Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Magnesium Limitation Is an Environmental Trigger of the Pseudomonas aeruginosa Biofilm Lifestyle

Magnesium Limitation Is an Environmental Trigger of the Pseudomonas aeruginosa Biofilm Lifestyle

  • Heidi Mulcahy, 
  • Shawn Lewenza


Biofilm formation is a conserved strategy for long-term bacterial survival in nature and during infections. Biofilms are multicellular aggregates of cells enmeshed in an extracellular matrix. The RetS, GacS and LadS sensors control the switch from a planktonic to a biofilm mode of growth in Pseudomonas aeruginosa. Here we detail our approach to identify environmental triggers of biofilm formation by investigating environmental conditions that repress expression of the biofilm repressor RetS. Mg2+ limitation repressed the expression of retS leading to increased aggregation, exopolysaccharide (EPS) production and biofilm formation. Repression of retS expression under Mg2+ limitation corresponded with induced expression of the GacA-controlled small regulatory RNAs rsmZ and rsmY and the EPS biosynthesis operons pel and psl. We recently demonstrated that extracellular DNA sequesters Mg2+ cations and activates the cation-sensing PhoPQ two-component system, which leads to increased antimicrobial peptide resistance in biofilms. Here we show that exogenous DNA and EDTA, through their ability to chelate Mg2+, promoted biofilm formation. The repression of retS in low Mg2+ was directly controlled by PhoPQ. PhoP also directly controlled expression of rsmZ but not rsmY suggesting that PhoPQ controls the equilibrium of the small regulatory RNAs and thus fine-tunes the expression of genes in the RetS pathway. In summary, Mg2+ limitation is a biologically relevant environmental condition and the first bonafide environmental signal identified that results in transcriptional repression of retS and promotes P. aeruginosa biofilm formation.


In the natural environment and during infection of susceptible hosts bacteria predominantly grow as biofilms. Biofilms are surface-associated, microbial communities, which are embedded in an extracellular matrix composed primarily of bacterial-derived exopolysaccharides (EPS) and DNA [1][3]. Biofilms have been intensively studied in recent years due to their significance in industrial, natural and medical settings. A multicellular biofilm lifestyle for bacteria has survival advantages compared to bacteria living as planktonic individual cells. During infection, growth as a biofilm confers resistance to multiple environmental stresses, antibiotics and the immune system [4][6].

The environmental bacterium and opportunistic human pathogen Pseudomonas aeruginosa PAO1 is one of the most widely used model organisms for studying bacterial biofilm formation. P. aeruginosa is a versatile Gram-negative bacterium that grows in soil and marine environments, as well as on plant and animal tissues [7]. P. aeruginosa has also emerged as a major opportunistic human pathogen during the past century [8] and individuals with the genetic disease Cystic Fibrosis (CF) are particularly susceptible [9]. A large body of in vitro and in vivo data in the literature indicates that P. aeruginosa exists as a biofilm in the lungs of CF patients [10][15].

Biofilm formation protects bacteria from harsh and stressful conditions [16] and the ability of bacteria to adapt to changing environmental conditions is essential for survival. P. aeruginosa is capable of sensing environmental conditions and adapting to changing conditions through modification of gene expression. A common mechanism of adaptation, which is both rapid and reversible, utilizes two-component systems (TCS) [17]. In many bacterial systems, TCS are involved in the regulation of biofilm formation. Typical TCS are comprised of a membrane-anchored histidine kinase sensor and a cytoplasmic response regulator (RR). After the sensor detects specific environmental signals, a signal transduction cascade is initiated that results in phosphorylation of a RR, which activates or represses necessary target genes. In P. aeruginosa multiple sensor proteins have been identified, mostly through genetic screens, which are important for P. aeruginosa biofilm formation. These include the sensor proteins GacS, RetS, LadS, SadS and PhoQ [18][23] as well as the BfiS, BfmS and MifS sensors, which control progression through distinct stages of biofilm maturation [24]. However, the exact signals detected by most of these environmental sensors are unknown.

RetS is required for expression of the virulence-associated type III secretion system (T3SS) and for repression of the pel and psl biofilm matrix EPS biosynthesis genes through both transcriptional and post-transcriptional regulation in P. aeruginosa [19], [20], [25], [26]. RetS directly interacts and forms heterodimers with the GacS sensor protein, preventing activation of the GacAS pathway and repressing biofilm formation [27], [28]. Reduced levels of RetS favors the formation of GacS homodimers, autophosphorylation of GacS and activation of the GacA-controlled the small regulatory RNAs (sRNA), rsmZ and rsmY [29], [30]. RsmZ and rsmY bind and sequester the post-transcriptional regulatory protein RsmA, which normally functions to bind other target mRNAs. RsmA-mediated regulation can be direct, through mRNA binding and preventing initiation of translation such as that observed for psl [26] or indirect, by interfering with the translation of specific regulatory factors [25]. However in addition post-transcriptional regulation of psl genes, genome-wide transcriptional profiling identified retS as a transcriptional regulator of the pel and psl EPS operons [19].

It has been proposed that the RetS sensor responds to environmental conditions encountered during acute infections maintaining P. aeruginosa in a planktonic growth state capable of Type III secretion [20]. In contrast to this model, we report here that Mg2+ limitation causes transcriptional repression of retS and promotes a switch from the planktonic to a biofilm lifestyle. Repression of retS occurred through direct repression by the cation sensing PhoPQ two-component system. Recent work from our laboratory has identified extracellular DNA as a chelator of divalent cations that activates the PhoPQ two-component system, resulting in the expression of antibiotic resistance genes [31]. To our knowledge this is the first demonstration of a specific environmental signal that promotes a switch to a biofilm mode of growth by acting through the RetS/LadS/GacS pathway. As DNA is abundant both in the natural and host environment [32][37], Mg2+ limitation is a relevant biological signal and is encountered ubiquitously by P. aeruginosa.


Identification of Mg2+ limitation as an environmental signal that represses retS expression

RetS is required for repression of EPS biosynthesis genes and preventing biofilm formation in P. aeruginosa [19]. Our strategy was to identify environmental signals that repressed the expression of retS and thus likely promoted biofilm formation in P. aeruginosa. To identify environmental signals that repressed the levels of RetS, we monitored expression of the retS promoter fused to the lux (bioluminescence) reporter in diverse growth conditions. The growth conditions tested were intended to mimic the conditions faced by P. aeruginosa during chronic lung infections (Table S1).

