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CELL AND MOLECULAR BIOLOGY OF MICROBES

A role for the Rcs phosphorelay in regulating expression of plant cell wall degrading enzymes in Pectobacterium carotovorum subsp. carotovorum

Liis Andresen, Erki Sala, Viia Kõiv, Andres Mäe

  • Department of Genetics, Institute of Molecular and Cell Biology, University of Tartu, Estonian Biocenter, 23 Riia Street, Tartu 51010, Estonia
  • Correspondence
    Andres Mäe
    amae{at}ebc.ee

    • Microbiology 2010; 156(5):1323–1334 · https://doi.org/10.1099/mic.0.033936-0

    View at publisher PubMed

    Abstract

    The Rcs phosphorelay is a signal transduction system that influences the virulence phenotype of several pathogenic bacteria. In the plant pathogen Pectobacterium carotovorum subsp. carotovorum (Pcc) the response regulator of the Rcs phosphorelay, RcsB, represses expression of plant cell wall degrading enzymes (PCWDE) and motility. The focus of this study was to identify genes directly regulated by the binding of RcsB that also regulate expression of PCWDE genes in Pcc. RcsB-binding sites within the regulatory regions of the flhDC operon and the rprA and rsmB genes were identified using DNase I protection assays, while in vivo studies using flhDC : : gusA, rsmB : : gusA and rprA : : gusA gene fusions revealed gene regulation. These experiments demonstrated that the operon flhDC, a flagellar master regulator, was repressed by RcsB, and transcription of rprA was activated by RcsB. Regulation of the rsmB promoter by RcsB is more complicated. Our results show that RcsB represses rsmB expression mainly through modulating flhDC transcription. Neverthless, direct binding of RcsB on the rsmB promoter region is possible in certain conditions. Using an rprA-negative mutant, it was further demonstrated that RprA RNA is not essential for regulating expression of PCWDE under the conditions tested, whereas overexpression of rprA increased protease expression in wild-type cells. Stationary-phase sigma factor, RpoS, is the only known target gene for RprA RNA in Escherichia coli; however, in Pcc the effect of RprA RNA was found to be rpoS-independent. Overall, our results show that the Rcs phosphorelay negatively affects expression of PCWDE by inhibiting expression of flhDC and rsmB.

    • The GenBank/EMBL/DDBJ accession numbers for the rprA, flhDC and rsmB genes are GQ339472, GQ344498, and GQ344499, respectively.

    • A supplementary table listing predicted RcsB-binding sites in Pectobacterium atrosepticum is available with the online version of this paper.

    Edited by: I. K. Toth

    INTRODUCTION

    The phytopathogenicity of Pectobacterium carotovorum subsp. carotovorum (Pcc) is largely due its capacity to synthesize and secrete plant cell wall degrading enzymes (PCWDE), including pectinases, cellulases and protease (Heikinheimo et al., 1995; Marits et al., 1999; Mäe et al., 1995; Pirhonen et al., 1991). Survival of infecting Pcc largely depends on production of PCWDE for the transition from latent infection to disease state. A number of global and specific regulatory genes contribute to expression of PCWDE in Pcc (Burr et al., 2006; Chatterjee et al., 1995; Cui et al., 1995, 1999, 2001, 2005; Eriksson et al., 1998; Flego et al., 2000; Harris et al., 1998; Hyytiäinen et al., 2003; Liu et al., 1998, 1999; Sjöblom et al., 2006). The post-transcriptional Rsm system plays a key role in regulating the expression of PCWDE (Chatterjee et al., 1995; Cui et al., 1995; Liu et al., 1998).The Rsm system consists of two regulators in Pcc: a small protein, RsmA, and RsmB RNA. RsmA is a post-transcriptional global regulator that is thought to bind mRNAs of PCWDE genes and affect their stability, whereas RsmB RNA binds multiple molecules of RsmA, thereby neutralizing its effect (Liu et al., 1998). Recently, the Rsm system was shown to be dependent on expression of a global regulator of flagellar genes, FlhDC. FlhDC activates the expression of PCWDE in Pcc strain 71 by modulating transcription of RsmB (Cui et al., 2008).

