuclear leukocytes is quorum sensing dependent. Microbiology 151, 373–383. Bjedov, I., Tenaillon, O., Gerard, B., Souza, V., Denamur, E., Radman, M., Taddei, F., and Matic, I. (2003). Stress-induced mutagenesis in bacteria. Science 300, 1404–1409. Boenigk, J., and Arndt, H. (2000). Comparative studies on the feeding behavior of two heterotrophic nanoflagellates: the filter-feeding choanoflagellate Monosiga ovata and the raptorial-feeding kinetoplastid Rhynchomonas nasuta. Aquat. Microb. Ecol. 22, 243–249. Bohannan, B.J.M., and Lenski, R.E. (2000). Linking genetic chance to community evolution: insights from studies of bacteria and bacteriophage. Ecol. Lett. 3, 362–377. Boles, B.R., hoendel, M., and Singh, P K. (2004). Self-generated diversity produces “insurance effects” in biofilm communities. Proc. Natl. Acad. Sci. USA 101, 16630–16635. Boraas, M.E., Seale, D.B., and Boxhorn, JE. (1998). Phagotrophy by a flagellate selects for colonial prey: A possible origin of multicellularity. Evol. Ecol. 12, 153–164. Brown, R.K., and Kelly, F.J. (1994). Evidence for increased oxidative damage in patients with cystic fibrosis. Pediatr. Res. 36, 487–493. Bylund, J., Burgess, L.A., Cescutti, P., Ernst, R.K., and Speert, D.P. (2006). Exopolysaccharides from Burkholderia cenocepacia inhibit neutrophil chemotaxis and scavenge reactive oxygen species. J. Biol. Chem. 281, 2526–2532. Caron, D.A., and Goldman, J.C. (1990). Protozoan nutrient regeneration. In: Ecology of Marine Protozoa, G.M. Capriulo, ed. (New York, Oxford University Press), pp. 283–306. Cavalier-Smith, T. (2002). he phagotrophic origin of eukaryotes and phylogenetic classification of protozoa. Int. J. Syst. Evol. Microbiol. 52, 297–354. Celli, J., and Finlay, B.B. (2002). Bacterial avoidance of phagocytosis. Trends Microbiol. 10, 232–237. Chan, C., Burrows, L.L., and Deber, C.M. (2004). Helix induction in antimicrobial peptides by alginate in biofilms. J. Biol. Chem. 279, 38749–38754.

|

209

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|

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Ciofu, O., Riis, B., Pressler, T., Poulsen, H.E., and Høiby, N. (2005). Occurrence of hypermutable Pseudomonas aeruginosa in cystic fibrosis patients is associated with the oxidative stress caused by chronic lung inflammation. Antimicrob. Agents Chemother. 49, 2276–2282. Clarholm, M. (1985). Interactions of bacteria, protozoa and plants leading to mineralization of soil nitrogen. Soil Biol. Biochem. 17, 181–187. Cosson, P., Zulianello, L., Join-Lambert, O., Faurisson, F., Gebbie, L., Benghezal, M., Van Delden, C., Curty, L.K., and Kohler, T. (2002). Pseudomonas aeruginosa virulence analyzed in a Dictyostelium discoideum host system. J. Bacteriol. 184, 3027–3033. Costerton, J.W., Cheng, K.J., Geesey, G.G., Ladd, T.I., Nickel, J.C., Dasgupta, M.K., and Marrie, T.J. (1987). Bacterial biofilms in nature and disease. Annu. Rev. Microbiol. 41, 435–464. Darby, C., Hsu, J.W., Ghori, N., and Falkow, S. (2002). Caenorhabditis elegans: Plague bacteria biofilm blocks food intake. Nature 417, 243–244. Davies, B., Chattings, L.S., and Edwards, S.W. (1991). Superoxide generation during phagocytosis by Acanthamoeba castellanii: similarities to the respiratory burst of immune phagocytes. J. Gen. Microbiol. 137, 705–710. Devreotes, P.N., and Zigmond, S.H. (1988). Chemotaxis in eukaryotic cells: A focus on leukocytes and Dictyostelium. Annu. Rev. Cell Biol. 4, 649–686. Duffy, J.E., and Hay, M.E. (1990). Seaweed adaptations to herbivory. BioScience 40, 368–375. Ernst, R.K., Yi, E.C., Guo, L., Lim, K.B., Burns, J.L., Hackett, M., and Miller, S.I. (1999). Specific lipopolysaccharide found in cystic fibrosis airway Pseudomonas aeruginosa. Science 286, 1561–1565. Greub, G., and Raoult, D. (2004). Microorganisms resistant to free-living amoebae. Clin. Microbiol. Rev. 17, 413–433. Hahn, M.W., Moore, E.R.B., and Höfle, M.G. (2000). Role of microcolony formation in the protistan grazing defense of the aquatic bacterium Pseudomonas sp. MWH1. Microb. Ecol. 39, 175–185. Hahn, M.W., Moore, E R.B., and Höfle, M.G. (1999). Bacterial filament formation, a defense mechanism against flagellate grazing, is growth rate controlled in bacteria of different phyla. Appl. Environ. Microbiol. 65, 25–35. Hammer, A., Grüttner, C., and Schumann, R. (1999). he effect of electrostatic charge of food particles on capture efficiency by Oxyrrhis marina Dujardin (dinoflagellate). Protist 150, 375–382. Hayashi, K., Lee, D.A., and Quie, P.G. (1986). Chemiluminescent response of polymorphonuclear leukocytes to Streptococcus pneumoniae and Haemophilus influenzae in suspension and adhered to glass. Infect. Immun. 58, 2383–2385. Heaton, K., Drinkall, J., Minett, A., Hunt, A., and Parry, J.D. (2001). Amoeboid grazing on surface associated prey. In: Biofilm Community Interactions: Chance or Necessity?, P. Gilbert, D.G. Allison, M. Brading, J. Verran, and J. Walker, eds. (Cardiff, Bioline Press), pp. 293–301. Herbst, R., Ott, C., Jacobs, T., Marti, T., Marciano-Cabral, F., and Leippe, M. (2002). Pore-forming polypeptides of the pathogenic protozoon Naegleria fowleri. J. Biol. Chem. 277, 22353–22360. Hoffmann, J.A., Kafatos, F.C., Janeway Jr, C.A., and Ezekowitz, R.A.B. (1999). Phylogenetic perspectives in innate immunity. Science 284, 1313–1318. Jackson, S.M., and Jones, E.B.G. (1991). Interactions within biofilms: the disruption of biofilm structure by protozoa. Kieler Meeresforsch. 8, 264–268. Jesaitis, A.J., Franklin, M.J., Berglund, D., Sasaki, M., Lord, C.I., Bleazard, J.B., Duffy, J.E., Beyenal, H., and Lewandowski, Z. (2003). Compromised host defense on Pseudomonas aeruginosa biofilms: Characterization of neutrophil and biofilm interactions. J. Immunol. 171, 4329–4339. Jürgens, K., and Matz, C. (2002). Predation as a shaping force for the phenotypic and genotypic composition of planktonic bacteria. Antonie Van Leeuwenhoek 81, 413–434. Kelly, N.M., Kluftinger, J.L., Pasloske, B.L., Paranchych, W., and Hancock, R.E.W. (1989). Pseudomonas aeruginosa pili as ligands for nonopsonic phagocytosis by fibronectin-stimulated macrophages. Infect. Immun. 57, 3841–3845. Kerr, B., Riley, M.A., Feldman, M.W., and Bohannan, B.J.M. (2002). Local dispersal promotes biodiversity in a real-life game of rock-paper-scissors. Nature 418, 171–174. Kharazmi, A. (1991). Mechanisms involved in the evasion of the host defence by Pseudomonas aeruginosa. Immunol. Lett. 30, 201–206. Kharazmi, A., Doring, G., Hoiby, N., and Valerius, N.H. (1984a). Interaction of Pseudomonas aeruginosa alkaline protease and elastase with human polymorphonuclear leukocytes in vitro. Infect. Immun. 43, 161–165.

Bioilms and Predation

Kharazmi, A., Hoiby, N., Doring, G., and Valerius, N.H. (1984b). Pseudomonas aeruginosa exoproteases inhibit human neutrophil chemiluminescence. Infect. Immun. 44, 587–591. Koch, C., and Hoiby, N. (1993). Pathogenesis of cystic fibrosis. Lancet 341, 1065–1069. Kocianova, S., Vuong, C., Yao, Y., Voyich, J.M., Fischer, E.R., DeLeo, F.R., and Otto, M. (2005). Key role of poly-G-DL-glutamic acid in immune evasion and virulence of Staphylococcus epidermidis. J. Clin. Invest. 115, 688–694. Lawrence, J.R., and Snyder, R.A. (1998). Feeding behaviour and grazing impacts of a Euplotes sp. on attached bacteria. Can. J. Microbiol. 44, 623–629. Learn, D.B., Brestel, E.P., and Seetharama, S. (1987). Hypochlorite scavenging by Pseudomonas aeruginosa alginate. Infect. Immun. 55, 1813–1818. Lee, D.A., Hoidal, J.R., Clawson, C.C., Quie, P.G., and Peterson, P.K. (1983). Phagocytosis by polymorphonuclear leukocytes of Staphylococcus aureus and Pseudomonas aeruginosa adherent to plastic, agar, or glass. J. Immunol. Methods 63, 103–114. Leid, J.G., Shirtliff, M.E., Costerton, J.W., and Stoodley, P. (2002). Human leukocytes adhere to, penetrate, and respond to Staphylococcus aureus biofilms. Infect. Immun. 70, 6339–6345. Leid, J.G., Willson, C.J., Shirtliff, M.E., Hassett, D J., Parsek, M.R., and Jeffers, A.K. (2005). he exopolysaccharide alginate protects Pseudomonas aeruginosa biofilm bacteria from IFN-G-mediated macrophage killing. J. Immunol. 175, 7512–7518. Levin, B.R., and Antia, R. (2001). Why we don’t get sick: the within-host population dynamics of bacterial infections. Science 292, 1112–1115. Lock, R., Öhman, L., and Dahlgren, C. (1987). Phagocytic recognition mechanisms in human granulocytes and Acanthamoeba castellanii using type 1 fimbriated Escherichia coli as phagocytic prey. FEMS Microbiol. Lett. 44, 135–140. Mahenthiralingam, E., and Speert, D P. (1995). Nonopsonic phagocytosis of Pseudomonas aeruginosa by macrophages and polymorphonuclear leukocytes requires the presence of the bacterial flagellum. Infect. Immun. 63, 4519–4523. Mai-Prochnow, A., Webb, J.S., Ferrari, B.C., and Kjelleberg, S. (2006). Ecological advantages of autolysis during the development and dispersal of Pseudoalteromonas tunicata biofilms. Appl. Environ. Microbiol. 72, 5414–5420. Marshall, K.C. (1986). Adsorption and adhesion in microbial growth at interfaces. Adv. Colloid Interface Sci. 25, 59–86. Mathee, K., Ciofu, O., Sternberg, C., Lindum, P.W., Campbell, J.I. A., Jensen, P., Johnsen, A.H., Givskov, M., Ohman, D.E., Molin, S., et al. (1999). Mucoid conversion of Pseudomonas aeruginosa by hydrogen peroxide: a mechanism for virulence activation in the cystic fibrosis lung. Microbiology 145, 1349–1357. Matz, C., Boenigk, J., Arndt, H., and Jürgens, K. (2002a). Role of bacterial phenotypic traits in selective feeding of the heterotrophic nanoflagellate Spumella sp. Aquat. Microb. Ecol. 27, 137–148. Matz, C., Deines, P., and Jürgens, K. (2002b). Phenotypic variation in Pseudomonas sp CM10 determines microcolony formation and survival under protozoan grazing. FEMS Microbiol. Ecol. 39, 57–65. Matz, C., Bergfeld, T., Rice, S.A., and Kjelleberg, S. (2004a). Microcolonies, quorum sensing and cytotoxicity determine the survival of Pseudomonas aeruginosa biofilms exposed to protozoan grazing. Environ. Microbiol. 6, 218–226. Matz, C., Deines, P., Boenigk, J., Arndt, H., Eberl, L., Kjelleberg, S., and Jurgens, K. (2004b). Impact of violacein-producing bacteria on survival and feeding of bacterivorous nanoflagellates. Appl. Environ. Microbiol. 70, 1593–1599. Matz, C., McDougald, D., Moreno, A.M., Yung, P.Y., Yildiz, F.H., and Kjelleberg, S. (2005). Biofilm formation and phenotypic variation enhance predation-driven persistence of Vibrio cholerae. Proc. Natl. Acad. Sci. USA 102, 16819–16824. McCann, K.S. (2000). he diversity-stability debate. Nature 405, 228–233. McClure, C.D., and Schiller, N.L. (1996). Inhibition of macrophage phagocytosis by Pseudomonas aeruginosa rhamnolipids in vitro and in vivo. Curr. Microbiol. 33, 109–117. Molin, S., and Tolker-Nielsen, T. (2003). Gene transfer occurs with enhanced efficiency in biofilms and induces enhanced stabilisation of the biofilm structure. Curr. Opin. Biotechnol. 14, 255–261. Molmeret, M., Horn, M., Wagner, M., Santic, M., and Abu Kwaik, Y. (2005). Amoebae as training grounds for intracellular bacterial pathogens. Appl. Environ. Microbiol. 71, 20–28.

