Search tips
Search criteria 


Logo of jdrHomeAboutSubmit a Manuscript
J Dent Res. 2010 March; 89(3): 205–218.
PMCID: PMC3318030

Biofilm Dispersal

Mechanisms, Clinical Implications, and Potential Therapeutic Uses


Like all sessile organisms, surface-attached communities of bacteria known as biofilms must release and disperse cells into the environment to colonize new sites. For many pathogenic bacteria, biofilm dispersal plays an important role in the transmission of bacteria from environmental reservoirs to human hosts, in horizontal and vertical cross-host transmission, and in the exacerbation and spread of infection within a host. The molecular mechanisms of bacterial biofilm dispersal are only beginning to be elucidated. Biofilm dispersal is a promising area of research that may lead to the development of novel agents that inhibit biofilm formation or promote biofilm cell detachment. Such agents may be useful for the prevention and treatment of biofilms in a variety of industrial and clinical settings. This review describes the current status of research on biofilm dispersal, with an emphasis on studies aimed to characterize dispersal mechanisms, and to identify environmental cues and inter- and intracellular signals that regulate the dispersal process. The clinical implications of biofilm dispersal and the potential therapeutic applications of some of the most recent findings will also be discussed.

Keywords: biofilm, detachment, dispersal, dispersion, erosion, matrix, plaque, seeding, sloughing, transmission


Surface-associated communities of bacteria known as biofilms play a role in the pathogenesis of many chronic infections. In the oral cavity, biofilms that form on teeth produce acids that cause dental caries, and biofilms that grow in the gingival sulcus contribute to the pathogenesis of periodontitis (Marsh, 2006). Biofilms that form in other organs of the body cause numerous, often life-threatening, infections such as cystic fibrosis pneumonia and catheter-related endocarditis (Costerton et al., 1999). All biofilms, regardless of their location, share several common features. These include the synthesis of an extracellular polymeric matrix that holds the bacterial cells together, and an increase in resistance to killing by host defenses and antimicrobial agents compared with the resistance exhibited by free-living or ‘planktonic’ cells (Mah and O’Toole, 2001). The inherent protective nature of the biofilm colony makes most biofilm-associated infections difficult or impossible to eradicate.

Biofilm development can be divided into three distinct stages: attachment of cells to a surface, growth of the cells into a sessile biofilm colony, and detachment of cells from the colony into the surrounding medium. The initial, reversible interaction between a bacterial cell and a surface is mediated by non-specific Lifshitz-van der Waals, Lewis acid-base, and electrostatic forces. This transient attachment is reinforced by host- and tissue-specific adhesins that are located on the bacterial cell surface or on cellular appendages such as pili and fimbriae (Rosan and Lamont, 2000). This results in the irreversible attachment of the bacterial cell to the surface. In the case of dental plaque, which can be comprised of hundreds of bacterial species, colonization of tooth surfaces follows an ordered progression, with initial adhesion of early ‘pioneer’ species to the enamel surface followed by attachment of later-colonizing species to the already-attached early colonizers (Marsh, 2004).

The second stage of biofilm development involves the multiplication of bacteria on the surface and the concomitant synthesis of an extracellular polymeric matrix. The matrix holds the bacterial cells together in a mass and firmly attaches the bacterial mass to the underlying surface. Some examples of polymeric biofilm matrix components produced by oral bacteria include the well-studied glucan polysaccharides of Streptococcus mutans (Banas and Vickerman, 2003), proteinaceous fimbriae produced by Aggregatibacter actinomycetemcomitans and Porphyromonas gingivalis (Kachlany et al., 2001; Lamont et al., 2002), and extracellular, double-stranded DNA in biofilms produced by A. actinomycetemcomitans, S. mutans, and S. intermedius (Inoue et al., 2003; Petersen et al., 2004, 2005). In addition to providing a structural ‘scaffold’ for the biofilm colony, the matrix also contributes to biofilm-mediated antimicrobial resistance, either by acting as a diffusion barrier, or by binding directly to antimicrobial agents and preventing their access to the biofilm cells (Mah and O’Toole, 2001).

Continued growth of bacterial cells on a surface leads to the development of mature biofilm colonies containing millions of tightly packed cells gathered into pillar- and mushroom-shaped masses that project outward into the surrounding medium for hundreds of microns (Hall-Stoodley et al., 2004). These structures are interspersed with fluid-filled channels which act as a primitive circulatory system, allowing for the exchange of nutrients and waste products with the bulk fluid phase. In addition, masses of biofilm cells often contain demarcated internal spaces that are devoid of cells. Thus, mature biofilm colonies are complex, highly differentiated structures. Numerous microenvironments that differ with respect to pH, oxygen concentration, nutrient availability, and cell density exist within the biofilm colony. This results in a great deal of heterogeneity in metabolic and reproductive activity among cells located in different parts of the colony. Metabolically inactive cells located in the interior of the colony may be resistant to the actions of antimicrobial agents that target actively growing cells (Fux et al., 2004).

The final stage of biofilm development is the detachment of cells from the biofilm colony and their dispersal into the environment. This is an essential stage of the biofilm life cycle that contributes to biological dispersal, bacterial survival, and disease transmission. Like other stages of biofilm development, dispersal can be a complex process that involves numerous environmental signals, signal transduction pathways, and effectors (Karatan and Watnick, 2009). No single mechanism of biofilm dispersal is utilized by all bacteria.

This article reviews the current literature on biofilm dispersal. Although dispersal is the least-understood stage of the biofilm life cycle, increasing numbers of studies on this process are being published. An important rationale for these studies is that understanding the mechanisms of biofilm dispersal is expected to lead to the development of clinically useful agents that inhibit biofilm formation or promote biofilm detachment. This review is divided into two main sections. The first section describes the known mechanisms of biofilm dispersal, and the second section describes the known chemical signals that regulate the dispersal process. The clinical implications of biofilm dispersal will also be discussed, as will the potential therapeutic applications of some of the most recent findings.

Mechanisms Of Biofilm Dispersal

Bacterial biofilm dispersal can be divided into three distinct phases: (i) detachment of cells from the biofilm colony; (ii) translocation of the cells to a new location; and (iii) attachment of the cells to a substrate in the new location. Thus, S. mutans cells that detach from dental plaque can be transported to the saliva of an infant by direct contact or by means of a vector such as a shared spoon, and then attach to the tooth surface and initiate colonization of the new host. Similarly, cells that detach from a Legionella biofilm growing in a cooling tower can be transported by means of air-borne water droplets to the lungs of a susceptible host, where they can attach to alveolar macrophages and initiate infection. In the literature and in this review, the terms ‘detachment’, ‘dispersal’, and ‘dispersion’ are used interchangeably to refer to the cell-detachment phase of the dispersal process. Studies on the movement of detached cells to a new location fall mostly under the discipline of disease transmission.

In general, mechanisms of biofilm dispersal can be divided into two broad categories: active and passive. Active dispersal refers to mechanisms that are initiated by the bacteria themselves, whereas passive dispersal refers to biofilm cell detachment that is mediated by external forces such as fluid shear, abrasion (collision of solid particles with the biofilm), predator grazing, and human intervention (Lawrence et al., 2002; Choi and Morgenroth, 2003; Ymele-Leki and Ross, 2007). In a complex community such as dental plaque, close relationships between species based on competition, mutualism, predation, or parasitism are likely to have resulted in the evolution of various other passive dispersal mechanisms. These may include interspecific antimicrobial compounds, quorum-sensing signals, or matrix-degrading enzymes. Phagocytosis, a form of predator grazing, may also contribute to the passive dispersal of oral biofilms (Erard et al., 1989).

At least three distinct modes of biofilm dispersal have been identified: erosion, sloughing, and seeding. Erosion refers to the continuous release of single cells or small clusters of cells from a biofilm at low levels over the course of biofilm formation. Sloughing refers to the sudden detachment of large portions of the biofilm, usually during the later stages of biofilm formation (Marshall, 1988; Lappin-Scott and Bass, 2001; Stoodley et al., 2001; Wilson et al., 2004). Seeding dispersal, also known as central hollowing, refers to the rapid release of a large number of single cells or small clusters of cells from hollow cavities that form inside the biofilm colony (Boles et al., 2005; Ma et al., 2009). Erosion and sloughing can be either active or passive processes, whereas seeding dispersal is always an active process. The following sections describe some of the mechanisms of active biofilm dispersal that have been described to date.

Enzymatic Degradation of the Biofilm Matrix

A basic mechanism of biofilm dispersal that is utilized by phylogenetically diverse bacteria is the production of extracellular enzymes that degrade adhesive components in the biofilm matrix. Since the biofilm matrix encases the bacterial cells within the biofilm colony, degradation of the matrix results in the detachment of cells from the colony and their release into the environment. Matrix-degrading enzymes implicated in active biofilm dispersal include glycosidases, proteases, and deoxyribonucleases (Table).

Bacterial Enzymes Implicated in Active Biofilm Dispersal

One well-studied biofilm-matrix-degrading enzyme is dispersin B, a glycoside hydrolase produced by the periodontopathogen A. actinomycetemcomitans (Kaplan et al., 2003b). Dispersin B degrades poly-N-acetylglucosamine (PNAG), a biofilm matrix polysaccharide that mediates attachment of A. actinomycetemcomitans cells to abiotic surfaces, intercellular adhesion (autoaggregation), and resistance to killing by detergents and human phagocytic cells (Kaplan et al., 2003b, 2004b; Izano et al., 2007, 2008b; Venketaraman et al., 2008). Evidence that dispersin B is involved in biofilm dispersal comes from studies utilizing mutant strains that do not produce the enzyme (Kaplan et al., 2003b). When cultured in broth, these mutant strains produce biofilm colonies that are similar in morphology to wild-type colonies, but the mutant colonies fail to release cells into the medium and disperse (Fig. 1). Further evidence that dispersin B is involved in biofilm dispersal comes from studies showing that purified dispersin B enzyme detaches pre-formed biofilm colonies produced by A. actinomycetemcomitans and other PNAG-producing bacteria (Itoh et al., 2005; Izano et al., 2007). In addition, over-expression of dispersin B in wild-type A. actinomycetemcomitans biofilms results in a hyper-dispersal phenotype (unpublished data). An orthologous dispersin B enzyme is produced by the porcine respiratory pathogen Actinobacillus pleuropneumoniae (Kaplan et al., 2004b), although its role in biofilm dispersal has not been investigated. Genes homologous to the A. actinomycetemcomitans dispersin B structural gene (dspB) are present in the genomes of several other bacteria, including the human oral commensal Aggregatibacter aphrophilus, and the bovine ruminal species Actinobacillus succinogenes and Mannheimia succiniciproducens. There is no evidence that these other species produce a functional dispersin B enzyme. A homologue of dspB was identified in the genome of Staphylococcus lugdunensis, although its role in biofilm dispersal was not investigated (Frank and Patel, 2007).