RetS expression was induced between 10- and 50-fold, or repressed between 7- and 125-fold in the conditions tested. Over the course of 20 h-growth, maximal repression (up to 125-fold) of retS was observed in limiting Mg2+ conditions (Figure 1A). We were particularly interested in retS gene expression under limiting Mg2+ growth conditions because we recently showed that extracellular DNA is an efficient chelator of divalent cations including Mg2+ [31]. Consistent with DNA acting as a chelator of Mg2+, retS expression was repressed up to 25-fold in media supplemented with exogenous DNA (Figure 1A). The expression of retS was repressed in a Mg2+ concentration-dependent manner (Figure 1B), with maximal repression observed at 0.02 mM Mg2+. Gene expression was normalized to growth as growth rates of cells grown under 2 mM or 0.02 mM Mg2+ were similar (6.2 h and 6.6 h, respectively).

Figure 1. Limiting Mg2+ repressed retS expression.

(A) The expression of retS was monitored in PAO1, using a promoter lux fusion (pMS402-lux), in different media and expressed as fold change in the medium indicated relative to BM2 2 mM Mg2+ during late log phase (11 h). (B) Expression of retS (CPS/OD600) was repressed by Mg2+ limitation in a concentration-dependent manner. Values are representative of at least 3 independent experiments. For each experiment the standard deviations were not greater that +/−10% of the mean value.

Mg2+ limitation results in a gene expression signature consistent with the switch to a biofilm lifestyle

Several studies examining the regulation of virulence factor production in P. aeruginosa identified a potential regulatory switch involving the RetS/LadS/GacS two component sensors, that controls the transition between acute and chronic infection related phenotypes [19], [20]. Identification of limiting Mg2+ as an environmental signal that represses expression of the known biofilm repressor gene retS prompted us to investigate the expression of a panel of genes controlled by the RetS/LadS/GacS pathway that are known to be important for either acute or chronic P. aeruginosa infections.

The data in Figure 2A represents the time course of fold induction (green) or repression (red) of genes in BM2 growth medium with limiting Mg2+ relative to high Mg2+ over 18 h of growth. In the cluster analysis depicted in Figure 2A, gene expression profiles cluster into two distinct groups: genes that are upregulated or downregulated in limiting Mg2+ conditions. The cluster of induced genes included genes previously identified as regulated by Mg2+ limitation by the PhoPQ or PmrAB TCS. These included PA3553 (arnC) from the LPS modification operon PA3552-3559, the outer membrane protein oprH, and a putative polyamine synthesis gene PA4774 [38][40]. Additionally the small regulatory RNAs encoded by rsmZ and rsmY, as well as genes from two independent EPS biosynthesis operons, pelD and pslA, were also induced by Mg2+ limitation (Figure 2). In contrast, genes encoding the retS biofilm regulator, type II secretion system genes (xcpR and aprA) and a type III secretion system effector, exoT, clustered together as genes repressed by Mg2+ limitation. To demonstrate the magnitude of induction or repression, Figure 2B illustrates the maximal change in gene expression over 18 h.

Figure 2. Limiting Mg2+ induced expression of EPS biosynthesis and small regulatory RNAs while retS expression was repressed.

(A) Gene expression data are expressed as fold induction (green) or fold repression (red) of bacteria cultured in BM2 0.02 mM Mg2+ relative to BM2 2 mM Mg2+. For each gene, each square represents relative expression values measured at 20 min intervals throughout growth. Gene expression profiles were grouped by hierarchical clustering using average linkage analysis (Cluster 3.0) and visualized using Treeview. (B) Maximal fold change in gene expression of transcriptional lux fusions in BM2 0.02 mM Mg2+ relative to BM2 2 mM Mg2+ over 18 h. Values are representative of at least 3 independent experiments and error bars represent the standard error of the mean (SEM).

According to the current model of reciprocal regulation of acute and chronic infection related traits by the RetS/LadS/GacS pathway, biofilms and chronic infections are promoted under conditions that induce EPS expression while simultaneously repressing the T2SS and T3SS [19], [20]. The expression profiles observed for the genes shown in Figure 2 are consistent with limiting Mg2+ acting as a signal that promotes a switch from the planktonic to an aggregative, biofilm mode of growth. We observed that maximal repression of the T2SS and T3SS genes occurred in the log phase of growth, prior to maximal repression of retS. This indicates that additional factors, the identification of which are beyond the scope of this study, may be involved in regulating these secretion system genes under Mg2+ limitation.

Mg2+ limitation promotes biofilm formation and aggregation

Pel and Psl EPS are essential biofilm matrix components that are important for adhesion and biofilm formation [41], [42]. As the expression of pel and psl EPS biosynthesis genes were strongly induced by Mg2+ limitation, we predicted that biofilm formation would be increased under these conditions. Mg2+ limitation strongly promoted biofilm formation, as measured by crystal violet (CV) staining of the total biomass adhered to the polystyrene pegs (Figure 3A) or to glass surfaces (Figure 3B). Additionally, large bacterial aggregates (50–70 µM diameter) (Figure 3C) were observed in mid-log phase planktonic cultures under Mg2+ limitation. No large aggregates of bacteria were visible when grown in high Mg2+ (Figure 3D).

Figure 3. Mg2+ limitation promoted biofilm formation and aggregation in P. aeruginosa.

(A) Attachment of cells to polystyrene pegs was monitored using crystal violet staining over time. Each point represents the average value obtained from twelve pegs and error bars represent the standard deviation. *, significant increase (p<0.05, ANOVA) in BM2 0.02 mM Mg2+ compared to that of BM2 2 mM Mg2+. (B) Attachment of cells to glass after 24 h measured using crystal violet staining. (C–D) Aggregation of PAO1 pCHAP6656 in liquid cultures was visualized by fluorescence microscopy in BM2 (C) 0.02 mM or (D) 2 mM Mg2+. Images are representative of data obtained in 3 independent experiments. Scale bars represent 10 µM.

The cation chelators DNA and EDTA induce biofilm formation

We have recently shown that extracellular DNA can function as a cation chelator that activates the cation-sensing PhoPQ TCS, leading to increased antimicrobial peptide resistance [31]. Since limiting Mg2+ is an environmental condition that also promoted biofilm formation, we proposed that cation chelation by extracellular DNA may ultimately impose a cation limitation on cells and promote biofilm formation.

To test this hypothesis, we cultivated ring biofilms in media with increasing concentrations of exogenous DNA. The addition of salmon sperm DNA promoted biofilm formation in a concentration-dependent manner (Figure 4A). To confirm that the chelating activity of DNA promoted biofilm formation, as opposed to the adhesive capacity of DNA, exogenous Mg2+ was added to cultures. The biofilm promoting effect of exogenous DNA was significantly reduced by addition of 10 mM excess Mg2+, to a final concentration of 12 mM Mg2+(Figure 4A). The cation chelator EDTA also caused a concentration-dependent increase in biofilm formation that was also completely neutralized by 10 mM excess Mg2+ (Figure 4B).