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Monger, B.C., Landry, M.R., and Brown, S.L. (1999). Feeding selection of heterotrophic marine nanoflagellates based on the surface hydrophobicity of their picoplankton prey. Limnol. Oceanogr. 44, 1917–1927. Navarre, W.W., and Schneewind, O. (1999). Surface proteins of gram-positive bacteria and mechanisms of their targeting to the cell wall envelope. Microbiol. Mol. Biol. Rev. 63, 174–229. Palma, M., DeLuca, D., Worgall, S., and Quadri, L.E.N. (2004). Transcriptome analysis of the response of Pseudomonas aeruginosa to hydrogen peroxide. J. Bacteriol. 186, 248–252. Parry, J.D. (2004). Protozoan grazing of freshwater biofilms. Adv. Appl. Microbiol. 54, 167–196. Parsek, M.R., and Greenberg, E.P. (2005). Sociomicrobiology: the connections between quorum sensing and biofilms. Trends Microbiol. 13, 27–33. Pedersen, K. (1982). Factors regulating microbial biofilm development in a system with slowly flowing seawater. Appl. Environ. Microbiol. 44, 1196–1204. Pessi, G., and Haas, D. (2000). Transcriptional control of the hydrogen cyanide biosynthetic genes hcnABC by the anaerobic regulator ANR and the quorum-sensing regulators LasR and RhlR in Pseudomonas aeruginosa. J. Bacteriol. 182, 6940–6949. Pukatzki, S., Kessin, R.H., and Mekalanos, J.J. (2002). he human pathogen Pseudomonas aeruginosa utilizes conserved virulence pathways to infect the social amoeba Dictyostelium discoideum. Proc. Natl. Acad. Sci. USA 99, 3159–3164. Rainey, P.B., and Travisano, M. (1998). Adaptive radiation in a heterogeneous environment. Nature 394, 69–72. Ritchie, A.J., Yam, A.O.W., Tanabe, K.M., Rice, S.A., and Cooley, M.A. (2003). Modification of in vivo and in vitro T- and B-cell-mediated immune responses by the Pseudomonas aeruginosa quorum-sensing molecule N-(3-oxododecanoyl)-L-homoserine lactone. Infect. Immun. 71, 4421–4431. Sauer, K., Camper, A.K., Ehrlich, G.D., Costerton, J.W., and Davies, D.G. (2002). Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J. Bacteriol. 184, 1140–1154. Simpson, J.A., Smith, S.E., and Dean, R.T. (1989). Scavenging by alginate of free radicals released by macrophages. Free Radical Biol. Med. 6, 347–353. Smith, R.S., Harris, S.G., Phipps, R., and Iglewski, B. (2002). he Pseudomonas aeruginosa quorum-sensing molecule N-(3-oxododecanoyl)homoserine lactone contributes to virulence and induces inflammation in vivo. J. Bacteriol. 184, 1132–1139. Smith, R.S., and Iglewski, B.H. (2003). Pseudomonas aeruginosa quorum-sensing systems and virulence. Curr. Opin. Microbiol. 6, 56–60. Speert, D.P., Loh, B.A., Cabral, D.A., and Salit, I.E. (1986). Nonopsonic phagocytosis of nonmucoid Pseudomonas aeruginosa by human neutrophils and monocyte-derived macrophages is correlated with bacterial piliation and hydrophobicity. Infect. Immun. 53, 207–212. Taddei, F., Halliday, J.A., Matic, I., and Radman, M. (1997). Genetic analysis of mutagenesis in aging Escherichia coli colonies. Mol. Gen. Genetics 256, 277–281. Tateda, K., Ishii, Y., Horikawa, M., Matsumoto, T., Miyairi, S., Pechere, J.C., Standiford, T.J., Ishiguro, M., and Yamaguchi, K. (2003). he Pseudomonas aeruginosa autoinducer N-3-oxododecanoyl homoserine lactone accelerates apoptosis in macrophages and neutrophils. Infect. Immun. 71, 5785–5793. Telford, G., Wheeler, D., Williams, P., Tomkins, P.T., Appleby, P., Sewell, H., Stewart, G.S.A.B., Bycroft, B.W., and Pritchard, D.I. (1998). he Pseudomonas aeruginosa quorum-sensing signal molecule N-(3oxododecanoyl)-L-homoserine lactone has immunomodulatory activity. Infect. Immun. 66, 36–42. Tollrian, R., and Harvell, C. (1999). he Ecology and Evolution of Inducible Defenses (Princeton, Princeton University Press). Vaitkevicius, K., Lindmark, B., Ou, G., Song, T., Toma, C., Iwanaga, M., Zhu, J., Andersson, A., Hammarström, M.L., Tuck, S., and Wai, S.N. (2006). A Vibrio cholerae protease needed for killing of Caenorhabditis elegans has a role in protection from natural predator grazing. Proc. Natl. Acad. Sci. USA 103, 9280–9285. van Oss, C.J. (1978). Phagocytosis as a surface phenomenon. Annu. Rev. Microbiol. 32, 19–39. van Putten, J.P., and Robertson, B.D. (1995). Molecular mechanisms and implications for infection of lipopolysaccharide variation in Neisseria. Mol. Microbiol. 16, 847–853. Verhagen, F.J.M., and Laanbroek, H.J. (1992). Effects of grazing by flagellates on competition for ammonium between nitrifying and heterotrophic bacteria in chemostats. Appl. Environ. Microbiol. 58, 1962–1969.

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Voyich, J.M., Braughton, K.R., Sturdevant, D.E., Whitney, A.R., Said-Salim, B., Porcella, S.F., Long, R.D., Dorward, D.W., Gardner, D.J., Kreiswirth, B.N., et al. (2005). Insights into mechanisms used by Staphylococcus aureus to avoid destruction by human neutrophils. J. Immunol. 175, 3907–3919. Vuong, C., Voyich, J.M., Fischer, E.R., Braughton, K.R., Whitney, A.R., DeLeo, F.R., and Otto, M. (2004). Polysaccharide intercellular adhesin (PIA) protects Staphylococcus epidermidis against major components of the human innate immune system. Cell. Microbiol. 6, 269–275. Walker, T.S., Tomlin, K.L., Worthen, G.S., Poch, K.R., Lieber, J.G., Saavedra, M.T., Fessler, M.B., Malcolm, K.C., Vasil, M.L., and Nick, J.A. (2005). Enhanced Pseudomonas aeruginosa biofilm development mediated by human neutrophils. Infect. Immun. 73, 3693–3701. Weitere, M., Bergfeld, T., Rice, S.A., Matz, C., and Kjelleberg, S. (2005). Grazing resistance of Pseudomonas aeruginosa biofilms depends on type of protective mechanism, developmental stage and protozoan feeding mode. Environ. Microbiol. 7, 1593–1601. Wildschutte, H., Wolfe, D.M., Tamewitz, A., and Lawrence, J.G. (2004). Protozoan predation, diversifying selection, and the evolution of antigenic diversity in Salmonella. Proc. Natl. Acad. Sci. USA 101, 10644–10649. Winzer, K., and Williams, P. (2001). Quorum sensing and the regulation of virulence gene expression in pathogenic bacteria. Int. J. Med. Microbiol. 291, 131–143. Yildiz, F.H., and Schoolnik, G.K. (1999). Vibrio cholerae O1 El Tor: Identification of a gene cluster required for the rugose colony type, exopolysaccharide production, chlorine resistance, and biofilm formation. Proc. Natl. Acad. Sci. USA 96, 4028–4033.

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Bioilms on Plant Surfaces Leo Eberl, Susanne B. von Bodman, and Clay Fuqua

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Abstract Land plants modify the terrestrial environment extensively by nutrient acquisition, water utilization, physical disruption and cohesion of the soil, and the release of complex exudate materials. Decaying plant matter is also a major source of organic material in soils. Large numbers of microorganisms associate with and flourish on, within, and around plants, colonizing virtually all exposed tissues. While some of these microbes may incite disease on certain plants, a large number are harmless or beneficial symbionts. Microbial populations multiply in response to the plant environment and often form multicellular complexes that range from small aggregates to expansive, highly structured biofilms. Plant-associated biofilms have important consequences for plant health and disease, as the microorganisms within these populations may provide benefits or, conversely, damage the host. he structure, activity and microbial diversity harbored within biofilms influence the plant interaction to varying degrees, dependent on plant type, growth stage and environmental conditions. Likewise, plants influence the bacterial population density, fostering communities that interact with each other and the plant through metabolic activity and cell–cell communication mechanisms that allow the microbes to coordinate their activities and optimize their competitive success. Introduction Plants support the growth of a wide variety of microorganisms on and within their tissues, including aerial portions of the plant, the vascular network, and root tissues below ground (Figure 12.1). Plant activity and phytoexudation dramatically modify the local environment, supplying certain relatively plentiful nutrients compared with the generally nutrient-limited terrestrial habitat (Walker et al., 2003). Plants also deplete other nutrients, such as phosphorous, from their local environment. Plant-associated microbes may establish commensal, mutualistic and pathogenic interactions with plants, or simply grow saprophytically on the nutrients released. Soil that immediately surrounds roots and that is influenced by plant activity is called the rhizosphere, and can be considered an extension of the plant-associated environment. Microbial populations that colonize plant tissues and the rhizosphere accumulate at specific physical locations and often multiply in response to the nutrients available. In many cases, multicellular assemblies of microbes result, and these

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Figure 12.1 Sites of bioilm formation on plants. A photograph of an axenically grown Arabidopsis thaliana seedling is shown. Bacteria colonizing phyllosphere/phylloplane on leaves, rhizosphere/rhizoplane on roots, and within the internal vascular system are depicted.

may be homogeneous structures or multispecies communities (Bloemberg et al., 2000; Watt et al., 2006). In some cases, the multicellular structures may have specific functions relevant to plant interactions, but in others may simply be the consequence of population expansion. In this chapter, we will consider the structure, composition and function of bacterial populations associated with terrestrial plants. Although their size, distribution and conformation may differ from biofilm populations formed in saturated aqueous environments, we will refer to them collectively as biofilms. Salient features of plant-associated microbial habitats Each type of plant tissue is chemically and physically unique and bacteria that colonize this tissue must adapt to these unique attributes. In addition to these intrinsic characteristics, local environmental conditions such as water saturation, temperature and pH can have a tremendous impact on microbial colonization. Finally, the adherent microbial populations can significantly alter the environment they inhabit, through growth, metabolism, resource competition and intercellular signaling. Different plant tissues present a range of microenvironments to colonizing microbes, and the density, size, species composition and activity of adherent microbial populations at different sites on the plant are reflective of this heterogeneity (Figure 12.1). he trichomes,

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stomates and veins of leaves provide physical and chemical heterogeneity in the phyllosphere that is exploited to the benefit of microbial epiphytes (Herrera and von Bodman, unpublished results, and Monier and Lindow, 2005a). he wax cuticle that often covers the leaf surface differs on the tops and undersides of leaves, and the undersides tend to accumulate more bacterial biomass (Krimm et al., 2005). Along the length of the root, cell type, surface chemistry and exudation activity vary considerably (Gregory, 2006). Actively growing root tissues typically exhibit higher exudation rates into the soil and root cap cells at the growing tip can be sloughed away. Older root tissue may elaborate root hairs, which again provide a distinct surface with which microbes can interact, most notably the symbiotic colonization of legumes by nitrogen-fixing rhizobia (Figure 12.2A) (Hirsch et al., 2001). Microbial populations flourish in response to nutrient release and exudation rates at different sites on roots, while conversely nutrient sequestration from the soil by the root may limit microbial growth. Recent examination of microbial populations on field grown wheat roots reveals high diversity that varies in a consistent way along the length of the root (Watt et al., 2006). he plant vascular network internally connects the different portions of the plant. he configuration of the vasculature, and the relative distribution of phloem and xylem elements differ between aerial and terrestrial plant tissues. Multiple cell and tissue types also exist within the vasculature and it is clear that pathogens which invade and colonize these systems have adapted to associate with specific structural components of the vasculature (Koutsoudis et al., 2006). Hydration is a critical attribute of all microbial systems, including those associated with terrestrial plants. Plants have adapted to a wide diversity of environments, ranging from extremely low saturation levels (e.g. deserts) to extremely high saturation (e.g. rainforests). he microbial populations associated with plants, accordingly, are also adapted to these conditions. Seasonal and other temporal changes in temperature, rainfall, and ground water levels can lead to short-term and long term fluctuations in soil hydration. Microbes that colonize different regions of the plant experience saturation levels characteristic of these tissues. Living plants must maintain a certain minimal fluid level within their vasculature, and saturation is relatively constant within these vessels, although osmolyte concentrations can

A

B

C

Figure 12.2 Colonization of plants by bacteria. (A) Fluorescence micrograph of root hairs of alfalfa colonized with GFP-expressing Sinorhizobium meliloti (unpublished image courtesy of D.J. Gage). (B) Fluorescence micrograph of GFP-expressing A. tumefaciens C58 bioilm on Arabidopsis roots. (Unpublished image from Tomlinson and Fuqua) (C) Electron micrograph of pea roots colonized by Burkholderia cenocepacia H111 (Picture is courtesy of Eshwar Mahenthiralingam, Cardiff University, UK).

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vary. In contrast, tissues on the plant exterior are exposed to greater variability in hydration, and adherent microbial populations may experience significant water limitation or matric stress (Chang and Halverson, 2003). On these exposed tissues, biofilms and aggregates can function to prevent dessication, and accordingly cells within these multicellular structures are more desiccation resistant than single cells (Monier and Lindow, 2005a). Roots maintain a water film on their surfaces, but this film varies considerably in thickness and uniformity with prevailing conditions (Gregory, 2006). Roots are also the site of water uptake by the plant, and provide a constant demand. Leaves and other aerial portions of the plant may be very dry because of exposure to air and sunlight, but can periodically be doused with fluid due to rainfall and accumulation of condensed water. Leaves typically elaborate a protective waxy cuticle in part to prevent water loss, and microbes that colonize these surfaces must adapt to this poorly wettable substratum. Deposition of the microbes that colonize plant tissues here are numerous routes, direct and indirect, by which microbes encounter plant tissues and thereby initiate the interactions that lead to biofilm formation. In the soil environment, microbial locomotion, through flagellar-driven swimming and swarming, twitching motility and gliding are all likely to play a role in colonization of plants. he relatively nutrient rich conditions generated by plant exudation into the soil provide a target for chemotaxis afforded through these mechanisms of motility (Walker et al., 2003). Motility and chemotaxis provide a competitive advantage and allow soil-borne microbes to effectively colonize productive sites on the plant (Lugtenberg et al., 2002). Accordingly, mutants that cannot swim or are defective in chemotaxis are often less able to compete during colonization of plant surfaces (Tans-Kersten et al., 2001; Turnbull et al., 2001; Vande Broeke and Venderleyden, 1995). In addition to self-propelled deposition there is ample opportunity for passive introduction of microbes onto plant tissues. Wind is a well established mechanism by which microbes are deposited on aerial plant surfaces (Pedgley, 1991). Rainwater, splatter and aerosols can also transfer microbes onto leaves and flowers. Water flow around root systems can act to inoculate cells and spores onto below ground tissues. Physical wounding of plants may introduce microbes from the external environment into internal tissues, including the vasculature. Insect and nematode activity can potentially inoculate microbes at any site on the plant, and sap-feeding insects are a common mode of transmission for vascular pathogens (Brlansky et al., 1983). Phyllosphere aggregates and bioilms Aerial surfaces such as leaves are stressful environments for colonizing bacteria, due to light, heat and desiccation. Although single cells can be deposited at any position, multicellular aggregates preferentially form around stomates and trichomes, or along leaf veins, responding to moisture and nutrient concentration (Monier and Lindow, 2004). he aggregates generally grow from one or a few colonizing cells that multiply at their site of deposition to form a multicellular structure (Figure 12.1). Fluorescent pseudomonads associated with cantaloupe and endive leaves were equally distributed in solitary or biofilm-associated populations (Morris and Monier, 2003). In contrast, the majority of Gram-positive epiphytes tended to localize within biofilms.