Figure 1.
Biofilm dispersal phenotypes of wild-type and dispersin B mutant strains of Aggregatibacter actinomycetemcomitans. (A) Dispersal of A. actinomycetemcomitans strain CU1000 (wild-type) and JK1023 (dispersin B mutant) in broth. (B) Biofilm formation by strains ...

The cariogenic bacterium S. mutans also produces an enzyme that mediates the release of cells from biofilms (Lee et al., 1996). This enzyme, referred to as surface-protein-releasing enzyme, or SPRE, degrades salivary receptor P1 (also known as antigen I/II or PAc), a 185-kDa surface protein that mediates attachment of S. mutans cells to the tooth surface (Vats and Lee, 2000). Degradation of P1 by exogenously added SPRE results in the detachment of a S. mutans monolayer formed on saliva-coated hydroxyapatite rods (Lee et al., 1996). In addition, a Tn917 SPRE-defective mutant strain was shown to detach from the rods at a significantly lower rate than the parental strain (Vats and Lee, 2000). Interestingly, active detachment of S. mutans biofilms occurs more rapidly as the pH of the medium drops (Tam et al., 2007). Since the production of SPRE is optimal at pH 5-6, this suggests that increased acidogenicity may trigger S. mutans biofilm detachment by the induction of SPRE activity. SPRE is also produced by S. gordonii, a pioneer colonizer and important endocarditis pathogen, and by several other non-oral pathogenic streptococci, including S. pneumoniae, S. pyogenes, and S. agalactiae (Vats and Lee, 2000).

Several other non-oral bacteria produce extracellular enzymes that degrade endogenous matrix components and mediate biofilm cell detachment. Mucoid strains of the human opportunistic pathogen Pseudomonas aeruginosa, for example, produce both alginate, a biofilm matrix polysaccharide composed of mannuronic and guluronic acids, and alginate lyase, an enzyme that degrades alginate. Increased expression of alginate lyase promotes the detachment of cells from P. aeruginosa biofilms cultured on agar surfaces (Boyd and Chakrabarty, 1994), and exogenously added alginate lyase increases the effectiveness of some antibiotics against P. aeruginosa biofilms cultured in broth (Alkawash et al., 2006; Alipour et al., 2009). Polysaccharide lyases that promote biofilm detachment are also produced by P. fluorescens and P. syringae (Allison et al., 1998; Preston et al., 2000). Other polysaccharide-degrading enzymes implicated in biofilm cell detachment include endo-β-1,4-mannanase, produced by the plant pathogen Xanthomonas campestris (Dow et al., 2003), and disaggregatase, produced by the archaeon Methanosarcina mazei (Xun et al., 1990).

Several extracellular proteases have also been implicated in biofilm cell detachment. In the plant saprophyte Pseudomonas putida, it has been shown that LapG protease cleaves a periplasmic protein (LapA) which anchors an unidentified biofilm matrix polysaccharide to the cell (Gjermansen et al., 2009). This process results in the release of cells from biofilms cultured in microplate wells or flow cells. In Staphylococcus aureus, deletion of the genes encoding the extracellular proteases aureolysin and Spl resulted in a significant increase in biofilm formation in the flow cells, and a concomitant decrease in planktonic cells present in the fluid effluent (Boles and Horswill, 2008). These findings indicate that S. aureus biofilm dispersal requires protease activity, although the targets for these proteases are not known.

In addition to the glycosidases and proteases mentioned above, the deoxyribonuclease known as thermonuclease or micrococcal nuclease has been implicated in cell detachment in S. aureus biofilms (Mann et al., 2009). S. aureus biofilms are readily detached from microplate wells by exogenously added deoxyribonucleases, including thermonuclease, indicating that extracellular DNA is a major biofilm matrix adhesin in this species (Izano et al., 2008a). It has been shown (Mann et al., 2009) that a thermonuclease-deficient mutant strain of S. aureus exhibited significantly increased biofilm formation in flow cells compared with the amount of biofilm exhibited by a wild-type strain. These findings suggest that thermonuclease may function as an endogenous mediator of biofilm dispersal in this species.

All of the matrix-degrading enzymes described above mediate biofilm dispersal by enabling bacteria to degrade their own biofilm matrix polymers. An alternative mechanism of enzyme-mediated biofilm dispersal may occur in multi-species biofilms such as dental plaque, where some bacteria may produce enzymes that degrade biofilm matrix polymers produced by other species. Interspecific matrix-degrading enzymes may have evolved to provide bacteria with a source of nutrients, or as a defense mechanism that detaches and displaces competing species. Zhang and Bishop (2003) provided evidence that this mechanism of biofilm dispersal may occur in complex biofilm communities found in wastewater treatment plants. Using a small-scale biofilm reactor that was seeded with wastewater and perfused with synthetic wastewater, they showed that bacteria present in activated sludge can degrade the extracellular matrix of biofilms produced by the wastewater bacteria, and then utilize the degradation products as an additional food source. Such action would likely result in the passive detachment of the wastewater biofilms.

Enzymatic Degradation of the Biofilm Substrate

Another mechanism of enzyme-mediated biofilm dispersal involves the production of extracellular enzymes that degrade the substrate on which the biofilm colony is growing. An interesting example of this type of dispersal may be found in the oral bacterium Streptococcus intermedius, which produces hyaluronidase, an enzyme that degrades the glycosaminoglycan hyaluronan (HA) found in the extracellular matrix of connective tissue. By breaking down the connective tissue, hyaluronidase may provide nutrients for the bacteria or allow the spread of bacteria and toxins deeper into the tissue. It was recently shown (Pecharki et al., 2008) that hyaluronidase may play a role in S. intermedius biofilm dispersal. These authors found that a hyaluronidase mutant strain formed significantly more biofilm in broth supplemented with HA than did a wild-type strain. Also, exogenous hyaluronidase dispersed S. intermedius biofilms grown in HA-supplemented medium. This mechanism may have clinical relevance to the dispersal of S. intermedius biofilms in the oral cavity.

In the intestinal pathogen Vibrio cholerae, a Zn-metalloprotease known as hemagglutinin protease (HAP) may promote detachment of vibrios from human intestinal epithelial cells by digesting epithelial cell receptors that bind to V. cholerae adhesins (Finkelstein et al., 1992). Evidence supporting this model comes from experiments showing that mutant vibrio strains lacking HAP remained attached to cultured intestinal cells, whereas wild-type strains readily detached (Finkelstein et al., 1992). In addition, pre-treatment of the cultured epithelial cells with purified HAP inhibited attachment of the vibrios in a time- and dose-dependent manner. Another example of this dispersal mechanism may be found in the marine bacteria Pseudoaltermonas sp. strain S91 and Vibrio furnissii (Yu et al., 1991; Baty et al., 2000). These bacteria form biofilms on solid chitin surfaces in marine environments, and also produce chitinase, which degrades the chitin for use as a food source. Chitin degradation results in detachment of the attached biofilm cells from artificially prepared chitin surfaces and from natural squid chitin surfaces in vitro (Baty et al., 2000).

Seeding Dispersal

Microscopic studies have shown that many bacteria produce biofilm colonies that contain hollow, internal cavities (Fig. 2). These cavities have been observed in biofilms produced by the human pathogens A. actinomycetemcomitans, P. aeruginosa, Serratia marcescens, and S. aureus (Kaplan et al., 2003a; Yarwood et al., 2004; Koh et al., 2007; Ma et al., 2009); by the plant-associated bacteria P. putida and Chromobacterium violacium (Tolker-Nielsen et al., 2000; Mai-Prochnow et al., 2008); and by the aquatic bacteria Caulobacter crescentus, Pseudoalteromonas tunicate, and Marinomonas mediterranea (Mai-Prochnow et al., 2004, 2008). Increasing evidence suggests that these hollow cavities play a role in biofilm cell detachment through a process termed ‘seeding dispersal’. In this process, the hollow cavities become filled with non-aggregated planktonic cells. Seeding dispersal results when a breach in the colony wall releases the planktonic cells from the cavities into the surrounding medium.

Figure 2.
Cross-sections of biofilm colonies produced by (A) A. actinomycetemcomitans, (B) P. aeruginosa, and (C) S. marcescens. Scale bars = 100 µm in (A), 50 µm in (B), and 200 µm in (C). Panels (B) and (C) are from Ma et al. (2009) and ...

Seeding dispersal has been well-characterized in the oral bacterium A. actinomycetemcomitans (Kaplan and Fine, 2002; Kaplan et al., 2003a,b). Two lines of evidence support the notion that A. actinomycetemcomitans biofilm colonies undergo seeding dispersal. First, microscopic analyses show that mature A. actinomycetemcomitans biofilm colonies cultured in broth develop internal, hollow cavities that are surrounded by a layer of non-aggregated cells (Fig. 2A) (Kaplan et al., 2003a). Second, when A. actinomycetemcomitans biofilms are cultured in broth, the release of cells from the biofilm into the broth over time is sudden rather than gradual, consistent with a seeding dispersal event (Fig. 1C). A. actinomycetemcomitans biofilm dispersal can be visualized by the culturing of biofilm colonies in the presence of buoyancy-driven convection currents under conditions of low vibration (Fig. 3) (Kaplan and Fine, 2002). Under these conditions, dispersed cells move along the surface by convection current, reattach to the surface, and then form new biofilm colonies. This results in the appearance of streamers of satellite colonies emanating from the dispersed biofilm colony. Biofilm colonies produced by mutant strains of A. actinomycetemcomitans deficient in the production of dispersin B still contain internal voids, but the void spaces are not surrounded by a layer of non-aggregated cells, and the colonies do not release cells into the medium and disperse (Figs. 1A, ,1C)1C) (Kaplan et al., 2003b). These findings indicate that depolymerization of PNAG by dispersin B is required for the production of the non-aggregated cell layer, but not for the formation of the hollow cavities themselves. Other oral bacteria that exhibit a similar mode of biofilm dispersal include Neisseria subflava and S. mutans (Fig. 3), and Aggregatibacter aphrophilus and S. mitis (Kaplan and Fine, 2002). In the case of N. subflava, a sudden spike in the number of CFU in the medium during biofilm formation was also observed (Kaplan and Fine, 2002).

Figure 3.
Dispersal of A. actinomycetemcomitans, N. subflava, and S. mutans biofilms in broth. Biofilms were stained with crystal violet. Scale bar = 1 mm. The panel on the left is from Kaplan and Fine (2002). Used with permission.

Seeding dispersal has been studied extensively in P. aeruginosa biofilms (Sauer et al., 2002; Hunt et al., 2004; Schooling et al., 2004; Boles et al., 2005; Purevdorj-Gage et al., 2005; Kirov et al., 2007; Pamp and Tolker-Nielsen, 2007; Ma et al., 2009). These studies have shown that the hollow cavities that form inside P. aeruginosa biofilm colonies are devoid of biofilm matrix polysaccharide, but contain numerous swimming bacterial cells (Ma et al., 2009). In some cases, motile cells in the hollow cavities can be seen swimming through openings in the colony wall and entering the bulk liquid (Sauer et al., 2002). Central hollowing and seeding dispersal in P. aeruginosa is evidently triggered by an increase in colony size, because a threshold colony diameter of > 80 µm is required for hollow cavity formation to occur (Purevdorj-Gage et al., 2005).

