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Department of Microbiology and Immunology, Room 505 Vail Building, Dartmouth Medical School, Hanover, NH 03755, USA
Correspondence
George A. O'Toole
georgeo{at}Dartmouth.edu
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ABSTRACT |
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A table of primers is available as supplementary data with the online version of this paper.
Present address: Center for Microbial Ecology, Room 540 Plant and Soil Science Building, Michigan State University, East Lansing, MI 48824, USA.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). To begin the formation of a biofilm, bacteria need to interact with the substratum (Korber et al., 1994; O'Toole & Kolter, 1998a). Each individual bacterium first undergoes reversible attachment, which involves contact of the pole of the cell with the surface – this interaction is relatively weak and can be easily disrupted. Eventually the cell attaches by its long axis, so-called irreversible attachment, in which the bacterium is very firmly attached to the surface, resulting in the formation of a monolayer of cells (Fletcher, 1996; Hinsa et al., 2003; Jensen et al., 1992; Lawrence et al., 1987; Marshall, 1979; van Loosdrecht et al., 1990; Zobell, 1943). Subsequent steps result in the formation of microcolonies leading to the development of a ‘mature’ biofilm (Davey & O'Toole, 2000; Reisner et al., 2005; Webb et al., 2003).
Previously we identified a number of transposon-generated mutations which rendered Pseudomonas fluorescens WCS365 unable to form a biofilm in the 96-well static assay system (O'Toole & Kolter, 1998b). Eight of the transposons had inserted into four adjacent genes (Fig. 1a) that we designated lapA, lapE, lapB and lapC (Hinsa et al., 2003). All the lap mutant strains were unable to make the transition from reversible attachment to the irreversible attachment stage of biofilm maturation. Based on sequence similarity, we predicted that LapE, LapB and LapC form an ATP-binding cassette (ABC) transporter and that LapA is a large adhesion protein. We demonstrated that the LapA protein was localized to the supernatant and cell-associated fractions of the wild-type bacterium and requires the LapEBC functions for this localization. Based on these data, we proposed that the LapEBC transporter is necessary to transport LapA to the cell-associated fraction and that LapA is an adhesin necessary for P. fluorescens to irreversibly attach to a surface (Hinsa et al., 2003).
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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) supplemented with 1 mM MgSO4 and either 0·2 % glucose +0·5 % Casamino acids (CAA) or 0·4 % sodium citrate. For flow cells, M63 medium with 1 mM MgSO4 and 1 mM citrate or 1 mM glucose was used. Escherichia coli was grown in LB at 37 °C. Antibiotics were added at the following concentrations: (i) E. coli: tetracycline (Tc), 15 µg ml–1; gentamicin (Gm), 10 µg ml–1; (ii) P. fluorescens: Tc, 45 µg ml–1; nalidixic acid (Nal), 20 µg ml–1; Gm, 35 µg ml–1. Arabinose at a final concentration of 0·2 % was added to induce gene expression from the pMQ72 vectors. To test for cellulose production, 0·02 % calcofluor was added to LB plates supplemented with 5 µM FeCl3, as described by Leigh et al. (1985). Congo red (40 µg ml–1) and Coomassie blue (20 µl ml–1) were added to LB agar plates to assess polysaccharide production (Friedman & Kolter, 2004). Alternatively, Congo red (40 µg ml–1) and Coomassie blue (20 µl ml–1) were added to M63 minimal medium plus glucose or citrate and supplemented with agar (at 0·8 %, 1 % and 1·5 % agar). After inoculation, these plates were incubated at either 30 °C or room temperature for up to several days. Sudan black staining for EPSII (galactoglucan) was performed according to Liu et al. (1998). Swimming was tested on LB solidified with 0·3 % agar, or M63 minimal medium plus citrate or glucose, solidified with 0·3 % agar. Swimming assay plates were incubated at either 30 °C or room temperature.
Molecular techniques.
Plasmids were constructed using standard molecular methods (Ausubel et al., 1990) or in Saccharomyces cerevisiae by in vivo recombination (Longtine et al., 1998). Plasmid constructs and vectors were transformed into chemically competent E. coli Top10 cells (Invitrogen) and then transferred into P. fluorescens by electroporation.
