- Research article
- Open Access
The RNA processing enzyme polynucleotide phosphorylase negatively controls biofilm formation by repressing poly-N-acetylglucosamine (PNAG) production in Escherichia coli C
- Thomas Carzaniga†1,
- Davide Antoniani†1,
- Gianni Dehò1,
- Federica Briani1Email author and
- Paolo Landini1Email author
© Carzaniga et al.; licensee BioMed Central Ltd. 2012
- Received: 19 June 2012
- Accepted: 1 October 2012
- Published: 21 November 2012
Transition from planktonic cells to biofilm is mediated by production of adhesion factors, such as extracellular polysaccharides (EPS), and modulated by complex regulatory networks that, in addition to controlling production of adhesion factors, redirect bacterial cell metabolism to the biofilm mode.
Deletion of the pnp gene, encoding polynucleotide phosphorylase, an RNA processing enzyme and a component of the RNA degradosome, results in increased biofilm formation in Escherichia coli. This effect is particularly pronounced in the E. coli strain C-1a, in which deletion of the pnp gene leads to strong cell aggregation in liquid medium. Cell aggregation is dependent on the EPS poly-N-acetylglucosamine (PNAG), thus suggesting negative regulation of the PNAG biosynthetic operon pgaABCD by PNPase. Indeed, pgaABCD transcript levels are higher in the pnp mutant. Negative control of pgaABCD expression by PNPase takes place at mRNA stability level and involves the 5’-untranslated region of the pgaABCD transcript, which serves as a cis-element regulating pgaABCD transcript stability and translatability.
Our results demonstrate that PNPase is necessary to maintain bacterial cells in the planktonic mode through down-regulation of pgaABCD expression and PNAG production.
- RNA processing
- Cell adhesion
Most bacteria can switch between two different lifestyles: single cells (planktonic mode) and biofilms, i.e., sessile microbial communities. Planktonic and biofilm cells differ significantly in their physiology and morphology and in their global gene expression pattern [1–3]. Extensive production of extracellular polysaccharides (EPS) represents a defining feature of bacterial biofilms; EPS are the major constituent of the so-called “biofilm matrix”, which also includes cell surface-associated proteins and nucleic acids [4, 5]. In addition to constituting the material embedding biofilm cells and to being a main determinant for surface attachment, the EPS are responsible for cell resistance to environmental stresses such as desiccation  and to predation by bacteriophages . In several bacterial species, EPS are also required for swarming motility [8, 9].
Expression of genes involved in EPS biosynthesis is controlled by complex regulatory networks responding to a variety of environmental and physiological cues, including stress signals, nutrient availability, temperature, etc. [10–13]. Regulation of EPS production can take place at any level, i.e., transcription initiation, mRNA stability, and protein activity. For instance, the vps genes, involved in EPS biosynthesis in Vibrio cholerae, are regulated at the transcription level by the CytR protein, in response to intracellular pyrimidine concentrations . The RsmA protein negatively regulates EPS production in Pseudomonas aeruginosa by repressing translation of the psl transcript . Finally, cellulose production in Gluconacetobacter xylinum and in various enterobacteria requires enzymatic activation of the cellulose biosynthetic machinery by the signal molecule cyclic-di-GMP (c-di-GMP) [16, 17], a signal molecule which plays a pivotal role as a molecular switch to biofilm formation in Gram negative bacteria . The great variety of regulatory mechanisms presiding to EPS biosynthesis, and the role of c-di-GMP as signal molecule mainly devoted to its control, underline the critical importance of timely EPS production for bacterial cells.
Polynucleotide phosphorylase (PNPase) plays an important role in RNA processing and turnover, being implicated in RNA degradation and in polymerization of heteropolymeric tails at the 3’-end of mRNA [19, 20]. PNPase is an homotrimeric enzyme that, together with the endonuclease RNase E, the DEAD-box RNA helicase RhlB, and enolase, constitute the RNA degradosome, a multiprotein machine devoted to RNA degradation [21, 22]. Despite the crucial role played by PNPase in RNA processing, the pnp gene is not essential; however, pnp inactivation has pleiotropic effects, which include reduced proficiency in homologous recombination and repair [23, 24], inability to grow at low temperatures  and inhibition of lysogenization by bacteriophage P4 . Moreover, lack of PNPase affects stability of several small RNAs, thus impacting their ability to regulate their targets .