    Motility is a key feature of many microbes' physiology, and flagella provide the necessary propulsion to the microbes in a liquid environment. FlhDC, which regulates the expression of flagellar genes (for a review, see Prüß, 2000), also regulates the consumption of glucose (Entner–Doudoroff pathway) and anaerobic respiration in Escherichia coli, as well as induction of purine and pyrimidine synthesis and repression of the urea cycle in Yersinia enterocolitica (Kapatral et al., 2004; Prüß et al., 2001, 2003). Flagellar motility is also a pathogenicity determinant in pectobacteria, and appears to be necessary for the successful invasion and infection of a plant host (Pirhonen et al., 1991; Pérombelon, 2002). Mutants defective in either flagellar synthesis or rotation exhibited a reduced ability to elicit soft-rot symptoms, despite the normal expression of PCWDE (Hossain et al., 2005).

    Previously, we demonstrated that both PCWDE synthesis and motility are regulated by the Rcs phosphorelay in Pcc (Andresen et al., 2007). The Rcs phosphorelay has been implicated in the synthesis of virulence factors in Pcc and other plant-pathogenic bacteria and also in human pathogens (Hinchliffe et al., 2008; Mouslim et al., 2004; Wang et al., 2009, 2007). RcsB activates the biosynthesis of capsular exopolysaccharides, transcription of the cell division gene, ftsZ, transcription of a cell envelope protein, osmC, and transcription of RprA RNA to increase expression of the stationary-phase sigma factor, RpoS, in E. coli (Carballes et al., 1999; Davalos-Garcia et al., 2001; Gervais et al., 1992; Gottesman et al., 1985; Majdalani et al., 2002). Francez-Charlot et al. (2003) demonstrated that the Rcs phosphorelay system represses motility through the direct binding of RcsB to the promoter of flhDC in E. coli.

    To gain further insight into the regulatory network that controls virulence gene expression in Pcc, we focused on the contributions of the Rcs phosphorelay to the expression of PCWDE. We purified RcsB from Pcc and investigated its mode of action in regulating the virulence phenotype: whether it directly binds to the promoters of PCWDE to repress the expression, indirectly represses transcription of virulence activator genes, or activates transcription of virulence repressor genes.

    METHODS

    Bacterial strains, vectors, and growth conditions.

    Strains and plasmids used in this study are listed in Table 1⇓. Pectobacterium carotovorum subsp. carotovorum (Pcc) strains were grown at 30 °C, and Escherichia coli was grown at 37 °C. Bacteria were grown on LB medium or M9 minimal salts medium containing 0.4 % glycerol (Miller, 1972; Sambrook & Russell, 2001). When required, media were supplemented with 0.2 % (w/v) polygalacturonic acid (PGA; Sigma), 150 μg ampicillin (Amp) ml−1, 100 μg kanamycin (Km) ml−1 or 25 μg chloramphenicol (Cm) ml−1.

    Table 1.

    Strains and plasmids used in this study

    Predicted RcsB-binding sites in Pectobacterium.

    Prediction of the RcsB regulon in Pectobacterium was performed using a web-based prediction program, Virtual Footprint Version 3.0 (Münch et al., 2005) on the genome sequence of Pectobacterium atrosepticum strain SCRI 1043 (Pba) (; Bell et al., 2004). A position-specific weight matrix was generated using RcsB-binding sites described by Wehland & Bernhard (2000), as well as RcsB-binding sequences identified in the promoters of flhDC, rprA, osmC and ftsZ genes in E. coli (Carballes et al., 1999; Davalos-Garcia et al., 2001; Francez-Charlot et al., 2003; Majdalani et al., 2002). The sensitivity threshold was set to 0.8–11.3, only RcsB-binding sites within 350 bp of the start of translation were considered, and the search was restricted to intergenic regions.

    PCR cloning and DNA sequencing.

    Standard DNA techniques described in Sambrook & Russell (2001) were used. Oligonucleotide primers used in this study for PCR amplification are listed in Table 2⇓. The genomic regions of the rprA, flhDC and rsmB genes of Pcc strain SCC3193 were cloned by using primers obtained from the genome sequence of Pba. The amplified fragments were cloned into plasmid pTZ57R/T, and sequenced. The GenBank accession numbers for the rprA, flhDC and rsmB genes are GQ339472, GQ344498, and GQ344499, respectively.

    Table 2.

    Oligonucleotide primers used in this study

    Overexpression and purification of histidine-tagged RcsB and RcsBD56E.