The sequence of the DNA flanking the transposon insertion was determined by cloning the lapD19 transposon insertion and determining the DNA sequence flanking the element, as described by O'Toole & Kolter (1998b). Further sequence surrounding the lapD19 insertion was obtained by sequence walking with primers designed from the vector oriented towards the transposon insertion (primers P2 and P3; see Table S1, available as supplementary data with the online version of this paper). This additional P. fluorescens sequence was used to design PCR primers (P268 and P269 within the lapD ORF and P265 within the lapA gene) to confirm the gene order inferred from sequence analysis. The resulting PCR product, obtained from primers P265-P269, was sequenced to verify the gene orientation. The lapD sequence was compared to two sequenced strains, P. fluorescens Pf0-1 (http://genome.jgi-psf.org/finished_microbes/psefl/psefl.home.html) and P. putida KT2440 (http://www.tigr.org/tigr-scripts/CMR2/GenomePage3.spl?database=gpp). PSORT analysis was carried out on the available internet site (http://www.psort.org/).
Construction of the lapD knockout mutant.
The primer pair P781-P782 was used to amplify an 
900 bp segment of chromosomal DNA from 630 bp upstream of the lapD gene to 270 bp downstream of the predicted translation start of the lapD gene. The primer pair P783-P784 was used to amplify an 670 bp segment of chromosomal DNA from 520 bp upstream of the lapD gene stop codon to 150 bp downstream of the stop codon. Primer P782 contained sequence that overlapped with the P783 primer, and vice versa for P783. The P781-P782 PCR product (1 µl) and the P783-P784 PCR product (1 µl) were added to another PCR reaction to serve as the template. Primers P781 and P784 were used to amplify the 1530 bp pair product by sewing PCR. The resulting product generates a deletion of 1670 nt internal to lapD. The PCR product was cloned into pGEM-T vector according to the manufacturer's instructions (Promega). Both the P781 and P784 primers contained an EcoRI site, which was not found in the knockout fragment. The knockout fragment was digested from the pGEM-T vector, using EcoRI, and ligated into pEX18-Tc, which had previously been digested by EcoRI and treated with bacterial alkaline phosphatase. The resulting plasmid, pEX18-DKO, was used to build a deletion of the lapD gene as previously described (Hoang et al., 1998) with the following modifications: the conjugation was performed at room temperature and the selection was performed at 30 °C. The presence of the deletion of the lapD gene was confirmed by PCR using the DKOrev and P265 primer pair.
Construction of the lapD-carrying plasmid.
The lapD gene was amplified with primers P744 and P745, which flank the gene and also contain 40 bp tails homologous to the MCS of plasmid pMQ72 (R. Shanks & G. O'Toole, unpublished data). This amplicon was recombined into linearized pMQ72 via yeast homologous recombination in S. cerevisiae (Longtine et al., 1998). In the pMQ72-based vectors the lapD gene is under the control of the arabinose-inducible promoter pBAD and induced with 0·2 % arabinose.
Biofilm assays.
Biofilm formation was measured using the microtitre dish assay system, performed as described previously (O'Toole & Kolter, 1998a, b; O'Toole et al., 1999), using minimal M63 medium supplemented with 0·2 % glucose and 0·5 % CAA or 0·4 % citrate as the growth substrate. We have reported previously that using either glucose and CAA or citrate as the growth substrate promotes biofilm formation that is dependent upon LapA (Hinsa et al., 2003). However, growth on citrate does promote a more robust biofilm than growth on glucose + CAA (Hinsa et al., 2003). To quantify biofilm formation in the 96-well format the crystal violet stain was solubilized in 150 µl 30 % glacial acetic acid and the absorbance determined at 550 nm; the absorbance from at least five wells was averaged for each assay and the standard deviation is represented by error bars.
Measurement of reversible and irreversible attached cells was performed as reported by Hinsa et al. (2003). Briefly, the bacterial strains were grown overnight in M63 supplemented with either glucose+CAA or 0·4 % citrate and subcultured into this same minimal medium, grown to mid-exponential phase, diluted 1 : 5 in the same medium, then 500 µl of this dilution was placed in a 24-well microtitre plate for microscopy. Movies were collected immediately at one frame per second for 60 s for analysis of reversible and irreversible attachment. Bacteria that were attached by a single pole of the cell and showed movement during the course of the 60 s movie were defined as ‘reversibly attached’ to the surface, as were bacteria that attached to the surface or detached from the surface during the course of the movie. Cells firmly attached to the surface by the long axis of the cell and which did not move during the 60 s interval in which the movie images were captured were defined as ‘irreversibly attached’. Three movies of 60 s each were captured for each strain from three independent wells.