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Among the best studied epiphytes is Pseudomonas syringae pv. syringae (Pss), that causes brown spot disease on bean (Table 12.1). Although many single Pss cells are observed to colonize the leaf surface, the bulk of the adherent biomass that results from colonization is contained in large aggregates comprised of thousands of cells (Monier and Lindow, 2004). hese aggregates survive desiccation stress better than solitary cells and colonizing cells that are introduced into aggregates have a better chance of survival than singly adherent Table 12.1 Examples of bioilm-forming, plant-associated bacteria1 Bacteria

Host plants

Interaction

Colonization site

Pseudomonas luorescens

Diverse

Commensal/ mutualist

Leaves

Pseudomonas syringae pv. syringae

Beans

Pathogen

Leaves

Methylobacterium spp.

Diverse

Commensal

Leaves (stomates)

Erwinia amylovora

Fruit trees

Pathogen

Fruit, leaves and lowers

Pantoea stewartii

Maize

Pathogen

Xylem vessels

Xylella fastidiosa

Citrus and grape

Pathogen

Xylem vessels

Xanthomonas campestris pv. campestres

Crucifers

Pathogen

Xylem vessels

Ralstonia solanacearum

Diverse

Pathogen

Roots to xylem

Clavibacter michiganensis pv. sepedonicus

Potato

Pathogen

Xylem vessels

Leifsonia xyli subsp. xyli

Sugar cane

Pathogen

Xylem vessels

Spiroplasma spp.

Diverse

Pathogen

Phloem vessels

Agrobacterium tumefaciens

Diverse dicots

Pathogen

Roots and crown tissue

Azospirillum brasilense

Cereals

Mutualist

Root hairs

Bacillus cereus

Diverse

Commensal

Roots

Burkholderia cenocepacia

Onions

Pathogen

Roots

Pseudomonas spp.

Diverse

Commensal/ mutualist

Roots

Rhizobium spp.

Legumes

Mutualist

Root hairs and nodules

Aerial tissue colonizers

Vascular colonizers

Root colonizers

1See

text for details

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cells (Monier and Lindow, 2005a). he Pss aggregates that form on leaf surfaces, have densely packed cells that are ideally suited for the population-density-sensitive signaling mechanism described as quorum sensing (QS). he fitness of adherent Pss populations requires a functional QS system (Quinones et al., 2004). Multispecies interactions between bacteria on aerial surfaces may also influence the populations that eventually develop. Analysis of strawberry plant leaves, revealed that colonization by a single type of epiphyte can significantly alter the wax cuticle and transpiration properties of the leaves, suggesting that this microbe might promote colonization by other microbes (Krimm et al., 2005). Inoculation of bean leaves with three different leaf epiphytes under high moisture conditions resulted in formation of large aggregates, but these aggregates comprised discrete sectors containing nearly homogeneous populations (Monier and Lindow, 2005b). he basis for this segregation remains unclear, although possible explanations are that each sector arises from clonal growth of sparsely colonizing bacteria, or that these bacteria release signaling compounds that ward off competing populations. Bioilm formation during plant vascular disease he xylem and phloem of plants represent a protected niche for the multiplication of biofilm-forming vascular pathogens. Microbes colonize the vasculature through active invasion, wounding and sap-feeding insect vectors. he best characterized systems are those pathogens that colonize the xylem. Phloem-infecting spiroplasma have been studied, but little is known regarding their interaction or growth within phloem vessels (Bove et al., 2003). Xylem-infecting pathogens often form biofilms within the vessels and produce copious polysaccharide that blocks the vasculature, damaging the infected tissue. In addition, degradative enzymes secreted by the infecting bacteria often augment tissue damage and aid in dispersal of the microbe through the vasculature. Several xylem pathogens appear to regulate colonization and subsequent dispersal, using QS mechanisms to gauge their population structure and appropriately time their dispersal (von Bodman et al., 2003). Pantoea stewartii (Ps) is the causative agent of Stewart’s wilt, a vascular disease of maize, introduced into the xylem by the corn flea beetle (Table 12.1). Recent work has revealed that Ps forms biofilms within infected vessels, preferentially associating with the annular rings and thickened areas of the protoxylem, synthesizing the polysaccharide stewartan and effectively disrupting transport processes (Koutsoudis et al., 2006). Although mutants that cannot produce stewartan are avirulent, derivatives that overproduce the polysaccharide do not continue to colonize the vasculature as efficiently as the wild type, suggesting that productive infection requires a balance of vasculature blockage and dispersal. Swarming motility is likely to play a role in the dispersal process (Herrera and von Bodman, unpublished results). he balance of colonization and dispersal is at least in part maintained through QS regulation of stewartan biosynthesis and motility (Minogue et al., 2005). Similar population dynamics within the plant vasculature are observed for xylem infection by the etiologic agent of citris variegated chlorosis and Pierce’s disease of grapevine (Table 12.1). Xylella fastidiosa (Xf) is a xylem-infecting, Gram-negative pathogen transmitted by leafhopper insects probing for xylem elements (Brlansky et al., 1983). Within the insect vector, Xf multiplies and forms polarly attached, palisade-like biofilms on foregut tis-

Plant Bioilms

sues (Newman et al., 2004). A cell signaling mechanism, that relies on a diffusible signaling factor (DSF) is important for insect colonization and subsequent transmission. Formation of these biofilms and the amplified pathogen population they embody is required for effective disease transmission to the plant. During transmission to the plant, single cells attach to xylem vessels elements (Newman et al., 2003). Production of Xf exopolysaccharide is required for disease, and areas with dense biofilms and high levels of exopolysaccharide are the sites of blockage. Movement of the pathogen through the vasculature occurs in the direction opposite to the flow of the transpiration stream, away from leaves, and has recently been shown to involve a type IV pilus and twitching motility (Meng et al., 2005). Effective attachment and biofilm formation, in contrast, requires a type I adhesive pilus structure and earlier evidence suggests that this may involve interaction with xylem elements via active thiol groups on the pilus or another cell surface structure (Leite et al., 2004). he related pathogen Xanthomonas campestris pv. campestris (Xcc) is introduced into the plant vasculature by wounding, and causes black rot on cruciferous plants (Table 12.1). Outside of the means of inoculation Xcc shares a very similar pattern of vascular infection with Xf, and also regulates this process with the DSF signaling system (Grossman and Dow, 2004). he polysaccharide xanthan gum and degradative exoenzymes are primary virulence determinants, governing vascular blockage and migration of the bacteria through the vasculature. Again, for both Xcc and Xf, the balance between biofilm formation and intravascular dispersal is a key determinant of disease progression. he wide host range pathogen Ralstonia solanacearum actively invades roots, infiltrating the vasculature and resulting in lethal wilt on diverse plants (Table 12.1). In the xylem, swimming and twitching motility are likely to facilitate surface colonization and biofilm formation (Kang et al., 2002; Tans-Kersten et al., 2001). It is again production of an exopolysaccharide that clogs the xylem vessels, and causes the wilting disease. A complex regulatory cascade, described as a confinement sensing mechanism, controls production of the exopolysaccharide and exoenzymes that are required for virulence (Schell, 2000). In addition to this array of Gram-negative pathogens, several Gram positive microbes also colonize the vasculature and damage plant hosts. Leifsonia xyli subsp. xyli causes ratoon stunting disease in sugar cane through blockage of the xylem (Weaver et al., 1977). Clavibacter michiganensis subsp. sepedonicus causes bacterial ring rot in potato, and attaches to xylem vessels in large aggregates cohered by exopolysaccharide (Marques et al., 2003). Filamentous actinobacteria form aggregates in the intracellular spaces of infected wheat, and it is possible that they are transmitted through the vasculature (Coombs and Franco, 2003). Bioilms on plant roots Plant roots are exposed to the tremendous microbial diversity of the soil environment. Microbial populations associated with soils may mobilize towards plant roots, attracted by exudates, and colonize different areas. he root cap and the zone of cell division is a productive colonization site as are mature root zones and root hairs (Semenov et al., 1999; Watt et al., 2006). he elongation zone of the root, immediately following the root cap, appears to support fewer adherent microbes. Although there are pathogens of plants that associate with roots, many of the adherent populations are benign or beneficial.

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Beneicial root colonizers here are a number of plant-growth promoting microbes that associate with roots. he best studied examples of beneficial plant-microbe interactions are the symbiotic rhizobia that drive the formation of nitrogen fixing nodules on legumes (Table 12.1). Rhizobia are attracted to flavonoid compounds released by legume root hairs, binding to root hair tips (Figure 12.2A). In the well studied formation of so-called indeterminate nodules, Nod factors produced by the rhizobia stimulate root hair curling and capture of the bacteria within the root hair, in which they proceed to migrate down an infection thread through which they invade cells in the root cortex, and establish endosymbiosis (Gage, 2004). Both the populations of rhizobia associated with root hairs and even the bacterial growth down the infection thread can be considered as specialized biofilms (Figure 12.2A). Rhizobia attach to specific host plants via polysaccharides binding to plant lectins and via calcium-binding proteins, initially called rhicadhesin and more recently proposed to be a group of proteins collectively called Rap adhesins (Laus et al., 2006; Russo et al., 2006; Smit et al., 1989). Adherent cells can multiply at the site of colonization to form multicellular assemblies. Rhizobial biofilms also form on abiotic surfaces (Fujishige et al., 2006; Russo et al., 2006). Another plant growth promoting rhizobacterium is Azospirillum brasilense, an Aproteobacterium that associates with the root systems of various cereals (Burdman et al., 2000). Exopolysaccharides, flagellar motility (swimming and swarming) and specific outer membrane proteins are required for effective colonization of cereal root systems. Root hairs and the elongation zone of the root appear to be favored colonization sites, and dense biofilms may be formed at these positions (Assmus et al., 1995). Once the adherent A. brasilense population is in place, they promote plant health by release of a number of different bioactive compounds that stimulate root hair proliferation and lateral root formation (Umali-Garcia et al., 1980). Coinoculation of Azospirillum with nitrogen fixing rhizobia provides enhanced benefits to plant production, suggesting possible synergism within mixed communities of these microbes (Burdman et al., 1998). Several Pseudomonas species and derivatives are effective plant growth promoting rhizobacteria, and some are biocontrol agents (Lugtenberg et al., 2002). On wheat roots, natural populations of pseudomonads comprise a significant proportion of the microbial community, residing within aggregates and biofilms (Watt et al., 2006). Roots are colonized along surface fissures when inoculated with plant growth-promoting pseudomonads, but this has not been commonly observed for natural populations, and thus may be a consequence of the inoculation strategy (Bloemberg et al., 2000). Flagellar motility and twitching are thought to play significant roles in root colonization, although they are not absolutely required for effective deposition (Turnbull et al., 2001; Vande Broek and Vanderleyden, 1995). he release of plant exudates into soils can stimulate rapid mobilization and chemotaxis of pseudomonads towards root systems (Espinosa-Urgel et al., 2002). he LapA cell surface protein (large adhesion protein A) identified in P. fluorescens is required for the transition from reversible to irreversible attachment on abiotic surfaces (Hinsa et al., 2003). A screen of Pseudomonas putida mutants for deficiencies in seed binding identified a lapA homologue in this microbe, and it manifested a deficiency in binding to roots as well (Espinosa-Urgel et al., 2000). hese findings suggest that the LapA protein may function as an adhesin during plant attachment and biofilm formation.

Plant Bioilms

Gram-positive microbes, specifically Bacillus species, can also be effective biocontrol agents (Ugoji et al., 2005). Bacillus species are abundant in the terrestrial environment, and can often colonize plants. Bacillus subtilis develops dense surface-associated populations, and these adherent structures have been correlated with effective biocontrol (Bais et al., 2004). Furthermore, it is clear that a number of functions involved in biocontrol of disease, also effect biofilm formation (Dunn et al., 2003). Root associations of plant pathogens In many cases, the manner in which pathogens associate with root systems is very similar to that of the plant growth promoting bacteria. However, pathogenic pseudomonads have been reported to form thicker, more confluent biofilms on root tissues, in contrast to the more heterogeneous colonization by beneficial pseudomonads (Bais et al., 2004; Walker et al., 2004). his difference may reflect the interactions that lead to disease compared to benign interactions, but may also be the consequence of different inoculation strategies, growth conditions and plant hosts. More studies are needed to compare pathogenic and commensal interactions on the same plant and perhaps in mixed populations. he ubiquitous plant disease called crown gall is caused by Agrobacterium tumefaciens, an A-proteobacterium and a close relative of symbiotic rhizobia (Escobar and Dandekar, 2003). Infection occurs at wound sites along roots and at the point where stem and root tissue converges (the crown), and leads to a horizontal genetic transfer from A. tumefaciens to the plant, directing uncontrolled proliferation of the tissue (the gall) and production of nutrients specific for the infecting microbe. Related species including A. rhizogenes and A. vitis cause hairy root disease and grape-specific necrosis, respectively. he mechanisms of plant attachment have remained elusive, although a two step model mediated initially by an as yet unidentified adhesin and followed by firmer attachment via cellulose fibril production has been widely espoused (Matthysse and Kijne, 1998). Once attached to root tissues, A. tumefaciens can form dense, structurally complex biofilms, extensively coating the epidermis and root hairs (Figure 12.2B) (Matthysse et al., 2005; Ramey et al., 2004). Comparable biofilms form on abiotic surfaces and several mutants and genetic variants affected for biofilm formation on these surfaces, show similar phenotypes on root tissues (Danhorn et al., 2004; Matthysse et al., 2005; Ramey et al., 2004). he role of biofilms during the disease process remains obscure, but may involve proximity to the appropriate infection site, or survival of the basal plant defense response. Oxygen limitation is a common condition in the rhizosphere and also within biofilms (Okinaka et al., 2002). An A. tumefaciens mutant disrupted for the FNR-type transcription factor SinR develops sparse, patchy biofilms on plant roots and abiotic surfaces (Ramey et al., 2004). his regulator is a component of an A. tumefaciens oxygen-limitation response pathway, suggesting a link between oxygen levels and biofilm structure. Similarly, limiting phosphorous is common in the rhizosphere due to plant sequestration. Limiting phosphorous enhances biofilm formation by A. tumefaciens, in contrast to the decreased biofilm formation reported for Pseudomonas aureofaciens in low phosphorous (Danhorn et al., 2004; Monds et al., 2001). It is intriguing that phosphorous resources can have such an opposite result on biofilm formation by two plant associated microbes. For A. tumefaciens, perhaps the prospect of acquiring phosphorous as a result of plant infection, provides a significant

