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. 2009 Jan 13;10(2):277–290. doi: 10.1111/j.1364-3703.2008.00531.x

The Rcs phosphorelay system is essential for pathogenicity in Erwinia amylovora

DONGPING WANG 1, SCHUYLER S KORBAN 2, YOUFU ZHAO 1,✉
PMCID: PMC6640219  PMID: 19236575

SUMMARY

The Rcs phosphorelay system is a modified two‐component signal transduction system found exclusively in Enterobacteriaceae. In this study, we characterized the roles of the Rcs system in Erwinia amylovora, a highly virulent and necrogenic enterobacterium causing fire blight disease on rosaceous plants. Our results showed that rcsB, rcsC, rcsD and rcsBD mutants were non‐pathogenic on immature pear fruit. The bacterial growth of these mutants was also greatly reduced compared with that of the wild‐type strain in immature pear fruit. In an in vitro amylovoran assay, rcsB and rcsD mutants were deficient in amylovoran production, whereas the rcsC mutant exhibited higher amylovoran production than that of the wild‐type. Consistent with amylovoran production, expression of the amylovoran biosynthetic gene amsG, using green fluorescent protein as a reporter, was not detectable in rcsB, rcsD and rcsBD mutants both in vitro and in vivo. The expression of amsG in vitro was higher in the rcsC mutant than in the wild‐type, whereas its expression in vivo was higher in the wild‐type than in the rcsC mutant. In addition, rcs mutants were more susceptible to polymyxin B treatment than the wild‐type, suggesting that the Rcs system conferred some level of resistance to polymyxin B. Furthermore, rcs mutants showed irregular and slightly reduced motility on swarming plates. Together, these results indicate that the Rcs system plays a major role in virulence and survival of E. amylovora in immature pear fruit.

INTRODUCTION

Fire blight, caused by the Gram‐negative bacterium Erwinia amylovora, is one of the most devastating bacterial diseases worldwide. Erwinia amylovora is capable of infecting blossoms, fruits, vegetative shoots, woody tissues and rootstock crowns, leading to blossom blight, shoot blight and rootstock blight symptoms (Vanneste, 2000). Erwinia amylovora is a member of the Enterobacteriaceae, which includes many important human and animal pathogens, such as Escherichia coli, Yersinia pestis, Salmonella enterica and Shigella flexneri. Like many other plant pathogenic bacteria, E. amylovora utilizes the capsular exopolysaccharide (EPS) to cause disease (Bellemann and Geider, 1992). In E. amylovora, the EPS amylovoran is an acidic heteropolymer composed of a pentasaccharide repeating unit containing four galactose residues and one glucuronic acid molecule (Nimtz et al., 1996). Although previous studies have established that amylovoran production is required for the pathogen to cause disease, the precise role of amylovoran in the disease process remains unclear, although a number of functions have been suggested. These functions include the plugging of plant vascular tissues, masking cell surface components to elude plant defences, movement through cortical tissue and retention of water and nutrients (Sjulin and Beer, 1978; Vanneste, 2000). Recently, amylovoran production has been shown to be necessary for biofilm formation in E. amylovora (Koczan et al., 2008).

In prokaryotes, the two‐component signal transduction (TCST) system is a major regulatory system that utilizes phosphorylation as a means for signal transduction (Chang and Stewart, 1998; Parkinson and Kofoid, 1992). Generally, a typical TCST system is composed of a cognate pair of proteins, a membrane‐bound histidine kinase (HK) sensor and a cytoplasmic response regulator (RR). TCST systems play critical roles in the sensing of and response to environmental conditions to regulate a wide array of cellular pathways, especially in bacterial pathogenesis (Flamez et al., 2007; Hoch and Silhavy, 1995). The Rcs phosphorelay system, first identified for its role in the transcriptional regulation of genes for capsular polysaccharide biosynthesis in Es. coli, is a very complex and well‐studied TCST system (Majdalani and Gottesman, 2005, 2007). The gene organization of the Rcs system is conserved among enterobacteria, in which the rcsBD operon is transcribed in one direction and the rcsC gene is transcribed in the opposite direction (Fig. 1a; Takeda et al., 2001). One unique feature of the Rcs system is that it is composed of three separate proteins containing four signalling domains (Fig. 1b). The RcsC protein is a hybrid sensor kinase that contains a conserved histidine kinase domain (HisKA) and a receiver domain (REC) (Rogov et al., 2006). RcsB is the RR and contains a conserved REC (Francez‐Charlot et al., 2003). In addition to RcsC and RcsB, a third protein RcsD (also known as YojN), containing a histidine phosphotransfer (HPt) domain, but lacking a conserved histidine residue in the HisKA domain, is also involved in the Rcs phosphorelay process (Huang et al., 2006; Takeda et al., 2001). On stimulation, RcsC is thought to transfer a phosphoryl group from its HisKA domain to its REC domain and then to the HPt domain of RcsD. The phosphoryl group from RcsD travels to the aspartate group of RcsB, and the phosphorylated RcsB further regulates the expression of downstream genes, such as colanic polysaccharide synthesis (cps) genes (Huang et al., 2006; Majdalani and Gottesman, 2005; Mizuno, 1998; Parkinson, 1993; Takeda et al., 2001).

Figure 1.

Figure 1

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Schematic maps of genes and proteins of the Rcs phosphorelay system in Erwinia amylovora. (a) Organization of the rcs genes in E. amylovora. (b) Domain organization of Rcs proteins. Domain limits for proteins are derived from the smart program (http://www.smart.embl‐heidelberg.de). Not drawn to scale. His‐kinase, histidine kinase domain; Hpt, histidine phosphotransferase domain; H‐T‐H, helix–turn–helix motif; TM, transmembrane domain; unlabelled box for RcsD, His‐Kinase domain lacking a conserved histidine residue.

In Es. coli, the Rcs phosphorelay system positively regulates cps gene expression (Jayaratne et al., 1993; Stevenson et al., 1996, Stout and Gottesman, 1990). The colanic acid capsule is produced primarily at low temperature in Es. coli, indicating that the colanic acid capsule does not contribute to virulence in Es. coli during infection of animals and humans (Allen et al., 1987a,b; Russo et al., 1995). Other studies also report that the Rcs system negatively regulates the flhDC operon (Wang et al., 2007), the products of which, in turn, positively act as the master regulator for flagella biosynthesis (Fraser and Hughes, 1999). Mutations in the Rcs system render the pathogen hypermotile (Francez‐Charlot et al., 2003). Recently, DNA microarray analysis of the rcsC mutant has revealed that more than 150 genes are controlled by rcsC; among them, 50% of genes encode proteins either localized to the envelope of Es. coli or with activities affecting properties of the bacterial surface (Ferrieres and Clarke, 2003).