The mechanism of central hollowing is not fully understood, but evidence suggests that it involves the death and lysis of a subpopulation of cells located in the center of the colony. In P. aeruginosa, for example, strains that are deficient in the production of the Cid/Lrg toxin-antitoxin system produce biofilm colonies that do not undergo central hollowing (Webb et al., 2003; Ma et al., 2009). The Cid/Lrg proteins are structurally and functionally related to bacteriophage-encoded holins, which regulate host cell lysis during the lytic cycle of infection by modulating the expression of murein hydrolase (Bayles, 2007). An orthologous Cid/Lrg system was shown to be responsible for cell death and DNA release in S. aureus biofilms, although the role of this system in biofilm dispersal has not been investigated (Rice et al., 2007; Mann et al., 2009). Genes that encode homologues of the Cid/Lrg system are present in many bacterial species, including the oral bacteria A. actinomycetemcomitans, Fusobacterium nucleatum, and S. mutans. Thus, the Cid/Lrg system may mediate cell death and biofilm dispersal in a variety of species (Bayles, 2007).

Several studies have shown that phage-mediated cell lysis can also lead to central hollowing and seeding dispersal in P. aeruginosa biofilms. A bacteriophage capable of superinfecting and lysing P. aeruginosa (bacteriophage Pf4) was detected in the fluid effluent from P. aeruginosa biofilms cultured in flow cells (Webb et al., 2003), and the amount of bacteriophage activity in the effluent paralleled the degree of cell death seen in the biofilm (Kirov et al., 2007). In addition, deletion of the entire Pf4 prophage genome from the P. aeruginosa chromosome resulted in the formation of biofilms that did not exhibit cell lysis, central hollowing, or seeding dispersal (Rice et al., 2009).

For motile bacteria, it has been shown that motility is: (i) required for the initial attachment of bacteria to surfaces (O’Toole and Kolter, 1998; Chiang and Burrows, 2003); (ii) repressed in mature biofilms (Sauer and Camper, 2001; Whiteley et al., 2001; Sauer et al., 2002, 2004); and (iii) induced upon biofilm dispersal (Jackson et al., 2002; Sauer et al., 2002; Purevdorj-Gage et al., 2005). These observations suggest that the induction of motility may itself represent a specific mechanism of biofilm dispersal, or may play a role in the initiation of seeding dispersal. However, studies have shown that seeding dispersal is not influenced by motility in P. aeruginosa and P. putida biofilms (Gjermansen et al., 2005; Morgan et al., 2006). In addition, non-motile species such as A. actinomycetemcomitans undergo seeding dispersal. Thus, other gene products that reduce the cohesiveness of the biofilm in the center of the colony evidently mediate seeding dispersal.

Seeding dispersal also occurs in biofilms produced by the marine bacterium Pseudoalteromonas tunicata (Mai-Prochnow et al., 2004). Dispersal involves the formation of central voids within the biofilm colony, extensive cell killing, and detachment of the biofilm from the substratum. Mutant strains deficient in the production of the autolytic protein AlpP did not exhibit cell death and biofilm dispersal. It was subsequently shown that AlpP acts as a lysine oxidase that generates hydrogen peroxide, which is directly responsible for cell death within the biofilm colony (Mai-Prochnow et al., 2008). Orthologous AlpP proteins mediate cell death and biofilm dispersal in C. crescentus, C. violaceum, and M. mediterranea (Mai-Prochnow et al., 2008).

Production of Rhamnolipids

Rhamnolipids are extracellular surfactants produced by P. aeruginosa (Soberón-Chávez et al., 2005). These compounds typically contain a dimer of 3-hydroxyfatty acids linked through a β-glycosidic bond to a mono- or di-rhamnose moiety. P. aeruginosa produces several rhamnolipids, including L-rhamnosyl-3-hydroxydecanoyl-3-hydroxydecanoate (mono-rhamnolipid) (Fig. 4A), and L-rhamnosyl-L-rhamnosyl-3-hydroxydecanoyl-3-hydroxydecanoate (di-rhamnolipid). Due to their amphipathic nature, rhamnolipids exhibit surface-acting properties that decrease the adhesiveness of cell-cell, cell-matrix, and cell-surface interactions (Neu, 1996). It has been shown (Boles et al., 2005) that inactivation of the rhamnolipid biosynthetic genes (rhaAB) in P. aeruginosa biofilms inhibits central hollowing and cell detachment, and that over-expression of rhaAB in wild-type biofilms increases cell detachment. Also, exogenous rhamnolipids induce central hollowing and biofilm detachment in wild-type P. aeruginosa biofilms (Boles et al., 2005; Dong et al., 2008). These findings suggest that rhamnolipids can mediate central hollowing in P. aeruginosa biofilms during seeding dispersal.

Figure 4.
Chemical signals implicated in biofilm dispersal.

The mechanism by which rhamnoloipids induce central hollowing is not known. These compounds most likely act by disrupting interactions among various cellular and matrix component within the biofilm colony. This model is supported by the fact that exogenously added rhamnolipids can inhibit P. aeruginosa biofilm formation on glass surfaces when added to the broth at the time of inoculation (Schooling et al., 2004). In addition, exogenously added P. aeruginosa rhamnolipids can disperse biofilms produced by other species of bacteria, including Bordetella bronchiseptica and Salmonella enterica serovar Typhimurium (Mireles et al., 2001; Irie et al., 2005). Interestingly, another surfactant, sodium dodecyl sulfate, can also induce central hollowing in P. aeruginosa biofilm colonies when exogenously added to the colonies (Boles et al., 2005). All of these observations support the notion that rhamnolipids disrupt biofilm cohesiveness in a non-specific manner. However, there is still no explanation for the fact that cells in the center of the biofilm colony are more susceptible to the actions of surfactants. Unknown phenotypic differences in the cells or matrix at the interior of the colony must account for the central hollowing induced by exogenous surfactants (Boles et al., 2005).

Modulation of Fimbrial Adherence

Two different pathogenic strains of E. coli, enteroaggregative E. coli (EAEC) and enteropathogenic E. coli (EPEC), have evolved unique mechanisms to achieve detachment from biofilms that form on intestinal epithelial cells. In EAEC, autoaggregation and adherence to the human intestinal mucosa are mediated by aggregative adherence fimbriae (AAFs). The adhesive properties of AAFs are modulated by a small protein named dispersin, which binds non-covalently to lipopolysaccharide (LPS) on the surface of the bacterium (Sheikh et al., 2002; Velarde et al., 2007). LPS is negatively charged, and AAFs, unlike most enteric pili, are positively charged. When dispersin is bound to LPS, it neutralizes the negative charge and allows the AAFs to extend away from the bacterial cell surface, where they can mediate their adhesive effects. In the absence of dispersin, the AAFs collapse onto the bacterial cell surface and become non-adhesive, due to their electrostatic interaction with the LPS. Thus, biofilm dispersal is achieved by down-regulation of dispersin protein.

A similar mechanism of dispersal is displayed by EPEC, which produces type IV bundle-forming pili (BFP) that mediate microcolony formation on human intestinal mucosa (Cleary et al., 2004). BFP can undergo structural alterations that result in the formation of either thin or thick BFP bundle structures (Knutton et al., 1999). Thin BFP bundles mediate microcolony formation, whereas thick BFP bundles are associated with a non-aggregative, planktonic phenotype. EPEC can therefore modulate microcolony cohesion and dispersal by modulating the structure of the BFP bundles, possibly thorough a mechanism that involves pilus retraction (Knutton et al., 1999).

Cell-division-mediated Biofilm Dispersal

Another possible mechanism of biofilm dispersal involves cell detachment due to cell division at the outer surface of the biofilm colony (Allison et al., 1990). When biofilm cells on the outer surface of the colony divide, one of the progeny cells may be located at a sufficient distance from the colony so that it is not subjected to the attractive forces of the biofilm matrix, and would therefore be liberated into the bulk fluid following cell separation (Gilbert et al., 1993). This dispersal mechanism has been exploited to obtain synchronous populations of bacteria from cells attached to a membrane filter and perfused with fresh medium (Helmstetter and Cummings, 1964; Gilbert et al., 1989). This mechanism of dispersal may account for biofilm erosion in some natural biofilms.

Regulation of Biofilm Dispersal

Many studies on the regulation of biofilm dispersal have focused on the identification of environmental conditions that trigger the dispersal process. Factors such as nutrient levels, oxygen tension, pH, and temperature have been shown to induce dispersal of biofilms produced by a variety of species (Karatan and Watnick, 2009). The rationale for these studies is the notion that dispersal of biofilm cells is a selective advantage when environmental conditions become unfavorable. However, dispersal is a selective advantage even when conditions are favorable. Biological dispersal has fundamental importance for the expansion, reproduction, and survival of all species. In addition to environmentally induced biofilm dispersal, therefore, it is likely that mechanisms of genetically programmed biofilm dispersal also exist. If this is true, then the life cycle of a biofilm may be analogous to that of a primitive, multi- cellular organism (Stoodley et al., 2002; Klausen et al., 2006). One example of programmed dispersal may be seen in A. actinomycetemcomitans biofilms, which undergo a reproducible, periodic detachment of biofilm biomass, even under conditions where nutrients are not limited (Fig. 1B). A similar reproducible pattern of biomass detachment is observed in biofilms produced by P. putida (Gjermansen et al., 2009) and Serratia marcescens (Rice et al., 2005).

The following sections describe some of the environmental cues and inter- and intracellular signals that have been shown to influence and regulate biofilm dispersal.

Nutrient Cues

Several studies have shown that sudden changes in nutrient availability can induce biofilm dispersal. Most work in this area has been performed with biofilms produced by pseudomonads. In experiments with P. aeruginosa, it was found that biofilms cultured in minimal medium in flow cells underwent dispersal in response to both a sudden decrease and a sudden increase in carbon substrate availability (Hunt et al., 2004; Sauer et al., 2004). Both of these responses may make sense from an ecological point of view. Cells may detach to escape unfavorable conditions when nutrients are scarce, or may choose to invest metabolic energy into reproduction and detachment when nutrients are plentiful. Nutrient starvation has also been shown to induce detachment of biofilms produced by P. fluorescens (Delaquis et al., 1989), P. putida (Gjermansen et al., 2009), and Pseudomonas sp. S9 (Wrangstadh et al., 1989). In P. fluorescens, starvation induced an increase in exopolysaccharide lyase production and biofilm dispersal (Allison et al., 1998), whereas in P. aeruginosa, alginate lyase activity was maximally induced in rapidly growing cells (Ott et al., 2001). Again, both of these observations suggest that increased biofilm dispersal rates may occur under both favorable and unfavorable conditions. Among non-pseudomonads, nutrient starvation was shown to increase biofilm dispersal in aquatic Aeromonas hydrophila (Sawyer and Hermanowicz, 1998), whereas high-nutrient conditions induced biofilm dispersal in environmental Acinetobacter sp. (James et al., 1995). In E. coli, exogenous glucose blocked biofilm dispersal induced by CsrA, a global regulator of central carbon flux, which further supports the hypothesis that nutrient cues can induce biofilm dispersal (Jackson et al., 2002).