Figure 4. Mg2+ chelation by DNA and EDTA induced biofilm formation.

Attachment of PAO1 to wells in a 96-well plate was assessed using crystal violet staining after 24 h growth in (A) BM2 2 or 12 mM Mg2+ with increasing concentration of salmon sperm DNA or (B) LB in the absence and presence of 10 mM Mg2+ with increasing concentrations of EDTA. Values are representative of at least 3 independent experiments and error bars represent the standard error of the mean (SEM). a, significant increase (p<0.05, ANOVA) in media alone compared to that of media with DNA or EDTA; b, significant increase (p<0.05, ANOVA) in media with DNA or EDTA compared to that of media with DNA or EDTA and excess (10 or 12 mM) Mg2+.

Mg2+ limitation induces production of EPS in the biofilm matrix

Biofilm formation is induced under limiting Mg2+ conditions (Figure 3). To confirm increased EPS production under limiting Mg2+ conditions, we compared EPS production in planktonic cultures grown in 0.02 and 2 mM Mg2+. Using the congo red assay for measuring EPS production [43], P. aeruginosa produced significantly more total EPS under Mg2+ limiting conditions relative to high Mg2+ conditions (Figure 5A). Single mutants in either the pel or psl EPS biosynthesis operons showed significantly decreased congo red binding, 40% of wild-type levels (Figure 5B). Furthermore, the double pel/psl mutant exhibited 60% reduced congo red binding relative to PAO1, confirming the requirement of Pel and Psl for EPS production (Figure 5B). Both rsmA and retS mutants (PAZH13, retS::lux) exhibited elevated EPS levels and were included as positive controls for EPS production.

Figure 5. Mg2+ limitation induced EPS production.

EPS production was quantified after 24 h growth of (A) PAO1 in BM2 2 or 0.02 mM Mg2+ or (B) Wild type and mutant strains grown in BM2 0.02 mM Mg2+ using congo red binding. (C) Ring biofilm formation in 96 well plates was quantitated in BM2 2 mM Mg2+ or 0.02 mM Mg2+ in the absence and presence of 5 mg/ml of cellulase. Values are representative of at least 3 independent experiments and error bars represent the standard error of the mean (SEM). (D) Fluorescence microscopy was used to visualize aggregation and EPS production. Bacteria were grown in BM2 liquid cultures with 2 or 0.02 mM Mg2+ supplemented with 200 µg/ml calcofluor. At 24 h cells were stained with 1 µM SYTO9 (live cells, green, upper panel) and 10 µM propidium iodide (dead cells and DNA, red, lower panel). Scale bars represent 10 µM. (E) For scanning electron microscopy, PAO1 was grown as biofilms on polystyrene pegs in BM2 containing 2 or 0.02 mM Mg2+ for 48 h. Scale bars represent 2 µM. Images are representative of data obtained in three independent experiments. (F) Expression of retS, measured using a promoter lux fusion (pMS402-lux), in BM2 2 mM Mg2+ or 0.02 mM Mg2+ in the absence and presence of EPS-degrading cellulase. Values are representative of at least 3 independent experiments. For each experiment the standard deviations were not greater that +/−10% of the mean value. *, significant difference (p<0.05, ANOVA) relative to control conditions or between wildtype and mutant strains. a, significant increase (p<0.05, ANOVA) in BM2 0.02 mM Mg2+ compared to that of BM2 2 mM Mg2+; b, significant difference (p<0.05, ANOVA) in BM2 0.02 mM compared to that of BM2 0.02 mM with 5 mg/ml of cellulase.

Cellulase has been used previously to degrade Psl EPS and effectively reduces P. aeruginosa biofilm formation [43]. The addition of cellulase during biofilm cultivation had no effect on biofilms cultivated in high Mg2+ conditions but caused a significant reduction in biofilm formation under Mg2+ limitation (Figure 5C). This data indicates the importance of EPS in biofilm formation under limiting Mg2+ conditions as degradation of the EPS matrix with cellulase results in significantly lower biofilm formation during growth in limiting Mg2+.

To correlate EPS production with surface attachment, we tested biofilm formation phenotypes of EPS synthesis mutants. Individual pel or psl and double pel/psl mutants exhibited significantly reduced biofilm formation (up to 80%) relative to PAO1 (Figure S1). Additionally, we found that biofilm formation under Mg2+ limitation was not significantly reduced in mutants lacking type IV pili or flagella production (Figure S1). This suggests that Mg2+-limited biofilm formation does not require the presence of pili or flagella, which are generally important adhesins for biofilm formation [44].

Calcofluor binds sugars with β-1,4 linkages and has previously been shown to positively correlate with EPS production in a number of bacterial species including P. aeruginosa [41], [43], [45], Salmonella enterica serovar Typhimurium, and Escherichia coli [46][49]. Microscopic analysis indicated calcofluor bound to P. aeruginosa aggregates cultured under limiting Mg2+ and the absence of both aggregation and calcofluor binding in high Mg2+ conditions (Figure 5D). Aggregates formed under limiting Mg2+-conditions were EPS-dependent as pelD, pslH and pel/psl mutants failed to aggregate or stain positively for EPS under these conditions (Figure S2). Scanning electron microscopy indicated that biofilms grown under high Mg2+ conditions resulted in diffuse cell clusters with no observable interconnecting matrix or conditioning layer (Figure 5E). However, biofilms grown in limiting Mg2+ conditions clearly produced an extracellular matrix that coated the plastic surface and connected cells in web-like patterns (Figure 5E). Taken together a combination of phenotypic assays and microscopic analysis confirmed EPS overproduction in limiting Mg2+ conditions and that increased expression of EPS promoted biofilm formation in limiting Mg2+ conditions.

Aggregation is not the signal that leads to retS repression under Mg2+ limiting conditions

Our data was consistent with the hypothesis that Mg2+ limitation led to increased EPS production and biofilm formation, as a result of repression of the biofilm repressor RetS. An alternative interpretation to this data could be that aggregation itself may serve as the environmental cue for repressing retS transcription. To test this possibility, we examined the effects of adding cellulase, which degrades EPS, on retS expression. Cellulase was added to limiting Mg2+ cultures at the beginning of growth, as cellulase is known to degrade the EPS matrix and reduce cell-cell aggregation [43] and biofilm formation (Figure 5C). Cellulase treatment did not result in an increase in retS expression (Figure 5F). Similarly, retS expression was still repressed in the non-aggregating pel/psl double mutant strain compared to wildtype levels (data not shown) indicating that Mg2+ limitation represses retS expression independent of aggregation.