For the flow-cell studies, the once-flow-through continuous culture flow-cell system was assembled as described by Christensen et al. (1999) and modified M63, as described above, was utilized as the growth medium.
Imaging.
Epifluorescent and phase-contrast microscopy were performed with a model DM IRBE microscope (Leica Microsystems) equipped with an Orca model C4742-5 CCD camera (Hamamatsu). Images were acquired and processed on a Macintosh G4 loaded with OpenLab software (Improvision).
Protein localization and Western analysis.
Inner- and outer-membrane protein samples were prepared according to Nunn & Lory (1993), with modifications. Twenty millilitre cultures grown in minimal M63 medium supplemented with citrate and MgSO4 were grown shaking at 30 °C for 16 h. The cultures were centrifuged for 10 min at 6000 g and the bacterial pellet was resuspended in 1·5 ml 200 mM Tris/HCl buffer (pH 7·5) and 20 mM EDTA. A Thermo Spectronic French press mini-cell was used to lyse the cells by processing twice at 20 000 p.s.i (1·38 MPa). Next, the samples were centrifuged at 12 200 g to pellet unbroken cells. The supernatant was removed and centrifuged at 100 000 g for 60 min at 4 °C. The pellet was resuspended in 500 µl Tris/HCl buffer (20 mM, pH 7·6) and then mixed with 500 µl 0·2 % Sarkosyl in 20 mM Tris/HCl (pH 7·6). The samples were placed on a shaker at room temperature for 20 min. The samples were then centrifuged at 100 000 g for 60 min at 4 °C. The supernatant fraction containing the inner membrane was removed and the pellet (outer membrane) was resuspended in 1 ml 20 mM Tris/HCl (pH 7·6). The combined inner- and outer-membrane fraction was prepared as described above with the modification that after centrifugation at 100 000 g for 60 min at 4 °C, the pellet was resuspended in 300 µl PBS and regarded as the total membrane fraction.
Cell-associated and cytoplasmic fractions were prepared as previously described (Hinsa et al., 2003). Briefly, the cell-associated fraction was generated by pelleting a 20 ml culture of bacterial cells, then resuspending the pellet in 400 µl PBS. The cell pellet was vortexed for 5 s to dislodge proteins weakly associated with the bacterial outer membrane, the bacteria were recentrifuged and the LapA-containing supernatant fraction was assayed by Western blotting. LapA was not detected in the inner- or outer-membrane fraction (Hinsa et al., 2003).
SDS loading buffer was mixed with each sample, followed by heat denaturation at 95 °C for 10 min. The protein samples for the SecY protein analysis were placed at 37 °C for 10 min instead of 95 °C (Duong & Wickner, 1999). The samples were resolved on a gradient polyacrylamide gel (4–15 %) at 25 mA. The protein was transferred to a nitrocellulose membrane in transblot buffer as described by Towbin et al. (1979). Western blots were developed using antibodies prepared to LapA, LapD, LapE and SecY. The LapA and LapE antibodies have been described previously (Hinsa et al., 2003). A LapD peptide (CSEKLRVESYQDNLTGLAN) was synthesized and used to generate antibodies against the LapD protein (Bio Synthesis Incorporated). A SecY peptide (CERGQRRIAVHYAKRQQGRKVFA), conserved in P. fluorescens and Pseudomonas aeruginosa, was synthesized and used to generate antibodies against the SecY protein (Bio Synthesis Incorporated). Western blots were developed with ECL Western detection reagents (Amersham).
Quantitative RT-PCR (qRT-PCR) analysis.