In this work, we show that deletion of the pnp gene results in strong cell aggregation and biofilm formation, due to overproduction of the EPS poly-N-acetylglucosamine. Increased biofilm formation was observed both in E. coli MG1655 and C-1a strains, being more pronounced in the latter. We demonstrate that PNPase negatively controls expression of the PNAG biosynthetic operon pgaABCD at post-transcriptional level, thus acting as a negative determinant for biofilm formation. Our observation that PNPase acts as an inhibitor of biofilm formation is consistent with previous findings highlighting the importance of regulation of EPS production and biofilm formation at mRNA stability level .
Bacteria and growth media
Bacterial strains and plasmids
Origin or reference
E. coli C, prototrophic
by P1 HTF AM72 transduction into C-1a
by P1 HTF AM72 transduction into C-5691
by P1 HTF AM70 transduction into C-1a
by P1 HTF AM70 transduction into C-5691
by P1 HTF AM56 transduction into C-1a
by P1 HTF AM56 transduction into C-5691
by P1 HTF AM105 transduction into C-1a
by P1 HTF AM105 transduction into C-5691
by P1 HTF JW1007 transduction into C-1a
by P1 HTF JW1007 transduction into C-5691
From C-1a by λ Red-mediated recombination; primers: FG2624 and FG2625
From C-1a by λ Red-mediated recombination; primers: FG2524 and FG2525
From C-5691 by λ Red-mediated recombination; primers: FG2524 and FG2525.
From C-1a by λ Red-mediated recombination; primers: FG2585 and FG2586.
From C-5691 by λ Red-mediated recombination; primers: FG2585 and FG2586.
by P1 HTF C-5940 transduction into C-5944
Δpnp-751 ΔcsrB::kan ΔcsrC::cat
by P1 HTF C-5940 transduction into C-5946
From C-1a by λ Red-mediated recombination; primers: PL674 and PL675.
From C-5691 by λ Red-mediated recombination; primers: PL674 and PL675.
From C-1a by λ Red-mediated recombination; primers: FG2755 and FG2756.
From C-5691 by λ Red-mediated recombination; primers: FG2755 and FG2756.
From MG1655 by λ Red-mediated recombination with a DNA fragment obtained by PCR of tet10 cassette of EB 1.3 with primers PL372 and PL373.
Plasmids and phage
pBAD24 derivative with a modified polylinker; carries an unique Nco I site overlapping the araBp transcription start
pBAD24 derivative; harbours an EcoRI-HindIII fragment of pEJ01 that carries the pnp gene
pBAD24 derivative; harbours an HindIII-XbaI fragment of pFCT6.9 that carries the rnb gene
pBAD24-Δ1 derivative; harbours the rnr gene (obtained by PCR on MG1655 DNA with FG2474-FG2475 oligonucleotides) between NcoI-HindIII sites
pJAMA8 derivative, harbours the -116 to +32 region relative to the pgaABCD transcription start site cloned into the SphI/XbaI sites
carries a His-tagged pnp allele
carries a His-tagged rnb allele
; received from Cecilia Arraiano
ori VColD; CamR
AmpR, ColE1; luxAB based promoter-probe vector.
pJAMA8 derivative, harbours the -116 to +234 region relative to the pgaABCD transcription start site cloned into the SphI/XbaI sites.
pJAMA8 derivative, harbours a translational fusion of pgaA promoter, regulatory region and first 5 codons of pgaA (-116 to +249 relative to transcription start site) with luxA ORF (Open Reading Frame).
pJAMA8 derivative, harbours ptac promoter of pGZ119HE cloned into the SphI/XbaI sites.
High transduction frequency phage P1 derivative
; received from Richard Calendar
Cell aggregation and adhesion assays
Cell aggregation was assessed as follows: overnight cultures grown in LD at 37°C on a rotatory device were diluted 50-fold in 50 ml of M9Glu/sup in a 250 ml flask. The cultures were then incubated at 37°C with shaking at 100 rpm. Cell adhesion to the flask walls was assessed in overnight cultures grown in M9Glu/sup medium at 37°C. Liquid cultures were removed and cell aggregates attached to the flask glass walls were stained with crystal violet for 5 minutes to allow for better visualization. Quantitative determination of surface attachment to polystyrene microtiter wells was carried out using crystal violet staining as previously described . Binding to Congo red (CR) was assessed in CR agar medium (1% casamino acid, 0.15% yeast extract, 0.005% MgSO4, 2% agar; after autoclaving, 0.004% Congo red and 0.002% Coomassie blue). Overnight cultures in microtiter wells were replica plated on CR agar plates, grown for 24 h at 30°C, and further incubated 24 h at 4°C for better detection of staining.