Overexpression of the RetS sensor prevents biofilm formation under Mg2+ limitation

Previous studies showed mutation of retS results in a hyperbiofilm phenotype due to hyperproduction of Pel and Psl exopolysaccharides [19], [20]. The expression of retS is repressed in Mg2+ limitation, which correlates with increased EPS production and biofilm formation. To definitively show that reduced levels of retS expression under Mg2+ limitation was essential for promoting biofilm formation, the retS gene was cloned (without its native promoter) under the control of a rhamnose-inducible promoter [50]. Biofilms were cultivated in the presence of increasing amounts of rhamnose to induce retS expression in a concentration-dependent manner. In the absence of rhamnose, expression of RetS from pSCRhaB2RetS caused a reduction in biofilm formation, most likely due to low, basal levels of expression of RetS from the plasmid (Figure 6). In support of our hypothesis, rhamnose-induced expression of RetS from pSCRhaB2RetS inhibited biofilm formation in limiting Mg2+ conditions (Figure 6), a condition which otherwise promotes biofilm formation (Figure 3), as a result of reduced retS expression (Figure 1).

Figure 6. Rhamnose-inducible expression of RetS prevented biofilm formation in Mg2+ limiting growth conditions.

Crystal violet staining of ring biofilms was performed in biofilms grown in BM2 0.02 mM Mg2+ with increasing rhamnose concentrations, where retS expression was induced using pSCRhaRetS. Values are representative of at least 3 independent experiments and error bars represent the standard error of the mean (SEM). ***, significant difference (p<0.001, ANOVA) in PAO1 or PAO1pSCRhaB2 (vector control) compared to that of PAO1pSCRhaRetS.

The Mg2+ sensing PhoPQ TCS influences the RetS regulatory pathway

Our results above identified a novel environmental signal that induced biofilm formation through repression of RetS. RetS acts as part of a complex signaling pathway that reciprocally regulates the expression of numerous virulence related genes [19]. It is unlikely that RetS itself responds to Mg2+ levels, since RetS is predicted to bind carbohydrates [51], [52]. Furthermore as PhoPQ is the only known TCS that is capable of sensing and responding to cation limitation in P. aeruginosa [53], [54], we hypothesized that PhoPQ repressed retS expression, and thus influenced the RetS/GacS/LadS pathway. If this model was true, it would be predicted that the expression profile of genes in the RetS pathway would be reversed in a phoP::xylE mutant relative to PAO1, and that high levels of retS expression in phoP::xylE would correspond to decreased levels of the small regulatory RNAs rsmZ and rsmY and the pel and psl EPS biosynthesis genes.

Expression of retS was derepressed in a phoP::xylE mutant compared to PAO1 in BM2 0.02 mM Mg2+, indicating that PhoP is required to repress retS (Figure 7A). No differences in retS expression were observed between PAO1 and phoP::xylE in BM2 2 mM Mg2+ (data not shown), a condition where the PhoPQ system is inactive. Expression of the psl EPS genes was reduced (60-fold), as predicted, in the phoP::xylE mutant (Figure 7A). In support of the link between PhoP and EPS production, a phoP::xylE mutant produced significantly less EPS as measured by the congo red binding assay and by direct visualization using transmission electron microscopy (Figure S3). Both rsmY and rsmZ had increased expression under limiting Mg2+conditions (Figure 2) and it was hypothesized that levels of both rsmY and rsmZ would decrease in a phoP::xylE mutant. This hypothesis was true for rsmY expression, which was repressed in the phoP::xylE mutant. In contrast, rsmZ expression was more highly expressed in a phoP::xylE mutant (Figure 7A). While rsmY and rsmZ are both induced under limiting Mg2+conditions in PAO1 (Figure 2), the relative levels of rsmY are higher (45-fold) (Figure 7A). In a phoP::xylE mutant this expression pattern is reversed, with higher relative levels of rsmZ observed (47-fold) (Figure 7A), indicating that the PhoPQ system differentially regulates rsmZ and rsmY expression.

Figure 7. Crosstalk between the Mg2+ sensing PhoPQ TCS and the RetS biofilm regulatory pathway.

(A) PAO1 and phoP::xylE were cultured in BM2 0.02 mM Mg2+. Expression of retS, rsmZ, rsmY and psl was analysed using plasmid encoded promoter-lux fusions. Gene expression data during log phase are shown. Values are representative of at least 3 independent experiments and error bars represent the standard error of the mean (SEM). (B) Identification of putative PhoP box (repeats indicated in bold) in the retS promoter with the predicted (BPROM) −10 and −35 sites (italicized). A full length (366 bp) and truncated (239 bp) retS promoter, with and without the predicted PhoP box (bold) (genome coordinates 5451891–5452257 and 5451891–5452131, respectively) were PCR amplified and labeled with DIG. Arrow indicates last nucleotide of reverse primer for truncated promoter, upstream of the GTG start codon (grey). (C) Gel shift assays of DIG-labeled full length and truncated retS promoter fragments with purified His6-PhoP protein. (D) RetS promoter activity (lux) in BM2 0.02 mM Mg2+ cultures of PAO1 and phoP::xylE using wildtype retS promoter sequences and in PAO1 following mutation (underlined) of the second repeat in the PhoP box in the retS promoter. Data shown are representative of 3 independent experiments and error bars represent the standard error of the mean (SEM). **, significant difference (p<0.01, ANOVA); ***, significant difference (p<0.001, ANOVA) in PAO1 compared to that of phoP::xylE.

PhoP directly represses retS and rsmZ expression

We previously characterized the PhoP regulon and identified PhoP binding sites in promoters of genes directly controlled by PhoPQ [40]. We examined the retS promoter for the presence of a PhoP consensus binding site (GTTCAGNNNNNRTTCAG) and found a candidate PhoP binding site between the start codon and the −10 promoter region (Figure 7B); a position consistent with PhoP acting as a repressor of gene expression. PhoP binding assays were performed using purified His6-PhoP with a 366 bp promoter fragment. PhoP bound this fragment causing a shift (Figure 7C). Using a truncated promoter fragment (255 bp), which excluded the predicted PhoP box, we observed that PhoP did not bind to the retS promoter lacking the predicted PhoP binding site (Figure 7C).

To confirm that PhoP repression of retS expression required the putative PhoP binding site, we constructed site-directed mutations in the PhoP box within the retS promoter. We initially constructed a retS promoter fragment that lacked the entire 18 bp PhoP box and this promoter-lux fusion had no promoter activity (data not shown), likely as a result of deleting the transcription start site. To ensure the mutation strategy did not interfere with transcription, we created site-directed changes in the second direct repeat of the PhoP box (Figure 7D). Substitution of the wildtype sequence cgttcc for cgggac, cgcctc or cgactc resulted in derepression of retS under limiting Mg2+ conditions in wildtype PAO1, thus confirming the identity of this sequence as the site of PhoP-mediated repression (Figure 7D).