    The rcsB gene was amplified using primers RcsBNcoI and RcsBHindIII and was cloned into the SmaI site of Bluescript SK+ to yield pLA1. The rcsB gene was subsequently subcloned using NcoI and HindIII into plasmid pET24d having a C-terminal His6 tag (pET24d-rcsB). RcsB was overexpressed from this plasmid as a fusion protein with a C-terminal His6 tag and purified as described by Hõrak et al. (2004). The protocol was modified to include buffers A/B [100 mM Tris/HCl (pH 7.5), 1 M NaCl, 25 mM imidazole, 10 mM DTT, 0.1 mM EDTA, 0.1 % Triton X-100, 5 % glycerol] and C [100 mM Tris/HCl (pH 7.5), 500 mM NaCl, 500 mM imidazole, 10 mM DTT, 0.1 mM EDTA, 0.1 % Triton X-100, and 50 % glycerol]. Purified protein was dialysed against storage buffer [10 mM Tris/HCl (pH 7.5), 200 mM KCl, 1 mM DTT, 0.5 mM EDTA, 50 % glycerol] and stored at −20 °C.

    Site-directed mutagenesis and purification of modified protein.

    The sequence of the rcsB in which the conserved aspartate codon (D56) was substituted with a codon for glutamic acid was generated as follows. Partial sequence of rcsB was amplified with primers rcsBalg and RcsBD56E from Pcc chromosomal DNA, purified with the UltraClean 15 DNA Purification kit (MO BIO), and subsequently used as a primer in a PCR paired with primer rcsCend, and chromosome of Pcc as a template. Amplified PCR product was cloned into the cloning vector pJET1 to yield pJET1-rcsBD56E, according to the supplied protocol (Fermentas). The rcsBD56E gene was amplified using primers RcsBNcoI and RcsBHindIII with plasmid pJET1-rcsBD56E as a template. The subsequent cloning, protein expression and purification procedures were the same as for histidine-tagged RcsB.

    DNase I footprinting.

    DNA fragments for DNase I footprinting assays were amplified by PCR from genomic DNA of Pcc SCC3193 and one of the primers end-labelled with [γ-32P]ATP by polynucleotide kinase (Fermentas). Amplified DNA fragments included: (i) flhDC (203 bp) having upstream sequences −374 bp to −171 bp using primers flhD1 and flhD2; (ii) rprA (159 bp) including upstream sequences −124 bp to +35 bp using primers rprAFP1 and rprAFP2; (iii) rsmB (325 bp) including upstream sequences −287 bp to +38 bp using primers rsmBpA and rsmBpY; and (iv) prtW (269 bp) including upstream sequences −55 bp to +214 bp using primers PROS and PRIMEX2. DNase I footprinting reactions were performed as described by Ilves et al. (2004), except that 0.01U DNase I (Fermentas) was used and the concentration of RcsB-6His or RcsBD56E-6His ranged from 0.5 to 20 μM. DNA sequencing reactions were performed using a Sequenase Version 2.0 kit (US Biochemicals). Footprinting and sequencing reactions were resolved and dried in 6.5 % denaturing polyacrylamide gels and scanned using a Typhoon phosphoimager screen (Amersham Biosciences).

    Construction of mutant strains.

    The rcsB : : Km fragment was amplified using primers rcsBP1 and rcsBP2, with plasmid, pKD4 as template. Replacement of chromosomal rcsB with rcsB : : Km was performed using the λ Red system (Datsenko & Wanner, 2000) to yield mutant SCC6030. The flhDC-negative mutant (SCC6028) was generated as described by Datsenko & Wanner (2000) using primers flhDP1 and flhDP2, which amplified the cat (CmR) gene from pKD3 with 49 bp of the flhD sequence on either side to position the insert +1 with respect to the start codon of flhD. The flhD/rcsB double mutant (SCC6031) was generated using the rcsB : : Km mutant (SCC6030) background and the same methods and primers as described above.

    To obtain a knockout mutant of rprA, plasmid pLA2 (pTZ57R/T containing a PCR fragment amplified with primers rprAY and rprAA2) was digested with restriction enzyme BplI (Fermentas) to disrupt the rprA gene sequence at position +25 relative to the transcription start site. The cat gene, obtained from plasmid pUT mini-Tn5 Cm as a BamHI fragment, was cloned into BplI-digested pLA2 to create pLA9. The disrupted rprA fragment was amplified from pLA9 using primers rprAY and rprAA2 and cloned into the EcoRV-digested vector, pGP704L, to yield pLA10. The rprA-negative mutant (SCC6029) was created using the λ Red system (Datsenko & Wanner, 2000), with primers rprAY and rprAA2 and pLA10 as a template.

    All of the mutants generated were verified by PCR amplification.