Gene expression determination
RNA extraction, Northern blot analysis and synthesis of radiolabelled riboprobes by in vitro transcription with T7 RNA polymerase were previously described [34, 35]. The DNA template for PGA riboprobe synthesis was amplified by PCR on C-1a genomic DNA with oligonucleotides FG2491/39 and FG2492/22. Autoradiographic images of Northern blots were obtained by phosphorimaging using ImageQuant software (Molecular Dynamics). Quantitative (real time) reverse transcriptase PCR (quantitative RT-PCR) was performed as described . Oligonucleotides PL101/21 and PL102/19 were used for 16S rRNA reverse transcription and PCR amplification. mRNA half-lives were estimated as described  by regression analysis of mRNA remaining (estimated by real time PCR) versus time after rifampicin addition. Luciferase assays were performed as in . Oligonucleotides utilized for Northern blot, real time PCR, and construction of reporter plasmids are listed in Additional file 1: Table S1.
PNAG production was determined as described . Bacteria were grown overnight in 3 ml of M9 Glu/sup medium at 37°C. Cells were collected by centrifugation and diluted in Tris-buffered saline [20 mM Tris–HCl, 150 mM NaCl (pH 7.4)] to an OD600 = 1.5. 1ml of suspension was centrifuged at 10,500 x g, resuspended in 300 μl of 0.5 M EDTA (pH 8.0), and incubated for 5 min at 100°C. Cells were removed by centrifugation at 10,500 x g for 6 min and 100 μl of the supernatant was incubated with 200 μg of proteinase K for 60 min at 60°C. Proteinase K was heat-inactivated at 80°C for 30 min. The solution was diluted 1:3 in Tris-buffered saline and 10 μl was spotted onto a nitrocellulose filter using a Dot-blot apparatus (Bio-Rad). The filter was saturated for about 2 hours in 0.1 M Tris–HCl (pH 7.5), 0.3 M NaCl, 0.1% Triton (Sigma Aldrich) and 5% milk and then incubated overnight at 4°C with a 1:1,000 dilution of purified PNAG antibodies (a kind gift from G.B. Pier ). PNAG antibodies were detected using a secondary anti-goat antibody (dilution 1:5,000) conjugated with horseradish peroxidase. Immunoreactive spots were revealed using ECL Western blotting reagent (Amersham Pharmacia Biotech).
When applicable, statistically significant differences among samples were determined using a t-test of analysis of variance (ANOVA) via a software run in MATLAB environment (Version 7.0, The MathWorks Inc.). Tukey’s honestly significant different test (HSD) was used for pairwise comparison to determine significance of the data. Statistically significant results were depicted by p-values <0.05.
Lack of PNPase induces cell aggregation in E. coli C
The aggregative phenotype of the C-5691 (Δpnp) strain was complemented by basal expression from a multicopy plasmid of the pnp gene under araBp promoter, indicating that low PNPase expression is sufficient to restore planktonic growth. Conversely, arabinose addition did not completely restore a wild type phenotype (Figure 1B, left panel), suggesting that PNPase overexpression may also cause aggregation. Ectopic expression of RNase II suppressed the aggregative phenotype of the pnp mutant (Figure 1B, right panel), thus suggesting that such a phenotype is controlled by the RNA degrading activity of PNPase. In contrast, however, RNase R overexpression did not compensate for lack of PNPase, indicating that different ribonucleases are not fully interchangeable in this process.
Inactivation of the pnp gene induces poly-N-acetylglucosamine (PNAG) production
The aggregative phenotype of the C-5691 (Δpnp) mutant, as determined by cell aggregation, surface adhesion, and Congo red binding experiments, was totally abolished by deletion of pgaC (Figure 2), which encodes the polysaccharide polymerase needed for biosynthesis of PNAG from UDP-N-acetylglucosamine . Deletion of pgaA, also part of the PNAG biosynthetic operon pgaABCD, produced identical effects as pgaC (data not shown). In contrast, no significant effects on either Congo red binding or cell aggregation and adhesion were detected in any Δpnp derivative unable to produce curli or colanic acid (Figure 2). Finally, deletion of the bcsA gene, which encodes cellulose synthase, led to a significant increase in cell adhesion to the flask glass walls (Figure 2A); this result is consistent with previous observations suggesting that, although cellulose can promote bacterial adhesion, it can also act as a negative determinant for cell aggregation, particularly in curli-producing E. coli strains [49, 50]. In the C-1a strain, carrying a wild type pnp allele, inactivation of genes involved in biosynthesis of curli, PNAG, cellulose and colanic acid did not result in any notable effects on cell aggregation (Additional file 2: Figure S1).