Although the Rcs phosphorelay system does not contribute to virulence in Es. coli, it plays a role in the regulation of virulence gene expression in other plant and animal pathogens. In the animal pathogen S. enterica, the type I capsule, which plays an important role in pathogenesis, is synthesized by genes similar to colanic acid genes of Es. coli, and these genes are regulated by the Rcs system (Detweiler et al., 2003; Whitfield and Roberts, 1999). In Es. coli and other enterobacteria, type I capsules are involved in protecting the bacteria from desiccation, phages and toxic chemicals, and in biofilm formation. Mutants of rcsC, rcsB and rcsD in S. enterica and Yersinia pseudotuberculosis poorly colonize host tissues (Hinchliffe et al., 2008; Mouslim et al., 2004). In addition, the Rcs phosphorelay system has been shown to be important for stress resistance in vitro, especially for resistance to the cationic peptide polymyxin B in S. enterica and resistance to bile in Y. pseudotuberculosis (Erickson and Detweiler, 2006; Hinchliffe et al., 2008). In the plant pathogen Erwinia (Pectobacterium) carotovora pv. carotovora, the causal agent of soft‐rot disease, the Rcs phosphorelay system negatively modulates the expression of plant cell wall‐degrading enzymes (Andresen et al., 2007). In Pantoea stewartii ssp. stewartii, causing Stewart's wilt disease of corn, the synthesis of the EPS stewartan is also dependent on Rcs (Minogue et al., 2005; Wehland et al., 1999). However, Rcs‐mediated activation of EPS synthesis is further controlled by a quorum sensing system (EsaI/R) in P. stewartii (Koutsoudis et al., 2006; Minogue et al., 2005). A mutation in esaI blocks EPS production, even in the presence of a functional Rcs system, thus rendering the mutant avirulent, whereas disruption of the esaR gene produces high levels of stewartan constitutively and the mutant is severely attenuated in virulence (Herrera et al., 2008; Koutsoudis et al., 2006).

In E. amylovora, the capsule polysaccharide biosynthesis is also regulated by the Rcs system (Bernhard et al., 1993; Wehland et al., 1999). In E. amylovora, amylovoran biosynthetic genes are encoded by the ams operon from amsA to amsL (Bugert and Geider, 1995). Previous studies have shown that amylovoran biosynthesis is strongly affected by rcsA (Bernhard et al., 1990; Coleman et al., 1990) and rcsB (Bereswill and Geider, 1997; Kelm et al., 1997). RcsB is a typical LuxR family RR with a helix–turn–helix motif in its C‐terminal DNA binding domain (Francez‐Charlot et al., 2003). However, RcsA is similar to RcsB but without the conserved aspartate residue in its N‐terminal REC (Bernhard et al., 1990). Genetic studies have shown that mutation in rcsB renders the pathogen non‐pathogenic and abolishes amylovoran production, whereas the rcsA mutant has reduced virulence but still produces about 10% of amylovoran (Bereswill and Geider, 1997; Bernhard et al., 1990). Further studies have shown that both RcsA and RcsB bind to an ‘RcsAB box’ (TaAGaatatTCctA) in the promoter of amsG and other genes, as RcsAB heterodimers or RcsB–RcsB homodimers (Wehland et al., 1999). However, the roles of RcsC and RcsD in amylovoran biosynthesis and bacterial virulence have not been studied previously.

The overall goal of this study was to determine the potential roles of the Rcs phosphorelay system in E. amylovora virulence and survival. In this study, we first investigated the function of the Rcs phosphorelay system in the virulence of E. amylovora. Our results showed that rcsB, rcsC, rcsD and rcsBD mutants were unable to grow and cause disease in immature pear tissue. In an in vitro assay, both RcsB and RcsD positively regulated the production of amylovoran, whereas RcsC acted as a negative regulator of amylovoran production. Consistent with amylovoran production, in vitro gene expression results, using an amsG promoter–green fluorescent protein (GFP) fusion as a reporter, demonstrated that the amsG gene was not expressed in rcsB, rcsD and rcsBD mutants, but was highly expressed in the rcsC mutant. However, expression of the amsG gene in the rcsC mutant in planta was lower than that in the wild‐type (WT). Moreover, it was found that the master regulator of flagella biosynthesis, the flhDC gene, was up‐regulated in all rcs mutants, as reported previously in other enterobacteria (Andresen et al., 2007; Francez‐Charlot et al., 2003; Hinchliffe et al., 2008). However, rcs mutants showed an irregular and partially reduced swarming motility phenotype on swarming plates when compared with WT. Our results also showed that the Rcs phosphorelay system conferred some level of resistance to polymyxin B in E. amylovora. Together, these findings indicate that the Rcs phosphorelay system plays a major role in the virulence and survival of E. amylovora.

RESULTS

Identification of the Rcs phosphorelay system in E. amylovora and generation of the rcs mutant

On the basis of the whole genomic sequence of E. amylovora (http://www.sanger.ac.uk/Projects/E_amylovora/) and with the aid of a previously reported rcsB gene sequence of E. amylovora (Bereswill and Geider, 1997; GenBank accession number Y09848), the rcs genes within the genome of E. amylovora were identified (Fig. 1a). A blast search has revealed that RcsB is the most conserved, sharing 92% and 99% amino acid identity and 97% and 100% amino acid similarity with those of Es. coli and E. tasmaniensis, respectively. RcsC shares 61% and 88% amino acid identity and 76% and 93% amino acid similarity with those of Es. coli and E. tasmaniensis, respectively. RcsD is the least conserved, sharing 52% and 84% amino acid identity and 70% and 91% amino acid similarity with those of Es. coli and E. tasmaniensis, respectively. These results indicate that rcs genes in E. amylovora are more homologous to those of E. tasmaniensis than to those of Es. coli (Kube et al., 2008).

In order to study the function of the Rcs system in the virulence of E. amylovora, we generated four non‐polar insertional mutants within the rcs genes, including rcsB, rcsC, rcsD and rcsBD double mutants, using the λ‐Red cloning technique reported previously (Zhao et al., 2006). In these mutants, the endogenous copy of the rcs genes was disrupted and replaced with an antibiotic resistance cassette, as described in ‘Experimental procedures’.

Mutations in the Rcs phosphorelay system render E. amylovora non‐pathogenic

To determine the role of RcsB, RcsC and RcsD in E. amylovora virulence, we conducted a virulence assay of the rcs mutants on immature pear fruits, which have been routinely used to test the pathogenicity of E. amylovora (Zhao et al., 2005, 2006). Mutants and WT strains were inoculated on immature pears as described, and disease development was assessed. At 2 days following inoculation, E. amylovora WT strain Ea1189 produced water‐soaked symptoms in pears with visible bacterial ooze. Four days after inoculation, Ea1189‐inoculated immature pears showed necrotic lesions and bacterial ooze formation. After 8 days, the necrosis turned black with more ooze production at the inoculation site. In contrast, no disease symptoms were observed on immature pear fruits inoculated with the rcs mutants (Fig. 2a). In another experiment, we inoculated immature pear fruits with an unusually large number [1010 colony‐forming units (cfu)/mL] of rcs mutants, and yet no symptoms were observed (data not shown). These results indicate that the Rcs system is required for E. amylovora to induce disease.