    Enzyme assays.

    For β-glucuronidase (GusA) assays, cells were grown in M9 minimal medium supplemented with 0.4 % glycerol with or without the addition of PGA. GusA activity was measured at 6, 8 and 10 h after inoculation. In the case of pMW119-flhDC : : gusA and pMW119-rprA : : gusA constructs, GusA activity was measured with p-nitrophenyl β-d-glucuronide as a substrate (Novel et al., 1974). The degradation product, p-nitrophenol (pNP), was detected at 405 nm, and the specific activity of GusA was expressed as nmol pNP liberated min−1 per OD600 unit. In the case of the pMW119-rsmB : : gusA construct, GusA activity was measured with 4-methylumbelliferyl β-d-glucuronide (MUG; DUCHEFA Biochemie) as substrate. Five microlitres of sample was taken at each time point and permeabilized in 90 μl CTAB buffer (0.05 % hexadecyltrimethylammonium bromide, 0.27 % β-mercaptoethanol and 1 mM EDTA in 50 mM sodium phosphate buffer pH 7) for 10 min. After addition of MUG (15 μmol l−1) to the permeabilized cells, accumulation of the florescent product 4-methylumbelliferone (MU) was measured with a fluorescence spectophotometer (GENios Plus; Tecan) using an excitation filter of 360 nm and an emission filter of 465 nm. The slope of relative fluorescence units per minute was calculated. GusA activity of the sample was expressed as nmol substrate hydrolysed h−1 ml−1 per OD600 unit, based on a standard curve of MU concentrations. All GusA assays were performed using three technical replications.

    To perform semiquantitative agarose plate assays of extracellular protease (Prt) production, cells were grown overnight in M9 medium and washed and suspended in the same medium. Drops of cell suspensions containing ∼106 cells were dotted onto milk-containing agar plates (Chatterjee et al., 1995), and the plates were photographed after 3 days of growth at 30 °C. The diameter of the halo produced was considered to be proportional to the amount of enzyme secreted. All assays were performed at least five times.

    To quantify the activity of pectate lyase and cellulase, cells were grown on M9 minimal medium supplemented with 0.2 % PGA. Three replications were used to measure enzyme activities as described previously (Pirhonen et al., 1991).

    Real-time RT-PCR.

    Wild-type cells harbouring the control plasmid, pMW119, or a plasmid overexpressing rprA, pMW119-rprA, were grown overnight on milk-containing agar plates at 30 °C. Cells were harvested and total RNA was isolated using the NucleoSpin RNA II kit (Macherey-Nagel). An additional treatment with DNase I (Fermentas) was performed according to the manufacturer's protocol. Real-time RT-PCR was performed using the Verso SYBR Green 1-Step QRT-PCR ROX kit (Thermo Scientific) using the Applied Biosystems 7900 HT real-time thermal cycler. Each reaction contained 2 ng total RNA and 100 nM forward and reverse primers. The sequences of primers used are given in Table 2⇑.

    The PCRs were run using the following programme: 50 °C for 15 min, 95 °C for 15 min to inactivate reverse transcriptase, and 40 cycles of 95 °C for 15 s and 60 °C for 30 s. Following PCR amplification, the reactions were subjected to temperature ramping to create the dissociation curve, measured as changes in fluorescence measurements as a function of temperature, by which the non-specific products can be detected. The dissociation programme was 95 °C for 1 min, 60 °C for 15 s, followed by a 2 % ramp from 60 °C to 95 °C. Results are presented as mean expression ratios calculated using the

    Figure image not available in archive
    method as described by Livak & Schmittgen (2001). The ffh gene (encoding a signal recognition particle involved in targeting and integration of inner-membrane proteins) was used for normalization of reverse transcriptase real-time PCR data according to Takle et al. (2007). All RNA isolations and reverse transcriptase real-time PCRs were performed using three replications.

    RESULTS

    The response regulator, RcsB, binds to the promoters of flhDC, rprA and rsmB

    We previously demonstrated that an rcsB-negative mutant expressed higher levels of PCWDE than wild-type Pcc (Andresen et al., 2007). To further investigate the regulatory effect of RcsB on PCWDE expression in Pcc, the promoter regions of the prtW, pehA, pelB, pelC and celV1 genes were analysed for RcsB-binding sites. Using the Virtual Footprint program (see Methods; Münch et al., 2005), regions upstream of these genes were not found to contain any putative RcsB-binding sites. Subsequently, the genome of Pba was analysed, and 16 putative RcsB-binding sites were found associated with promoter regions of 16 genes (see Supplementary Table S1, available with the online version of this paper). Of these genes, RcsB-binding sites in flhDC and rprA were previously characterized in E. coli, and rsmB codes for a global regulator of PCWDE in Pcc (Francez-Charlot et al., 2003; Liu et al., 1998; Majdalani et al., 2002). Therefore, flhDC, rprA and rsmB were selected for further analysis.