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enough benefit to offset the short term cost. Consistent with this speculation, other aspects of virulence are enhanced by low phosphorous levels (Winans, 1990). Bioilms on seeds and juvenile plants Many seed stocks are deliberately coated with rhizobacteria, such as nitrogen-fixing rhizobia, to promote the health of the juvenile plants that will grow. he biofilms that develop on the seeds during germination inoculate the growing plant and the developing rhizosphere. Bacterial adherence to seeds is therefore of great practical and environmental relevance. Human consumption of seeds and sprouts is also a significant route of infection, and thus of medical importance (Fett and Cooke, 2003). Examination of seed colonization of alfalfa seeds and sprouts from natural sources revealed rods and cocci embedded within exopolysaccharide matrix material (Matos et al., 2002). Quorum sensing and bioilm formation As is a theme of this volume, it has become evident that bacteria within biofilms can coordinate their activities and act in a concerted manner similar to multicellular organisms. hese interactions require cell–cell communication systems to distribute and modulate the various functions within a bacterial community. In support of this conjecture, it is now clear that many bacteria produce and respond to signal molecules, often utilized to monitor their own population densities in a process known as “quorum sensing” or QS (Fuqua et al., 1994). QS is a regulatory mechanism by which bacteria can perceive and respond to their relative numbers and their spatial distribution, a particularly valuable mechanism for gene regulation in densely populated biofilms. Moreover, in several cases QS was shown to be required for the formation and morphogenesis of biofilms (Davies et al., 1998; Huber et al., 2001). Detailed accounts of QS systems and the signaling molecules used to facilitate communication between physically separated cells are provided in Yarwood, Wood and Bentley, and Atkinson et al., this volume. Quorum sensing among plant-associated microbes To date, AHL-dependent quorum sensing circuits have been identified in more than 70 species of Gram-negative bacteria, in which they regulate diverse functions, including bioluminescence, plasmid conjugative transfer, synthesis of antibiotics and extracellular hydrolytic enzymes, motility, and production of virulence factors (Whitehead et al., 2001). In an extensive survey Cha et al. (1998) demonstrated that the majority of plant-associated bacteria produce AHL signal molecules. Specifically, they showed that almost all tested isolates of the genera Agrobacterium, Erwinia, Pantoea, and Rhizobium, and about half of the erwinias and pseudomonads tested synthesized detectable levels of AHLs. By contrast, only a few AHL producers could be identified among Xanthomonas isolates. Elasri et al. (2001) screened 137 soil-borne and plant-associated strains belonging to different Pseudomonas species for their ability to synthesize AHLs, and identified 54 that were positive for AHL production. In this study it was also observed that plant-associated and plant-pathogenic bacteria produced AHLs more frequently than soilborne strains. On the basis of these results it was speculated that the more intimate the relationship of the bacteria with the host plant, the higher the probability that it produces AHLs. Steidle et al. (2001) screened

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over 300 bacterial strains, isolated from the rhizosphere of tomato on standard laboratory media, and approximately 12% of the isolates produced detectable AHLs. Pierson et al. (1998) screened 700 wheat root-associated bacteria using different AHL biosensors and found that about 8% of the strains were able to activate at least one of the sensors. It should be considered that all of the surveys described evaluated culturable isolates, and that even among these bacteria AHL synthesis may not be active under the cultivation conditions employed. Even so, it is clear that AHL QS is common among plant-associated communities. Quorum sensing within plant bioilms he involvement of an AHL-based QS system in the regulation of biofilm formation was first reported for Pseudomonas aeruginosa, an opportunistic human pathogen that causes severe nosocomial infections in immunocompromised individuals and is responsible for the chronic lung infections of patients with cystic fibrosis (Davies et al., 1998; Govan and Deretic, 1996). P. aeruginosa is also capable of causing serious infections in non-mammalian host animals (Mahajan-Miklos et al., 2000) and plants (Rahme et al., 1995) and is an effective colonizer of plant roots (Pandey et al., 2005; Walker et al., 2004). In the study of Davies et al. it was shown that a lasI mutant of P. aeruginosa, defective in synthesis of the AHL N-3-oxo-dodecanoyl- -homoserine lactone (3OC12-HSL), formed biofilms that were flat, densely packed, and undifferentiated (Davies et al., 1998). he las system is one of two quorum-sensing systems that have been identified in P. aeruginosa and consists of the transcriptional activator LasR and the AHL synthase LasI, which directs the synthesis of 3OC12-HSL (Gambello et al., 1993; Pearson et al., 1994, Williams et al., this volume). he second system, designated rhl, consists of RhlR and RhlI, which directs the synthesis of N-butanoyl- -homoserine lactone (C4-HSL) (Ochsner and Reiser, 1995; Pearson et al., 1995). he two systems do not operate independently as the las system positively regulates expression of both rhlR and rhlI (Whiteley et al., 1999). Moreover, the QS circuitry is subject to various additional layers of regulation both at the transcriptional and post-transcriptional level ( Juhas et al., 2005). his complex regulatory network, integrating various environmental parameters, may be the reason that the influence of QS on biofilm formation has been observed to be highly dependent on the experimental conditions (Heydorn et al., 2002; Stoodley et al., 1999, Purevdorj et al., 2002). Most interestingly in the context of this chapter are the findings of Walker et al. (2004), who showed that P. aeruginosa strains are capable of infecting the roots of Arabidopsis and sweet basil (Ocimum basilicum) and cause plant mortality seven days post-inoculation. Prior to plant mortality, P. aeruginosa cells colonize the roots of Arabidopsis and sweet basil and form a biofilm as observed by various microscopic techniques. In accordance with a study by Heydorn et al. (2002) who found no differences between biofilms formed by the wild type and signal negative mutants on inanimate surfaces, Walker et al. (2004) demonstrated that the biofilm of a lasI mutant on the root surface is nearly indistinguishable from the one formed by the wild-type strain. However, QS-deficient mutants were significantly attenuated in their ability to infect sweet basil, confirming the importance of QS in controlling expression of pathogenic traits in P. aeruginosa. QS-regulated functions are also employed by P. aeruginosa to compete with

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other microbes during biofilm formation, and are therefore very likely influencing the composition and activity of the plant-associated community (An et al., 2006). A role for AHL-mediated quorum sensing in biofilm formation has also been demonstrated for several other bacteria that are commonly associated with plants: several Burkholderia species (Huber et al., 2001; Wopperer et al., 2006), P. putida (Steidle et al., 2002), and Serratia liquefaciens (Labbate et al., 2004). Similar to P. aeruginosa, AHL-negative mutants of B. cenocepacia are defective in the late stages of biofilm development and thus were unable to develop biofilms with typical mushroom- and stalk-shaped microcolonies separated by void spaces on abiotic surfaces (Figure 12.2C) (Huber et al., 2001). Employing a quorum quenching approach (i.e. the enzymatic degradation of AHL signal molecules), it was shown that QS not only regulates biofilm formation in B. cenocepacia but also in the large majority of strains from nine other Burkholderia species (Wopperer et al., 2006). he biofilm formed by the P. putida strain IsoF, a plant growth promoting rhizobacterium, on abiotic surfaces is very homogenous with limited structure. In contrast to B. cenocepacia and P. aeruginosa, QS mutants of P. putida IsoF formed structured biofilms, indicating that in this organism QS promotes formation of structurally homogeneous biofilms. In a recent study Dubern et al. (2006) demonstrated that in P. putida PCL1445 the production of the cyclic lipopeptides putisolvin I and II is QS-regulated. As these compounds possess surface tension-reducing abilities and are able to inhibit biofilm formation, a link between QS and biofilm formation could be established. S. liquefaciens forms biofilms through a series of defined stages that lead to a highly porous biofilm composed of cell chains, filaments, and cell clusters. QS plays important roles in the regulation of several stages of the biofilm life cycle of this organism (Labbate et al., 2004; Rice et al., 2005). However, the dependence of biofilm formation on QS is not absolute, as altered nutrient conditions can override QS control in S. liquefaciens. In the multicellular aggregates of the phyllosphere, QS is also an important process. P. syringae regulates motility, exopolysaccharide production and overall virulence during bean leaf interactions via AHL regulation (Quinones et al., 2005). Initial disease symptoms were more severe in AHL-mutants, but the tissue maceration typical of later infection stages, was abolished. Several aspects of epiphytic fitness of P. syringae are strongly influenced by QS, and cell–cell signaling is clearly engaged within the dense multicellular aggregates distributed at various locations on the leaf surface (Quinones et al., 2004). Transgenic bean plants that produce AHLs were less severely damaged than wild type and supported smaller populations of the pathogen. Other epiphytic pathogens such as Erwinia chrysanthemi also employ QS to control virulence (Reverchon et al., 1998). In vascular pathogens, QS plays an important role in regulating virulence factors including exopolysaccharides and degradative enzymes. Interestingly, similar mechanisms of adherence, vascular damage and dispersal are employed by different pathogens, and in many cases QS regulates aspects of these transitions. However, the actual signals employed and their underlying mechanisms fall into several distinct classes. he EsaI-EsaR system from Ps regulates exopolysaccharide synthesis in response to N-3-oxo-hexanoyl- -homoserine lactone (3OC6-HSL) (Koutsoudis et al., 2006; von Bodman et al., 1998). Null mutations in esaR result in stewartan overproduction, attenuated surface adherence, and reduced virulence, probably through non-productive aggregate formation, and limited spreading of the

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pathogen through the vasculature (Koutsoudis et al., 2006). Conversely, esaI null mutants are hyperadherent, and appear to be locked in an adhesive phase of infection, spreading poorly in the vasculature. he related vascular pathogens Xanthomonas campestris pv. campestris (Xcc) and Xylella fastidiosa (Xf) both employ the DSF diffusible signaling compound to control virulence and host interactions (see also Dow et al., this volume). DSF from Xcc is known to be the novel AB unsaturated fatty acid cis-11-methyl-2-dodecenoic acid and the signal in Xf is thought to be similar or identical (Wang et al., 2004). In Xcc DSF is perceived through the RpfC sensor kinase which controls activity of the RpfG response regulator (Dow et al., 2003). RpfG contains a HD-GYP domain, and directs turnover of cyclic diguanosine monophosphate (c-di-GMP), known to regulate many aspects of the cell surface, and hence influencing biofilm formation (Ryan et al., 2006). As with many c-di-GMP regulated processes, the precise connection between the second messenger and the known regulatory targets is not yet clear. In Xf, the DSF signal and Rpf homologues are also involved in disease transmission, although they appear to function in the insect vector, as well as the eventual plant host (Newman et al., 2004). In fact, mutants that cannot synthesize DSF are more virulent when manually inoculated onto plants than the wild type, but are not transmitted effectively from the insect vector. Finally, many studies have demonstrated the degradation of AHL signals by quorum quenching bacteria that produce lactonases and acylases (Zhang and Dong, 2004). Host plants can apparently perceive AHLs and respond to infecting populations (Mathesius et al., 2003). Recent work demonstrated that the presence of AHL-producing bacteria in the tomato rhizosphere increases the salicylic acid levels in the leaves and enhances its systemic resistance against the fungal leaf pathogen Alternaria alternata (Schuhegger et al., 2006). Macroarray analyses showed that synthetic AHLs systemically induce salicylic acid- and ethylene-dependent defense genes, suggesting that AHLs can play an important role in the biocontrol activity of rhizobacteria. Conversely, plants produce compounds that may mimic or inhibit AHL quorum sensing by microbial colonists (Teplitski et al., 2000). It is clear that the biofilms and aggregates of cells colonizing plants are producing and responding to a variety of signal inputs and these systems are certain to influence the microbial activity and persistence in the plant environment. Signal range and amplitude in the rhizosphere Using GFP-based AHL monitor strains it was possible to visualize communication between bacteria belonging to different genera in the rhizosphere of tomato plants (Steidle et al., 2001). hese experiments supported the view that AHL signal molecules can serve as a universal language for communication between different bacterial populations on the root surface. More recently, computer-assisted microscopy in combination with a geostatistical modeling approach was used to quantify the in situ spatial scale of AHL-mediated cell-tocell communication of P. putida on tomato and wheat root surfaces (Gantner et al., 2006). his analysis indicated that the effective “calling distance” on root surfaces was most frequent at 4–5 Mm, extended to 37 Mm in the root tip/elongation zone and further out to 78 Mm in the root hair zone. Interestingly, communication was found to occur not only within dense populations, but also in very small groups and over long ranges between individual bacteria.

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On the basis of this observation it was proposed that communication in the rhizosphere is governed more by the in situ spatial proximity of cells within AHL-gradients than the requirement for a quorum group of high population density. Hence, bacteria appear to use AHL gradients for sensing their positions relative to each other in the rhizosphere, an ability that may be particularly important for the formation of biofilms. Conclusions Bacteria interact extensively with plants and develop into complex multicellular populations. he relevance of these populations to plant-health and disease is just beginning to be understood and appreciated. he coming years of research should help to establish the generalities and specific facets that connect and distinguish the plant-associated biofilms of important pathogens, mutualists and commensals that inhabit the environment of the plant. hese diverse populations are not only physically and metabolically linked to the plant and each other, but also in communication via a diversity of molecular signals that microbiologists and plant researchers are discovering at a rapid pace. Understanding the complexities and diversity of the biofilms formed during plant-microbe interactions will continue to benefit from and contribute to the larger efforts to elucidate biofilm form and function in other host-associated and free-living environments. Acknowledgments Biofilm research is supported by the United States Department of Agriculture (NRI 2002-35319-12636) and the Indiana University META-Cyt program in the Fuqua lab, in the von Bodman lab by the USDA (NRI 2002-35319-1237, and USDA Agricultural Experiment Station grant CNSOO712), and in the Eberl lab by the Swiss National Fond (3100A0–104215). References An, D., Danhorn, T., Fuqua, C., and Parsek, M.R. (2006). Quorum sensing and motility mediate interactions between Pseudomonas aeruginosa and Agrobacterium tumefaciens in biofilm cocultures. Proc. Natl. Acad. Sci. USA 103, 3828–3833. Assmus, B., Hutzler, P., Kirchhof, G., Amann, R., Lawrence, J.R., and Hartmann, A. (1995). In-situ localization of Azospirillum brasilense in the rhizosphere of wheat with fluorescently labeled, ribosomal-RNA-targeted oligonucleotide probes and scanning confocal laser microscopy. Appl. Environ. Microbiol. 61, 1013–1019. Bais, H.P., Fall, R., and Vivanco, J.M. (2004). Biocontrol of Bacillus subtilis against infection of Arabidopsis roots by Pseudomonas syringae is facilitated by biofilm formation and surfactin production. Plant Physiol. 134, 307–319. Bloemberg, G.V., Wij es, A.H.M., Lamers, G.E.M., Stuurman, N., and Lugtenberg, B.J.J. (2000). Simultaneous imaging of Pseudomonas fluorescens WCS365 populations expressing three different autofluorescent proteins in the rhizosphere: new perspectives for studying microbial communities. Mol. Plant Microbe Interact. 13, 1170–1176. Bove, J.M., Renaudin, J., Saillard, C., Foissac, X., and Garnier, M. (2003). Spiroplasma citri, a plant pathogenic molligute: relationships with its two hosts, the plant and the leafhopper vector. Annu. Rev. Phytopathol. 41, 483–500 Epub 2003 Apr 2018. Brlansky, R.H., Timmer, L.W., French, W.J., and McCoy, R.E. (1983). Colonization of the sharpshooter vectors, Oncometopia nigricans and Homalodisca coagulata, by xylem-limited bacteria. Phytopathology 73, 530–535.