To establish whether induction of PNAG-dependent cell aggregation in the absence of PNPase is unique to E. coli C-1a or it is conserved in other E. coli strains, we performed adhesion assays comparing the standard laboratory strain MG1655 to its Δpnp derivative KG206. Similar to what observed for the E. coli C strains, deletion of the pnp gene in the MG1655 background resulted in a significant increase in adhesion to solid surfaces, which was totally abolished by pgaA deletion (Additional file 3: Figure S2). However, cell aggregation was not observed in KG206 liquid cultures (data not shown), suggesting that the effect of pnp deletion is less pronounced in the MG1655 background.
PNPase downregulates pgaABCD operon expression at post-transcriptional level
Enhanced stability of pgaABCD mRNA may account for (or at least contribute to) the increase in pgaABCD expression. Indeed, RNA degradation kinetics experiments performed by quantitative RT-PCR showed a small, but reproducible 2.5-fold half-life increase of pgaA mRNA in the Δpnp mutant (from 0.6 min in C-1a to 1.5 min in the pnp mutant; Additional file 4: Figure S3). A comparable effect was elicited by deletion of the csrA gene (estimated mRNA half-life, 1.5 min; Additional file 4: Figure S3), known to regulate pgaABCD mRNA stability in E. coli K12 [38, 51].
Post-transcriptional regulation of the pgaABCD operon by the CsrA protein targets its 234 nucleotide-long 5’-UTR. Therefore, we tested whether this determinant was also involved in pgaABCD control by PNPase. To this aim, we constructed several plasmids (see Table 1) harboring both transcriptional and translational fusions between different elements of the pgaABCD regulatory region and the luxAB operon, which encodes the catalytic subunits of Vibrio harveyi luciferase, as a reporter . Luciferase expression in both pnp+ and Δpnp strains was tested using the transcriptional fusion plasmids pΔLpga and pLpga1, which harbor the pgaABCD promoter region (pgaAp) alone (−116 to +32 relative to the transcript start site) and a region encompassing pgaAp and the entire pgaA leader (without its ATG start codon), respectively. In these constructs, translation of the luxAB transcript depends on the vector translation initiation region (TIR). Conversely, pLpga2 carries a translational fusion of the whole 5’-UTR and the first 5 codons of pgaA with luxA. A plasmid expressing luxAB from Ptac promoter (pTLUX) and the vector TIR was also tested as a control of PNPase effects on luciferase mRNA expression. The results of a typical experiment and relative luciferase activity (Δpnp vs. pnp+) are reported in Figure 4B. In agreement with the role of the 5’-UTR as a strong determinant for negative regulation of pgaABCD expression , luciferase activity was much higher in cells carrying the construct lacking the pgaABCD 5’-UTR (pΔLpga) regardless of the presence of PNPase. The small increment in luciferase expression from the pΔLpga plasmid detected in the Δpnp was not due to increased pgaAp promoter activity as it was observed also with pTLUX control plasmid. Conversely, luciferase expression by pLpga1 and pLpga2 was strongly affected by PNPase, as it increased 4.3- and 12.8-fold, respectively, in the PNPase defective strain (Figure 4B). The difference in relative luciferase activity between the pLpga1 and pLpga2 constructs might be explained by higher translation efficiency for the pLpga2 construct in the Δpnp strain. Altogether, the results of luciferase assays (Figure 4B) and mRNA decay experiments (Additional file 4: Figure S3) suggest that PNPase regulates pgaABCD mRNA decay by interacting with cis-acting determinants located in the 5’-UTR. PNPase has been recently shown to play a pivotal role in sRNA stability control [27, 56] and has been involved in degradation of CsrB and CsrC in Salmonella. We hypothesized that PNPase may act as a negative regulator of pgaABCD operon by promoting the degradation of the positive regulators CsrB and/or CsrC . To test this idea, we combined the Δpnp 751 mutation with other deletions of genes either encoding sRNAs known to affect pgaABCD expression (namely, csrB, csrC and mcaS), or csrD, whose gene product favors CsrB and CsrC degradation . We also readily obtained the ΔcsrA::kan mutation in C-1a (pnp+), indicating that, unlike in K-12 strains , csrA is not essential in E. coli C. Conversely, in spite of several attempts performed both by λ Red mediated recombination  and by P1 reciprocal transductions, we could not obtain a Δpnp ΔcsrA double mutant, suggesting that the combination of the two mutations might be lethal.