Oxygen tension may be another environmental signal that modulates biofilm dispersal. In P. putida, oxygen-limited biofilms exhibited significantly lower shear removal rates and significantly greater sloughing dispersal when compared with biofilms cultured under oxygen-rich conditions (Applegate and Bryers, 1991). It was hypothesized that this difference may be due to a higher amount of extracellular polymer present in oxygen-limited biofilms. In contrast, a sudden downshift in molecular oxygen induced a rapid and efficient dispersal of biofilms produced by Shewanella oneidensis, an anaerobic, metal-reducing bacterium found in deep sea sediments (Thormann et al., 2005). Carbon limitation did not induce S. oneidensisis biofilm dispersal.

Acyl Homoserine Lactones

The expression of biofilm-specific genes is often regulated by quorum-sensing, a regulatory mechanism that involves the synthesis, secretion, and sensing of small chemical signals called autoinducers (Irie and Parsek, 2008). As the cell density and autoinducer concentration increase, a threshold concentration of autoinducer triggers an increase in the transcription of biofilm-specific genes by activating transcription factors that bind to sequences upstream from these genes. Autoinducers have been shown to control several stages of biofilm formation, including surface attachment, matrix synthesis, the formation of fluid channels and pillar-like architecture, and dispersal (Hall-Stoodley et al., 2004; Stanley and Lazazzera, 2004). N-acyl-homoserine lactones (AHLs), produced by Gram-negative bacteria, are one of the best-studied classes of autoinducers (Fuqua and Greenberg, 2002). AHLs implicated in biofilm dispersal include 7,8-cis-N-(tetradecenoyl)homoserine lactone, produced by Rhodobacter sphaeroides (Fig. 4B) (Puskas et al., 1997), N-3-oxo-dodecanoyl homoserine lactone (3-oxo-C12-HSL), produced by P. aeruginosa (Fig. 4C) (Wilson et al., 2004; Purevdorj-Gage et al., 2005), and N-butanoyl-L-homoserine lactone (C4-HSL), produced by Serratia marcescens andP. aeruginosa (Fig. 4D) (Schooling et al., 2004; Rice et al., 2005).

In P. aeruginosa, biofilms formed by mutant strains deficient in the production of C4-HSL (ΔlasI/rhlI) do not undergo central hollowing (Purevdorj-Gage et al., 2005), and the addition of exogenous C4-HSL to wild-type biofilms induces biofilm dispersal (Schooling et al., 2004; Dong et al., 2008). C4-HSL may induce dispersal by up-regulation of the rhamnolipid biosynthetic gene rhaA (Davey et al., 2003). C4-HSL signaling is also required for sloughing in Serratia marcescens biofilms (Rice et al., 2005). In contrast, treatment of wild-type P. aeruginosa biofilms with furanone 56 (Fig. 4E), a synthetic inhibitor of AHL signaling, induces biofilm dispersal (Hentzer et al., 2002). These findings suggest that P. aeruginosa biofilm dispersal is regulated by multiple AHL signaling networks.

In R. sphaeroides, a mutant strain deficient in the production of 7,8-cis-N-(tetradecenoyl)homoserine lactone formed large aggregates of cells in broth, and clumping was reversed by the addition of the AHL signal (Puskas et al., 1997). In P. aeruginosa, however, the detachment rates and size distributions of detached cell clumps were the same in wild-type and 3-oxo-C12-HSL-mutant biofilms (Wilson et al., 2004). These findings suggest that AHL signaling regulates biofilm dispersal differently in different species.

Pseudomonas Quinolone Signal

Another autoinducer produced by P. aeruginosa is 2-heptyl-3-hydroxy-4-quinolone, also known as Pseudomonas quinolone signal, or PQS (Fig. 4F) (Pesci et al., 1999). It has been shown (Dong et al., 2008) that exogenous PQS induces dispersal of wild-type P. aeruginosa biofilms cultured in microplate wells. Previous studies showed that PQS mediates cell death and DNA release in P. aeruginosa biofilms (Allesen-Holm et al., 2006), which suggests that PQS may induce central hollowing.


The cholera bacterium V. cholerae produces the autoinducer furanosylborate, also known as AI-2 (Fig. 4G). V. cholerae AI-2 mutants form thicker biofilms than wild-type strains on glass coverslips (Hammer and Bassler, 2003), and are also deficient in biofilm detachment (Liu et al., 2007). Thus, AI-2 may regulate V. cholerae biofilm dispersal.


Glycopeptidolipids (GPLs) are monoglycosylated, fatty acylated peptides that are further modified by small variable oligosaccharides (Fig. 4H). GPLs are the dominant immunogenic glycolipids of many mycobacteria (Brennan et al., 1981). It has been shown (Freeman et al., 2006) that GPLs mediate biofilm dispersal in the opportunistic pathogen Mycobacterium avium. Using a recirculating water biofilm reactor meant to simulate the conditions of a drinking water distribution system, these authors found that wild-type and GPL-mutant strains bound equally well to stainless steel coupons in the reactor, but that the mutant cells were present in relatively small numbers in the recirculating water phase compared with the number of wild-type cells. It was hypothesized that the mutants detached inefficiently from the biofilm due to enhanced cell-to-cell interactions.


Phenazines are tricyclic pyrazines that are produced by various bacteria, including members of the genera Pseudomonas and Streptomyces (Laursen and Nielsen, 2004). Phenazines have been shown to increase the survival of bacteria in natural environments, possibly due to their antimicrobial activity against other micro-organisms. Pseudomonas chloroaphis, a biological control bacterium, produces two major phenazines, phenazine-1-carboxylic acid (PCA) and 2-hydroxy-PCA (2OH-PCA) (Fig. 4I). It has been shown (Maddula et al., 2008) that a P. chloroaphis strain deficient in the production of 2OH-PCA and a strain that overproduces 2OH-PCA both exhibited lower biofilm dispersal rates than a wild-type strain when cultured in a flow cell biofilm reactor. The mechanisms by which phenazines regulate biofilm dispersal are not known.

Fatty Acid Signals

Several bacteria secrete unsaturated fatty acids that function as intra- and interspecies cell-to-cell communication signals (Wang et al., 2004). In P. aeruginosa, the fatty acid cis-2-decenoic acid (Fig. 4J) acts as a positive signal that is sensed by bacteria, thereby inducing a cascade of event that results in degradation of the biofilm matrix and biofilm dispersal (Davies and Marques, 2009). Exogenous cis-2-decenoic acid has been shown to induce biofilm dispersal in P. aeruginosa and other phylogenetically diverse bacteria and some fungi (Davies and Marques, 2009). Another unsaturated fatty acid, cis-11-methyl-2-dodecenoic acid, also known as diffusible signal factor or DSF, causes biofilm dispersal in X. campestris by up-regulating expression of the biofilm-matrix-degrading enzyme endo-β-1,4-mannanase (Dow et al., 2003). Numerous other bacteria secrete fatty acid signals, although their role in biofilm dispersal has not been investigated (Ryan and Dow, 2008).

Peptide Signals

Staphylococci produce and secrete a number of peptide signals that accumulate in the extracellular environment. Evidence suggests that two of these peptides—δ-toxin produced by S. epidermidis and S. aureus (Fig. 4K), and autoinducing peptide I (AIP-I) produced by S. aureus (Fig. 4L)—may play a role in biofilm dispersal.

Mutant strains of S. epidermidis that do not produce δ-toxin form significantly more biofilm biomass in microplate wells compared with the amount produced by wild-type strains (Vuong et al., 2003). In addition, exogenous δ-toxin decreases biofilm attachment in δ-toxin mutant strains of both S. epidermidis and S. aureus (Vuong et al., 2000, 2003). Microarray analysis has shown that the genes encoding δ-toxin and other related peptide signals are highly down-regulated in S. epidermidis biofilms (Yao et al., 2005). It is possible that δ-toxin contributes to biofilm dispersal through a physical mechanism that involves its detergent-like properties (Vuong et al., 2003).

Autoinducing peptides comprise a family of extracellular cyclic peptide signals produced by different strains of S. aureus (Ji et al., 1997). Exogenous autoinducing peptide AIP-I induces sloughing of S. aureus biofilms cultured in flow cells (Boles and Horswill, 2008; Lauderdale et al., 2009). AIP-I may mediate biofilm dispersal by up-regulating expression of the matrix-degrading proteases aureolysin and Spl (Boles and Horswill, 2008).

Nitric Oxide

Nitric oxide (NO), an endogenous product of anaerobic metabolism, has been shown to induce dispersal of P. aeruginosa biofilms (Barraud et al., 2006). A P. aeruginosa strain lacking nitrate reductase, the only enzyme capable of generating metabolic NO through anaerobic respiration, does not disperse, whereas a NO reductase mutant exhibited a hyper-dispersal phenotype. In addition, exogenously added sodium nitroprusside, a NO donor, induces detachment of pre-formed P. aeruginosa biofilm colonies. In these experiments, NO was used at sublethal concentrations (25 to 500 nm), suggesting that NO functions as an extracellular signal to mediate the dispersal effect (Romeo, 2006).

Cyclic Diguanyl Monophosphate

Cyclic dimeric GMP (c-di-GMP) is an intracellular signal that regulates the transition from sessile to planktonic growth in a variety of bacteria (Yildiz, 2008). Increased levels of c-di-GMP generally result in an increase in exopolysaccharide and fimbriae production, and a decrease in motility, whereas decreased levels of c-di-GMP exert the opposite effects and induce biofilm dispersal. c-di-GMP-regulated biofilm dispersal has been observed in many species, including P. aeruginosa, P. fluorescens, Salmonella enterica serovar Typhimurium, E. coli, Shewanella oneidensis, and various vibrios (Simm et al., 2004; Thormann et al., 2005, 2006; Morgan et al., 2006; Boehm et al., 2009; Gjermansen et al., 2009; Newell et al., 2009; Yildiz and Visick, 2009). It has been shown that c-di-GMP up-regulates the biofilm-matrix-degrading protease LapG in P. fluorescens and P. putida (Gjermansen et al., 2009; Newell et al., 2009), up-regulates biofilm matrix polysaccharide production in E. coli (Boehm et al., 2009), and down-regulates motility in P. putida (Gjermansen et al., 2006).