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Burdman, S., Jurkevitch, E., Schwartsburd, B., Hampel, M., and Okon, Y. (1998). Aggregation in Azospirillum brasilense: effects of chemical and physical factors and involvement of extracellular components. Microbiology 144, 1989–1999. Burdman, S., Okon, Y., and Jurkevitch, E. (2000). Surface characteristics of Azospirillum brasilense in relation to cell aggregation and attachment to plant roots. Crit. Rev. Microbiol. 26, 91–110. Cha, C., Gao, P., Chen, Y.-C., Shaw, P.D., and Farrand, S.K. (1998). Production of acyl-homoserine lactone quorum-sensing signals by gram-negative plant-associated bacteria. Mol. Plant. Microbe. Interact. 11, 1119–1129. Chang, W.S., and Halverson, L.J. (2003). Reduced water availability influences the dynamics, development, and ultrastructural properties of Pseudomonas putida biofilms. J. Bacteriol. 185, 6199–6204. Coombs, J.T., and Franco, C.M. (2003). Isolation and identification of actinobacteria from surface-sterilized wheat roots. Appl. Environ. Microbiol. 69, 5603–5608. Danhorn, T., Hentzer, M., Givskov, M., Parsek, M., and Fuqua, C. (2004). Phosphorous limitation enhances biofilm formation of the plant pathogen Agrobacterium tumefaciens through the PhoR-PhoB regulatory system. J. Bacteriol. 186, 4492–4501. Davies, D.G., Parsek, M.R., Pearson, J.P., Iglewski, B.H., Costerton, J.W., and Greenberg, E.P. (1998). he involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280, 295–298. Dow, J.M., Crossman, L., Findlay, K., He, Y.Q., Feng, J.X., and Tang, J.L. (2003). Biofilm dispersal in Xanthomonas campestris is controlled by cell–cell signaling and is required for full virulence to plants. Proc. Natl. Acad. Sci. USA 100, 10995–11000 Epub 12003 Sep 10995. Dubern, J.F., Lugtenberg, B.J., and Bloemberg, G.V. (2006). he ppuI-rsaL-ppuR quorum-sensing system regulates biofilm formation of Pseudomonas putida PCL1445 by controlling biosynthesis of the cyclic lipopeptides putisolvins I and II. J. Bacteriol. 188, 2898–2906. Dunn, A.K., Klimowicz, A.K., and Handelsman, J. (2003). Use of a promoter trap to identify Bacillus cereus genes regulated by tomato seed exudate and a rhizosphere resident, Pseudomonas aureofaciens. Appl. Environ. Microbiol. 69, 1197–1205. Elasri, M., Delorme, S., Lemanceau, P., Stewart, G., Laue, B., Glickmann, E., Oger, P.M., and Dessaux, Y. (2001). Acyl-homoserine lactone production is more common among plant-associated Pseudomonas spp. than among soilborne Pseudomonas spp. Appl. Environ. Microbiol. 67, 1198–1209. Escobar, M.A., and Dandekar, A.M. (2003). Agrobacterium tumefaciens as an agent of disease. Trends Plant Sci. 8, 380–386. Espinosa-Urgel, M., Kolter, R., and Ramos, J.-L. (2002). Root colonization by Pseudomonas putida: love at first site. Microbiology 148, 341–343. Espinosa-Urgel, M., Salido, A., and Ramos, J.L. (2000). Genetic analysis of functions involved in adhesion of Pseudomonas putida to seeds. J. Bacteriol. 182, 2363–2369. Fett, W.F., and Cooke, P.H. (2003). Reduction of Escherichia coli O157:H7 and Salmonella on laboratoryinoculated alfalfa seed with commercial citrus-related products. J. Food Prot. 66, 1158–1165. Fujishige, N.A., Kapadia, N.N., De Hoff, P.L., and Hirsch, A.M. (2006). Investigations of Rhizobium biofilm formation. FEMS Microbiol. Ecol. 56, 195–206. Fuqua, W.C., Winans, S.C., and Greenberg, E.P. (1994). Quorum sensing in bacteria: he LuxR/LuxI family of cell density-responsive transcriptional regulators. J. Bacteriol. 176, 269–275. Gage, D.J. (2004). Infection and invasion of roots by symbiotic, nitrogen-fixing rhizobia during nodulation of temperate legumes. Microbiol. Mol. Biol. Rev. 68, 280–300. Gambello, M.J., Kaye, S., and Iglewski, B.H. (1993). LasR of Pseudomonas aeruginosa is a transcriptional activator of the alkaline protease gene (apr) and an enhancer of exotoxon A expression. Infect. Immun. 61, 1180–1184. Gantner, S., Schmid, M., Durr, C., Schuhegger, R., Steidle, A., Hutzler, P., Langebartels, C., Eberl, L., Hartmann, A., and Dazzo, F.B. (2006). In situ quantitation of the spatial scale of calling distances and population density-independent N-acylhomoserine lactone-mediated communication by rhizobacteria colonized on plant roots. FEMS Microbiol. Ecol. 56, 188–194. Govan, J.R.W., and Deretic, V. (1996). Microbial pathogenesis in cystic fibrosis: mucoid Pseudomonas aeruginosa and Burkholderia cepacia. Microbiol. Rev. 60, 539–574. Gregory, P. (2006). Plant Roots: Growth, Activity and Interaction with Soils (Oxford, Blackwell Publishing). Grossman, M., and Dow, J.M. (2004). Biofilm formation and dispersal in Xanthomonas campestris. Microbes Infect. 6, 623–629.

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Heydorn, A., Ersboll, B., Kato, J., Hentzer, M., Parsek, M.R., Tolker-Nielsen, T., Givskov, M., and Molin, S. (2002). Statistical analysis of Pseudomonas aeruginosa biofilm development: impact of mutations in genes involved in twitching motility, cell-to-cell signaling, and stationary-phase sigma factor expression. Appl. Environ. Microbiol. 68, 2008–2017. Hinsa, S.M., Espinosa-Urgel, M., Ramos, J.L., and O’Toole, G.A. (2003). Transition from reversible to irreversible attachment during biofilm formation by Pseudomonas fluorescens WCS365 requires an ABC transporter and a large secreted protein. Mol. Microbiol. 49, 905–918. Hirsch, A.M., Lum, M.R., and Downie, J.A. (2001). What makes the rhizobia-legume symbiosis so special? Plant Physiol. 127, 1484–1492. Huber, B., Riedel, K., Hentzer, M., Heydorn, A., Gotschlich, A., Givskov, M., Molin, S., and Eberl, L. (2001). he cep quorum-sensing system of Burkholderia cepacia H111 controls biofilm formation and swarming motility. Microbiology 147, 2517–2528. Juhas, M., Eberl, L., and Tummler, B. (2005). Quorum sensing: the power of cooperation in the world of Pseudomonas. Environ. Microbiol. 7, 459–471. Kang, Y., Liu, H., Genin, S., Schell, M.A., and Denny, T.P. (2002). Ralstonia solanacearum requires type 4 pili to adhere to multiple surfaces and for natural transformation and virulence. Mol. Microbiol. 46, 427–437. Koutsoudis, M.D., Tsaltas, D., Minogue, T.D., and von Bodman, S.B. (2006). Quorum-sensing regulation governs bacterial adhesion, biofilm development, and host colonization in Pantoea stewartii subspecies stewartii. Proc. Natl. Acad. Sci. USA 103, 5983–5988 Epub 2006 Apr 5983. Krimm, U., Abanda-Nkpwatt, D., Schwab, W., and Schreiber, L. (2005). Epiphytic microorganisms on strawberry plants (Fragaria ananassa cv. Elsanta): identification of bacterial isolates and analysis of their interaction with leaf surfaces. FEMS Microbiol. Ecol. 53, 483–492. Labbate, M., Queck, S.Y., Koh, K.S., Rice, S.A., Givskov, M., and Kjelleberg, S. (2004). Quorum sensingcontrolled biofilm development in Serratia liquefaciens MG1. J. Bacteriol. 186, 692–698. Laus, M.C., Logman, T.J., Lamers, G.E., Van Brussel, A.A., Carlson, R.W., and Kijne, J.W. (2006). A novel polar surface polysaccharide from Rhizobium leguminosarum binds host plant lectin. Mol. Microbiol. 59, 1704–1713. Leite, B., Andersen, P.C., and Ishida, M.L. (2004). Colony aggregation and biofilm formation in xylem chemistry-based media for Xylella fastidiosa. FEMS Microbiol. Lett. 230, 283–290. Lugtenberg, B.J.J., Chin-A-Woeng, T.F., and Bloemberg, G.V. (2002). Microbe-plant interactions: principles and mechanisms. Antonie Van Leeuwenhoek 81, 373–383. Mahajan-Miklos, S., Rahme, L.G., and Ausubel, F.M. (2000). Elucidating the molecular mechanisms of bacterial virulence using non-mammalian hosts. Mol. Microbiol. 37, 981–988. Marques, L.L.R., De Boer, S.H., Ceri, H., and Olsen, M.E. (2003). Evaluation of biofilms formed by Clavibacter michiganensis subsp. sepedonicus. Phytopathology 93, S57. Mathesius, U., Mulders, S., Gao, M., Teplitski, M., Caetano-Anolles, G., Rolfe, B.G., and Bauer, W.D. (2003). Extensive and specific responses of a eukaryote to bacterial quorum-sensing signals. Proc. Natl. Acad. Sci. USA 100, 1444–1449 Epub 2003 Jan 1402. Matos, A., Garland, J.L., and Fett, W.F. (2002). Composition and physiological profiling of sprout-associated microbial communities. J. Food Prot. 65, 1903–1908. Matthysse, A.G., and Kijne, J.W. (1998). Attachment of Rhizobiaceae to plant cells. In: he Rhizobiaceae: Molecular Biology of Model Plant-Associated Bacteria, H.P. Spaink, A. Kondorosi, and P.J.J. Hooykaas, eds. (Dordrecht/Boston/London, Kluwer Academic Publishers), pp. 235–249. Matthysse, A.G., Marry, M., Krall, L., Kaye, M., Ramey, B.E., Fuqua, C., and White, A.R. (2005). he effect of cellulose overproduction on binding and biofilm formation on roots by Agrobacterium tumefaciens. Mol. Plant. Microbe Interact. 18, 1002–1010. Meng, Y., Li, Y., Galvani, C.D., Hao, G., Turner, J.N., Burr, T.J., and Hoch, H.C. (2005). Upstream migration of Xylella fastidiosa via pilus-driven twitching motility. J. Bacteriol. 187, 5560–5567. Minogue, T.D., Carlier, A.L., Koutsoudis, M.D., and von Bodman, S.B. (2005). he cell density-dependent expression of stewartan exopolysaccharide in Pantoea stewartii ssp. stewartii is a function of EsaRmediated repression of the rcsA gene. Mol. Microbiol. 56, 189–203. Monds, R.D., Silby, M.W., and Mahanty, H.K. (2001). Expression of the Pho regulon negatively regulates biofilm formation by Pseudomonas aureofaciens PA147-2. Mol. Microbiol. 42, 415–426. Monier, J.M., and Lindow, S.E. (2004). Frequency, size, and localization of bacterial aggregates on bean leaf surfaces. Appl. Environ. Microbiol. 70, 346–355.