    To identify the DNA-binding sites for RcsB in the promoter regions of flhDC, rprA and rsmB, DNase I footprinting assays were performed using purified, histidine-tagged RcsB. The DNA footprinting assay of flhD upstream sequences (−374 to −171) revealed an RcsB-binding site that protected a region spanning 26 bp from −262 to −236 (Fig. 1a⇓). In the rprA promoter, bp −124 to +35 and bp −51 to −27 were protected by binding of RcsB (Fig. 1b⇓). In contrast, neither the rsmB promoter, nor the promoter of an extracellular protease, PrtW, used as a negative control, were protected from DNase I cleavage in the presence of histidine-tagged RcsB (data not shown). The putative RcsB-binding site in the rsmB promoter was identified using purified histidine-tagged RcsBD56E. Replacement of conserved aspartate residue in position 56 with glutamic acid is known to make the RcsB protein constitutively active in E. coli, without needing RcsC, RcsF or RcsA for its activity (Gupte et al., 1997). DNase I footprinting revealed that the protected regions extended from position −192 to position −211 and from position −213 to position −230 upstream of the start codon of rsmB (Fig. 1c⇓). The prtW promoter fragment, used as a negative control, was not protected from DNase I cleavage in the presence of histidine-tagged RcsBD56E (data not shown). The nucleotide sequences of the protected regions identified in this study are similar to known RcsB target sites (Fig. 1d⇓).

    Figure image not available in archive
    Fig. 1.

    (a–c) DNase I footprinting assays of histidine-tagged RcsB, histidine-tagged RcsBD56E, and promoter regions of flhDC (a), rprA (b) and rsmB (c). DNA probes labelled on the coding strand were incubated with varying concentrations (0–20 μM) of histidine-tagged RcsB (RcsB-6H) or histidine-tagged RcsBD56E (RcsBD56E-6His) prior to treatment with DNase I. Sequencing gels are shown and the letters A, T, G, and C indicate the DNA sequences of fragments digested by DNase I. (d) The RcsB-binding sites of the flhDC, rprA and rsmB promoters of Pcc are aligned with known RcsB-binding sequences in E. coli, predicted RcsB-binding sequences in Pba, as well as the RcsAB box described by Wehland & Bernhard (2000). Lower-case letters in the RcsAB box indicate nucleotides that have a conservation rate of 50–70 % relative to the consensus sequence, and upper-case letters indicate a conservation rate of >70 %.

    RcsB modulates expression of flhDC, rprA and rsmB in Pcc

    To further characterize the influence of RcsB binding on flhDC, rprA and rsmB expression, fusions of gusA with flhDC, rprA and rsmB were created, and gene expression for the three constructs was compared between the wild-type strain and an rcsB-negative mutant. In the rcsB-negative mutant, a fourfold increase in expression of the flhDC : : gusA fusion was detected, while only low levels of expression of the rprA : : gusA fusion were detected compared to the wild-type strain. These results indicate that flhDC and rprA are directly regulated by RcsB in Pcc, with RcsB positively regulating rprA expression, and negatively affecting flhDC expression.

    The expression of the rsmB : : gusA fusion was increased 15–18-fold in an rcsB-negative mutant as compared to the wild-type strain (Fig. 2a⇓). We also found that the effect of the rcsB mutation on rsmB : : gusA expression is mediated through flhDC in these conditions. We compared the expression of rsmB : : gusA fusion in an flhDC-negative mutant and in an flhDC/rcsB double-negative mutant. No significant differences were found in rsmB : : gusA expression between these strains in M9 minimal salts medium supplemented with glycerol throughout the entire growth curve (data not shown). When the same mutants were tested for rsmB : : gusA expression in M9 minimal salts medium supplemented with PGA, GusA activity in flhDC/rcsB double-negative mutant was similar to that in flhDC-negative strain, except at one time point (6 h after inoculation) in the early exponential growth phase, where the flhDC/rcsB double mutant expressed 3.4 times higher GusA activity than the flhDC-negative strain (Fig. 2b⇓).