In this report, we have shown that PNPase negatively regulates the production of the adhesion factor PNAG, thus maintaining the bacterial cells in a planktonic state (Figures 13) when grown at 37°C in supplemented minimal medium. Our results are in line with previous works by other groups connecting PNPase to regulation of outer membrane proteins in E. coli and curli production in Salmonella . Thus, PNPase seems to play a pivotal role in regulating the composition of cell envelope and the production of adhesion surface determinants. PNPase-dependent regulation of PNAG production requires its ribonuclease activity, as suggested by the observation that overexpression of RNase II can compensate for lack of PNPase (Figure 1B). Cell aggregation in the absence of PNPase is suppressed by RNase II, but not by RNase R. This reminds what previously showed for cold sensitivity in pnp mutants, which is also solely suppressed by RNase II  and reinforces the notion that, albeit partially redundant, RNA degradation pathways possess a certain degree of specificity and are not fully interchangeable .
The precise mechanistic role played by PNPase in regulation of pgaABCD expression, as well as the physiological signals to which it responds, remain elusive. PNPase activity is modulated (at least in vitro) by cyclic-di-GMP , a signal molecule implicated in biofilm formation . However, deletion of the dos gene, encoding a c-di-GMP phosphodiesterase which co-purifies with the RNA degradosome , did not affect pgaABCD expression (data not shown). Key molecules in energy metabolism and carbon flux, such as ATP and citrate also influence PNPase activity [64, 65]. Thus, it can be speculated that environmental or physiological signals might regulate pgaABCD expression by controlling the level of specific metabolites that could directly modulate PNPase activity.
Our data clearly indicate that PNPase controls PNAG production by negatively regulating the pgaABCD operon at post-transcriptional level and that it targets the 5’-UTR of the pgaABCD transcript, thus similar to the translational repressor CsrA (Figures 45 and Additional file 4: Figure S3). This would suggest that the two proteins might belong to the same regulatory network. However, probing this hypothesis is complicated by the observation that in E. coli C, the mechanisms of CsrA-dependent gene expression regulation and its modulation by small RNAs might be more complex than in E. coli K-12, where the current model for CsrA regulation has been developed. This notion is somehow suggested by the fact that, while deletion of the csrA gene is lethal for E. coli K-12 when grown on glucose-based media , this is not the case for E. coli C. Moreover, to our surprise, the lack of putative positive regulators such as CsrB, CsrC and McsA resulted in an increase of pgaABCD expression levels both in the Δpnp and in its parental strain C-1a, which would suggest a negative role of these sRNAs in pgaABCD control (Figure 5). Genes encoding cell surface-associated structures seem to constitute a “hotspot” for post-transcriptional regulation involving small non coding RNAs. For instance, multiple control of gene expression by sRNAs has already been demonstrated for csgD, which encodes the master regulator for the biosynthesis of thin aggregative fimbriae (curli), one of the major adhesion factors in E. coli[28, 55, 66, 67]. It is thus possible that, in E. coli C, increased pgaABCD expression in mutant strains carrying deletions of sRNA-encoding genes might be due to feedback induction of yet unidentified factors which might play a role in CsrA-dependent regulation. This possibility is supported by the observation that CsrB, CsrC and McaS mutually control their transcript level both in E. coli K and C  (T. Carzaniga and F. Briani, unpublished data). pgaABCD operon regulation appears to be an intriguing model system for the study of post-transcriptional modulation of gene expression in bacteria.
In this work, we have unravelled a novel role for PNPase as a negative regulator of pgaABCD expression and PNAG biosynthesis. Thus, PNPase activity contributes to keeping E. coli cells in the planktonic state. Our findings underline the importance of post-transcriptional regulation for genes encoding cell surface-associated structures and factors involved in biofilm formation and suggest the existence of strain-specific variability in these regulatory mechanisms. Indeed, small RNA-dependent post-transcriptional regulation of pgaABCD expression in E. coli C is more complex than the model proposed for E. coli K-12, possibly connected to a central role played by PNAG as a determinant for biofilm formation in the former strain.
We thank Gerald B. Pier (Harvard Medical School, Boston, USA) for his kind gift of anti-PNAG antibodies, Cecilia Arraiano for sending pFCT6.9 plasmid, Maria Pasini for the microscope images, Michela Casali for technical assistance and Michela Gambino for the statistical analysis. This study was supported by PRIN (Project 2008K37RHP) Research Programs of the Italian Ministry for University and Research.
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