Our data suggested that PhoP controlled multiple genes in this pathway by repressing both retS and rsmZ (Figure 7A). Analysis of the rsmZ promoter revealed a candidate PhoP binding site, between the start codon and the −10 promoter region (Figure 8A), similar to the PhoP-repressing binding site identified in the retS promoter (Figure 7B). Binding assays indicated that His6-PhoP was capable of direct binding to the rsmZ promoter (Figure 8B). Taken together, these data indicate that Mg2+ limitation is sensed by the PhoPQ two-component system, which acts as a direct negative regulator of both retS and rsmZ expression, resulting in enhanced biofilm formation.

Figure 8. PhoP repressed rsmZ expression through direct binding.

(A) Identification of putative PhoP box (bold) in rsmZ promoter (genome coordinates 4057874–4057855); −10 and −35 indicated in italics. (B) Gel shift assay of DIG-labeled rsmZ promoter fragment with purified His6-PhoP protein. Data shown are representative of 3 independent experiments.


Previous studies of P. aeruginosa have identified multiple signaling networks that control biofilm formation. These include the sensor kinases GacS, RetS and LadS that comprise a sophisticated regulatory network involved in reciprocal regulation of biofilm-associated and classical virulence-associated genes [19], [20]. However, the central question regarding the role of TCS in biofilm formation remains the identification of the actual environmental signals that are sensed.

The gene expression profiling approach described here, successfully identified Mg2+ limitation as an environmental condition that simultaneously repressed retS expression and induced expression of the pel/psl EPS biosynthesis genes. Our observation of transcriptional effects on the EPS biosynthesis genes that correlated with decreased levels of retS expression is consistent with a previous microarray study that detected transcriptional regulation of pel and psl in a retS mutant [19]. An additional study has identified post-transcriptional regulation of psl through RsmA [26], which lies downstream of RetS in the regulatory pathway. Thus, RetS is capable of regulating EPS production by both transcriptional and post-transcriptional mechanisms.

The kinetics of gene expression in Figure 2 revealed three gene expression patterns during the early, middle and late stages of growth. The known PmrA- and PmrA/PhoP-regulated genes PA4774 and PA3553 (arnC) were induced early in growth at 1.5 and 5 h, respectively. The exclusively PhoP-controlled gene oprH was maximally induced at the middle stage of growth, 13.5 h. Maximal induction of rsmZ, rsmY, pel and psl occurred at later stages of growth following repression of retS between 7.5 and 14 h (Figure 2A). Other sensor/regulator genes tested but which did not show a significant fold change under high and limiting Mg2+ conditions included rocR, rocA1, ladS, gacS, fleSR and pilSR. In agreement with the gene expression profile, growth under Mg2+ limitation caused a switch to the biofilm mode with increased aggregation (Figure 3), EPS production (Figure 5) and a hyperbiofilm phenotype (Figure 3).

Aggregation effects observed during growth in limiting Mg2+ conditions were controlled by PhoP-mediated repression of retS and fine-tuning of rsmZ/rsmY levels. Analysis of rsmZ and rsmY expression in phoP::xylE indicated that PhoP acts differently on the expression of these two sRNA-encoding genes, directly controlling the expression of rsmZ and indirectly controlling the levels of rsmY. This is similar to previous studies where rsmY and rsmZ were differentially regulated by other TCS [55], [56] and further suggests that rsmZ and rsmY are not functionally redundant but may exert different effects depending on the environmental conditions. In the wildtype PAO1, both rsmZ and rsmY are induced under Mg2+ limitation (Figure 2). This is likely a consequence of reduced RetS levels and activation of the GacAS pathway [19]. However the relative level of rsmY in PAO1 is significantly higher compared to that of rsmZ (45-fold) (Figure 7A), suggesting that expression of rsmZ is limited by PhoP repression (Figure 8). It may be that direct PhoP regulation of rsmZ serves to fine-tune the relative levels of the small regulatory RNAs, in order to maintain higher relative levels of rsmY, which corresponds to increased biofilm formation. This is consistent with a recent report showing that the overexpression of rsmZ, but not rsmY, impairs P. aeruginosa biofilm formation [56].

Extracellular DNA has also been shown to be important for the early stages of biofilm formation [37] and Mg2+ sequestration by DNA may play a role in maturation of P. aeruginosa biofilms by inducing EPS matrix production. Extracellular DNA, of both bacterial and eukaryotic origin, can be found at high concentrations (up to 20 mg/ml) in the CF lung [33], [35], [36]. DNA also accumulates in other environmental niches of P. aeruginosa, including soil ([57] and aquatic environments [34], and therefore likely contributes to P. aeruginosa biofilm formation in the natural environment.

Cation chelation by both DNA and EDTA can induce biofilm formation (Figure 4B). This observation is in contrast to a previous study, which showed that EDTA is a potent biofilm disrupter in P. aeruginosa [58]. This apparent contradiction can be explained by opposite effects of subinhibitory and lethal concentrations of chelators. Since EDTA is lethal at a concentration of 1 mM by disrupting membrane integrity [31], it is not surprising that 50 mM EDTA disrupts the biofilm matrix structure and kills biofilm cells [58].

This study identifies Mg2+ limitation as an important environmental trigger of P. aeruginosa biofilm development and increases our understanding of the potential role for extracellular DNA and cation chelation in the regulation of antibiotic resistant and aggregative biofilms. Novel approaches to block the PhoPQ and/or RetS signaling pathways, or to neutralize extracellular DNA, may be an effective and novel treatment strategy to prevent or reduce biofilm formation during infections and in the natural environment.

Materials and Methods

Strains, plasmids and media conditions

All strains and plasmids are listed in Tables 1 and 2, respectively. For details on construction of promoter fusions see Methods S1 primer Table S2. P. aeruginosa strains were routinely grown and maintained on Luria-Bertani (LB) plates or LB broth at 37°C and cultured in defined basal medium 2 (BM2) media [59] containing 0.02 mM Mg2+ (limiting) and 2 mM Mg2+ (high). Trimethoprim (Sigma-Aldrich) was added at 300 µg/ml for selection and 150 µg/ml for plasmid maintenance. Unless otherwise stated succinate (20 mM) was used as the carbon source. The source of DNA was fish sperm DNA-potassium salt (USB, Cleveland, OH).

Gene expression assays

Gene expression assays were carried out as previously described [31]. Gene expression values (counts per second, CPS) were normalized to cell number (optical density, OD). Gene expression profiles were grouped by hierarchical clustering using complete linkage. Analysis was performed using Cluster 3.0 and visualized using Treeview [60].