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Monier, J.M., and Lindow, S.E. (2005a). Aggregates of resident bacteria facilitate survival of immigrant bacteria on leaf surfaces. Microb. Ecol. 49, 343–352 Epub 2005 Jul 2007. Monier, J.M., and Lindow, S.E. (2005b). Spatial organization of dual-species bacterial aggregates on leaf surfaces. Appl. Environ. Microbiol. 71, 5484–5493. Morris, C.E., and Monier, J.-M. (2003). he ecological significance of biofilm formation by plant-associated bacteria. Annu. Rev. Phytopathol. 41, 429–453. Newman, K.L., Almeida, R.P., Purcell, A.H., and Lindow, S.E. (2003). Use of a green fluorescent strain for analysis of Xylella fastidiosa colonization of Vitis vinifera. Appl. Environ. Microbiol. 69, 7319–7327. Newman, K.L., Almeida, R.P., Purcell, A.H., and Lindow, S.E. (2004). Cell–cell signaling controls Xylella fastidiosa interactions with both insects and plants. Proc. Natl. Acad. Sci. USA 101, 1737–1742 Epub 2004 Jan 1730. Ochsner, U.A., and Reiser, J. (1995). Autoinducer-mediated regulation of rhamnolipid biosurfactant synthesis in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 92, 6424–6428. Okinaka, Y., Yang, C.-H., Perna, N.T., and Keen, N.T. (2002). Microarray profiling of Erwinia chrysanthemi 3937 genes that are regulated during plant infection. Mol. Plant Microbe Interact. 15, 619–629. Pandey, P., Kang, S.C., Gupta, C.P., and Maheshwari, D.K. (2005). Rhizosphere competent Pseudomonas aeruginosa GRC (1) produces characteristic siderophore and enhances growth of Indian mustard (Brassica campestris). Curr. Microbiol. 51, 303–309. Pearson, J.P., Gray, K.M., Passador, L., Tucker, K.D., Eberhard, A., Iglewski, B.H., and Greenberg, E.P. (1994). Structure of the autoinducer required for expression of Pseudomonas aeruginosa virulence genes. Proc. Natl. Acad. Sci. USA 91, 197–201. Pearson, J.P., Passador, L., Iglewski, B.H., and Greenberg, E.P. (1995). A second N-acylhomoserine lactone signal produced by Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 92, 1490–1494. Pedgley, D.E. (1991). Aerobiology: he atmosphere as a source and sink for microbes. In: Microbial Ecology of Leaves, A.J.H., and S.S. Hirano, eds. (New York, Springer-Verlag). Pierson, E.A., Wood, D.W., Cannon, J.A., Blachere, F.M., and Pierson, L.S. (1998). Interpopulation signaling via N-acyl-homoserine lactones among bacteria in the wheat rhizosphere. Mol. Plant Microbe Interact. 11, 1078–1084. Purevdorj, B., Costerton, J.W., and Stoodley, P. (2002). Influence of hydrodynamics and cell signaling on the structure and behavior of Pseudomonas aeruginosa biofilms. Appl. Environ. Microbiol. 68, 4457–4464. Quinones, B., Dulla, G., and Lindow, S.E. (2005). Quorum sensing regulates exopolysaccharide production, motility, and virulence in Pseudomonas syringae. Mol. Plant Microbe Interact. 18, 682–693. Quinones, B., Pujol, C.J., and Lindow, S.E. (2004). Regulation of AHL production and its contribution to epiphytic fitness in Pseudomonas syringae. Mol. Plant Microbe Interact. 17, 521–531. Rahme, L.G., Stevens, E.J., Wolfort, S.F., Shao, J., Tompkins, R.G., and Ausubel, F.M. (1995). Common virulence factors for bacterial pathogenicity in plants and animals. Science 268, 1899–1902. Ramey, B.E., Matthysse, A.G., and Fuqua, C. (2004). he FNR-type transcriptional regulator SinR controls maturation of Agrobacterium tumefaciens biofilms. Mol. Microbiol. 52, 1495–1511. Reverchon, S., Bouillant, M.L., Salmond, G., and Nasser, W. (1998). Integration of the quorum-sensing system in the regulatory networks controlling virulence factor synthesis in Erwinia chrysanthemi. Mol. Microbiol. 29, 1407–1418. Rice, S.A., Koh, K.S., Queck, S.Y., Labbate, M., Lam, K.W., and Kjelleberg, S. (2005). Biofilm formation and sloughing in Serratia marcescens are controlled by quorum sensing and nutrient cues. J. Bacteriol. 187, 3477–3485. Russo, D.M., Williams, A., Edwards, A., Posadas, D.M., Finnie, C., Dankert, M., Downie, J.A., and Zorreguieta, A. (2006). Proteins exported via the PrsD-PrsE type I secretion system and the acidic exopolysaccharide are involved in biofilm formation by Rhizobium leguminosarum. J. Bacteriol. 188, 4474–4486. Ryan, R.P., Fouhy, Y., Lucey, J.F., Crossman, L.C., Spiro, S., He, Y.W., Zhang, L.H., Heeb, S., Cámara, M., Williams, P., et al. (2006). Cell–cell signaling in Xanthomonas campestris involves an HD-GYP domain protein that functions in cyclic di-GMP turnover. Proc. Natl. Acad. Sci. USA 103, 6712–6717 Epub 2006 Apr 6712. Schell, M.A. (2000). Control of virulence and pathogenicity genes of Ralstonia solanacearum by an elaborate sensory network. Annu. Rev. Phytopathol. 38, 263–292.

|

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|

Eberl et al.

Schuhegger, R., Ihring, A., Gantner, S., Bahnweg, G., Knappe, Vogg, G., Hutzler, P., Schmid, M., Van Breusegem, M., Eberl, L., Hartmann, A., and Langebartels C. (2006). Induction of systemic plant resistance by N-acylhomoserine lactone-producing rhizosphere bacteria. Plant Cell Environ. 29, 909–918. Semenov, A.M., van Bruggen, A.H.C., and Zelenev, V.V. (1999). Moving waves of bacterial populations and total organic carbon along roots of wheat. Microb. Ecol. 37, 116–128. Smit, G., Logman, T.J.J., Boerrigter, M.E.T.I., Kijne, J.W., and Lugtenberg, B.J.J. (1989). Purification and partial characterization of the Ca2+ dependent adhesin from Rhizobium leguminosarum biovar viciae, which mediates the first step in attachment of Rhizobiaceae cells to plant root hair tips. J. Bacteriol. 171, 4054–4062. Steidle, A., Allesen-Holm, M., Riedel, K., Berg, G., Givskov, M., Molin, S., and Eberl, L. (2002). Identification and characterization of an N-acylhomoserine lactone-dependent quorum-sensing system in Pseudomonas putida strain IsoF. Appl. Environ. Microbiol. 68, 6371–6382. Steidle, A., Sigl, K., Schuhegger, R., Ihring, A., Schmid, M., Gantner, S., Stoffels, M., Riedel, K., Givskov, M., Hartmann, A., et al. (2001). Visualization of N-acylhomoserine lactone-mediated cell–cell communication between bacteria colonizing the tomato rhizosphere. Appl. Environ. Microbiol. 67, 5761–5770. Stoodley, P., Lewandowski, Z., Boyle, J.D., and Lappin-Scott, H.M. (1999). Structural deformation of bacterial biofilms caused by short-term fluctuations in fluid shear: an in situ investigation of biofilm rheology. Biotechnol. Bioeng. 65, 83–92. Tans-Kersten, J., Huang, H., and Allen, C. (2001). Ralstonia solanacearum needs motility for invasive virulence on tomato. J. Bacteriol. 183, 3597–3605. Teplitski, M., Robinson, J.B., and Bauer, W.D. (2000). Plants secrete substances that mimic bacterial N-acyl homoserine lactone signal activities and affect population density-dependent behaviors in associated bacteria. Mol. Plant Microbe Interact. 13, 637–648. Turnbull, G.A., Morgan, J.A.W., Whipps, J.M., and Saunders, J.R. (2001). he role of bacterial motility in the survival and spread of Pseudomonas fluorescens in soil and in the attachment in bacterial-plant interactions. FEMS Microb. Ecol. 35, 57–65. Ugoji, E.O., Laing, M.D., and Hunter, C.H. (2005). Colonization of Bacillus spp. on seeds and in plant rhizoplane. J. Environ. Biol. 26, 459–466. Umali-Garcia, M., Hubbell, D.H., Gaskins, M.H., and Dazzo, F.B. (1980). Association of Azospirillum with grass roots. Appl. Environ. Microbiol. 39, 219–226. Vande Broek, A., and Vanderleyden, J. (1995). he role of bacterial motility, chemotaxis, and attachment in bacterial-plant interactions. Mol. Plant Microbe Interact. 8, 800–810. von Bodman, S.B., Bauer, W.D., and Coplin, D.L. (2003). Quorum sensing in plant-pathogenic bacteria. Annu. Rev. Phytopathol. 41, 455–482 Epub 2003 Apr 2029. von Bodman, S.B., Majerczak, D.R., and Coplin, D.L. (1998). A negative regulator mediates quorumsensing control of exopolysaccharide production in Pantoea stewartii subsp. stewartii. Proc. Natl. Acad. Sci. USA 95, 7687–7692. Walker, T.S., Bais, H.P., Deziel, E., Schweizer, H.P., Rahme, L.G., Fall, R., and Vivanco, J.M. (2004). Pseudomonas aeruginosa-plant root interactions. Pathogenicity, biofilm formation, and root exudation. Plant Physiol. 134, 320–331 Epub 2003 Dec 2030. Walker, T.S., Bais, H.P., Grotewold, E., and Vivanco, J.M. (2003). Root exudation and rhizosphere biology. Plant Physiol. 132, 44–51. Wang, L.H., He, Y., Gao, Y., Wu, J.E., Dong, Y.H., He, C., Wang, S.X., Weng, L.X., Xu, J.L., Tay, L., et al. (2004). A bacterial cell–cell communication signal with cross-kingdom structural analogues. Mol. Microbiol. 51, 903–912. Watt, M., Hugenholtz, P., White, R., and Vinall, K. (2006). Numbers and locations of native bacteria on field-grown wheat roots quantified by fluorescence in situ hybridization (FISH). Environ. Microbiol. 8, 871–884. Weaver, L., Teakle, D.S., and Hayward, A.C. (1977). Ultrastructural studies on the bacterium associated with the ratoon stunting disease of sugar cane. Aust. J. Agric. Res. 28, 843–852. Whitehead, N.A., Barnard, A.M.L., Slater, H., Simpson, N.J.L., and Salmond, G.P.C. (2001). Quorumsensing in gram-negative bacteria. FEMS Microbiol. Rev. 25, 365–404. Whiteley, M., Lee, K.M., and Greenberg, E.P. (1999). Identification of genes controlled by quorum sensing in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. USA 96, 13904–13909.

Plant Bioilms

Winans, S.C. (1990). Transcriptional induction of an Agrobacterium regulatory gene at tandem promoters by plant-released phenolic compounds, phosphate starvation, and acidic growth media. J. Bacteriol. 172, 2433–2438. Wopperer, J., Cardona, S.T., Huber, B., Jacobi, C.A., Valvano, M.A., and Eberl, L. (2006). A quorumquenching approach to investigate the conservation of quorum-sensing-regulated functions within the Burkholderia cepacia complex. Appl. Environ. Microbiol. 72, 1579–1587. Zhang, L.H., and Dong, Y.H. (2004). Quorum sensing and signal interference: diverse implications. Mol. Microbiol. 53, 1563–1571.

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Deborah A. Hogan, Matthew J. Wargo, and Nancy Beck

Abstract Bacterial biofilm formation on fungi participates in the synergistic degradation of substrates, antagonism of fungal growth, bacterial utilization of fungi as nutrient sources, and the formation of more complex synergistic associations for the purposes of nutrient acquisition. While bacterial biofilm formation has been described in many systems, the molecular mechanisms that govern these interactions are not yet well understood. Analysis of physical interactions between Pseudomonas aeruginosa and the dimorphic opportunistic fungal pathogen Candida albicans has provided insights into factors involved in attachment and matrix production, and has demonstrated a role for the bacterial quorum sensing molecule, 3-oxo-C12-homoserine lactone, within bacterial-fungal biofilms. Subsequent to P. aeruginosa biofilm formation on the fungus, extracellular bacterial products contribute to the death of the fungal hyphae. Studies focused on the interactions between bacteria and fungi in the phytosphere have illustrated additional processes that contribute to bacterial biofilm formation on fungi including bacterial chemotaxis towards fungal cells, the crossfeeding of nutrients between interacting species, and the expression of specific genes upon contact with fungal cells. By understanding bacterial biofilm formation on fungi, we will gain insight into economically important interactions, such as those involved in the bacterial biocontrol of fungal plant pathogens. Furthermore, using tractable bacterial-fungal biofilm model systems, we may uncover important elements of bacterial biofilms on other living surfaces such as plant and animal tissues. Overview: bacterial bioilms on fungal surfaces From the perspective of the bacterium, there are a number of ways in which biofilm formation on fungi can be beneficial. First, bacterial colonization of a fungal surface may enable the bacteria to exploit the fungus as a source of nutrients (Hogan and Kolter, 2002). Bacteria may scavenge nutrients from the fungal cell wall, consume fungi-secreted products, or induce lysis of the fungal cells thereby liberating the intracellular contents for consumption by the local bacterial population. Second, in communities where bacteria and fungi are in competition for nutrients, biofilm formation on fungal cells could enhance bacterial antagonism of fungi by concentrating bacterially derived antifungal compounds. hird, biofilm formation on the surface of fungal hyphae would enable bacteria to “travel” with fungi as they extend into new areas in search of nutrients. Fourth, bacterial attachment to the hyphal surfaces may enhance synergistic actions of bacteria and fungi needed to breakdown complex substrates. Lastly, bacterial colonization of a fungal surface may be a first step in

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more complex bacterial-fungal endosymbiont interactions such as those that are critical in the root rhizosphere (Dorr et al., 1998). he wide array of different interactions between bacteria and fungi illustrates the potential importance of these relationships in many freeliving and host-associated ecosystems. In this chapter, we focus on the molecular factors involved in bacterial biofilm formation on fungi. To facilitate future research in this area, this chapter aims to highlight a number of different techniques and concepts that are important in the study of these specialized interspecies interactions. We mainly focus on biofilm formation by Pseudomonads on the surfaces of fungal hyphae, though the biofilm interactions between fungi and a handful of other Gram-positive and Gram-negative bacteria are also discussed. he chapter is divided into two sections. First, the interactions between two opportunistic pathogens (Pseudomonas aeruginosa and the fungus Candida albicans) are discussed. Second, we address interactions between bacteria and fungi in the rhizosphere and phyllosphere. Pseudomonas aeruginosa bioilm formation on Candida albicans Biofilm interactions between the Gram-negative bacterium P. aeruginosa and the fungus C. albicans may have relevance to the study of infections associated with cystic fibrosis (CF). Individuals with CF, a genetic disease that results from mutations in the CFTR transmembrane conductance regulator, are highly susceptible to chronic, progressive pulmonary infections that severely damage lung tissues and most often lead to respiratory failure in early adulthood (Rajan and Saiman, 2002). Several lines of evidence suggest that the microorganisms in CF sputum are in a biofilm-like state (Costerton et al., 1999; Hoiby et al., 2001; Singh et al., 2000). While the predominant colonizer of the CF lung is P. aeruginosa, C. albicans, a dimorphic fungus, and Aspergillus fumigatus, another opportunistic fungal pathogen, are also commonly observed (Bakare et al., 2003; Bauernfeind et al., 1987; Bhargava et al., 1989; Burns et al., 1999; Cheng et al., 1990; Haase et al., 1991; Hughes and Kim, 1973; Navarro et al., 2001). he effects of mixed bacterial-fungal infections on the host lung are not yet known. In vitro analysis of the relationship between P. aeruginosa and C. albicans has shown that P. aeruginosa attaches to and forms biofilms on the surface of C. albicans (Hogan and Kolter, 2002). Within the P. aeruginosa biofilms, the fungal hyphae are killed. Studies using different P. aeruginosa and C. albicans strains have yielded data that support the hypothesis that P. aeruginosa biofilm formation is necessary, but not sufficient, for killing (Hogan and Kolter, 2002). hus, the P. aeruginosa–C. albicans biofilm interaction may enable us to study some of the links between biofilm formation and virulence. Below we describe several aspects of the P. aeruginosa–C. albicans relationship including attachment, biofilm development, quorum sensing regulation, and P. aeruginosa killing of fungal cells. Each of these topics will be discussed separately. Attachment When P. aeruginosa and C. albicans are co-cultured in carbon-limited minimal medium at 37oC, the bacteria readily attach to the fungal filaments (Figure 13.1A) (Hogan and Kolter, 2002). Subsequently, the attached cells form a dense biofilm over the course of

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A

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C

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F

Figure 13.1 Attachment and bioilm formation on fungi by different Pseudomonas species. A-D Bioilm formation by P. aeruginosa PA14 on C. albicans hyphae. (A) P. aeruginosa initial colonization of the fungus as shown by phase contrast microscopy to demonstrate attachment by one bacterial pole. (B) P. aeruginosa bioilm formation at 24h is shown by differential interference contrast microscopy (DIC). (C-D) DIC images of P. aeruginosa bioilms to show colonization of C. albicans hyphae, but not yeast (C) and the development of dense bacterial bioilms between fungal hyphae (D). (E) Confocal laser scanning microscopy analyses of the attachment and colonization of Fusarium oxysporum. f.sp. radicis lycopersici (tagged with CFP) by Pseudomonas luorescens WCS365 (tagged with E-GFP). A colonized hyphae (E) and a colonized spore (E-inset) are shown. (F) Pseudomonas syringae pv. syringae B728a, constitutively expressing gfp, colonizing the surface of an unlabeled Neurospora crassa hypha grown over on water-agar. Images 1A-D captured by N.B. and D.A.H. Figure 13.1E is courtesy of S. de Weert and G.V. Bloemberg. Figure 13.1F is courtesy of G. Wichmann and S. Lindow.