Figure 2.

Figure 2

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Pathogenicity tests of Erwinia amylovora wild‐type (WT) strain, rcs mutants and complemented strains of rcs mutants on immature pears. (a) Symptoms caused by WT, ΔrcsB, ΔrcsC, ΔrcsD and ΔrcsBD mutants in immature pears. (b) Symptoms caused by WT and complementation strains of rcs mutants. 1, ΔrcsB (pWDP3); 2, ΔrcsD (pWDP3); 3, ΔrcsBD (pWDP3); 4, ΔrcsC (pWDP2). DPI, days post‐inoculation.

To ensure that a lack of disease symptoms of rcs mutants was specifically related to mutations in the rcs genes, we tested the virulence of rcs mutants complemented with cloned original rcs genes either in high‐copy (for rcsC gene) or low‐copy (for rcsBD genes) vectors. Complemented strains produced similar symptoms and comparable disease progression to the WT strain on immature pear fruits (Fig. 2b). As shown in Fig. 2b, black lesions appeared 4 days following infection and spread systemically in complemented rcs mutants by 7 days. These complemented strains also produced ooze, an indicator of amylovoran production. These results indicate that complementation of rcs mutants rescues their virulence. Interestingly, the rcsB, rcsD and rcsBD mutants cannot be complemented by a high‐copy‐number vector containing the rcsBD operon (data not shown). Our results clearly demonstrate that the Rcs phosphorelay system is required for virulence in E. amylovora.

The quantification of bacterial growth in infected immature pears was determined for both WT and rcs mutants at 4 days post‐inoculation. Disease symptoms caused by the WT strain on immature pear fruits were correlated with high levels of bacterial growth in pear tissues (Fig. 3a). In contrast with WT, the bacterial number of rcs mutants decreased rapidly following inoculation, representing an approximately 106‐fold reduction relative to that of WT (Fig. 3a). These results indicate that rcs mutants are unable to colonize and survive in immature pears. The growth of complemented strains of rcs mutants was also monitored, and these complemented strains were fully restored in growth when compared with the mutant strains by day 3 (Fig. 3b). In summary, these experiments suggest that the Rcs phosphorelay system plays a critical role in the virulence and survival of E. amylovora on pears.

Figure 3.

Figure 3

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Bacterial growth of Erwinia amylovora wild‐type (WT), rcs mutants and complemented strains of rcs mutants on immature pear fruits. (a) Growth of E. amylovora WT, ΔrcsB, ΔrcsC, ΔrcsD and ΔrcsBD during infection of immature pears. (b) Growth of WT and complemented strains of rcs mutants during infection of immature pears. 1, ΔrcsB (pWDP3); 2, ΔrcsD (pWDP3); 3, ΔrcsBD (pWDP3); 4, ΔrcsC (pWDP2). Growth of bacterial strains was monitored at 0, 1, 2, 3 and 4 days after inoculation. Data points represent the means of three replicates ± standard errors. Similar results were obtained in repeated independent experiments. CFU, colony‐forming unit.

The Rcs phosphorelay system regulates amylovoran biosynthesis in E. amylovora

To determine whether the rcs phosphorelay system of E. amylovora affects amylovoran biosynthesis, bacterial cells were grown in amylovoran‐inducing medium, and quantitative spectrophotometric assays were performed to determine the levels of amylovoran production in vitro. As shown in Fig. 4a, amylovoran production was almost undetectable in rcsB, rcsD and rcsBD mutants, as well as in the ams operon deletion mutant (as a control) (Zhao and Sundin, 2008). In contrast, the rcsC mutant produced a 6.2‐fold increase in amylovoran production compared with that of WT 3 days after inoculation (Fig. 4a).

Figure 4.

Figure 4

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(a) Amylovoran production of Erwinia amylovora wild‐type (WT), rcs mutants and complemented strains of rcs mutants in vitro. Bacterial strains were grown in MBMA (3 g KH2PO4, 7 g K2HPO4, 1 g (NH4)2SO4, 2 mL glycerol, 0.5 g citric acid, 0.03 g MgSO4) medium with 1% sorbitol for 72 h at 28 °C with shaking. The amount of amylovoran was measured with the cetylpyrimidinium chloride (CPC) assay and normalized to a cell density of unity. Data points represent the means of three replicates ± standard deviation. Similar results were obtained in three independent experiments. Δams, an amylovoran biosynthesis operon (amsA‐L) deletion mutant (Zhao and Sundin, 2008). (b) Expression of the amsG gene in WT and rcs mutants in vitro and in vivo. The green fluorescent protein (GFP) intensity in WT strain and rcs mutants containing the amsG promoter–GFP fusion plasmid was measured by flow cytometry. The experiment was repeated at least three times and similar results were obtained. Full line, in vivo on immature pear fruits; broken line, in vitro in MBMA plus 1% sorbitol. Count, number of bacterial cells; GFP‐A, green fluorescent protein absorbance.

To verify whether a lack or overproduction of amylovoran in vitro in rcs mutants was specifically related to mutations in rcs genes, amylovoran biosynthesis was measured by the complementation of rcs mutants. We found that amylovoran biosynthesis in the rcsC mutant was partially complemented by introducing the original rcsC gene in a high‐copy‐number vector. As shown in Fig. 4a, the complemented strain of the rcsC mutant produced less than one‐half of the amount of amylovoran relative to that of the rcsC mutant, but produced at a level of approximately twofold higher than that of WT. Complemented strains of rcsB, rcsD and rcsBD mutants by a low‐copy‐number vector containing the rcsBD operon also produced relatively higher levels of amylovoran, with optical density at 600 nm (OD600) values of 0.24, 0.32 and 0.30, respectively, compared with 0.16 for WT. These results indicate that the Rcs system regulates amylovoran biosynthesis either as a negative regulator (RcsC) or as a positive regulator (RcsBD) in vitro.