The wild-type and the 
lapD mutant strains were subcultured from overnight planktonic cultures grown in M63 minimal medium supplemented with MgSO4 and sodium citrate, subcultured and grown to late-exponential phase in this same medium. RNA was obtained using the Qiagen RNA kit as instructed with the following modification that lysozyme digestion was performed for 5 min. Following elution, a second DNase treatment was performed with 50 U DNase I (Roche) for 1 h at room temperature. The RNA was then purified with the Qiagen kit following the manufacturer's instructions. The RNA concentrations were determined and normalized before cDNA was generated. First-strand cDNA synthesis was carried out with 3 µg RNA using the First-Strand cDNA synthesis kit according to the manufacturer's instructions (Amersham Biosciences). For a positive normalization control, primers were designed to the constitutively expressed rplU gene (P780 and P779). The primer set (P771 and P772) was used to quantify lapA gene expression in the wild-type and lapD mutant strains. The SYBR Green PCR kit from Applied Biosystems was used for qRT-PCR reactions, according to the manufacturer's instructions, and the reactions were run in an ABI Prism 7700 Sequencing Detection System. Statistical analysis was performed using a Student's t-test in Excel.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). We identified the gene disrupted by the transposon by sequencing the region adjacent to the cloned transposon fragment. The DNA sequence obtained was then compared with the P. fluorescens Pf0-1 and P. putida KT2440 genome sequences. The transposon was found to map to an ORF we designated lapD; therefore the mutant allele was designated lapD19. The transposon had inserted just past the midpoint of the lapD ORF. The lapD gene is 88 % identical at the nucleotide level to ORF93 (contig 483) of P. fluorescens Pf0-1 and 71 % identical to ORF PP0165 of P. putida KT2440.
Based on the P. fluorescens Pf0-1 genome we predicted that the lapD gene would map close to the previously identified lap region (Fig. 1a
) (Hinsa et al., 2003). To determine the chromosomal location of lapD, sequence walking was performed from the cloned lapD19 : : Tn5 transposon and PCR was performed to verify the orientation of this gene relative to that of lapA (data not shown). Based on these data we determined that lapD is adjacent to and transcribed in the opposite direction from lapA (Fig. 1a). The organization of the lap genes in P. fluorescens WCS365 is identical to that found in the sequenced strain P. fluorescens Pf0-1.
The mass of LapD is predicted to be 71 kDa, and based on PSORT analysis, is an inner-membrane protein containing two transmembrane domains. Based on the sequence of the lapD gene we predict that this ORF would encode a protein containing HAMP, GGDEF and EAL domains. A HAMP domain is commonly found in the cytoplasmic portion of a membrane-bound protein and is hypothesized to play a role in sending a signal from the periplasmic N-terminus to the cytoplasmic C-terminus of a protein (Aravind & Ponting, 1999). LapD also has a predicted GGDEF domain that has 36 % identity and 52 % similarity to the consensus sequence across 152 amino acids. Some GGDEF-domain-containing proteins have variable motifs (Spiers et al., 2003), including LapD, which contains a shortened GGEF sequence instead of the typical GGDEF residues. This same unusual sequence is found in the predicted LapD proteins encoded by P. fluorescens Pf0-1, P. putida KT4220 and P. aeruginosa PAO1 (PA1433). LapD also has a predicted EAL domain, but although LapD has 37 % identity and 51 % similarity across 233 amino acids of the EAL domain sequence (Galperin et al., 2001), LapD has a KVL sequence instead of the hallmark EAL sequence. Proteins containing the GGDEF and EAL domains are predicted to possess diguanylate cyclase and phosphodiesterase activities, respectively, thus controlling the intracellular levels of the nucleotide cyclic di-GMP (Galperin et al., 2001; Simm et al., 2004).
The lapD gene is conditionally required for biofilm formation
To verify that lapD is important for biofilm formation we disrupted this gene in P. fluorescens WCS365, resulting in an unmarked knockout mutation (lapD). The disruption of the gene was confirmed by PCR analysis (data not shown). The lapD mutant, like the lapD19 transposon mutation, was defective for biofilm formation in the static 96-well biofilm plate assay, with an approximately 43 % reduction in crystal violet staining compared to the wild-type (Fig. 1b). We used the lapD mutant in all subsequent studies. There was no difference in growth rate between the wild-type and the lapD mutant in minimal medium supplemented with glucose+CAA or citrate (data not shown).
We carried out complementation analysis to confirm that the lapD gene was responsible for the biofilm-defective phenotype of the lapD strain. The lapD gene was amplified from P. fluorescens DNA and cloned into pMQ72 via homologous recombination in S. cerevisiae (Longtine et al., 1998), resulting in plasmid pMQ72-lapD. In the pMQ72-lapD construct the lapD gene is under the control of the arabinose-inducible promoter derived from the pBAD plasmid (Guzman et al., 1995). Both pMQ72 and pMQ72-lapD were introduced into the wild-type strain and lapD mutant and these strains were analysed for biofilm formation (Fig. 1c).