    Figure image not available in archive
    Fig. 2.

    Expression of promoters fused with reporter gene gusA coding for β-glucuronidase (GusA). GusA activity was measured at different times after inoculation and experiments were performed in triplicate. Error bars represent sd. (a) Expression of flhDC : : gusA, rprA : : gusA and rsmB : : gusA fusions was measured in wild-type (wt; SCC3193) and rcsB-negative (rcsB : : Km; SCC6030) strains grown in M9 minimal medium supplemented with glycerol. In the case of the flhDC and rprA promoter constructs (pMW119-flhDC : : gusA and pMW119-rprA : : gusA) GusA activity was measured with p-nitrophenyl β-d-glucuronide as a substrate (left axis), whereas in the case of the rsmB promoter fusion (pMW119-rsmB : : gusA) GusA activity was measured with 4-methylumbelliferyl β-d-glucuronide as a substrate (right axis); (b) Expression of rsmB : : gusA fusion in wild-type (▪), flhDC-negative mutant (•) and flhDC/rcsB double mutant (▴) in minimal medium supplemented with 0.2 % PGA.

    RcsB modulates expression of PCWDE through flhDC

    To determine whether the Rcs phosphorelay modulates expression of PCWDE by regulating flhDC, flhDC mutants were created in Pcc strain SCC3193. As shown in Fig. 3⇓, inactivation of flhDC resulted in a dramatic decrease in levels of pectate lyases, cellulase and protease compared to the wild-type strain. When expression of flhDC was restored using a low-copy plasmid (pMW119-flhDC), expression of PCWDE increased in the wild-type strain, and was restored in the flhDC-negative strain. An rcsB null mutation in a wild-type background was also previously shown to increase expression of PCWDE (Andresen et al., 2007). To investigate whether rcsB inactivation could restore expression of PCWDE in an flhDC mutant, enzyme production in an flhDC/rcsB double mutant was analysed. The double mutant did not regain enzyme production when compared to the rcsB mutant strain, indicating that the effect of rcsB inactivation on expression of PCWDE is dependent on FlhDC.

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    Fig. 3.

    Influence of FlhDC and RprA RNA on the expression of PCWDE. (a, b) For assays of pectate lyases (a) and cellulase (b), enzyme activities were measured 10 h after inoculation and the means±sd of three independent experiments are shown. (c) Production of protease was observed after 3 days and the diameter of the halo (in mm) was measured from at least five independent experiments. ND, not detectable; wt, wild-type strain SCC3193; flhDC : : Cm, flhDC-negative strain SCC6028; rcsB : : Km, rcsB-negative strain SCC6030; flhDC : : Cm rcsB : : Km, flhDC- and rcsB- negative double mutant; rprA : : Cm, rprA-negative strain SCC6029; flhDCpMW119, flhDC overexpression plasmid; rprApMW119, rprA overexpression plasmid; pMW119, control vector.

    Overexpression of RprA RNA modulates rpoS-independent expression of an extracellular protease and has only a minor effect on expression of pectate lyases and cellulase

    As shown in Fig. 2(a)⇑, transcription of the rprA gene is positively regulated by RcsB. To assess the capacity of RprA RNA to regulate expression of PCWDE, an rprA-negative mutant was constructed and enzyme assays were performed. The activity of pectate lyases, cellulase and protease in both mutant and wild-type strains was detected. Inactivation of rprA did not affect expression of these enzymes (Fig. 3⇑). When rprA was overexpressed in the wild-type strain, protease activity increased (Fig. 3⇑). Cellulase activity of the strain overexpressing rprA was 1.7 times higher than that of the wild-type strain, while activity of pectate lyase was only slightly elevated (Fig. 3⇑).

    RprA RNA was initially discovered and characterized in E. coli as a positive regulator of rpoS translation (Majdalani et al., 2001). The effect of RpoS on RprA RNA modulation of PCWDE in Pcc was therefore analysed using plate assays. As shown in Fig. 4⇓, overexpression of rprA in an rpoS-negative mutant stimulated protease expression, indicating that the effect of rprA on protease synthesis is independent of rpoS.

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    Fig. 4.

    Effect of rprA overexpression on protease expression in wild-type (wt; strain SCC3193) and RpoS-mutant (rpoS : : Km; strain SCC8002) strains. Wild-type containing vector pMW119 was used as a control (wt+pMW119). Assay plates were photographed after 3 days and the diameter of the halo (in mm) was measured from at least five independent experiments.