Clinical Relevance Of Biofilm Dispersal

Biofilm formation is the primary mode of growth for bacteria in most natural and clinical environments. For many pathogenic bacteria, therefore, biofilm dispersal plays a critical role in the transmission of bacteria from environmental reservoirs to human hosts, in the transmission of bacteria between hosts, and in the exacerbation and spread of infection within a single host.

Many pathogens are transmitted to human hosts from environmental reservoirs. The opportunistic pathogen P. aeruginosa, for example, lives in soil, water, vegetation, sinks, faucets, respiratory therapy equipment, and on the hands of healthcare workers (Foca et al., 2000). Other important pathogens that colonize environmental reservoirs include Legionella, Vibrio, Mycobacterium, and Listeria. Biofilm formation plays a key role in the ability of these bacteria to colonize most environmental niches, and biofilm dispersal is their primary means of escaping the biofilm to be translocated to their human hosts. Sloughing dispersal may be an important factor in the transmission of some environmental pathogens, because sloughing can result in the detachment of a sufficient number of cells for an infective dose that is not typically found in bulk fluid (Hall-Stoodley and Stoodley, 2005).

Biofilm dispersal also plays a key role in the communicable transmission of many pathogens. For example, S. mutans can detach from dental biofilms in a mother’s mouth and be transmitted to an infant by direct or indirect contact (Berkowitz and Jones, 1985). Similarly, A. actinomycetemcomitans can be transmitted from person to person by means of a shared toothbrush (Stabholz et al., 1998). This type of dispersal may contribute to the host-to-host transmission of many respiratory pathogens (Morris, 2007), and of V. cholerae during cholera epidemics (Nielsen et al., 2008).

The intra-host spread and persistence of bacteria are also mediated by biofilm dispersal. For example, detached S. mutans cells can be translocated to adjacent or opposing teeth by means of salivary flow (Svanberg and Loesche, 1978), and transient bacteremias are frequently detected following dental procedures (Kinane et al., 2005). Other examples of intra-host spread include: hospital-acquired pneumonia, caused by bacteria detached from biofilms in a patient’s endotracheal tube (Adair et al., 1999); infectious kidney stones, caused by bacteria detached from a biofilm in a patient’s bladder (Mathoera et al., 2000); and embolic events in endocarditis (Parsek and Singh, 2003). Twitching motility, a mechanism of surface translocation and a potential mode of biofilm dispersal, may contribute to persistence of P. aeruginosa in the lungs (Chiang and Burrows, 2003).

Although biofilm dispersal clearly plays a major role in disease transmission, few studies have examined the role of biofilm dispersal in pathogenesis. It has been shown (Bieber et al., 1998) that biofilm dispersal is required for full virulence of enteropathogenic E. coli in human volunteers. Using a model that measures diarrhea following oral inoculation, these authors found that a mutant strain deficient in the production of type IV bundle-forming pili, which are required for biofilm dispersal, was 200-fold less virulent than a wild-type strain. In the plant pathogen X. campestris, biofilm dispersal mediated by production of the matrix-degrading enzyme endo-β-1,4-mannanase was required for full virulence of the bacterium in plants (Dow et al., 2003).

Potential Therapeutic Uses

An anticipated offshoot of research on biofilm dispersal is the development of novel therapeutic and prophylactic approaches for the treatment of biofilm infections. Some classes of agents that may have clinical utility are biofilm-matrix-degrading enzymes, quorum-sensing signals, surfactants, and small molecule inhibitors of bacterial diguanylate cyclases.

Among the biofilm-matrix-degrading enzymes, dispersin B of A. actinomycetemcomitans has received the most attention. In vitro, exogenously added dispersin B has been shown to inhibit biofilm formation, detach pre-formed biofilms, and sensitize pre-formed biofilms to killing by antimicrobial agents, bacteriophages, and host defenses in phylogenetically diverse bacteria (Kaplan et al., 2004a; Itoh et al., 2005; Lu and Collins, 2007; Izano et al., 2008a,b; Venketaraman et al., 2008). In vivo, dispersin B completely eliminated S. aureus port-related bloodstream infections in catheterized sheep when used in combination with teicoplanin as a catheter lock solution (Jose del Pozo, personal communication), and reduced the rate of S. aureus catheter colonization from 97% to 3% when used in combination with triclosan as a catheter coating in a rabbit model of subcutaneous implant infections (Darouiche et al., 2009). One of the main advantages of dispersin B and other matrix-degrading enzymes is that they do not kill bacteria or inhibit their growth. This reduces the chances for the evolution of resistance to these agents.

Alginate lyase is another matrix-degrading enzyme that has therapeutic potential against biofilm-related pulmonary infections caused by P. aeruginosa. In vitro, alginate lyase reduces the viscoelasticity of purulent sputum from individuals with cystic fibrosis (Mrsny et al., 1994) and enhances the antibiotic killing of mucoid P. aeruginosa in biofilms (Alkawash et al., 2006; Alipour et al., 2009). In vivo, alginate lyase increases the effectiveness of amikacin against mucoid strains of P. aeruginosa in a rabbit model of infective endocarditis (Bayer et al., 1992). Alginate lyase, when used in combination with deoxyribonuclease, may be useful for the treatment of alginate polysaccharide build-up in the lungs of individuals with cystic fibrosis (Wong et al., 2000; VanDevanter and Van Dalfsen, 2005).

Many small molecules have been shown to induce biofilm cell detachment in vitro (Fig. 4), and some of these may have clinical applications. The NO donor sodium nitroprusside, for example, induces detachment of pre-formed P. aeruginosa biofilm colonies in vitro and greatly enhances the efficacy of antibiotics in the removal of the biofilms (Barraud et al., 2006). However, one study showed that sodium nitroprusside increased biofilm formation by P. aeruginosa and Burkholderia cenocepacia in vitro (Zaitseva et al., 2009). Quorum-sensing autoinducers, analogues, and antagonists are another class of promising anti-biofilm agents, but their utility in the clinic has still not been demonstrated. This may be because quorum-sensing circuitry is extremely complex, and the biological activity of these compounds is difficult to predict from in vitro studies. Unsaturated fatty acids are another example of signaling molecules that exhibit broad-spectrum biofilm-detaching activity in vitro (Davies and Marques, 2009), but their effectiveness in vivo has not been evaluated.

The c-di-GMP signaling pathways have received considerable attention over the past few years. Diguanylate cyclase enzymes are a very attractive target for antimicrobial therapy, because they are found only in bacteria and not in eukaryotic cells. Since decreased levels of intracellular c-di-GMP induce biofilm dispersal, inhibitors of these enzymes should inhibit biofilm formation or promote biofilm dispersal. However, diguanylate cyclases comprise a large super-family of enzymes, with many bacteria having dozens of homologues. The biological effect of diguanylate cyclase inhibition, therefore, is not easy to predict (Yildiz, 2008). The recent reporting of the three-dimensional structures of two guanylate cyclases, PleD from Caulobacter crescentus and FimX from P. aeruginosa (Wassmann et al., 2007; Navarro et al., 2009), should facilitate the discovery of small-molecule diguanylate cyclase inhibitors by rational drug design.


Research on biofilm dispersal is in its infancy. Virtually all dispersal studies have been performed in vitro under the controlled conditions of a laboratory, and most of these were performed with monospecies biofilms. It is extremely difficult to extrapolate these results to any environmental biofilm, especially a complex biofilm community such as dental plaque. Although numerous potential dispersal-inducing agents have been identified, it remains to be seen whether any of these agents will have clinical significance.

Sessile organisms such as plants and fungi have evolved a multitude of ingenious strategies to disperse seeds and spores into the environment to colonize new sites. In fact, burrs, helicopter seeds, and spore-shooting fungi represent some of the most remarkable evolutionary adaptations in nature. It is likely that biofilm bacteria have evolved similarly diverse dispersal mechanisms that are waiting to be discovered. There is a good chance that continued advances in biofilm dispersal research will soon lead to the development of novel therapies based on these findings.


Biofilm research in the author’s laboratory is funded by the National Institute of Allergies and Infectious Diseases and by Genentech, Inc.