In order to correlate amylovoran production with amylovoran biosynthesis gene expression, the promoter of the amsG gene, the first gene in the amylovoran operon, was cloned in a promoterless GFP reporter vector. The GFP intensity was measured by flow cytometry in WT and rcs mutants containing the amsG promoter–GFP fusion plasmid in liquid medium. As shown in Fig. 4b, the amsG gene was expressed at a basal level in the WT strain, with a GFP intensity value of 1.70 (geometric mean), relative to the vector control (GFP value of about 1.50). The GFP intensity level of the amsG promoter in rcsB, rcsD and rcsBD mutants was measured to be 1.47, 1.48 and 1.46, respectively (Table 1). However, in the rcsC mutant, the GFP intensity (geometric mean of 5.64) of the amsG promoter was more than three times higher than that of WT (Table 1). The GFP intensity was also measured for both WT and rcs mutants containing the amsG promoter–GFP fusion following inoculation onto immature pear fruits for up to 2 days. The expression of amsG in rcsB, rcsD and rcsBD mutants in vivo was similar to that in vitro. The GFP intensity of the amsG gene in WT increased by more than twofold from in vitro (1.70) to in vivo (3.95). In contrast, amsG expression in the rcsC mutant was lower in vivo (2.75) than in vitro (5.64) (Fig. 4b; Table 1). These results indicate that the expression of amsG in vitro and in vivo is correlated with amylovoran production in WT and rcs mutants. Moreover, the amsG gene is induced in WT in planta.

Table 1.

Green fluorescent protein (GFP) fluorescence intensity in wild‐type (WT) and rcs mutants containing amsG and flhDC promoter–GFP fusion plasmid.

Bacterial strain GFP intensity in vitro (geometric mean ± standard deviation) GFP intensity in vivo (geometric mean ± standard deviation)
WT (pWDP4) 1.70 ± 0.04 3.95 ± 0.12
ΔrcsB (pWDP4) 1.47 ± 0.02 1.52 ± 0.02
ΔrcsC (pWDP4) 5.64 ± 0.05 2.75 ± 0.15
ΔrcsD (pWDP4) 1.48 ± 0.04 1.53 ± 0.05
ΔrcsBD (pWDP4) 1.46 ± 0.01 1.51 ± 0.03
WT (pSN1) 2.21 ± 0.08 ND
ΔrcsB (pSN1) 8.23 ± 0.24 ND
ΔrcsC (pSN1) 8.86 ± 0.62 ND
ΔrcsD (pSN1) 9.24 ± 0.15 ND
ΔrcsBD (pSN1) 9.66 ± 0.36 ND
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in vitro, liquid broth; in vivo, pear slices; ND, not determined.

The Rcs phosphorelay system negatively regulates flhDC gene expression, but swarming motility is only partially affected

In Es. coli, mutations in the Rcs system render Es. coli hypermotile (Francez‐Charlot et al., 2003; Takeda et al., 2001; Wang et al., 2007). Swarming motility was assessed for both WT and rcs mutants by inoculating bacterial cells on a swarming plate (0.3% agar) and measuring the diameter of the circle covered by bacterial cells 48 h after inoculation. The rcs mutants exhibited irregular and partially reduced motility compared with that of the WT strain, which showed a round circle of 3.01 cm in diameter 2 days following inoculation (data not shown). However, the diameters for the rcsB, rcsC, rcsD and rcsBD mutants were about 2.51 ± 0.17, 2.45 ± 0.2, 2.48 ± 0.28 and 2.43 ± 0.11 cm (longest part of the irregular movement), respectively. The irregular movement could only be partially restored in rcs mutants complemented with their corresponding genes (data not shown). These results indicate that mutation in the Rcs system in E. amylovora does not confer hypermotility as in Es. coli, but instead renders them less motile.

To determine whether the Rcs phosphorelay system regulates flhDC gene expression in E. amylovora, we also constructed a flhDC promoter–GFP fusion and introduced it into the WT and rcs mutants. The GFP intensity values of the flhDC promoter in rcsB, rcsC, rcsD and rcsBD mutants were about 8.6, 8.8, 9.2 and 9.6, respectively, approximately four times higher than that detected for WT (2.2) (Table 1). Together, these findings indicate that the flhDC gene is up‐regulated in rcs mutants when bacterial cells are grown on swarming plates, and that the Rcs system acts as a negative regulator for flhDC gene expression in E. amylovora.

The Rcs phosphorelay system in E. amylovora regulates resistance to polymyxin B

To determine whether the Rcs phosphorelay system regulates resistance to polymyxin B in E. amylovora, WT and rcs mutant strains were tested for their sensitivity to polymyxin B exposure (killing effect), and the survival number of each strain was counted by serial dilution plating. After 1 h of polymyxin B treatment, the survival rate of WT was about 3.2%, whereas the survival rates of rcs mutants were about 0.52%, 0.45%, 0.48% and 0.36% for rcsB, rcsC, rcsD and rcsBD mutants, respectively (Fig. 5). We also observed slower recovery [smaller colony forming on Luria–Bertani (LB) plates] of rcs mutants. The colony of the WT strain was visible after 16 h on LB plates, whereas it took about 25 h for rcs mutants to become visible. Complementation of rcsB, rcsC, rcsD and rcsBD mutants rescued the survival rates to 3.3%, 2.7%, 2.9% and 2.8%, respectively (Fig. 5). On polymyxin B treatment, complemented strains were visible 18 h after plating on LB plates. These results indicate that the Rcs system confers some level of resistance to polymyxin B in E. amylovora and plays a role in the survival of the pathogen.

Figure 5.

Figure 5

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Polymyxin B killing assay. Bacterial strains were grown overnight in Luria–Bertani (LB) broth and treated with an equal volume of polymyxin B (3 µg/mL) for 1 h. Surviving bacterial cells were counted by plating on LB plates. Data points represent the means of three replicates ± standard deviations. The experiment was repeated at least three times. CFU, colony‐forming unit.

DISCUSSION

Fire blight studies have contributed greatly to the elucidation of the genetic, molecular and physiological basis of E. amylovora pathogenesis. Early studies revealed that E. amylovora utilizes an essential virulence system—the hrp type III secretion system (T3SS)—to deliver effector proteins into host plants and to cause disease (Oh and Beer, 2005; Wei et al., 1992). In addition to T3SS and its associated effector proteins, other major virulence determinants identified in E. amylovora, contributing to pathogenesis and plant colonization, include the acidic extracellular polysaccharide amylovoran (Bernhard et al., 1993; Steinberger and Beer, 1988).

In this study, our results have demonstrated that the Rcs phosphorelay system not only regulates amylovoran biosynthesis, but also renders the pathogen more resistant to the antimicrobial peptide (AMP) polymyxin B. This suggests that the Rcs phosphorelay system controls one of the virulence factors in E. amylovora, and may also play a major role in the survival of E. amylovora.