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24–72 hours (Figure 13.1B) (Hogan and Kolter, 2002). here is a significant degree of specificity to the factors that are involved in the physical interactions between P. aeruginosa and C. albicans. While P. aeruginosa quickly colonizes C. albicans hyphae, it cannot attach to, nor form biofilms on, yeast-form C. albicans cells (Hogan and Kolter, 2002) (Figure 13.1C). C. albicans hyphal growth is stimulated by numerous factors including nutrient limitation, serum and certain amino acids; hyphae induced under any of these conditions support P. aeruginosa attachment. he reason for the differential attachment to hyphae and yeast is not yet known and could be due to the presence of epitopes that are unique to hyphae, or to factors that obstruct bacterial attachment on yeast cells. Numerous differences between the protein profiles, carbohydrate composition and surface charge of hyphae versus yeast have been reported (Chaffin et al., 1998). Because P. aeruginosa is able to form biofilms on fungi other than C. albicans, such as Aspergillus nidulans and Alternaria alternato (Hogan unpublished data), it appears that either P. aeruginosa recognizes surface structures that are common across a wide range of ascomycete fungi or that P. aeruginosa can recognize multiple fungal surface structures. hese two possibilities are not mutually exclusive. he bacterial factors that participate in fungal attachment are under regulatory control. P. aeruginosa attachment is enhanced by unknown factors that are regulated by cell density-dependent acylhomoserine lactone signals and nutrient availability (Hogan and Kolter, 2002). Interestingly, some P. aeruginosa strains, including strain PAO1, do not readily attach to C. albicans hyphae when grown under the conditions that promote P. aeruginosa strain PA14 attachment (Hogan and Kolter, 2002). hese data illustrate that P. aeruginosa strains have different bacterial surface characteristics or structures that will impact their ability to physically interact with different cell types. As has been described for bacterial attachment to abiotic surfaces such as glass or plastic (Marshall et al., 1971) (see MacEachran and O’Toole, this volume) and to plant cells (Hendrickson et al., 2001), P. aeruginosa PA14 attachment to the fungus occurs by one bacterial pole (Hogan and Kolter, 2002). Time-lapse microscopy shows that bacterial attachment to fungal hyphae is at first reversible with P. aeruginosa cells rapidly attaching to and detaching from the hyphal surface (data not shown). At some frequency, cells remain attached to the hyphal cell and initiate the formation of biofilms (Figure 13.1B–D). Bioilm development After individual P. aeruginosa cells have colonized the surfaces of the fungal hyphae, the formation of mature biofilms on the fungal surfaces is observed. he extent of the similarities between biofilm formation on abiotic surfaces and biofilm formation on fungi are not yet known. A number of P. aeruginosa mutants that are defective in P. aeruginosa attachment to plastic are also defective in biofilm formation on fungi. For example, flagellar mutants are delayed in biofilm formation on both abiotic surfaces and C. albicans hyphae; and P. aeruginosa mutants defective in the retractile type IV pili, which form weak, undifferentiated biofilms on plastic, produce biofilms on fungi that are thicker and less tightly packed compared to those formed by the wild type (Hogan and Kolter, 2002; O’Toole and Kolter, 1998). An extracellular matrix surrounds the bacterial and fungal cells within P. aeruginosa biofilms on C. albicans hyphae (Figure 13.1C), and over the course of 72 hours, the biofilms continue to grow to fill in the spaces between C. albicans filaments (Figure 13.1D).

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Since both organisms are capable of producing an extracellular matrix, it is unclear if the matrix is of bacterial or fungal origin or if it is comprised of a mixture of the materials from both organisms (Chandra et al., 2001; Friedman and Kolter, 2004a; Friedman and Kolter, 2004b; Hawser et al., 1998; Jackson et al., 2004; Matsukawa and Greenberg, 2004) For more information on biofilm matrix, see Pamp et al., this volume. P. aeruginosa antagonism towards C. albicans Using both vital staining techniques and viable counts, it has been shown that the fungal hyphae within P. aeruginosa biofilms are killed within 24–48 hours of biofilm formation (Hogan and Kolter, 2002). To identify those factors that participate in this antagonistic interaction, the rate of fungal killing by different P. aeruginosa mutants were compared to the wild type using a quantitative, plate-count assay to monitor fungal viability of a constitutively filamentous C. albicans tup1/tup1 mutant (Braun and Johnson, 1997; Hogan and Kolter, 2002). Under the nutrient-limiting conditions of the assay, biofilm formation is necessary for fungal killing. P. aeruginosa mutants that lack a functional flagellum, such as the flgK mutant, are delayed in biofilm formation on the fungal surface and are delayed in fungal killing. Mutants lacking certain global regulators, such as RpoN, or quorum sensingrelated transcription factors, such as LasR and RhlR, were both defective in biofilm formation (see Atkinson et al., this volume) and decreased in their ability to kill C. albicans hyphae (Hogan and Kolter, 2002). hough these mutations in global regulators are pleiotropic, the correlation between the ability to form biofilms and fungal killing towards C. albicans is consistent with other data that link biofilm formation and virulence. Virulence factors that have been implicated in human disease, such as the secreted phospholipase C, which degrades eukaryotic membrane lipids, also participate in biofilmrelated fungal killing (Hogan and Kolter, 2002; Hollsing et al., 1987; Lanotte et al., 2003; Woods et al., 1997). he fact that biofilm formation enhances, and may be required for, killing suggests that these genes may be differentially regulated within biofilms on biotic surfaces, or that these secreted factors are more efficacious when they are produced in close proximity to the target cells. Because P. aeruginosa biofilm formation on C. albicans occurs much more readily under nutrient limiting conditions, it has been speculated that the bacteria are using the fungal hyphae as a source of nutrients and that biofilm formation allows for the synergistic action of degradative enzymes and for the capture of nutrients released upon fungal lysis. Quorum sensing molecules in bacterial bioilms on fungi When P. aeruginosa and C. albicans were co-cultured, the presence of the bacteria promoted growth as yeast-form cells despite conditions that would normally stimulate C. albicans hyphal growth. Because P. aeruginosa can neither colonize, nor kill yeast cells, this change in morphology allows C. albicans cells to survive in the presence of P. aeruginosa. hrough a genetic screen, it was found that P. aeruginosa mutants defective in the production of 3oxo-C12 homoserine lactone, a cell–cell signaling molecule, are unable to inhibit C. albicans filamentation (Hogan et al., 2004). he addition of 3OC12HSL alone to inducing medium at concentrations between 20–200 MM is sufficient to block hyphae formation with no effects on overall growth. he accumulation of 3OC12HSL in P. aeruginosa biofilms or in

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colonies likely contributes to this interaction (Charlton et al., 2000) as supernatants from planktonic cultures do not contain sufficient quantities of 3OC12HSL to affect C. albicans morphology. hese findings indicate the possibility that there are chemical interactions within biofilms that are difficult or impossible to detect in planktonic co-cultures (Hogan et al., 2004). Pseudomonas spp. bioilm interactions with fungi in association with plants In soils and in association with plant surfaces, there are many opportunities for physical interactions between bacteria and fungi. Fungi-associated bacterial communities appear to be distinct from the microflora in bulk soil in terms of species composition and relative abundance with Pseudomonas and Burkholderia frequently being detected as fungal colonizers (Frey-Klett et al., 2005; Rangel-Castro et al., 2002; Timonen and Hurek, 2006). hese bacterial–fungal interactions likely play very important roles in microbial community ecology. Some bacterial species can serve as biocontrol agents that protect plants from pathogenic fungi (Whipps, 2001). For example, P. fluorescens can effectively protect tomato plants against infection by Fusarium oxysporum and its ability to colonize both the roots and fungal cells (Figure 13.1E) likely aids in this interaction (Bolwerk et al., 2003). Recent unpublished work by G. Wichmann and S. Lindow has found that Pseudomonas syringae readily colonizes Neurospora crassa hyphae under a variety of conditions. Like P. aeruginosa attachment to C. albicans (described above) initial colonization of fungal hyphae by P. syringae involves polar attachment to the fungal surface followed by biofilm development (Figure 13.1F) (G. Wichmann and S. Lindow, unpublished data). Two different pathovars, P. syringae pv. syringae B728a and pv. tomato DC3000 are able to colonize the fungal hyphae, and contact between P. syringae and N. crassa correlates with fungal cell death. As P. syringae is commonly found on plant surfaces where it interacts with numerous microbial species including yeasts and filamentous fungi (Lindow and Brandl, 2003), the antagonistic interactions demonstrated between P. syringae and N. crassa may reflect a role for bacterial biofilm formation in competition with fungi in the phyllosphere. Other bacterial-fungal interactions that occur in association with plants are those within mycorrhizas. Mycorrhizas are symbiotic associations between fungi and plants for the purposes of nutrient acquisition; these associations also often contain helper bacteria that promote the formation of these beneficial structures (Aspray et al., 2006; Garbaye, 1994). Like the antagonistic interactions described above, the formation of mycorrhiza also involves a number of physical interactions between bacteria and fungi. As described for P. aeruginosa interactions with C. albicans above, the interactions can be broken down into several stages including detection of the fungal host, attachment to the fungal cells, and the response of the bacterium to growth on a biotic, fungal surface. Examples of chemotaxis towards fungal products, colonization of fungal surfaces, and induction of antifungal activities are discussed in detail below. Bacterial detection of the fungus here are many types of motility that may enable bacteria, such as Pseudomonas spp. and Burkholderia spp. to reach the fungus. Studies by Martinez-Granero et al. have found that the

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rhizosphere, which contains abundant fungal hyphae, selects for P. fluorescens phenotypic variants that are highly motile suggesting that motility plays a critical role in accessing and exploiting this environment (Martinez-Granero et al., 2006). Several examples illustrate that soil Pseudomonads exhibit chemotaxis towards plant and fungal exudates thereby allowing bacteria to congregate around populations of fungi in soils (Chet et al., 1971; Lugtenberg et al., 2001). Specific compounds that have been shown to elicit chemotaxis are fusaric acid, a secreted mycotoxin (de Weert et al., 2004) and trehalose, a sugar commonly found in fungi (hevelein, 1984; P. Frey-Klett, personal communication). In some cases, the viability of the fungus affects bacterial attachment. In one study, Pseudomonas fluorescens was shown to attach fifty-two times better to the live hyphae of a specific Glomus sp. than to dead hyphae, while Bacillus cereus was seen to attach better to the dead Glomus spp. hyphae (Toljander et al., 2006). Bacillus has also been shown to attach preferentially to damaged hyphae in an in vivo analysis (Artursson and Jansson, 2003). he factors that contribute to the differentiation between live and dead hyphae are not yet known. Selective colonization of live hyphae may reflect a selection for bacteria capable of exploiting fungal metabolites, while attachment to only dead hyphae may imply the role of some bacteria in succession within the community. Furthermore, in some instances, bacteria colonization of only dead hyphae may suggest that some fungi have active mechanisms to resist bacterial colonization. Bacterial attachment and bioilm formation Recent studies have begun to examine the molecular features of soil bacteria and fungi that allow for bacterial colonization of the fungal surface. Different bacteria can recognize a number of fungal species in a variety of morphological states including hyphae, spores, and fruiting bodies (Levy et al., 2003; Malajczuk et al., 1977; Rangel-Castro et al., 2002) as well as on and in mycorrhizas (Frey-Klett et al., 1997). Furthermore, fungal-associated bacteria can be selective in terms of which fungal species they attach to, and in the particular region of the fungus to be colonized. For example, a variety of unidentified, but morphologically diverse, bacteria preferentially colonize the mycorrhiza instead of the radial hyphae network of fungi associated with pine roots (Nurmiaho-Lassila et al., 1997). Electron microscopic analysis of Burkholderia sp. colonization of an arbuscular mycorrhizal fungus, Gigaspora decipiens, shows the involvement of fibrillar structures of unknown composition (Levy et al., 2003). Dorr et al. (1998) showed that an Azoracus sp. uses type IV pili, and that pili-mediated interactions were critical for adhesion and establishment of mycorrhizal associations with the fungus. As these relationships are studied further, the extent of biofilm maturation on the fungal surface will be examined and the role of other previously identified biofilm development genes will be explored. Bacterial response within fungal bioilms One of the most exciting aspects of research on bacterial biofilm formation on fungi is the analysis of how microbial physiologies are affected during these intimate and dynamic interactions between bacterium and fungus. Using an in vivo expression technology (IVET) approach, Lee and Cooksey (2000) identified four Pseudomonas putida genes that are specifically induced during the growth of this biocontrol bacterium on the surface of the

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fungus Phytophthora parasitica. he genes associated with fungal colonization included an uncharacterized transcription factor, an ABC transporter, and a porin (Lee and Cooksey, 2000). Separate studies have shown that trehalose may prove to be an important molecule during bacterial interactions with fungi. While trehalose utilization is not common among bacteria, many Pseudomonas spp. can grow on this fungally derived substrate. It has been proposed that the ability for Pseudomonas spp. to utilize trehalose may be an important part of bacterial growth in association with fungi such as the edible ectomycorrhizal fungus, Cantharellus cibarius, without causing fungal cell damage (Rangel-Castro et al., 2002). A different P. fluorescens sp. has been shown to induce its trehalose utilization genes when exposed to fungal culture supernatant, and the presence of trehalose enhances inhibition of Pythium debaryanum in a radial growth assay (Gaballa et al., 1997; Rincon et al., 2005). here is evidence that other uncharacterized chemical and physical signals likely also participate in the bacterial response to growth on fungi. Close association between Burkholderia sp. and fungi has been shown to promote mycorrhiza formation (Aspray et al., 2006), and Azoarcus sp., which uses type IV pili to attach to the fungal surface, only develops the intracellular structures necessary for efficient nitrogen fixation when the bacterial and fungal species are grown together (Dorr et al., 1998). Summary and future directions he study of bacterial biofilm formation on fungi could lead in a number of exciting directions. First, the identification of factors involved in the attachment and colonization of specific bacterial–fungal pairs could greatly aid in the development of more effective strains for biocontrol applications or degradation of complex substrates. he identification of the attachment factors themselves, along with an understanding of their regulation in accordance with environmental conditions may also provide insight into those environments where bacterial-fungal interactions are important. A second interesting area of research is the response of the fungus to bacterial biofilm formation on its surface. he identification of fungal factors that can block or disrupt bacterial biofilm formation, or that can kill bacterial biofilm cells could have important practical applications. Bacterial biofilms on implanted medical devices are highly problematic and costly in the clinic, and the high levels of antibiotic resistance exhibited by biofilm bacteria is thought to contribute to the inability to treat some biofilm-associated human infections. Lastly, the molecular analysis of bacterial factors that are important for beneficial and antagonistic biofilm formation on microscopic eukaryotes, including fungi, may provide new insight into those genes and factors that participate in the colonization of other eukaryotic cells including those belonging to plant and animals. Acknowledgments his work was funded by the Pew Biomedical Scholars Program (D.A.H) and the National Institutes of Health P20-RR018787 from the IDeA Program of the National Center for Research Resources (D.A.H.), T32 AI07519 (M.J.W.), and T32 DK007301 (N.B.). We would like to thank Dr. G. Bloemberg and Dr. S. de Weert, University of Leiden, for contributing images of P. fluorescens on Fusarium cells. Special thanks to G. Wichmann and S. Lindow, Univ. of California, Berkeley, for their contribution of unpublished findings.