Biofilm formation by the wild-type strain is not altered when carrying either pMQ72 or pMQ72-lapD and grown in the presence of arabinose (first three bars, Fig. 1c). These data suggest that expression of the lapD gene from this plasmid does not lead to changes in biofilm formation by the wild-type strain. Biofilm formation by the lapD strain carrying the pMQ72 vector is not affected when growing under inducing conditions; however, biofilm formation by the lapD strain is restored to wild-type levels when the pMQ72-lapD construct is grown in the presence of arabinose (last three bars in Fig. 1c). Western blots show that the level of LapD in the lapD/pMQ72-lapD is slightly higher than the wild-type, and LapD production is induced in the presence of arabinose (Fig. 1d). These data confirm that the lapD gene plays a role in biofilm formation by P. fluorescens.
We showed previously that mutations in lapA rendered cells defective in the transition from reversible, polar attachment to irreversible attachment wherein cells are firmly adhered to the substratum by the long axis of the cell (Hinsa et al., 2003). We performed similar studies with the lapD mutant strain grown on glucose+CAA (Fig. 2a). The wild-type strain shows 90 % irreversibly attached bacteria versus the lapA mutant, wherein only 5 % of the bacteria are irreversibly attached. These findings correspond to the phenotype reported previously (Hinsa et al., 2003). The lapD mutant shows an intermediate phenotype, with 40 % of the cells irreversibly attached. The partial defect in irreversible attachment of the lapD mutant is consistent with the partial biofilm defect observed for this strain in the 96-well plate assay (Fig. 1b).
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). In glucose+CAA medium, the wild-type strain forms a significantly better biofilm compared to the lapD strain over the first 8 h, with approximately 45 % less crystal violet staining observed for the lapD wells. By 24 h, the lapD strain is able to form a biofilm comparable to the wild-type strain (Fig. 2b). These data suggest that the lapD mutant is delayed in biofilm formation in glucose+CAA medium.
Surprisingly, in citrate medium there is no significant difference between the wild-type and the mutant over the 24 h period (Fig. 2c). Biofilm formation by the lapD mutant is markedly different from that of the lapA, lapE, lapB and lapC mutants, which were unable to form a biofilm under any conditions tested, including when grown on either glucose+CAA or citrate medium (Hinsa et al., 2003). Therefore we propose that LapD is conditionally required for biofilm formation in the static biofilm system.
Analysis of biofilm formation in a flow cell
To analyse the development of a mature biofilm, we grew the wild-type and the lapD mutant in a flow-cell system using minimal medium supplemented with 1 mM glucose. Flow cells provide a constant influx of fresh nutrients, thus sustaining the continued development of the biofilm over many days. Biofilms grown in the flow cell form the characteristic architecture composed of large macrocolonies surrounded by fluid-filled channels after 2 days incubation. The wild-type bacteria established a biofilm consisting of microcolonies and macrocolonies by day 2, while the lapD mutant bacteria were only able to attach individually or in small microcolonies to the surface (data not shown).
We also examined biofilm formation at 1 and 2 days in a flow-cell system using minimal medium supplemented with 1 mM sodium citrate. As shown in Fig. 3, the wild-type bacteria establish microcolonies at day 1 and then form microcolonies and macrocolonies surrounded by channels at day 2. In contrast, for the lapD strain, only single cells were seen to attach and only a few small microcolonies formed at day 1. More microcolonies were apparent at day 2 but no larger macrocolonies were formed by the lapD strain. These results suggest that, while no defect was observed in the static system, under the more stringent flow conditions the lapD mutant is unable to form a mature biofilm in minimal citrate medium.
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70 kDa band in the inner membrane of the wild-type strain, which matches the predicted 71 kDa size for LapD (Fig. 4a). This band was absent in both the inner- and outer-membrane fractions of the lapD strain (Fig. 4a).
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) (Hinsa et al., 2003). Furthermore, the LapE protein fractionates to the outer membrane in the lapD mutant at wild-type levels, suggesting that the LapD protein is not required for the synthesis or the proper localization of the LapE protein.
As an additional control for the inner-membrane fractionation protocol, we generated a peptide antibody against SecY, a well-characterized inner-membrane protein (Akiyama & Ito, 1985). The 48 kDa SecY protein fractionated almost exclusively to the inner-membrane fraction (Fig. 4b). Taken together, these data strongly support the conclusion that the LapD protein localizes to the inner membrane of P. fluorescens.