TCST systems are complicated regulatory systems used by microorganisms to sense and respond to environmental signals which result in gene expression (Stock et al., 1990). As one of the most studied TCST systems, the Rcs phosphorelay system has been shown to contribute to the virulence of many pathogenic bacteria, including Salmonella and P. stewartii (Dominguez‐Bernal et al., 2004; Mouslim et al., 2004; Wehland et al., 1999). Previous studies have shown that RcsB contributes to virulence in E. amylovora (Kelm et al., 1997). Our studies confirm this and, for the first time, further prove that RcsC and RcsD are also essential for virulence. In another experiment, rcs mutants remained capable of inducing a hypersensitive response in tobacco (Y. Zhao, unpublished data), thus indicating that T3SS remains functional in rcs mutants. These findings suggest that the Rcs system contributes to virulence in E. amylovora in part by specifically regulating amylovoran biosynthesis. Interestingly, when the complementation of rcs mutants was attempted, we failed to complement rcsB, rcsD and rcsBD mutants when rcsBD genes were cloned in a high‐copy‐number vector (pGEM T‐easy), but succeeded when rcsBD genes were cloned in a low‐copy‐number vector (pWSK29). It is likely that an overabundance of unphosphorylated RcsB may provide dominant negative regulation by competing for binding to the amsG promoter or by acting as a repressor.

It seems that amylovoran biosynthesis in the E. amylovora WT strain is tightly controlled by the Rcs system. In this study, the rcsC mutant produced more amylovoran than did WT, and the amsG gene was highly expressed in vitro in this mutant. At first, it was difficult to explain these findings, as the mutant did not cause disease on immature pear fruits and no ooze was observed. However, similar observations have been reported in S. enterica, wherein mutations in rcsC resulted in constitutive capsule synthesis, and rcsC mutants were severely attenuated in virulence (Garcia‐Calderon et al., 2005). Furthermore, the P. stewartii strain that mutated in esaR, having a dominant effect on Rcs‐mediated activation of stewartan synthesis, constitutively synthesized stewartan, and was significantly reduced in virulence (Koutsoudis et al., 2006; Minogue et al., 2005).

The above observations have raised questions as to why mutations in rcsC result in the overproduction of EPS, and why the overproduction of EPS renders the pathogen less virulent, in spite of the fact that EPS is a virulence factor. One possibility is that RcsC may be needed to sense an unknown signal in vivo, and thus specifically phosphorylates RcsBD in vivo; yet RcsBD could be cross‐phosphorylated by cross‐talking kinases or small phospho‐donors in vitro. Recent studies have demonstrated that HKs have significant kinetic preferences for their cognate RRs to prevent unwanted cross‐talk in vivo. In other words, HKs have intrinsic abilities to recognize their cognate substrates in vivo (Skerker et al. 2005). Further studies have shown that a subset of co‐evolving residues around H‐boxes is sufficient to control the specificity of HKs (Skerker et al., 2008). Nevertheless, given the complex nature of the Rcs phosphorelay system, it has also been proposed that additional input signals may be sensed by RcsD in liquid medium (Majdalani and Gottesman, 2005). Therefore, another possible explanation is that these additional input signals may trigger the phosphorylation of RcsD or RcsB by cross‐phosphorylation, thus activating amylovoran biosynthesis in vitro in the rcsC mutant background, but not in vivo. Furthermore, it has been shown that the RcsC sensor kinase in Es. coli controls the expression of a regulon in response to growth on a solid surface (Ferrieres and Clarke, 2003). The expression of cpsB increases in an RcsC‐dependent manner when cells are grown on a solid surface (Ferrieres and Clarke, 2003). This suggests that a solid surface can serve as an environmental signal to activate the RcsC kinase. Our in vivo gene expression data support this, thus indicating that a solid surface can be an environmental factor in inducing the expression of the amsG gene, and this may depend on RcsC through sensing of this solid surface signal.

It has been reported that RcsC has a dual function in transferring a phosphoryl group to RcsB (as a kinase) or by removing a phosphate from RcsB (as a phosphatase) (Majdalani and Gottesman, 2005). The first evidence for a dual role of RcsC is provided by the expression of the promoter for another target gene of RcsB, the small regulatory RNA RprA (Majdalani et al., 2002, 2005). In Es. coli, the expression of rprA is off in the WT strain, but a null mutation of rcsC leads to higher levels of rprA expression. This suggests that the loss of RcsC (phosphatase) prevents the removal of the phosphate from RcsB, which may be cross‐phosphorylated from unknown sources, resulting in the activation of RcsB activity. Further evidence that RcsC acts as a phosphatase is provided by the observation that cells carrying a point mutation in rcsC137 in Es. coli constitutively express cps genes at high levels (Clarke et al., 2002). Recessive Salmonella rcsC mutants with constitutive activation of the Rcs system provide further evidence that RcsC has both positive and negative regulatory effects on EPS biosynthesis (Garcia‐Calderon et al., 2005). In this study, the amsG gene was slightly expressed in the WT strain in vitro, but expressed at higher levels in a liquid medium in the rcsC mutant in vitro, suggesting that the activation of RcsB from an unknown source in liquid medium is not dephosphorylated by RcsC.

In P. stewartii, the hypermucoid esaR mutant is highly attenuated in virulence (Minogue et al., 2005). It has been proposed that this bacterium may predominantly require an EPS‐free state to perform functions such as surface adhesion at low cell density (Herrera et al., 2008; Koutsoudis et al., 2006). Consequently, premature synthesis of EPS interferes with the normal disease process. Further evidence indicates that both spatiotemporal gene expression and EPS production greatly influence the degree of bacterial adhesion during in vitro biofilm formation and dissemination within a host plant (Herrera et al., 2008; Koutsoudis et al., 2006). In addition, reduced virulence of Salmonella rcsC mutants with constitutive activation of the Rcs system could be partially restored by a double mutation in either the cps or rcsA gene (Garcia‐Calderon et al., 2005). These results suggest that the overproduction of type I EPS is one of the reasons for virulence attenuation in Salmonella rcsC mutants. Thus, a similar scenario may be involved in the findings observed in this study, and, indeed, biofilm formation has recently been implied in the pathogenicity of E. amylovora (Koczan et al., 2008). However, one of the differences pertains to the fact that the rcsC mutant in E. amylovora has shown the overproduction of amylovoran in vitro, although this was not constitutive. Further studies are needed to determine the roles of amylovoran production, bacterial adhesion and biofilm formation in the virulence of E. amylovora.