Biofilm, aggregation, congo red and calcofluor binding assays

Biofilm formation was quantified by crystal violet (CV) staining (OD600) as previously described [44]. For details see supplementary information. Bright field microscopy was used to assess aggregation in mid-log cultures grown in BM2 media with 2 mM or 0.02 mM Mg2+. Congo red binding assays were performed as previously described [43] with minor modifications (see supplementary information). Bacterial aggregates were stained with calcofluor (EPS), 1 µM SYTO9 (live cells) and 10 µM propidium iodide (dead cells and extracellular DNA), mounted on agarose beds. All microscopy was performed using a Leica DMIREB2 inverted microscope equipped with an ORCA-ER digital camera and Openlab software (Improvision) and analysed using Adobe Photoshop.

Scanning electron microscopy

P. aeruginosa peg-adhered biofilms were grown as previously described in high or limiting Mg2+ BM2 for 48 h, fixed as previously described [61], gold coated and visualized using a XL30 environmental scanning electron microscope.

Overexpression of RetS

The coding region of retS was PCR amplified, digested with XbaI and HindIII and ligated into the pSCrhaB2 vector under the control of a rhamnose-inducible promoter (pSCrhaB2RetS) [50]. Biofilms were cultured in BM2 media with 2 mM or 0.02 mM Mg2+ in the presence of 0–2% rhamnose.

PhoP protein purification and gel shift assays

PhoP protein was purified from Escherichia coli BL21 (GE Healthcare) containing His6-PhoP [40] cloned into pET28a as previously described [40]. A full length rsmZ and retS and truncated version of the retS promoter construct were PCR amplified (primers, Table S2) and digoxigenin (DIG)-labeled using the DIG Gel Shift Kit, 2nd Generation (Roche), according to manufacturer's instructions. Samples were separated by electrophoresis on 6% native polyacrylamide gels. transferred onto nylon membranes, probed with anti-DIG antibodies and chemiluminescence detected on the ChemiDoc XRS system (Bio-Rad).

Statistical Analysis

Statistical analysis was performed using GraphPad Prism 5 software. 2-way ANOVA was used to calculate significant differences between PAO1 and mutant strains.

Supporting Information

Figure S1.

Limiting magnesium-induced biofilm formation is dependent on EPS production but independent of pili or flagella production. Attachment of PAO1 and relevant mutants to polystyrene pegs was assessed by crystal violet staining and OD600 measurement in BM2 2 mM Mg2+ or 0.02 mM Mg2+ at 24 h. Bars represent the average values obtained from eight pegs and the error bars represent the standard deviation. Significant differences were observed between strains grown in BM2 2 mM Mg2+ and BM2 0.02 mM Mg2+ (a, p<0.05, ANOVA) and between PAO1 and mutant strains grown in BM2 0.02 mM Mg2+ (b, p<0.05, ANOVA). PAO1w, (Wozniak laboratory strain, Ohio State University) is the parent strain of the pslH and pel/pslH double mutant.


Figure S2.

EPS mutants grown in BM2 0.02 mM Mg2+ failed to aggregate or stain with calcofluor. Bacteria were grown in BM2 0.02 mM Mg2+ supplemented with 200 µg/ml calcofluor (blue, EPS stain). At 24 h cells were removed, stained with 1 µM syto9 (green, live cells) and visualized on agarose beds by fluorescence microscopy. Merged blue/green fluorescence images are representative of three independent experiments.


Figure S3.

PhoP regulates EPS production in P. aeruginosa. (A) Quantification of EPS production using congo red binding (corrected A490) in PAO1 and phoP::xylE grown in BM2 0.02 mM Mg2+ at 24 h. (B) Transmission electron microscopy of PAO1 and phoP::xylE. Bacteria were grown at 37°C overnight on BM2 0.02 mM Mg2+ 0.5% agar plates. Cells were prepared and stained as described by Hyland et al. 2006 [61] and examined using a Hitachi S-7000 transmission electron microscope.


Table S1.

List of media used to assess retS expression.


Table S2.

Primers used in this study. XhoI restriction sites are bolded; BamHI restriction site are underlined; overlap regions for SOE PCR are bolded and italicized; the modified nucleotides of the PhoP box in the retS promoter are capitalized.


Methods S1.

Additional information on strains and methods.



The authors would like to thank R. DeVinney for the use of her fluorescence microscope, M. Schoel for help with electron microscopy, C.D. Sibley, M.G. Surette and D. Storey (University of Calgary) and R.E. Hancock (University of British Columbia) for providing strains and D.J. Wozniak (Ohio State University) for supplying the pslH mutant (WFPA818) and the pel/psl double mutant prior to publication. The authors acknowledge E.P. O'Grady, J.B. McPhee and W.J. Gooderham for helpful comments and critical reading of the manuscript.

Author Contributions

Conceived and designed the experiments: HM SL. Performed the experiments: HM. Analyzed the data: HM. Wrote the paper: HM SL.