  • Adair CG, Gorman SP, Feron BM, Byers LM, Jones DS, Goldsmith CE, et al. (1999). Implications of endotracheal tube biofilm for ventilator-associated pneumonia. Intensive Care Med 25:1072-1076 [PubMed]
  • Alipour M, Suntres ZE, Omri A. (2009). Importance of DNase and alginate lyase for enhancing free and liposome encapsulated aminoglycoside activity against Pseudomonas aeruginosa. J Antimicrob Chemother 64:317-325 [PubMed]
  • Alkawash MA, Soothill JS, Schiller NL. (2006). Alginate lyase enhances antibiotic killing of mucoid Pseudomonas aeruginosa in biofilms. APMIS 114:131-138 [PubMed]
  • Allesen-Holm M, Barken KB, Yang L, Klausen M, Webb JS, Kjelleberg S, et al. (2006). A characterization of DNA release in Pseudomonas aeruginosa cultures and biofilms. Mol Microbiol 59:1114-1128 [PubMed]
  • Allison DG, Evans DJ, Brown MRW, Gilbert P. (1990). Possible involvement of the division cycle in dispersal of Escherichia coli from biofilms. J Bacteriol 172:1667-1669 [PMC free article] [PubMed]
  • Allison DG, Ruiz B, SanJose C, Jaspe A, Gilbert P. (1998). Extracellular products as mediators of the formation and detachment of Pseudomonas fluorescens biofilms. FEMS Microbiol Lett 167:179-184 [PubMed]
  • Applegate DH, Bryers JD. (1991). Effects of carbon and oxygen limitations and calcium concentrations on biofilm removal process. Biotechnol Bioeng 37:17-25 [PubMed]
  • Banas JA, Vickerman MM. (2003). Glucan-binding proteins of the oral streptococci. Crit Rev Oral Biol Med 14:89-99 [PubMed]
  • Barraud N, Hassett DJ, Hwang SH, Rice SA, Kjelleberg S, Webb JS. (2006). Involvement of nitric oxide in biofilm dispersal of Pseudomonas aeruginosa. J Bacteriol 188:7344-7353 [PMC free article] [PubMed]
  • Baty AM, 3rd, Eastburn CC, Techkarnjanaruk S, Goodman AE, Geesey GG. (2000). Spatial and temporal variations in chitinolytic gene expression and bacterial biomass production during chitin degradation. Appl Environ Microbiol 66:3574-3585 [PMC free article] [PubMed]
  • Bayer AS, Park S, Ramos MC, Nast CC, Eftikhar F, Schiller NL. (1992). Effect of alginase on the natural history and antibiotic therapy of experimental endocarditis caused by mucoid Pseudomonas aeruginosa. Infect Immun 60:3979-3985 [PMC free article] [PubMed]
  • Bayles KW. (2007). The biological role of death and lysis in biofilm development. Nature Rev Microbiol 5:721-726 [PubMed]
  • Berkowitz RJ, Jones P. (1985). Mouth-to-mouth transmission of the bacterium Streptococcus mutans between mother and child. Arch Oral Biol 30:377-379 [PubMed]
  • Bieber D, Ramer SW, Wu C-Y, Murray WJ, Tobe T, Fernandez R, et al. (1998). Type IV pili, transient bacterial aggregates, and virulence of enteropathogenic Escherichia coli. Science 280:2114-2118 [PubMed]
  • Boehm A, Steiner S, Zaehringer F, Casanova A, Hamburger F, Ritz D, et al. (2009). Second messenger signaling governs Escherichia coli biofilm induction upon ribosomal stress. Mol Microbiol 72:1500-1516 [PubMed]
  • Boles BR, Horswill AR. (2008). Agr-mediated dispersal of Staphylococcus aureus biofilms. PLoS Pathog 4:e1000052. [PMC free article] [PubMed]
  • Boles BR, Thoendel M, Singh PK. (2005). Rhamnolipids mediate detachment of Pseudomonas aeruginosa from biofilms. Mol Microbiol 57:1210-1223 [PubMed]
  • Boyd A, Chakrabarty AM. (1994). Role of alginate lyase in cell detachment of Pseudomonas aeruginosa. Appl Environ Microbiol 60:2355-2359 [PMC free article] [PubMed]
  • Brennan PJ, Aspinall GO, Shin JE. (1981). Structure of the specific oligosaccharides from the glycopeptidolipid antigens of serovars in the Mycobacterium avium-Mycobacterium intracellulare-Mycobacterium scrofulaceum complex. J Biol Chem 256:6817-6822 [PubMed]
  • Chiang P, Burrows LL. (2003). Biofilm formation by hyperpiliated mutants of Pseudomonas aeruginosa. J Bacteriol 185:2374-2378 [PMC free article] [PubMed]
  • Choi YC, Morgenroth E. (2003). Monitoring biofilm detachment under dynamic changes in shear stress using laser-based particle size analysis and mass fractionation. Water Sci Technol 47:69-76 [PubMed]
  • Cleary J, Lai LC, Shaw RK, Straatman-Iwanowska A, Donnenberg MS, Frankel G, et al. (2004). Enteropathogenic Escherichia coli (EPEC) adhesion to intestinal epithelial cells: role of bundle-forming pili (BFP), EspA filaments and intimin. Microbiology 150(Pt 3):527-538 [PubMed]
  • Costerton JW, Stewart PS, Greenberg EP. (1999). Bacterial biofilms: a common cause of persistent infections. Science 284:1318-1322 [PubMed]
  • Darouiche RO, Mansouri MD, Gawande PV, Madhyastha S. (2009). Antimicrobial and antibiofilm efficacy of triclosan and DispersinB¯ combination. J Antimicrob Chemother 64:88-93 [PubMed]
  • Davey ME, Caiazza NC, O’Toole GA. (2003). Rhamnolipid surfactant production affects biofilm architecture in Pseudomonas aeruginosa PAO1. J Bacteriol 185:1027-1036 [PMC free article] [PubMed]
  • Davies DG, Marques CN. (2009). A fatty acid messenger is responsible for inducing dispersion in microbial biofilms. J Bacteriol 191:1393-1403 [PMC free article] [PubMed]
  • Delaquis PJ, Caldwell DE, Lawrence JR, McCurdy AR. (1989). Detachment of Pseudomonas fluorescens from biofilms on glass surfaces in response to nutrient stress. Microb Ecol 18:199-210 [PubMed]
  • Dong YH, Zhang XF, An SW, Xu JL, Zhang LH. (2008). A novel two-component system BqsS-BqsR modulates quorum sensing-dependent biofilm decay in Pseudomonas aeruginosa. Commun Integr Biol 1:88-96 [PMC free article] [PubMed]
  • Dow JM, Crossman L, Findlay K, He YQ, Feng JX, Tang JL. (2003). Biofilm dispersal in Xanthomonas campestris is controlled by cell-cell signalling and is required for full virulence in plants. Proc Natl Acad Sci USA 100:10995-11000 [PubMed]
  • Erard JC, Miyasaki KT, Wolinsky LE. (1989). Detachment of oral bacteria from saliva-coated hydroxyapatite by polymorphonuclear leukocytes. J Periodontol 60:211-216 [PubMed]
  • Finkelstein RA, Boesman-Finkelstein M, Chang Y, Häse CC. (1992). Vibrio cholerae hemagglutinin/protease, colonial variation, virulence, and detachment. Infect Immun 60:472-478 [PMC free article] [PubMed]
  • Foca M, Jakob K, Whittier S, Latta PD, Factor S, Rubenstein D, et al. (2000). Endemic Pseudomonas aeruginosa infection in a neonatal intensive care unit. New Engl J Med 343:695-700 [PubMed]
  • Frank KL, Patel R. (2007). Poly-N-acetylglucosamine is not a major component of the extracellular matrix in biofilms formed by icaADBC-positive Staphylococcus lugdunensis isolates. Infect Immun 75:4728-4742 [PMC free article] [PubMed]
  • Freeman R, Geier H, Weigel KM, Do J, Ford TE, Cangelosi GA. (2006). Roles for cell wall glycopeptidolipids in surface adherence and planktonic dispersal of Mycobacterium avium. Appl Environ Microbiol 72:7554-7558 [PMC free article] [PubMed]
  • Fuqua C, Greenberg EP. (2002). Listening in on bacteria: acyl-homoserine lactone signalling. Nat Rev Mol Cell Biol 3:685-695 [PubMed]
  • Fux CA, Wilson S, 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 [PMC free article] [PubMed]
  • Gilbert P, Allison DG, Evans DJ, Handley PS, Brown MR. (1989). Growth rate control of adherent bacterial populations. Appl Environ Microbiol 55:1308-1311 [PMC free article] [PubMed]
  • Gilbert P, Evans DJ, Brown MR. (1993). Formation and dispersal of biofilms in vivo and in situ. J Appl Bacteriol 74(Suppl):67S-78S [PubMed]
  • Gjermansen M, Ragas P, Sternberg C, Molin S, Tolker-Nielsen T. (2005). Characterization of starvation-induced dispersion in Pseudomonas putida biofilms. Environ Microbiol 7:894-906 [PubMed]
  • Gjermansen M, Ragas P, Tolker-Nielsen T. (2006). Proteins with GGDEF and EAL domains regulate Pseudomonas putida biofilm formation and dispersal. FEMS Microbiol Lett 265:215-224 [PubMed]
  • Gjermansen M, Nilsson M, Yang L, Tolker-Nielsen T. (2009). Characterization of starvation-induced dispersion in Pseudomonas putida biofilms: genetic elements and molecular mechanisms. Mol Microbiol (in press). doi:10.1111/j.1365-2958.2009.06793.x10.1111/j.1365-2958.2009.06793.x [PubMed] [Cross Ref]
  • Hall-Stoodley L, Stoodley P. (2005). Biofilm formation and dispersal and the transmission of human pathogens. Trends Microbiol 13:7-10; erratum in Trends Microbiol 3:300-301, 2005. [PubMed]
  • Hall-Stoodley L, Costerton JW, Stoodley P. (2004). Bacterial biofilms: from the natural environment to infectious diseases. Nature Rev Microbiol 2:95-108 [PubMed]
  • Hammer BK, Bassler BL. (2003). Quorum sensing controls biofilm formation in Vibrio cholerae. Mol Microbiol 50:101-114 [PubMed]
  • Helmstetter CE, Cummings DJ. (1964). An improved method for the selection of bacterial cells at division. Biochim Biophys Acta 82:608-610 [PubMed]
  • Hentzer M, Riedel K, Rasmussen TB, Heydorn A, Andersen JB, Parsek MR, et al. (2002). Inhibition of quorum sensing in Pseudomonas aeruginosa biofilm bacteria by a halogenated furanone compound. Microbiology 148(Pt 1):87-102 [PubMed]
  • Hunt SM, Werner EM, Huang B, Hamilton MA, Stewart PS. (2004). Hypothesis for the role of nutrient starvation in biofilm detachment. Appl Environ Microbiol 70:7418-7425 [PMC free article] [PubMed]
  • Inoue T, Shingaki R, Sogawa N, Sogawa CA, Asaumi J, Kokeguchi S, et al. (2003). Biofilm formation by a fimbriae-deficient mutant of Actinobacillus actinomycetemcomitans. Microbiol Immunol 47:877-881 [PubMed]
  • Irie Y, Parsek MR. (2008). Quorum sensing and microbial biofilms. Curr Top Microbiol Immunol 322:67-84 [PubMed]
  • Irie Y, O’Toole GA, Yuk MH. (2005). Pseudomonas aeruginosa rhamnolipids disperse Bordetella bronchiseptica biofilms. FEMS Microbiol Lett 250:237-243 [PubMed]
  • Itoh Y, Wang X, Hinnebusch BJ, Preston JF, 3rd, Romeo T. (2005). Depolymerization of β-1,6-N-acetyl-D-glucosamine disrupts the integrity of diverse bacterial biofilms. J Bacteriol 187:382-387 [PMC free article] [PubMed]