Several environmental conditions have been reported to activate the Rcs phosphorelay system, including growth on a solid surface, desiccation and changes in osmolarity (Majdalani and Gottesman, 2005). One of the examples for the observed responses to solid surfaces is swarming motility. In both Es. coli and S. enterica, the RcsCDB phosphorelay system has been shown to negatively regulate swarming motility (Takeda et al., 2001; Wang et al., 2007). Mutations in rcsC, rcsD and rcsB lead to a hypermotile phenotype and higher expression of the flhDC operon, the master regulator of flagella biosynthesis. Products of flhDC genes, in turn, positively regulate second (such as fliA) or third (such as fliC) class genes for flagella biosynthesis (McCarter, 2006). This negative regulation of the flhDC operon by the Rcs system is in agreement with other supporting evidence for reduced motility under conditions inducing cps synthesis in Es. coli. Consistent with previous findings, our results demonstrate that the expression of the flhDC operon is higher in all rcs mutants than in WT in E. amylovora. Surprisingly, rcs mutants have shown irregular and reduced swarming motilities on swarming plates when compared with those of WT, exhibiting circular movement. It is worth noting here that we have identified several other TCST mutants that are either hypermotile (circle) or non‐motile on swarming plates. However, all of these mutants have shown even higher expression of the flhDC operon than that in rcs mutants (Y. Zhao, unpublished data). These data further indicate that, in addition to flagella, other factors also control swarming motility in E. amylovora. Irregular movements in rcs mutants may be attributed to the remodelling of bacterial surfaces during growth on a solid surface, as reported recently for Es. coli (Ferrieres and Clarke, 2003). Indeed, DNA microarray analysis of the rcsC mutant has revealed that more than 150 genes are controlled by rcsC, and 50% of these genes encode proteins that are either localized to the envelope of Es. coli or have activities affecting properties of the bacterial surface, e.g. the production of colanic acid. Further studies are needed to elucidate why Es. coli and E. amylovora rcs mutants show different motility behaviour, even though flhDC genes are up‐regulated in both cases.

Endogenous AMPs produced by plants are known to play important roles in the host defence system and innate immunity (Oard and Enright, 2006). Most AMPs are cationic and are believed to interact with their primary target, the cell membrane. They are reported to kill bacteria by binding to lipopolysaccharides (LPSs) through ionic bonds with negatively charged phosphoryl groups of the lipid A (Rosenfeld and Shai, 2006). Several TCST systems, including PhoP/Q, PmrA/B and the Rcs system, have been reported to regulate resistance to AMPs, including the peptide antibiotic polymyxin B (Erickson and Detweiler, 2006; Hancock and McPhee, 2005; Lee et al., 2004). These regulatory factors are required for the modification of the LPS structure, leading to a reduction of negative charges along the surface, thus altering interactions with AMPs. Many TCST system‐targeted genes, including ugd and pbgP, are responsible for mediating the modifications of LPS necessary for resistance to polymyxin B (Kato et al., 2007, Mouslim and Groisman, 2003). It has been shown that the rcsC gene in S. typhimurium regulates the expression of ugd, a gene that is responsible for the synthesis and incorporation of l‐aminoarabinose into LPS to induce resistance to polymyxin B (Mouslim and Groisman, 2003). Our results clearly indicate that the Rcs system in E. amylovora confers some level of resistance to polymyxin B. It is probable that the Rcs system in E. amylovora also affects properties of the bacterial surface, which leads to resistance to polymyxin B. Although the mechanism remains unknown, it is tempting to speculate that the Rcs phosphorelay system increases E. amylovora resistance to AMPs, thus playing a role in the survival of the pathogen during the infection of plant tissues. It will be interesting to investigate whether there are any cross‐talks between different TCST systems in E. amylovora that regulate resistance to AMPs, as reported in other enterobacteria.

In summary, we have confirmed previous reports and further demonstrated that the Rcs system is essential for E. amylovora to cause disease. Although the Rcs systems in Es. coli and E. amylovora both control the biosynthesis of extracellular polysaccharide and negatively regulate the flhDC operon, the phenotypes of rcs mutants in these two bacteria are quite different. The Rcs system does not contribute to the virulence of Es. coli, but mutations of the rcs system render it hypermotile. However, the Rcs system does contribute to the virulence of E. amylovora, but mutations of the rcs system only partially affect motility. It is tempting to speculate that, during evolution, the Rcs system maintains its basic function to regulate gene expression; however, it may have adapted to specific host niches and sense specific host signals to control many important cellular processes, including motility and virulence. Our future work will focus on the identification of genes that are regulated by the Rcs system globally, the study of how the Rcs system interacts with other TCST systems, such as PhoP/Q and PmrA/B, and the determination of the roles of other components of the Rcs system, such as rcsF (Majdalani et al., 2005), during the infection of host plants.

EXPERIMENTAL PROCEDURES

Bacterial strains and culture media

The bacterial strains and plasmids used in this study are listed in Table 2. LB medium is used routinely for culturing E. amylovora. When necessary, the following antibiotics were added to the medium: 50 µg/mL kanamycin (Km) and 100 µg/mL ampicillin (Ap). Amylovoran production was determined by the growth of bacteria in MBMA medium (3 g KH2PO4, 7 g K2HPO4, 1 g (NH4)2SO4, 2 mL glycerol, 0.5 g citric acid, 0.03 g MgSO4) plus 1% sorbitol (Bellemann et al., 1994).

Table 2.

Bacterial strains, plasmids and primers used in this study.