  1. 1. Branda SS, Vik S, Friedman L, Kolter R (2005) Biofilms: The matrix revisited. Trends Microbiol 13(1): 20–26.
  2. 2. Hall-Stoodley L, Stoodley P (2009) Evolving concepts in biofilm infections. Cell Microbiol 11(7): 1034–1043.
  3. 3. Lopez D, Vlamakis H, Kolter R (2010) Biofilms. Cold Spring Harb Perspect Biol 2(7): a000398.
  4. 4. Mah TF, O'Toole GA (2001) Mechanisms of biofilm resistance to antimicrobial agents. Trends Microbiol 9(1): 34–39.
  5. 5. Matz C, Kjelleberg S (2005) Off the hook–how bacteria survive protozoan grazing. Trends Microbiol 13(7): 302–307.
  6. 6. Anderson GG, O'Toole GA (2008) Innate and induced resistance mechanisms of bacterial biofilms. Curr Top Microbiol Immunol 322: 85–105.
  7. 7. Hardalo C, Edberg SC (1997) Pseudomonas aeruginosa: Assessment of risk from drinking water. Crit Rev Microbiol 23(1): 47–75.
  8. 8. Bodey GP, Bolivar R, Fainstein V, Jadeja L (1983) Infections caused by Pseudomonas aeruginosa. Rev Infect Dis 5(2): 279–313.
  9. 9. Ramsey BW, Pepe MS, Quan JM, Otto KL, Montgomery AB, et al. (1999) Intermittent administration of inhaled tobramycin in patients with cystic fibrosis. cystic fibrosis inhaled tobramycin study group. N Engl J Med 340(1): 23–30.
  10. 10. Lamarche MG, Dozois CM, Daigle F, Caza M, Curtiss R 3rd, et al. (2005) Inactivation of the pst system reduces the virulence of an avian pathogenic Escherichia coli O78 strain. Infect Immun 73(7): 4138–4145.
  11. 11. Davies DG, Parsek MR, Pearson JP, Iglewski BH, Costerton JW, et al. (1998) The involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280(5361): 295–298.
  12. 12. Costerton JW, Stewart PS, Greenberg EP (1999) Bacterial biofilms: A common cause of persistent infections. Science 284(5418): 1318–1322.
  13. 13. Singh PK, Schaefer AL, Parsek MR, Moninger TO, Welsh MJ, et al. (2000) Quorum-sensing signals indicate that cystic fibrosis lungs are infected with bacterial biofilms. Nature 407(6805): 762–764.
  14. 14. Costerton JW (2001) Cystic fibrosis pathogenesis and the role of biofilms in persistent infection. Trends Microbiol 9(2): 50–52.
  15. 15. Bjarnsholt T, Jensen PO, Fiandaca MJ, Pedersen J, Hansen CR, et al. (2009) Pseudomonas aeruginosa biofilms in the respiratory tract of cystic fibrosis patients. Pediatr Pulmonol 44(6): 547–558.
  16. 16. Klebensberger J, Lautenschlager K, Bressler D, Wingender J, Philipp B (2007) Detergent-induced cell aggregation in subpopulations of Pseudomonas aeruginosa as a preadaptive survival strategy. Environ Microbiol 9(9): 2247–2259.
  17. 17. Stock AM, Robinson VL, Goudreau PN (2000) Two-component signal transduction. Annu Rev Biochem 69: 183–215.
  18. 18. Parkins MD, Ceri H, Storey DG (2001) Pseudomonas aeruginosa GacA, a factor in multihost virulence, is also essential for biofilm formation. Mol Microbiol 40(5): 1215–1226.
  19. 19. Goodman AL, Kulasekara B, Rietsch A, Boyd D, Smith RS, et al. (2004) A signaling network reciprocally regulates genes associated with acute infection and chronic persistence in Pseudomonas aeruginosa. Dev Cell 7(5): 745–754.
  20. 20. Ventre I, Goodman AL, Vallet-Gely I, Vasseur P, Soscia C, et al. (2006) Multiple sensors control reciprocal expression of Pseudomonas aeruginosa regulatory RNA and virulence genes. Proc Natl Acad Sci U S A 103(1): 171–176.
  21. 21. Kuchma SL, Connolly JP, O'Toole GA (2005) A three-component regulatory system regulates biofilm maturation and type III secretion in Pseudomonas aeruginosa. J Bacteriol 187(4): 1441–1454.
  22. 22. Ramsey MM, Whiteley M (2004) Pseudomonas aeruginosa attachment and biofilm development in dynamic environments. Mol Microbiol 53(4): 1075–1087.
  23. 23. Gooderham WJ, Hancock RE (2009) Regulation of virulence and antibiotic resistance by two-component regulatory systems in Pseudomonas aeruginosa. FEMS Microbiol Rev 33(2): 279–294.
  24. 24. Petrova OE, Sauer K (2009) A novel signaling network essential for regulating Pseudomonas aeruginosa biofilm development. PLoS Pathog 5(11): e1000668.
  25. 25. Brencic A, Lory S (2009) Determination of the regulon and identification of novel mRNA targets of Pseudomonas aeruginosa RsmA. Mol Microbiol 72(3): 612–632.
  26. 26. Irie Y, Starkey M, Edwards AN, Wozniak DJ, Romeo T, et al. (2010) Pseudomonas aeruginosa biofilm matrix polysaccharide psl is regulated transcriptionally by RpoS and post-transcriptionally by RsmA. Mol Microbiol.
  27. 27. Goodman AL, Merighi M, Hyodo M, Ventre I, Filloux A, et al. (2009) Direct interaction between sensor kinase proteins mediates acute and chronic disease phenotypes in a bacterial pathogen. Genes Dev 23(2): 249–259.
  28. 28. Workentine ML, Harrison JJ, Weljie AM, Tran VA, Stenroos PU, et al. (2010) Phenotypic and metabolic profiling of colony morphology variants evolved from Pseudomonas fluorescens biofilms. Environ Microbiol.
  29. 29. Kay E, Humair B, Denervaud V, Riedel K, Spahr S, et al. (2006) Two GacA-dependent small RNAs modulate the quorum-sensing response in Pseudomonas aeruginosa. J Bacteriol 188(16): 6026–6033.
  30. 30. Brencic A, McFarland KA, McManus HR, Castang S, Mogno I, et al. (2009) The GacS/GacA signal transduction system of Pseudomonas aeruginosa acts exclusively through its control over the transcription of the RsmY and RsmZ regulatory small RNAs. Mol Microbiol 73(3): 434–445.
  31. 31. Mulcahy H, Charron-Mazenod L, Lewenza S (2008) Extracellular DNA chelates cations and induces antibiotic resistance in Pseudomonas aeruginosa biofilms. PLoS Pathog 4(11): e1000213.
  32. 32. Hoskins LC (1978) Host and microbial DNA in the gut lumen. J Infect Dis 137(5): 694–698.
  33. 33. Ranasinha C, Assoufi B, Shak S, Christiansen D, Fuchs H, et al. (1993) Efficacy and safety of short-term administration of aerosolised recombinant human DNase I in adults with stable stage cystic fibrosis. Lancet 342(8865): 199–202.
  34. 34. Lorenz MG, Wackernagel W (1994) Bacterial gene transfer by natural genetic transformation in the environment. Microbiol Rev 58(3): 563–602.
  35. 35. Brandt T, Breitenstein S, von der Hardt H, Tummler B (1995) DNA concentration and length in sputum of patients with cystic fibrosis during inhalation with recombinant human DNase. Thorax 50(8): 880–882.
  36. 36. Ulmer JS, Herzka A, Toy KJ, Baker DL, Dodge AH, et al. (1996) Engineering actin-resistant human DNase I for treatment of cystic fibrosis. Proc Natl Acad Sci U S A 93(16): 8225–8229.