    Regulation of PCWDE by FlhDC and RprA RNA does not overlap

    The results shown in Fig. 3⇑ demonstrate that both flhDC and rprA are able to enhance protease expression in Pcc. To determine whether flhDC and rprA can functionally replace each other, an flhDC overexpression construct was introduced into an rprA-negative mutant and an rprA overexpression construct was introduced into a flhDC-negative mutant. When flhDC was exogenously expressed from pMW119-flhDC in a rprA-negative background, protease expression was increased. However, when rprA was exogenously expressed from pMW119-rprA in a flhDC-negative background, protease expression did not increase (Fig. 5⇓). To determine whether the positive effect of rprA overexpression on protease expression is dependent on an increase in flhDC expression, reverse transcription real-time PCR was used to evaluate flhDC mRNA levels in a wild-type strain, an rcsB-negative mutant, and a strain overexpressing rprA. In the rcsB-negative mutant, flhDC mRNA levels were 1.36±0.15 times higher than in the wild-type strain. In contrast, overexpression of rprA decreased flhDC mRNA levels compared with wild-type (rprA overexpression to wild-type ratio 0.84±0.22). These results demonstrate that activation of protease transcription associated with overexpression of rprA is not caused by increased flhDC transcription.

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    Fig. 5.

    Effects of overexpressing flhDC and rprA on protease expression in rprA-negative (rprA : : Cm; strain SCC6029) and flhDC-negative (flhDC : : Cm; strain SCC6028) mutants, respectively. Assay plates were photographed after 3 days of incubation and the diameter of the halo (in mm) was measured from at least five independent experiments. ND, not detectable.

    DISCUSSION

    Genes directly regulated by the Rcs phosphorelay response regulator, RcsB, have been shown to have RcsB-binding sites in their promoters (Wehland & Bernhard, 2000). However, no RcsB-binding sites were identified in any of the PCWDE promoters analysed, although sequences similar to RcsB sites were found in the regulatory regions of the rsmB, flhDC and rprA genes (Supplementary Table S1). In vitro, RcsB was shown to directly bind the promoter regions of flhDC, rprA and rsmB. The regulatory regions of flhDC and rprA were bound by the unphosphorylated form of RcsB, whereas the rsmB promoter was bound only by RcsBD56E, a constitutively active form of RcsB (Gupte et al., 1997). These results indicate that the binding sites of RcsB in the rsmB promoter region may have lower affinity for unphosphorylated RcsB when compared to the binding sites in the rprA and flhDC promoters. This would suggest that rsmB is directly regulated by RcsB in response to a signal activating the Rcs phosphorelay, whereas flhDC and rprA are regulated by RcsB even in the absence of the signal.

    In vivo, RcsB positively affected rprA transcription and negatively affected flhDC and rsmB transcription (Fig. 2a⇑), demonstrating that direct transcriptional regulation of rprA and flhDC expression by the Rcs phosphorelay is not exclusive to E. coli and Salmonella (Francez-Charlot et al., 2003; Majdalani et al., 2002; Wang et al., 2007). The flagella master regulator, FlhDC, has been previously shown to positively regulate RsmB RNA levels in Pcc (Cui et al., 2008; Fig. 2b⇑); thus RcsB may regulate transcription of rsmB both directly and indirectly through modulating the expression of flhDC. Our experiments with an flhDC-negative mutant and an flhDC/rcsB double-negative mutant indicate that in minimal medium the effect of RcsB on rsmB transcription is mediated by FlhDC. When these mutants were grown on minimal medium supplemented with PGA the expression of the rsmB : : gusA fusion was 3.4 times higher in the flhDC/rcsB double-negative mutant than in the flhDC-negative mutant at one time point in the early exponential growth phase (Fig. 2b⇑). These results indicate that rsmB expression is mainly regulated by RcsB via flhDC also in the presence of PGA, but can be directly regulated by RcsB at certain stages of growth. Why the difference in rsmB : : gusA expression between flhDC-negative and flhDC/rcsB double-negative mutant appears only in the presence of PGA and at a certain time is currently under investigation.