Strain, plasmid or primer Relevant characters or sequences (5′–3′)* Reference or source
Strain
E. amylovora
Ea1189 Wild‐type, isolated from apple Burse et al. (2004)
Z3205ΔrcsB rcsB::Km; KmR‐insertional mutant of rcsB of Ea1189, KmR This study
Z3207ΔrcsC rcsC::Km; KmR‐insertional mutant of rcsC of Ea1189, KmR This study
Z3206ΔrcsD rcsD::Km; KmR‐insertional mutant of rcsD of Ea1189, KmR This study
Z3205‐06ΔrcsBD rcsBD::Km; KmR‐insertional mutant of rcsDB of Ea1189, KmR This study
Z0118Δams KmR‐insertional mutant of ams operon (15.8 kb) of Ea1189, KmR Zhao and Sundin (2008)
Es. Coli
DH10B F− mcrA Δ(mrr‐hsdRMS‐mcrBC) Φ80lacZΔM15 ΔlacX74 recA1 endA1 araΔ139 Δ(ara, leu)7697 galU galK λ—rpsL (StrR) nupG Invitrogen (Carlsbad, CA, USA)
Plasmid
pKD46 ApR, PBAD gam bet exo pSC101 oriTS Datsenko and Wanner (2000)
pKD13 KmR, FRT cat FRT PS1 PS2 oriR6 K rgbN Datsenko and Wanner (2000)
pGEM® T‐easy ApR, PCR cloning vector Promega
pFPV25 ApR, GFP‐based promoter trap vector containing a promoterless gfpmut3a gene Valdivia & Falkow (1997)
pWSK29 ApR; cloning vector, low copy number Wang and Kushner (1991)
pWDP1 4.54‐kb PCR fragment containing rcsBD gene in pGEM T‐easy vector This study
pWDP2 3.88‐kb PCR fragment containing rcsC gene in pGEM T‐easy vector This study
pWDP3 4.54‐kb EcoRI‐BamHI fragment containing rcsBD gene in pWSK29 This study
pWDP4 721‐bp KpnI‐XbaI DNA fragment containing promoter sequence of amsG gene in pFPV25 This study
pSN1 906‐bp EcoRI‐BamHI DNA fragment containing promoter sequence of flhD gene in pFPV25 This study
Primer†
 B3205F ATGTTGCCATATAAATTTCCGCTAACTTCCGGCAATGTAACACGATTTTTGCGATTGTGTAGGCTGGAGCT
 B3205R TTATTGGTTACCTTGCTGCAGGAGATCTTTGACATAAGTGTCAACTTCACATTCCGGGGATCCGTCGACC
 B3206F GTGAAGTTGACACTTATGTCAAAGATCTCCTGCAGCAAGGTAACCAATAAGCGATTGTGTAGGCTGGAGCT
 B3206R TTATTTATCTACCGGCGTCATGCTTACTGATGACAGGTAGTTAAGCAAAGATTCCGGGGATCCGTCGACC
 B3207F ATGGCCGGGATAACTGGCTTTTTCTGCCTCCCGGCGCTACTGTATCGGCAGCGATTGTGTAGGCTGGAGCT
 B3207R TTACCGCAACCTGTTACTTAGCGCTTCCCGGCTCTTTCTTACCCGCCCGGATTCCGGGGATCCGTCGACC
 GFP1 ATGAGTAAAGGAGAAGAACTTTTCAC
 GFP2 TTATTTGTATAGTTCATCCATGC
 AmsG1 CGGGGTACCGTATCGCTTAAAGGGGAAACG (KpnI)
 AmsG2 CTAGTCTAGACACCTGGAAAGCCATTAATCA (XbaI)
 flhD1 CCGGAATTCCGTTGTTGCCGATGCTAATA (EcoRI)
 flhD2 CGCGGATCCCAATACCCAGGCGAAACATC (BamHI)
 RcsC1 GATCAAGGTAACCGAGCGTAAC
 RcsC2 AGCTTAACCGCAGCATTAAAAC
 RcsBD1 CCGGAATTCCTAGCACAATTCACAAGGTTGG (EcoRI)
 RcsBD2 CGCGGATCCCTCCTAATGAACTGCCGCTACT (BamHI)
 Km1 CAGTCATAGCCGAATAGCCT
 Km2 CGGTGCCCTGAATGAACTGC
 Z3205F GCCAACGCCCGACTGATTACAAG
 Z3206R GCCGGGAAGCGCTAAGTAACAGGT
 Z3207F GTTCGCCGCTAAGCCGTGGTCA
 Z3207R GAGAAAATCAGCGCGGGTGGTTAC
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*

KmR, ApR, kanamycin and ampicillin resistance, respectively.

†

Italic nucleotides are restriction sites added and the restriction enzymes are indicated at the end of the primers.

DNA manipulation and sequence analysis

Plasmid DNA purification, polymerase chain reaction (PCR) amplification of genes, isolation of fragments from agarose gels, cloning, and restriction enzyme digestion and T4 DNA ligation were performed using standard molecular procedures (Sambrook et al., 1989). DNA sequencing was performed at the Keck Center for Functional and Comparative Genomics (University of Illinois at Urbana‐Champaign, IL, USA). Sequence management and contig assembly were conducted using Sequencher 4.7 software. Database searches were conducted using the blast programs at the National Center for Biotechnology Information (NCBI) (http://www.ncbi.nlm.nih.gov/BLAST) (Altschul et al., 1997).

Construction of rcs mutants in E. amylovora by λ‐Red recombinase cloning

Erwinia amylovora stable mutants were generated using λ phage recombinases, as described previously (Datsenko and Wanner, 2000; Zhao et al., 2006). Briefly, E. amylovora Ea1189 was transformed with plasmid pKD46 expressing recombinases redα, redβ and redγ. The transformant Ea1189 (pKD46) was grown overnight at 28 °C, reinoculated in LB broth containing 0.1% arabinose and grown to exponential phase OD600 = 0.8. Cells were collected, made electrocomponent and stored at –80 °C. Recombination fragments consisting of a kan gene with its own promoter, flanked by a 50‐nucleotide homology arm, were generated by PCR using plasmid pKD13 as a template. For the primers used for the generation of rcs mutants, the 5′ end of each primer contained 50‐nucleotide homology arms corresponding to the first and last 50 bases of the corresponding gene(s), whereas the remaining 21 or 20 nucleotides of each primer at the 3′ end were designed to amplify the kan cassette from pKD13. For the generation of the rcsB, rcsC, rcsD and rcsBD mutants, the primer pairs B3205F‐R, B3207F‐R, B3206F‐R and B3205F‐B3206R were used to amplify kan gene fragments from the pKD13 vector. To confirm rcsB, rcsC, rcsD and rcsBD mutants by PCR, primers Z3205F, Z3206R, Z3207F‐R and internal primer pairs km1 and km2 of the kan gene were used. PCR products were gel purified using a gel purification kit from Promega (Madison, WI, USA). Electroporation was performed according to standard protocols, and transformants were plated on LB with Ap and Km. For the resulting mutants, the majority of the coding region of each gene was replaced by the KmR marker, except for the first and last 50 nucleotides. The rcsB, rcsC, rcsD and rcsBD mutants were designated as Z3205ΔrcsB, Z3207ΔrcsC, Z3206ΔrcsD and Z3205‐06ΔrcsBD, respectively.

Cloning rcs genes for complementation of the rcs mutants

For complementation of the rcs mutants, the regulatory and gene sequences of the rcsBD operon and rcsC open reading frame (ORF) were used to design primers to amplify fragments of the gene and/or operon and their promoter sequences. Primer pairs rcsBD1‐rcsBD2, without restriction sites, and rcsC1‐rcsC2 were used to amplify 4.54‐kb and 3.88‐kb DNA fragments from the E. amylovora WT strain, which contain upstream and downstream sequences of the rcsBD operon and the rcsC gene, respectively. The two PCR fragments were cloned into pGEM T‐easy vector through A–T ligation. The final plasmids were designated pWDP1 and pWDP2, respectively.

The rcsBD operon was also cloned into a low‐copy‐number vector pWSK29. Primer pair rcsBD1‐rcsBD2, containing EcoRI and BamHI restriction sites, was used to amplify the rcsBD operon and its flanking sequences. Following amplification, DNA fragments and the vector were both digested by EcoRI and BamHI, and ligated together. The final plasmid was designated as pWDP3. All plasmids were introduced into E. amylovora strain by electroporation. Transformants were selected on LB plates supplemented with Km and Ap. Their genotypes were confirmed by both enzymatic digestion and sequencing.