  37. 37. Whitchurch CB, Tolker-Nielsen T, Ragas PC, Mattick JS (2002) Extracellular DNA required for bacterial biofilm formation. Science 295(5559): 1487.
  38. 38. Nicas TI, Hancock RE (1980) Outer membrane protein H1 of Pseudomonas aeruginosa: Involvement in adaptive and mutational resistance to ethylenediaminetetraacetate, polymyxin B, and gentamicin. J Bacteriol 143(2): 872–878.
  39. 39. McPhee JB, Lewenza S, Hancock RE (2003) Cationic antimicrobial peptides activate a two-component regulatory system, PmrA-PmrB, that regulates resistance to polymyxin B and cationic antimicrobial peptides in Pseudomonas aeruginosa. Mol Microbiol 50(1): 205–217.
  40. 40. McPhee JB, Bains M, Winsor G, Lewenza S, Kwasnicka A, et al. (2006) Contribution of the PhoP-PhoQ and PmrA-PmrB two-component regulatory systems to Mg2+-induced gene regulation in Pseudomonas aeruginosa. J Bacteriol 188(11): 3995–4006.
  41. 41. Friedman L, Kolter R (2004) Two genetic loci produce distinct carbohydrate-rich structural components of the Pseudomonas aeruginosa biofilm matrix. J Bacteriol 186(14): 4457–4465.
  42. 42. Ryder C, Byrd M, Wozniak DJ (2007) Role of polysaccharides in Pseudomonas aeruginosa biofilm development. Curr Opin Microbiol 10(6): 644–648.
  43. 43. Ma L, Jackson KD, Landry RM, Parsek MR, Wozniak DJ (2006) Analysis of Pseudomonas aeruginosa conditional psl variants reveals roles for the psl polysaccharide in adhesion and maintaining biofilm structure postattachment. J Bacteriol 188(23): 8213–8221.
  44. 44. O'Toole GA, Kolter R (1998) Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol Microbiol 30(2): 295–304.
  45. 45. Shih P, Huang C (2002) Effects of quorum-sensing deficiency on Pseudomonas aeruginosa biofilm formation and antibiotic resistance. J Antimicrob Chemother 49(2): 309–314.
  46. 46. Leigh JA, Signer ER, Walker GC (1985) Exopolysaccharide-deficient mutants of rhizobium meliloti that form ineffective nodules. Proc Natl Acad Sci U S A 82(18): 6231–6235.
  47. 47. Zogaj X, Nimtz M, Rohde M, Bokranz W, Romling U (2001) The multicellular morphotypes of Salmonella typhimurium and Escherichia coli produce cellulose as the second component of the extracellular matrix. Mol Microbiol 39(6): 1452–1463.
  48. 48. Solano C, Garcia B, Valle J, Berasain C, Ghigo JM, et al. (2002) Genetic analysis of Salmonella enteritidis biofilm formation: Critical role of cellulose. Mol Microbiol 43(3): 793–808.
  49. 49. Ledeboer NA, Jones BD (2005) Exopolysaccharide sugars contribute to biofilm formation by Salmonella enterica serovar typhimurium on HEp-2 cells and chicken intestinal epithelium. J Bacteriol 187(9): 3214–3226.
  50. 50. Cardona ST, Valvano MA (2005) An expression vector containing a rhamnose-inducible promoter provides tightly regulated gene expression in Burkholderia cenocepacia. Plasmid 54(3): 219–228.
  51. 51. Anantharaman V, Aravind L (2003) Application of comparative genomics in the identification and analysis of novel families of membrane-associated receptors in bacteria. BMC Genomics 4(1): 34.
  52. 52. Webb JS, Thompson LS, James S, Charlton T, Tolker-Nielsen T, et al. (2003) Cell death in Pseudomonas aeruginosa biofilm development. J Bacteriol 185(15): 4585–4592.
  53. 53. Lesley JA, Waldburger CD (2001) Comparison of the Pseudomonas aeruginosa and escherichia coli PhoQ sensor domains: Evidence for distinct mechanisms of signal detection. J Biol Chem 276(33): 30827–30833.
  54. 54. Prost LR, Daley ME, Bader MW, Klevit RE, Miller SI (2008) The PhoQ histidine kinases of Salmonella and Pseudomonas spp. are structurally and functionally different: Evidence that pH and antimicrobial peptide sensing contribute to mammalian pathogenesis. Mol Microbiol 69(2): 503–519.
  55. 55. Bordi C, Lamy MC, Ventre I, Termine E, Hachani A, et al. (2010) Regulatory RNAs and the HptB/RetS signalling pathways fine-tune Pseudomonas aeruginosa pathogenesis. Mol Microbiol 76(6): 1427–1443.
  56. 56. Petrova OE, Sauer K (2010) The novel two-component regulatory system BfiSR regulates biofilm development by controlling the small RNA rsmZ through CafA. J Bacteriol 192(20): 5275–5288.
  57. 57. Nielsen KM, Johnsen PJ, Bensasson D, Daffonchio D (2007) Release and persistence of extracellular DNA in the environment. Environ Biosafety Res 6(1–2): 37–53.
  58. 58. Banin E, Brady KM, Greenberg EP (2006) Chelator-induced dispersal and killing of Pseudomonas aeruginosa cells in a biofilm. Appl Environ Microbiol 72(3): 2064–2069.
  59. 59. Mulcahy H, Charron-Mazenod L, Lewenza S (2010) Pseudomonas aeruginosa produces an extracellular deoxyribonuclease that is required for utilization of DNA as a nutrient source. Environ Microbiol 12(6): 1621–1629.
  60. 60. Eisen MB, Spellman PT, Brown PO, Botstein D (1998) Cluster analysis and display of genome-wide expression patterns. Proc Natl Acad Sci U S A 95(25): 14863–14868.
  61. 61. Hyland RM, Griener TP, Mulvey GL, Kitov PI, Srivastava OP, et al. (2006) Basis for N-acetyllactosamine-mediated inhibition of enteropathogenic Escherichia coli localized adherence. J Med Microbiol 55(Pt 6): 669–675.
  62. 62. Lewenza S, Mhlanga MM, Pugsley AP (2008) Novel inner membrane retention signals in Pseudomonas aeruginosa lipoproteins. J Bacteriol 190(18): 6119–6125.
  63. 63. Macfarlane EL, Kwasnicka A, Ochs MM, Hancock RE (1999) PhoP-PhoQ homologues in Pseudomonas aeruginosa regulate expression of the outer-membrane protein OprH and polymyxin B resistance. Mol Microbiol 34(2): 305–316.
  64. 64. Lewenza S, Falsafi RK, Winsor G, Gooderham WJ, McPhee JB, et al. (2005) Construction of a mini-Tn5-luxCDABE mutant library in Pseudomonas aeruginosa PAO1: A tool for identifying differentially regulated genes. Genome Res 15(4): 583–589.
  65. 65. Pessi G, Williams F, Hindle Z, Heurlier K, Holden MT, et al. (2001) The global posttranscriptional regulator RsmA modulates production of virulence determinants and N-acylhomoserine lactones in Pseudomonas aeruginosa. J Bacteriol 183(22): 6676–6683.
  66. 66. Sibley CD, Duan K, Fischer C, Parkins MD, Storey DG, et al. (2008) Discerning the complexity of community interactions using a Drosophila model of polymicrobial infections. PLoS Pathog 4(10): e1000184.
  67. 67. Duan K, Dammel C, Stein J, Rabin H, Surette MG (2003) Modulation of Pseudomonas aeruginosa gene expression by host microflora through interspecies communication. Mol Microbiol 50(5): 1477–1491.