A mutation in lapD decreases extracellular levels of LapA
We previously reported that the LapA protein is localized to the supernatant and the cell-associated fraction of P. fluorescens. As described in Methods, the cell-associated fraction represents proteins weakly associated with the cell surface. We hypothesized that the LapD protein might have an impact on the ability of the LapA protein to localize to the cell-associated fraction. To test this hypothesis, we collected the cell-associated fraction and the cytoplasmic fractions from wild-type and lapD mutant bacteria that had been grown overnight in minimal medium with citrate. There is very little LapA produced when bacteria are grown in glucose medium (data not shown); therefore, we used citrate-grown bacteria to quantify the extent of LapA secreted in the wild-type versus lapD mutant.
As seen in Fig. 5, there was less LapA protein in the cell-associated fraction of the lapD strain compared to the wild-type strain. Using densitometry to quantify this difference revealed 7·3-fold less LapA in the cell-associated fraction of the lapD mutant compared to the wild-type strain (P<0·009, n=8). As expected, the LapA-specific band was not detected in the lapA mutant. There was no observed difference in LapA level in the supernatant fractions between the wild-type and lapD mutant (data not shown); however, LapA was difficult to detect by Western blot in the supernatant fraction.
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lapD strain could be attributable to a defect in the secretion of the LapA protein or in the production of the LapA protein. There was no detectable difference in the amount of LapA in the cytoplasmic fraction between the wild-type and the lapD mutant (Fig. 5, right panel) as determined by densitometry (0·99-fold change, P>0·49, n=4).
To determine whether the LapD protein has an impact on the transcription of the lapA gene, qRT-PCR analysis was performed. RNA was collected from late-exponential-phase wild-type and lapD mutant cells grown in minimal medium plus citrate. Primers to the constitutively expressed rplU gene were used as a positive normalization control (Kuchma et al., 2005). An RNA-only control (lacking cDNA) was also used to ensure there was no DNA contamination in the RNA preparations – no PCR products were obtained from these control reactions (data not shown). Expression levels of lapA were quantified in picograms of input cDNA by using a standard curve method for absolute quantification as previously described by Kuchma et al. (2005).
There was no difference in the expression levels of the lapA transcript between the wild-type and lapD mutant bacteria. The relative expression of lapA in the wild-type is 0·0173+±0·0019 versus 0·0216+±0·0049 in the lapD mutant strain (P>0·02, n=12).
A mutation in lapD does not affect several factors known to influence biofilm formation
Proteins containing a GGDEF domain have been shown to regulate cellulose production in bacteria including Gluconacetobacter, Salmonella and Pseudomonas (Garcia et al., 2004; Romling, 2005; Ross et al., 1991; Spiers et al., 2003; Tal et al., 1998). Furthermore, cellulose synthesis has been shown to be important for adhesion to surfaces and biofilm formation by a variety of micro-organisms (Gal et al., 2003; Garcia et al., 2004; Matthysse & McMahan, 1998; Spiers et al., 2003). To test for a defect in cellulose production, wild-type and lapD strains were spotted on LB medium supplemented with 0·02 % calcofluor and 5 µM FeCl3 (Leigh et al., 1985). Iron was added to suppress production of fluorescent siderophores. There was no difference in calcofluor-dependent fluoresence between the wild-type and lapD mutant strain.
We also investigated if there were changes in polysaccharide production between the wild-type and lapD mutant. Polysaccharide production was monitored on LB medium and minimal medium plus citrate or glucose supplemented with Congo red and Coomassie blue using a range of agar concentrations (Friedman & Kolter, 2004). There was no difference in Congo red binding between the strains under the variety of conditions we tested. Finally, the ability of the lapD mutant to produce EPSII was monitored by Sudan black staining (Liu et al., 1998). In Rhizobium meliloti, mutants unable to produce EPSII or succinolglycan are stained by Sudan black, whereas the wild-type remains unstained (Liu et al., 1998). P. fluorescens and P. putida have been shown to produce galactoglucans which have been implicated in biofilm formation (Read & Costerton, 1987). There was no difference in Sudan black staining between the wild-type and the lapD mutant.