    Our results are consistent with previous studies that have shown that the flagella master regulator, FlhDC, contributes to the virulence of Pcc (Cui et al., 2008). The results of this study demonstrate that inactivation of rcsB in an flhDC-negative mutant does not restore expression of PCWDE. Since the flhDC mutant and the flhDC/rcsB double mutant exhibited the same phenotype, a model where the Rcs phosphorelay targets PCWDE genes indirectly via FlhDC is supported. Cui et al. (2008) reported that FlhDC activates expression of PCWDE indirectly by reducing the activity of the Rsm regulatory system. Based on these data and our results, we hypothesize that the Rcs phosphorelay regulates the expression of PCWDE by modulating the activity of the Rsm system directly (only in certain conditions) by regulating rsmB expression and indirectly through flhDC (Fig. 6⇓). Furthermore, our results suggest that the effect of the Rcs phosphorelay on motility of Pcc presented in our previous paper (Andresen et al., 2007) was most likely due to elevated expression of flhDC in the rcs-negative mutants.

    Figure image not available in archive
    Fig. 6.

    A proposed model for regulated expression of PCWDE by the Rcs phosphorelay in Pcc. The steps of the model indicated on the shaded background are based on data from this study. RcsB affects the expression of PCWDE by directly binding the promoters of rprA, rsmB and flhDC. RcsB positively controls the transcription of small regulatory RprA RNA and negatively affects the transcription of rsmB and flhDC. A question mark indicates that the exact conditions in which these interactions become important in PCWDE regulation are not yet known. Expression of FlhDC activates gacA expression and represses hexA expression, which in turn increases the level of RsmB RNA in Pcc (Cui et al., 2008) and decreases the pool of free RsmA, resulting in enhanced expression of PCWDE (Liu et al., 1998). Not all of the molecular mechanisms encompassed by this model are fully characterized (indicated with dashed lines). Furthermore, RsmC acts as an anti-FlhDC factor by binding to FlhDC and interfering with FlhDC action (Chatterjee et al., 2009).

    We were also interested in dissecting the contribution of rprA to expression of PCWDE in Pcc. In contrast to flhDC, which has a significant effect on expression of PCWDE, only increased expression of an extracellular protease was detected in response to exogenous expression of rprA in the wild-type strain. Our data further exclude the possibility that RprA RNA is dependent on flhDC for regulation of protease expression. Moreover, although exogenous expression of rprA did not enhance flhDC mRNA levels as detected by reverse transcription real-time PCR assays, it is intriguing to hypothesize that RprA RNA may affect the translation or proteolysis of FlhDC.

    In E. coli and Salmonella, RprA RNA positively affects translation of RpoS by direct interaction with rpoS mRNA, which is currently the only known target for RprA RNA (Jones et al., 2006; Majdalani et al., 2001, 2002). In both organisms RprA RNA has an effect on rpoS translation in the absence of DsrA RNA (another RNA regulator of rpoS translation) under conditions of osmotic shock (Jones et al., 2006; Majdalani et al., 2001). The results of these studies indicate that RprA RNA needs specific conditions for its activity on rpoS translation. In this study, rprA overexpression was shown to affect protease expression, suggesting that transcription of rprA could stimulate protease expression in the wild-type strain under physiological conditions where the expression level of rprA is elevated. Correspondingly, in E. coli, rprA expression has been shown to increase when the Rcs phosphorelay is artificially activated (Majdalani et al., 2005). Further studies are needed to address the role of RprA RNA as a regulator of protease synthesis in Pcc under conditions that activate the Rcs phosphorelay.

    In the current study, rpoS was not necessary for protease expression induced by RprA RNA (Fig. 4⇑), and this is the first evidence for a role for RprA RNA that is independent of RpoS. It remains unclear whether RprA RNA directly binds the target protease mRNA, or whether RprA RNA indirectly induces protease expression via another regulatory gene.

    In summary, we have demonstrated that the Rcs phosphorelay mediates expression of PCWDE mainly through the direct binding of RcsB to the flhDC promoter, which in turn causes a subsequent decrease in rsmB transcription. Another way to repress rsmB transcriptsion is by direct binding of RcsB on the promoter region of rsmB, but the conditions in which that becomes important in PCWDE regulation are not yet clear. Lower RsmB RNA levels increase the levels of active RsmA in the cell, which effectuates the degradation of PCWDE gene mRNAs (Liu et al., 1998; Fig. 6⇑). Further study is needed to identify the direct target genes for FlhDC in the regulatory pathway controlling expression of PCWDE genes, the physiological conditions that stimulate the regulatory role of RprA, and the complex relationship between these regulatory factors.

    Acknowledgments

    This research was supported by the Estonian Science Foundation (GLOMR7082 and SF0180088s08). We thank Dr T. Alamäe for critical reviewing of the manuscript.

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