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References Artursson, V., and Jansson, J.K. (2003). Use of bromodeoxyuridine immunocapture to identify active bacteria associated with arbuscular mycorrhizal hyphae. Appl. Environ. Microbiol. 69, 6208–6215. Aspray, T.J., Eirian Jones, E., Whipps, J.M., and Bending, G.D. (2006). Importance of mycorrhization helper bacteria cell density and metabolite localization for the Pinus sylvestris-Lactarius rufus symbiosis. FEMS Microbiol. Ecol. 56, 25–33. Bakare, N., Rickerts, V., Bargon, J., and Just-Nubling, G. (2003). Prevalence of Aspergillus fumigatus and other fungal species in the sputum of adult patients with cystic fibrosis. Mycoses 46, 19–23. Bauernfeind, A., Bertele, R.M., Harms, K., Horl, G., Jungwirth, R., Petermuller, C., Przyklenk, B., and Weisslein-Pfister, C. (1987). Qualitative and quantitative microbiological analysis of sputa of 102 patients with cystic fibrosis. Infection 15, 270–277. Bhargava, V., Tomashefski, J.F., Jr., Stern, R.C., and Abramowsky, C.R. (1989). he pathology of fungal infection and colonization in patients with cystic fibrosis. Hum. Pathol. 20, 977–986. Bolwerk, A., Lagopodi, A.L., Wij es, A.H.M., Lamers, G.E.M., Chin-A-Woeng, T.F.C., Lugtenberg, B.J.J., and Bloemberg, G.V. (2003). Interactions in the tomato rhizosphere of two Pseudomonas biocontrol strains with the phytopathogenic fungus Fusarium oxysporum f. sp. radicis-lycopersici. Mol. Plant Microbe Interact. 16, 983–993. Braun, B.R., and Johnson, A.D. (1997). Control of filament formation in Candida albicans by the transcriptional repressor TUP1. Science 277, 105–109. Burns, J.L., Van Dalfsen, J.M., Shawar, R.M., Otto, K.L., Garber, R.L., Quan, J.M., Montgomery, A.B., Albers, G.M., Ramsey, B.W., and Smith, A.L. (1999). Effect of chronic intermittent administration of inhaled tobramycin on respiratory microbial flora in patients with cystic fibrosis. J. Infect. Dis. 179, 1190–1196. Chaffin, W.L., Lopez-Ribot, J.L., Casanova, M., Gozalbo, D., and Martinez, J.P. (1998). Cell wall and secreted proteins of Candida albicans: identification, function, and expression. Microbiol. Mol. Biol. Rev. 62, 130–180. Chandra, J., Kuhn, D.M., Mukherjee, P.K., Hoyer, L.L., McCormick, T., and Ghannoum, M.A. (2001). Biofilm formation by the fungal pathogen Candida albicans: development, architecture, and drug resistance. J. Bacteriol. 183, 5385–5394. Charlton, T.S., de Nys, R., Netting, A., Kumar, M., Givskov, M., and Kjellerberg, S. (2000). A novel and sensitive method for the quantification of N-3-oxoacyl homoserine lactones using gas chromatography-mass spectrometry: application to a model bacterial biofilm. Environ. Microbiol. 2, 530–541. Cheng, S.H., Gregory, R.J., Marshall, J., Paul, S., Souza, D.W., White, G.A., O’Riordan, C.R., and Smith, A.E. (1990). Defective intracellular transport and processing of CFTR is the molecular basis of most cystic fibrosis. Cell 63, 827–834. Chet, I., Fogel, S., and Mitchell, R. (1971). Chemical detection of microbial prey by bacterial predators. J. Bacteriol. 106, 863–867. Costerton, J.W., Stewart, P.S., and Greenberg, E.P. (1999). Bacterial biofilms: a common cause of persistent infections. Science 284, 1318–1322. de Weert, S., Kuiper, I., Lagendijk, E.L., Lamers, G.E.M., and Lugtenberg, B.J.J. (2004). Role of chemotaxis towards fusaric acid in colonization of hyphae of Fusarium oxysporum f. sp. radicis-lycopersici by Pseudomonas fluorescens WCS365. Mol. Plant Microbe Interact. 16, 1185–1191. Dorr, J., Hurek, T., and Reinhold-Hurek, B. (1998). Type IV pili are involved in plant-microbe and fungus-microbe interactions. Mol. Microbiol. 30, 7–17. Frey-Klett, P., Chavatte, M., Clausse, M.L., Courrier, S., Le Roux, C., Raaijmakers, J., Martinotti, M.G., Pierrat, J.C., and Garbaye, J. (2005). Ectomycorrhizal symbiosis affects functional diversity of rhizosphere fluorescent pseudomonads. New Phytol. 165, 317–328. Frey-Klett, P., Pierrat, J.C., and Garbaye, J. (1997). Location and survival of mycorrhiza helper Pseudomonas fluorescens during establishment of ectomycorrhizal symbiosis between Laccaria bicolor and Douglas Fir. Appl. Environ. Microbiol. 63, 139–144. Friedman, L., and Kolter, R. (2004a). Genes involved in matrix formation in Pseudomonas aeruginosa PA14 biofilms. Mol. Microbiol. 51, 675–690. Friedman, L., and Kolter, R. (2004b). Two genetic loci produce distinct carbohydrate-rich structural components of the Pseudomonas aeruginosa biofilm matrix. J. Bacteriol. 186, 4457–4465.

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Gaballa, A., Abeysinghe, P.D., Urich, G., Matthijs, S., De Greve, H., Cornelis, P., and Koedam, N. (1997). Trehalose induces antagonism towards Pythium debaryanum in Pseudomonas fluorescens ATCC 17400. Appl. Environ. Microbiol. 63, 4340–4345. Garbaye, J. (1994). Helper bacteria: a new dimension to the mycorrhizal symbiosis. New Phytol. 128, 197–210. Haase, G., Skopnik, H., Groten, T., Kusenbach, G., and Posselt, H.G. (1991). Long-term fungal cultures from sputum of patients with cystic fibrosis. Mycoses 34, 373–376. Hawser, S.P., Baillie, G.S., and Douglas, L.J. (1998). Production of extracellular matrix by Candida albicans biofilms. J. Med. Microbiol. 47, 253–256. Hendrickson, E.L., Plotnikova, J., Mahajan-Miklos, S., Rahme, L.G., and Ausubel, F.M. (2001). Differential roles of the Pseudomonas aeruginosa PA14 rpoN gene in pathogenicity in plants, nematodes, insects, and mice. J. Bacteriol. 183, 7126–7134. Hogan, D.A., and Kolter, R. (2002). Pseudomonas-Candida interactions: An ecological role for virulence factors. Science 296, 2229–2232. Hogan, D.A., Vik, A., and Kolter, R. (2004). A Pseudomonas aeruginosa quorum-sensing molecule influences Candida albicans morphology. Mol. Microbiol. 54, 1212–1223. Hoiby, N., Krogh Johansen, H., Moser, C., Song, Z., Ciofu, O., and Kharazmi, A. (2001). Pseudomonas aeruginosa and the in vitro and in vivo biofilm mode of growth. Microbes Infect. 3, 23–35. Hollsing, A.E., Granstrom, M., Vasil, M.L., Wretlind, B., and Strandvik, B. (1987). Prospective study of serum antibodies to Pseudomonas aeruginosa exoproteins in cystic fibrosis. J. Clin. Microbiol. 25, 1868–1874. Hughes, W.T., and Kim, H.K. (1973). Mycoflora in cystic fibrosis: some ecologic aspects of Pseudomonas aeruginosa and Candida albicans. Mycopathol. Mycol. Appl. 50, 261–269. Jackson, K.D., Starkey, M., Kremer, S., Parsek, M.R., and Wozniak, D.J. (2004). Identification of psl, a locus encoding a potential exopolysaccharide that is essential for Pseudomonas aeruginosa PAO1 biofilm formation. J. Bacteriol. 186, 4466–4475. Lanotte, P., Mereghetti, L., Lejeune, B., Massicot, P., and Quentin, R. (2003). Pseudomonas aeruginosa and cystic fibrosis: correlation between exoenzyme production and patient’s clinical state. Pediatr. Pulmonol. 36, 405–412. Lee, S.W., and Cooksey, D.A. (2000). Genes expressed in Pseudomonas putida during colonization of a plant-pathogenic fungus. Appl. Environ. Microbiol. 66, 2764–2772. Levy, A., Chang, B.J., Abbott, L.K., Kuo, J., Harnett, G., and Inglis, T.J. (2003). Invasion of spores of the arbuscular mycorrhizal fungus Gigaspora decipiens by Burkholderia spp. Appl. Environ. Microbiol. 69, 6250–6256. Lindow, S.E., and Brandl, M.T. (2003). Microbiology of the phyllosphere. Appl. Environ. Microbiol. 69, 1875–1883. Lugtenberg, B.J., Dekkers, L., and Bloemberg, G.V. (2001). Molecular determinants of rhizosphere colonization by Pseudomonas. Annu. Rev. Phytopathol. 39, 461–490. Malajczuk, N., Nesbitt, H.J., and Glenn, A.R. (1977). A light and electron microscope study of the interaction of soil bacteria with Phytophthora cinnamomi Rands. Can. J. Microbiol. 23, 1518–1525. Marshall, K.C., Stout, R., and Mitchell, R. (1971). Selective sorption of bacteria from seawater. Can. J. Microbiol. 17, 1413–1416. Martinez-Granero, F., Rivilla, R., and Martin, M. (2006). Rhizosphere selection of highly motile phenotypic variants of Pseudomonas fluorescens with enhanced competitive colonization ability. Appl. Environ. Microbiol. 72, 3429–3434. Matsukawa, M., and Greenberg, E.P. (2004). Putative exopolysaccharide synthesis genes influence Pseudomonas aeruginosa biofilm development. J. Bacteriol. 186, 4449–4456. Navarro, J., Rainisio, M., Harms, H.K., Hodson, M.E., Koch, C., Mastella, G., Strandvik, B., and McKenzie, S.G. (2001). Factors associated with poor pulmonary function: cross-sectional analysis of data from the ERCF. European Epidemiologic Registry of Cystic Fibrosis. Eur. Respir. J. 18, 298–305. Nurmiaho-Lassila, E.L., Timonen, S., Haahtela, K., and Sen, R. (1997). Bacterial colonization patterns of intact Pinus sylvestris mycorrhizospheres in dry pine forest soil: an electron microscopy study. Can. J. Microbiol. 43, 1017–1035. O’Toole, G.A., and Kolter, R. (1998). Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol. Microbiol. 30, 295–304.

Bacterial Bioilms on Fungal Surfaces

Rajan, S., and Saiman, L. (2002). Pulmonary infections in patients with cystic fibrosis. Semin. Respir. Infect. 17, 47–56. Rangel-Castro, J.I., Levenfors, J.J., and Danell, E. (2002). Physiological and genetic characterization of fluorescent Pseudomonas associated with Cantharellus cibarius. Can. J. Microbiol. 48, 739–748. Rincon, A., Ruiz-Diez, B., Garcia-Fraile, S., Garcia, J.A., Fernandez-Pascual, M., Pueyo, J.J., and de Felipe, M.R. (2005). Colonisation of Pinus halepensis roots by Pseudomonas fluorescens and interaction with the ectomycorrhizal fungus Suillus granulatus. FEMS Microbiol. Ecol. 51, 303–311. Singh, P.K., Schaefer, A.L., Parsek, M.R., Moninger, T.O., Welsh, M.J., and Greenberg, E.P. (2000). Quorum-sensing signals indicate that cystic fibrosis lungs are infected with bacterial biofilms. Nature 407, 762–764. hevelein, J.M. (1984). Regulation of trehalose mobilization in fungi. Microbiol. Rev. 48, 42–59. Timonen, S., and Hurek, T. (2006). Characterization of culturable bacterial populations associating with Pinus sylvestris: Suillus bovinus mycorrhizospheres. Can. J. Microbiol. 52, 769–778. Toljander, J.F., Artursson, V., Paul, L.R., Jansson, J.K., and Finlay, R.D. (2006). Attachment of different soil bacteria to arbuscular mycorrhizal fungal extraradical hyphae is determined by hyphal vitality and fungal species. FEMS Microbiol. Lett. 254, 34–40. Whipps, J.M. (2001). Microbial interactions and biocontrol in the rhizosphere. J. Exp. Bot. 52, 487–511. Woods, D.E., Lam, J.S., Paranchych, W., Speert, D.P., Campbell, M., and Godfrey, A.J. (1997). Correlation of Pseudomonas aeruginosa virulence factors from clinical and environmental isolates with pathogenicity in the neutropenic mouse. Can. J. Microbiol. 43, 541–551.

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