Construction of promoter–GFP fusions for gene expression assays

For gene expression assays, flanking sequences of the amsG and flhD ORFs were used to design primers to amplify DNA fragments. Primer pairs amsG1‐amsG2 and flhD1‐flhD2 with restriction sites were used to amplify 721‐bp and 906‐bp DNA fragments from the E. amylovora WT strain, containing promoter sequences of amsG and flhD genes, respectively. PCR products and the promoter trapping vector pFPV25 were both digested with BamHI and EcoRI for the amsG gene and with PstI and KpnI for the flhDC gene, respectively. The resulting fragments were gel purified, ligated together and cloned to the upstream of the promoterless gfp gene. The final plasmids were designated as pWDP4 and pSN1 for the amsG and flhD genes, respectively, and were confirmed by restriction enzyme digestion and sequencing.

Immature pear virulence assays

For E. amylovora WT and mutant strains, bacterial suspensions were grown overnight in LB broth, harvested by centrifugation and resuspended in 0.5 × sterile phosphate‐buffered saline (PBS) with bacterial cells adjusted to concentrations of ~1 × 103 to 1 × 104 cfu/µL (OD600 = 0.1 and then diluted 100 times) in PBS. Immature fruits of pear (Pyrus communis L. cv. ‘Bartlett’) were surface sterilized and pricked with a sterile needle, as described previously (Zhao et al., 2005, 2006). Wounded fruits were inoculated with 2 µL of cell suspensions, and incubated in a humidified chamber at 26 °C. Symptoms were recorded at 2, 4, 6, 7 and 8 days post‐inoculation.

For bacterial population studies, fruit tissues surrounding the inoculation site were excised using a #4 cork borer, as described previously (Zhao et al., 2005, 2006), and homogenized in 0.5 mL of 0.5 × PBS. Bacterial growth was monitored by dilution plating of the ground material on LB medium amended with the appropriate antibiotics. For each strain tested, fruits were assayed in triplicate, and each experiment was repeated at least twice.

Cetylpyrimidinium chloride (CPC) assay for the determination of amylovoran concentration

The amylovoran concentration in supernatants of bacterial cultures was determined quantitatively by a turbidity assay with CPC, as described previously (Bellemann et al., 1994; Hildebrand et al., 2006). Briefly, for E. amylovora WT, mutants and complemented strains, bacterial suspensions were grown overnight in LB broth with or without appropriate antibiotics, harvested by centrifugation and washed with PBS three times. After the final wash, the bacterial pellet was resuspended in 200 µL of PBS. A total of 100 µL of bacterial suspension was inoculated into 10 mL of MBMA medium with 1% sorbitol. One millilitre of bacterial cells was pelleted 2–3 days after inoculation at 28 °C with shaking. Following centrifugation, 50 µL of CPC at 50 mg/mL was added to 1 mL of supernatant. After 10 min of incubation at room temperature, the amylovoran concentration was determined by measuring OD600 turbidity. The final concentration of amylovoran production was normalized for a cell density of 1.0. For each strain tested, the experiment was repeated at least three times.

Swarming motility assay

For E. amylovora WT and mutant strains, bacterial suspensions were grown overnight in LB broth with or without appropriate antibiotics, harvested by centrifugation, washed with PBS once and resuspended in 200 µL of PBS. Then, bacterial suspensions were diluted 10 times in water, and 5 µL of the diluted bacterial suspension was plated onto the centre of swarming agar plates (10 g tryptone, 5 g NaCl, 3 g agar per litre), as described previously (Hildebrand et al., 2006; Skerker et al., 2005). Swarming diameters were determined following incubation at 28 °C for up to 3 days. The experiments were repeated at least three times.

Polymyxin B killing assay

Bacterial strains were grown overnight in LB broth without antibiotics, collected by centrifugation at 3220 g for 10 min and washed with PBS three times. The pellets were resuspended in 200 µL of PBS, and 10‐fold serial dilutions of bacterial suspensions were made on 96‐well plates. Equal volumes of polymyxin B (3 µg/mL) were added to 100 µL of bacterial suspensions, and incubated at room temperature for 1 h. Then, the bacterial numbers for each dilution were counted by plating on LB medium with or without appropriate antibiotics. The percentage survival was calculated by dividing polymyxin B‐treated versus untreated cfus (Erickson and Detweiler, 2006). Dilution of each strain was carried out in triplicate. The experiment was repeated at least three times.

GFP reporter gene assay by flow cytometry

The FACSCanto flow cytometer (BD Biosciences, San Jose, CA, USA) was used to monitor the GFP intensity of WT and mutant strains containing the corresponding promoter–GFP construct (Jacobi et al., 1998). For in vitro amsG gene expression, WT and rcs mutants containing the amsG promoter–GFP fusion plasmid were grown in LB overnight, harvested and resuspended in water. Bacterial suspensions were re‐inoculated in MBMA broth with 1% sorbitol and grown at 28 °C with shaking for up to 3 days. Bacterial cultures were then harvested by centrifugation, washed once with PBS and then resuspended in PBS for flow cytometry assay. For in vivo gene expression assay, bacterial strains were inoculated onto immature pear fruits by cutting the fruit in half, and adding the bacterial suspensions directly to the surface. For rcs mutants, several pear slices were inoculated in order to recover sufficient bacterial cells for flow cytometry assay. After incubation at 26 °C for up to 2 days, bacterial cells were collected by either washing or centrifugation, washed twice with PBS and resuspended in PBS for flow cytometry assay.

For flhDC gene expression, WT and rcs mutants containing the flhDC promoter–GFP fusion plasmid were grown on swarming plates as described above. Following incubation for 2 days at 28 °C, bacterial cells were collected, washed and resuspended in PBS for flow cytometry assay. Flow cytometry was performed on LSRII 10‐parameter multilaser analysers (BD Biosciences). For both cases, data were collected for a total of 100 000 events and statistically analysed by gating using the flow cytometry software FCS Express V3 (De Novo Software, Los Angeles, CA, USA). A geometric mean was calculated for each sample. Each treatment was performed in triplicate and each experiment was repeated three times.

ACKNOWLEDGEMENTS

We would like to thank Sridevi Nakka for constructing the flhDC promoter–GFP fusion plasmid. We would also like to thank Dr George W. Sundin and Gayle McGhee (Michigan State University, East Lansing, MI, USA) and Dr Virginia Stockwell (Oregon State University, Corvallis, OR, USA) for providing the immature pear fruits used in this experiment. We are grateful to Dr S. Kushner (University of Georgia, Athens, GA, USA) and Dr R. Valdivia (Duke University, Durham, NC, USA) for providing the pWSK29 and pFPV25 plasmids, respectively. We are also grateful to Dr Steve Beer (Cornell University and the Sanger Institute, Ithaca, NY, USA) for the genome sequence of E. amylovora strain Ea273. This work was funded by the United States Department of Agriculture and Illinois Agricultural Experiment Station.

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