  • Izano EA, Wang H, Ragunath C, Ramasubbu N, Kaplan JB. (2007). Detachment and killing of Aggregatibacter actinomycetemcomitans biofilms by dispersin B and SDS. J Dent Res 86:618-622 [PubMed]
  • Izano EA, Amarante MA, Kher WB, Kaplan JB. (2008a). Differential roles of poly-N-acetylglucosamine surface polysaccharide and extracellular DNA in Staphylococcus aureus and Staphylococcus epidermidis biofilms. Appl Environ Microbiol 74:470-476 [PMC free article] [PubMed]
  • Izano EA, Sadovskaya I, Wang H, Vinogradov E, Ragunath C, Ramasubbu N, et al. (2008b). Poly-N-acetylglucosamine mediates biofilm formation and detergent resistance in Aggregatibacter actinomycetemcomitans. Microb Pathogen 44:52-60 [PMC free article] [PubMed]
  • Jackson DW, Suzuki K, Oakford L, Simecka JW, Hart ME, Romeo T. (2002). Biofilm formation and dispersal under the influence of the global regulator CsrA of Escherichia coli. J Bacteriol 184:290-301 [PMC free article] [PubMed]
  • James GA, Korber DR, Caldwell DE, Costerton JW. (1995). Digital image analysis of growth and starvation responses of a surface-colonizing Acinetobacter sp. J Bacteriol 177:907-915 [PMC free article] [PubMed]
  • Ji G, Beavis R, Novick RP. (1997). Bacterial interference caused by autoinducing peptide variants. Science 276:2027-2030 [PubMed]
  • Kachlany SC, Planet PJ, Desalle R, Fine DH, Figurski DH, Kaplan JB. (2001). flp-1, the first representative of a new pilin gene subfamily, is required for non-specific adherence of Actinobacillus actinomycetemcomitans. Mol Microbiol 40:542-554 [PubMed]
  • Kaplan JB, Fine DH. (2002). Biofilm dispersal of Neisseria subflava and other phylogenetically diverse oral bacteria. Appl Environ Microbiol 68:4943-4950 [PMC free article] [PubMed]
  • Kaplan JB, Meyenhofer MF, Fine DH. (2003a). Biofilm growth and detachment of Actinobacillus actinomycetemcomitans. J Bacteriol 185:1399-1404 [PMC free article] [PubMed]
  • Kaplan JB, Ragunath C, Ramasubbu N, Fine DH. (2003b). Detachment of Actinobacillus actinomycetemcomitans biofilm cells by an endogenous β-hexosaminidase activity. J Bacteriol 185:4692-4698 [PMC free article] [PubMed]
  • Kaplan JB, Velliyagounder K, Ragunath C, Fine DH, Ramasubbu N. (2004a). Enzymatic detachment of Staphylococcus epidermidis biofilms. Antimicrob Agents Chemother 48:2633-2636 [PMC free article] [PubMed]
  • Kaplan JB, Velliyagounder K, Ragunath C, Rohde H, Mack D, Knobloch JK, et al. (2004b). Genes involved in the synthesis and degradation of matrix polysaccharide in Actinobacillus actinomycetemcomitans and Actinobacillus pleuropneumoniae biofilms. J Bacteriol 186:8213-8220 [PMC free article] [PubMed]
  • Karatan E, Watnick P. (2009). Signals, regulatory networks, and materials that build and break bacterial biofilms. Microbiol Molec Biol Rev 73:310-347 [PMC free article] [PubMed]
  • Kinane DF, Riggio MP, Walker KF, MacKenzie D, Shearer B. (2005). Bacteraemia following periodontal procedures. J Clin Periodontol 32:708-713 [PubMed]
  • Kirov SM, Webb JS, O’May CY, Reid DW, Woo JK, Rice SA, et al. (2007). Biofilm differentiation and dispersal in mucoid Pseudomonas aeruginosa isolates from patients with cystic fibrosis. Microbiology 153(Pt 10):3264-3274 [PubMed]
  • Klausen M, Gjermansen M, Kreft JU, Tolker-Nielsen T. (2006). Dynamics of development and dispersal in sessile microbial communities: examples from Pseudomonas aeruginosa and Pseudomonas putida model biofilms. FEMS Microbiol Lett 261:1-11 [PubMed]
  • Knutton S, Shaw RK, Anantha RP, Donnenberg MS, Zorgani AA. (1999). The type IV bundle-forming pilus of enteropathogenic Escherichia coli undergoes dramatic alterations in structure associated with bacterial adherence, aggregation and dispersal. Mol Microbiol 33:499-509 [PubMed]
  • Koh KS, Lam KW, Alhede M, Queck SY, Labbate M, Kjelleberg S, et al. (2007). Phenotypic diversification and adaptation of Serratia marcescens MG1 biofilm-derived morphotypes. J Bacteriol 189:119-130 [PMC free article] [PubMed]
  • Lamont RJ, El-Sabaeny A, Park Y, Cook GS, Costerton JW, Demuth DR. (2002). Role of the Streptococcus gordonii SspB protein in the development of Porphyromonas gingivalis biofilms on streptococcal substrates. Microbiology 148:1627-1636 [PubMed]
  • Lappin-Scott HM, Bass C. (2001). Biofilm formation: attachment, growth, and detachment of microbes from surfaces. Am J Infect Control 29:250-251 [PubMed]
  • Lauderdale KJ, Malone CL, Boles BR, Morcuende J, Horswill AR. (2009). Biofilm dispersal of community-associated methicillin-resistant Staphylococcus aureus on orthopedic implant material. J Orthop Res 28:55-61 [PubMed]
  • Laursen JB, Nielsen J. (2004). Phenazine natural products: biosynthesis, synthetic analogues and biological activity. Chem Rev 104:1663-1686 [PubMed]
  • Lawrence JR, Scharf B, Packroff G, Neu TR. (2002). Microscale evaluation of the effects of grazing by invertebrates with contrasting feeding modes on river biofilm architecture and composition. Microb Ecol 44:199-207 [PubMed]
  • Lee SF, Li YH, Bowden GH. (1996). Detachment of Streptococcus mutans biofilm cells by an endogenous enzymatic activity. Infect Immun 64:1035-1038 [PMC free article] [PubMed]
  • Liu Z, Stirling FR, Zhu J. (2007). Temporal quorum-sensing induction regulates Vibrio cholerae biofilm architecture. Infect Immun 75:122-126 [PMC free article] [PubMed]
  • Lu TK, Collins JJ. (2007). Dispersing biofilms with engineered enzymatic bacteriophage. Proc Natl Acad Sci USA 104:11197-11202 [PubMed]
  • Ma L, Conover M, Lu H, Parsek MR, Bayles K, Wozniak DJ. (2009). Assembly and development of the Pseudomonas aeruginosa biofilm matrix. PLoS Pathog 5:e1000354. [PMC free article] [PubMed]
  • Maddula VS, Pierson EA, Pierson LS., 3rd (2008). Altering the ratio of phenazines in Pseudomonas chlororaphis (aureofaciens) strain 30-84 effects on biofilm formation and pathogen inhibition. J Bacteriol 190:2759-2766 [PMC free article] [PubMed]
  • Mah TF, O’Toole GA. (2001). Mechanisms of biofilm resistance to antimicrobial agents. Trends Microbiol 9:34-39 [PubMed]
  • Mai-Prochnow A, Evans F, Dalisay-Saludes D, Stelzer S, Egan S, James S, et al. (2004). Biofilm development and cell death in the marine bacterium Pseudoalteromonas tunicata. Appl Environ Microbiol 70:3232-3238 [PMC free article] [PubMed]
  • Mai-Prochnow A, Lucas-Elio P, Egan S, Thomas T, Webb JS, Sanchez-Amat A, et al. (2008). Hydrogen peroxide linked to lysine oxidase activity facilitates biofilm differentiation and dispersal in several gram-negative bacteria. J Bacteriol 190:5493-5501 [PMC free article] [PubMed]
  • Mann EE, Rice KC, Boles BR, Endres JL, Ranjit D, Chandramohan L, et al. (2009). Modulation of eDNA release and degradation affects Staphylococcus aureus biofilm maturation. PLoS One 4:e5822. [PMC free article] [PubMed]
  • Marsh PD. (2004). Dental plaque as a microbial biofilm. Caries Res 38:204-211 [PubMed]
  • Marsh PD. (2006). Dental plaque as a biofilm and a microbial community—implications for health and disease. BMC Oral Health 6(Suppl 1):14. [PMC free article] [PubMed]
  • Marshall KC. (1988). Adhesion and growth of bacteria at surfaces in oligotrophic habitats. Can J Microbiol 34:503-506
  • Mathoera RB, Kok DJ, Nijman RJ. (2000). Bladder calculi in augmentation cystoplasty in children. Urology 56:482-487 [PubMed]
  • Mireles JR, Toguchi A, Harshey RM. (2001). Salmonella enterica serovar Typhimurium swarming mutants with altered biofilm-forming abilities: surfactin inhibits biofilm formation. J Bacteriol 183:5848-5854 [PMC free article] [PubMed]
  • Morgan R, Kohn S, Hwang SH, Hassett DJ, Sauer K. (2006). BdlA, a chemotaxis regulator essential for biofilm dispersion in Pseudomonas aeruginosa. J Bacteriol 188:7335-7343 [PMC free article] [PubMed]
  • Morris DP. (2007). Bacterial biofilm in upper respiratory tract infections. Curr Infect Dis Rep 9:186-192 [PubMed]
  • Mrsny RJ, Lazazzera BA, Daugherty AL, Schiller NL, Patapoff TW. (1994). Addition of a bacterial alginate lyase to purulent CF sputum in vitro can result in the disruption of alginate and modification of sputum viscoelasticity. Pulm Pharmacol 7:357-366 [PubMed]
  • Navarro MV, De N, Bae N, Wang Q, Sondermann H. (2009). Structural analysis of the GGDEF-EAL domain-containing c-di-GMP receptor FimX. Structure 17:1104-1116 [PMC free article] [PubMed]
  • Neu TR. (1996). Significance of bacterial surface-active compounds in interaction of bacteria with interfaces. Microbiol Rev 60:151-166 [PMC free article] [PubMed]
  • Newell PD, Monds RD, O’Toole GA. (2009). LapD is a bis-(3′,5′)-cyclic dimeric GMP-binding protein that regulates surface attachment by Pseudomonas fluorescens Pf 0-1. Proc Natl Acad Sci USA 106:3461-3466 [PubMed]
  • Nielsen AT, Dolganov NA, Otto G, Miller MC, Wu CY, Schoolnik GK. (2008). RpoS controls the Vibrio cholerae mucosal escape response. PLoS Pathog 2:e109. [PMC free article] [PubMed]
  • O’Toole GA, Kolter R. (1998). Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol Microbiol 30:295-304 [PubMed]
  • Ott CM, Day DF, Koenig DW, Pierson DL. (2001). The release of alginate lyase from growing Pseudomonas syringae pathovar phaseolicola. Curr Microbiol 42:78-81 [PubMed]
  • Pamp SJ, Tolker-Nielsen T. (2007). Multiple roles of biosurfactants in structural biofilm development by Pseudomonas aeruginosa. J Bacteriol 189:2531-2539 [PMC free article] [PubMed]
  • Parsek MR, Singh PK. (2003). Bacterial biofilms: an emerging link to disease pathogenesis. Annu Rev Microbiol 57:677-701 [PubMed]
  • Pecharki D, Petersen FC, Scheie AA. (2008). Role of hyaluronidase in Streptococcus intermedius biofilm. Microbiology 154(Pt 3):932-938 [PubMed]
  • Pesci EC, Milbank JB, Pearson JP, McKnight S, Kende AS, Greenberg EP, et al. (1999) Quinolone signaling in the cell-to-cell communication system of Pseudomonas aeruginosa. Proc Natl Acad Sci USA 96:11229-11234 [PubMed]