We also tested swimming motility, which has been shown to be important for bacterial colonization of surfaces (de Weger et al., 1987; DeFlaun et al., 1990; O'Toole & Kolter, 1998b; Sauer & Camper, 2001). Swimming was monitored on LB medium solidified with 0·3 % agar or M63 medium supplemented with citrate or glucose (plus 0·3 % agar) – no differences in swimming motility were observed between wild-type and the lapD mutant under any conditions tested.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). In the studies presented here, we identified the LapD protein, which has a possible role in controlling LapA protein secretion and thus biofilm formation.
We determined that the sad-19 mutation, which confers a biofilm defect, mapped to an ORF that we have designated lapD. Using PCR and sequence analysis, we determined that lapD is located adjacent to lapA on the chromosome of P. fluorescens WCS365. The lapD mutant is distinct from the other lap mutants because it can form a biofilm in minimal medium supplemented with citrate in the static 96-well assay at 8 h (O'Toole & Kolter, 1998b). The lapD strain forms a less robust biofilm at 8 h in minimal medium supplemented with glucose+CAA compared to the wild-type strain; however, it is able to form a biofilm comparable to the wild-type by 24 h, suggesting that the lapD strain is only delayed (and not completely blocked) for biofilm formation in glucose medium. Based on these data we propose that in the static 96-well biofilm assay the LapD protein is not absolutely required for biofilm formation. The ability of the lapD strain to form a biofilm in the more stringent flow-cell system was tested and under these conditions it was only able to form microcolonies after 2 days incubation and could not progress to form a mature biofilm comparable to the wild-type strain. These results suggest that the lapD mutation causes a conditional biofilm phenotype on citrate medium that is only apparent in the more rigorous flow-cell assay. Recent work by Gjermansen et al. (2005) also found that a mutation in the lapD homologue in P. putida (PP0165) is unable to form a biofilm comparable to the wild-type strain in flow-cell experiments. The differential requirement for LapD in glucose+CAA versus citrate may reflect the fact that more LapA is produced when bacteria are grown in citrate than when grown on glucose+CAA (not shown). LapD may only be required in the static assay to facilitate the secretion of LapA when lower levels of LapA are produced.
We hypothesized that the LapD protein might affect the synthesis or function of the previously identified ABC transporter or the large adhesin LapA. The LapE protein is produced at wild-type levels and able to properly localize in a lapD strain, suggesting the LapD protein is not required for production or assembly of this component of the ABC transporter. Further localization studies determined that less LapA protein is found in the cell-associated fraction in the lapD strain compared to the wild-type. However, an equal amount of LapA protein is found in the cytoplasm of both strains (Fig. 5). One caveat of interpreting the data presented in Fig. 5, based on the smear below the LapA band in the Western blot, is that LapA in the cytoplasm appears to be proteolysed. Therefore, although our quantification of the Western blots indicates no change in LapA level in the cytoplasm, it is difficult to measure small changes in LapA cytoplasmic levels. The qRT-PCR analysis also showed similar lapA transcript levels for the wild-type and lapD strain. Finally, we showed that there was no detectable effect of the lapD mutation on several other biofilm-associated phenotypes, including motility and matrix production.
Based on our current data we propose that the LapD protein modulates the secretion of the LapA protein and thus biofilm formation. A low level of the LapA protein is able to reach the cell-associated fraction in the absence of the LapD protein and we predict that in the static biofilm assay system this is enough of the LapA adhesin for the bacteria to attach to the surface, but more LapA protein is required to initiate biofilm formation in the more stringent flow-cell system.
The precise mechanism by which the LapD protein effects secretion of the LapA protein is not known. Perhaps the LapD protein, which localizes to the inner membrane, can affect the function of the LapEBC ABC transporter and thus control secretion of the LapA protein. However, we did not see an accumulation of LapA in the cytoplasm as might be expected if a lapD mutant has decreased LapA secretion. However, as shown in Fig. 5, LapA in the cytoplasm seems to be susceptible to proteolysis as indicated by the smear below the main LapA band on the Western blot. This technical difficulty makes it difficult to accurately measure small changes in LapA levels in the cytoplasm. Thus it is possible that LapD could also affect production of LapA, or alternatively, the turnover of LapA in the cytoplasm. The focus of ongoing studies is to continue to improve our understanding of the effects of LapD on LapA localization, production and stability.
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ACKNOWLEDGEMENTS |
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Received 18 November 2005;
revised 20 January 2006;
accepted 23 January 2006.
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