  • Petersen FC, Pecharki D, Scheie AA. (2004). Biofilm mode of growth of Streptococcus intermedius favored by a competence-stimulating signaling peptide. J Bacteriol 186:6327-6331 [PMC free article] [PubMed]
  • Petersen FC, Tao L, Scheie AA. (2005). DNA binding-uptake system: a link between cell-to-cell communication and biofilm formation. J Bacteriol 187:4392-4400 [PMC free article] [PubMed]
  • Preston LA, Wong TY, Bender CL, Schiller NL. (2000). Characterization of alginate lyase from Pseudomonas syringae pv. syringae. J Bacteriol 182:6268-6271 [PMC free article] [PubMed]
  • Purevdorj-Gage B, Costerton WJ, Stoodley P. (2005). Phenotypic differentiation and seeding dispersal in non-mucoid Pseudomonas aeruginosa biofilms. Microbiology 151(Pt 5):1569-1576 [PubMed]
  • Puskas A, Greenberg EP, Kaplan S, Schaefer AL. (1997). A quorum-sensing system in the free-living photosynthetic bacterium Rhodobacter sphaeroides. J Bacteriol 179:7530-7537 [PMC free article] [PubMed]
  • Rice KC, Mann EE, Endres JL, Weiss EC, Cassat JE, Smeltzer MS, et al. (2007). The cidA murein hydrolase regulator contributes to DNA release and biofilm development in Staphylococcus aureus. Proc Natl Acad Sci USA 104:8113-8118 [PubMed]
  • Rice SA, Koh KS, Queck SY, Labbate M, Lam KW, Kjelleberg S. (2005). Biofilm formation and sloughing in Serratia marcescens are controlled by quorum sensing and nutrient cues. J Bacteriol 187:3477-3485 [PMC free article] [PubMed]
  • Rice SA, Tan CH, Mikkelsen PJ, Kung V, Woo J, Tay M, et al. (2009). The biofilm life cycle and virulence of Pseudomonas aeruginosa are dependent on a filamentous prophage. ISME J 3:271-282 [PMC free article] [PubMed]
  • Romeo T. (2006). When the party is over: a signal for dispersal of Pseudomonas aeruginosa biofilms. J Bacteriol 188:7325-7327 [PMC free article] [PubMed]
  • Rosan B, Lamont RJ. (2000). Dental plaque formation. Microbes Infect 2:1599-1607 [PubMed]
  • Ryan RP, Dow JM. (2008). Diffusible signals and interspecies communication in bacteria. Microbiology 154(Pt 7):1845-1858 [PubMed]
  • Sauer K, Camper AK. (2001). Characterization of phenotypic changes in Pseudomonas putida in response to surface-associated growth. J Bacteriol 183:6579-6589 [PMC free article] [PubMed]
  • Sauer K, Camper AK, Ehrlich GD, Costerton JW, Davies DG. (2002). Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J Bacteriol 184:1140-1154 [PMC free article] [PubMed]
  • Sauer K, Cullen MC, Rickard AH, Zeef LAH, Davies DG, Gilbert P. (2004). Characterization of nutrient-induced dispersion in Pseudomonas aeruginosa PAO1 biofilms. J Bacteriol 186:7312-7326 [PMC free article] [PubMed]
  • Sawyer LK, Hermanowicz SW. (1998). Detachment of biofilm bacteria due to variations in nutrient supply. Wat Sci Tech 37:211-214
  • Schooling S, Charaf UK, Allison DG, Gilbert P. (2004). A role for rhamnolipid in biofilm dispersion. Biofilms 1:91-99
  • Sheikh J, Czeczulin JR, Harrington S, Hicks S, Henderson IR, Le Bouguénec C, et al. (2002). A novel dispersin protein in enteroaggregative Escherichia coli. J Clin Invest 110:1329-1337 [PMC free article] [PubMed]
  • Simm R, Morr M, Kader A, Nimtz M, Römling U. (2004). GGDEF and EAL domains inversely regulate cyclic di-GMP levels and transition from sessility to motility. Mol Microbiol 53:1123-1134 [PubMed]
  • Soberón-Chávez G, Lépine F, Déziel E. (2005). Production of rhamnolipids by Pseudomonas aeruginosa. Appl Microbiol Biotechnol 68:718-725 [PubMed]
  • Stabholz A, Mann J, Agmon S, Soskolne WA. (1998). The description of a unique population with a very high prevalence of localized juvenile periodontitis. J Clin Periodontol 25(11 Pt 1):872-878 [PubMed]
  • Stanley NR, Lazazzera BA. (2004). Environmental signals and regulatory pathways that influence biofilm formation. Mol Microbiol 52:917-924 [PubMed]
  • Stoodley P, Wilson S, Hall-Stoodley L, Boyle JD, Lappin-Scott HM, Costerton JW. (2001). Growth and detachment of cell clusters from mature mixed-species biofilms. Appl Environ Microbiol 67:5608-5613 [PMC free article] [PubMed]
  • Stoodley P, Sauer K, Davies DG, Costerton JW. (2002). Biofilms as complex differentiated communities. Annu Rev Microbiol 56:187-209 [PubMed]
  • Svanberg ML, Loesche WJ. (1978). Intraoral spread of Streptococcus mutans in man. Arch Oral Biol 23:557-561 [PubMed]
  • Tam K, Kinsinger N, Ayala P, Qi F, Shi W, Myung NV. (2007). Real-time monitoring of Streptococcus mutans biofilm formation using a quartz crystal microbalance. Caries Res 41:474-483 [PMC free article] [PubMed]
  • Thormann KM, Saville RM, Shukla S, Spormann AM. (2005). Induction of rapid detachment in Shewanella oneidensis MR-1 biofilms. J Bacteriol 187:1014-1021 [PMC free article] [PubMed]
  • Thormann KM, Duttler S, Saville RM, Hyodo M, Shukla S, Hayakawa Y, et al. (2006). Control of formation and cellular detachment from Shewanella oneidensis MR-1 biofilms by cyclic di-GMP. J Bacteriol 188:2681-2691 [PMC free article] [PubMed]
  • Tolker-Nielsen T, Brinch UC, Ragas PC, Andersen JB, Jacobsen CS, Molin S. (2000). Development and dynamics of Pseudomonas sp. biofilms. J Bacteriol 182:6482-6489 [PMC free article] [PubMed]
  • VanDevanter DR, Van Dalfsen JM. (2005). How much do Pseudomonas biofilms contribute to symptoms of pulmonary exacerbation in cystic fibrosis? Pediatr Pulmonol 39:504-506 [PubMed]
  • Vats N, Lee SF. (2000). Active detachment of Streptococcus mutans cells adhered to epon-hydroxylapatite surfaces coated with salivary proteins in vitro. Arch Oral Biol 45:305-314 [PubMed]
  • Velarde JJ, Varney KM, Inman KG, Farfan M, Dudley E, Fletcher J, et al. (2007). Solution structure of the novel dispersin protein of enteroaggregative Escherichia coli. Mol Microbiol 66:1123-1135 [PubMed]
  • Venketaraman V, Lin AK, Le A, Kachlany SC, Connell ND, Kaplan JB. (2008). Both leukotoxin and poly-N-acetylglucosamine surface polysaccharide protect Aggregatibacter actinomycetemcomitans biofilm cells from macrophage killing. Microb Pathogen 45:173-180 [PMC free article] [PubMed]
  • Vuong C, Saenz HL, Götz F, Otto M. (2000). Impact of the agr quorum-sensing system on adherence to polystyrene in Staphylococcus aureus. J Infect Dis 182:1688-1693 [PubMed]
  • Vuong C, Gerke C, Somerville GA, Fischer ER, Otto M. (2003). Quorum-sensing control of biofilm factors in Staphylococcus epidermidis. J Infect Dis 188:706-718 [PubMed]
  • Wang LH, He Y, Gao Y, Wu JE, Dong YH, He C, et al. (2004). A bacterial cell-cell communication signal with cross-kingdom structural analogues. Mol Microbiol 51:903-912 [PubMed]
  • Wassmann P, Chan C, Paul R, Beck A, Heerklotz H, Jenal U, et al. (2007). Structure of BeF3–modified response regulator PleD: implications for diguanylate cyclase activation, catalysis and feedback inhibition. Structure 15:915-927 [PubMed]
  • Webb JS, Thompson LS, James S, Charlton T, Tolker-Nielsen T, Koch B, et al. (2003). Cell death in Pseudomonas aeruginosa biofilm development. J Bacteriol 185:4585-4592 [PMC free article] [PubMed]
  • Whiteley M, Bangera MG, Bumgarner RE, Parsek MR, Teitzel GM, Lory S, et al. (2001). Gene expression in Pseudomonas aeruginosa biofilms. Nature 413:860-864 [PubMed]
  • Wilson S, Hamilton MA, Hamilton GC, Schumann MR, Stoodley P. (2004). Statistical quantification of detachment rates and size distributions of cell clumps from wild-type (PAO1) and cell signaling mutant(JP1) Pseudomonas aeruginosa biofilms. Appl Environ Microbiol 70: 5847-5852 [PMC free article] [PubMed]
  • Wong TY, Preston LA, Schiller NL. (2000). Alginate lyase: review of major sources and enzyme characteristics, structure-function analysis, biological roles, and applications. Annu Rev Microbiol 54:289-340 [PubMed]
  • Wrangstadh M, Conway PL, Kjelleberg S. (1989). The role of an extracellular polysaccharide produced by the marine Pseudomonas sp. S9 in cellular detachment during starvation. Can J Microbiol 35:309-312
  • Xun LY, Mah RA, Boone DR. (1990). Isolation and characterization of disaggregatase from Methanosarcina mazei LYC. Appl Environ Microbiol 56:3693-3698 [PMC free article] [PubMed]
  • Yao Y, Sturdevant DE, Otto M. (2005). Genomewide analysis of gene expression in Staphylococcus epidermidis biofilms: insights into the pathophysiology of S. epidermidis biofilms and the role of phenol-soluble modulins in formation of biofilms. J Infect Dis 191:289-298 [PubMed]
  • Yarwood JM, Bartels DJ, Volper EM, Greenberg EP. (2004). Quorum sensing in Staphylococcus aureus biofilms. J Bacteriol 186:1838-1850 [PMC free article] [PubMed]
  • Yildiz FH. (2008). Cyclic dimeric GMP signaling and regulation of surface associated developmental programs. J Bacteriol 190:781-783 [PMC free article] [PubMed]
  • Yildiz FH, Visick KL. (2009). Vibrio biofilms: so much the same yet so different. Trends Microbiol 17:109-118 [PMC free article] [PubMed]
  • Ymele-Leki P, Ross JM. (2007). Erosion from Staphylococcus aureus biofilms grown under physiologically relevant fluid shear forces yields bacterial cells with reduced avidity to collagen. Appl Environ Microbiol 73:1834-1841 [PMC free article] [PubMed]
  • Yu C, Lee AM, Bassler BL, Roseman S. (1991). Chitin utilization by marine bacteria. J Biol Chem 266:24260-24267 [PubMed]
  • Zaitseva J, Granik V, Belik A, Koksharova O, Khmel I. (2009). Effect of nitrofurans and NO generators on biofilm formation by Pseudomonas aeruginosa PAO1 and Burkholderia cenocepacia 370. Res Microbiol 160:353-357 [PubMed]
  • Zhang X, Bishop PL. (2003). Biodegradability of biofilm extracellular polymeric substances. Chemosphere 50:63-69 [PubMed]

Articles from Journal of Dental Research are provided here courtesy of International and American Associations for Dental Research