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. 2021 Nov 27;23(3):355–369. doi: 10.1111/mpp.13164

The RavA/VemR two‐component system plays vital regulatory roles in the motility and virulence of Xanthomonas campestris

Maojuan Lin 1, Kejian Wu 1, Zhaohong Zhan 1, Duo Mi 1, Yingying Xia 1, Xiaolei Niu 1, Shipeng Feng 1, Yinhua Chen 1, Chaozu He 1, Jun Tao 1, Chunxia Li 1,✉
PMCID: PMC8828458  PMID: 34837306

Abstract

Xanthomonas campestris pv. campestris (Xcc) can cause black rot in cruciferous plants worldwide. Two‐component systems (TCSs) are key for bacterial adaptation to various environments, including hosts. VemR is a TCS response regulator and crucial for Xcc motility and virulence. Here, we report that RavA is the cognate histidine kinase (HK) of VemR and elucidate the signalling pathway by which VemR regulates Xcc motility and virulence. Genetic analysis showed that VemR is epistatic to RavA. Using bacterial two‐hybrid experiments and pull‐down and phosphorylation assays, we found that RavA can interact with and phosphorylate VemR, suggesting that RavA is the cognate HK of VemR. In addition, we found that RpoN2 and FleQ are epistatic to VemR in regulating bacterial motility and virulence. In vivo and in vitro experiments demonstrated that VemR interacts with FleQ but not with RpoN2. RavA/VemR regulates the expression of the flagellin‐encoding gene fliC by activating the transcription of the rpoN2‐vemR‐fleQ and flhF‐fleN‐fliA operons. In summary, our data show that the RavA/VemR TCS regulates FleQ activity and thus influences the expression of motility‐related genes, thereby affecting Xcc motility and virulence. The identification of this novel signalling pathway will deepen our understanding of Xcc–plant interactions.

Keywords: motility, RavA/VemR, signalling pathway, two‐component system, virulence

RavA phosphorylates VemR and might affect the interaction of VemR with FleQ, which regulates Xanthomonas campestris pv. campestris motility and virulence via FliA and Clp.

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1. INTRODUCTION

Xanthomonas campestris pv. campestris (Xcc), a gram‐negative bacterium with a unipolar flagellum, can cause black rot in cruciferous plants (Vicente & Holub, 2013). The polar flagellum of Xcc allows the bacterium to swim in liquid environments and gather on semisolid surfaces (Bardy et al., 2003). Flagella play multiple roles in the infection of host plants, including in bacterial motility, biofilm formation, biological attachment, and colonization of host tissues (Li & Wang, 2011; Malamud et al., 2011; Ottemann & Miller, 1997; Viducic et al., 2017). In addition, flagella are pathogen‐associated molecular patterns that can induce the host innate immune response (Zipfel et al., 2004). Therefore, flagella are considered potential targets of anti‐infection drugs. Bacterial flagella are molecular machines driven by electric motors that facilitate the rotation of long curved filaments (Berg, 2003). In Xcc, flagellum biosynthesis is controlled by a cascade of three‐level transcriptional regulation systems (Figure S1) involving the housekeeping sigma factor σ70 and two alternative sigma factors, σ54 and σ28 (Hu et al., 2005; Yang et al., 2009). σ70 (RpoD) regulates the expression of class I genes such as σ54 (rpoN2), flgM, and fleQ. Class I proteins are the main regulators of class II genes, such as F‐T3SS, rod and hook genes, and σ28 (fliA). FlgM, an anti‐sigma factor, binds FliA (σ28) and inhibits its activity. Free FliA initiates the expression of class III genes (including flagellin‐related genes) (Al Mamun et al., 1996; Liu & Matsumura, 1994; Yang et al., 2009).

In Xanthomonas, there are two rpoN genes encoding σ54, named rpoN1 and rpoN2. In an rpoN2 deletion mutant, the expression of flgG, flhB, and fliC was significantly down‐regulated (Hu et al., 2005). RpoN2 is thus closely related to the transcription of flagellum biosynthesis genes (Li, Wu, et al., 2020). FleQ is the only key activator of all RpoN‐dependent flagellar promoters, and the FleQ protein has a central RpoN interaction domain (G140–L357) (pfam00158). It is speculated that the six FleQ‐dependent promoters of flhF, fliL, fliF, flgG, fliQ, and flgB also have RpoN‐dependent sequences (Hu et al., 2005). In another transcriptional regulation system, flhF, fleN, and fliA form an operon whose expression depends on the alternative sigma factor FliA (Hu et al., 2005). The polymer of flagellin subunits encoded by the fliC gene constitutes the flagellar filaments extending outward from the cell body (Chilcott & Hughes, 2000; Macnab, 1992). After mutation of fliC, many bacteria were shown to lose their flagella and motility (Macnab, 1992).

In addition to the above genes, VemR, RavA, and Clp also play important roles in flagellum synthesis and bacterial movement. rpoN2, vemR, and fleQ form a transcriptional operon (Wu et al., 2019). RpoN2 and FleQ regulate the expression of flagellum biosynthesis genes in Xcc strain Xc17 (Yang et al., 2009), and VemR regulates cell motility in Xcc 8004 (Tao & He, 2010). Bacterial virulence, extracellular polysaccharide (EPS) production, and motility were decreased after vemR mutation in Xcc 8004. Recently, it was found that VemR interacts with RpoN2 to regulate the transcription of flgG, influencing the motility of Xanthomonas citri subsp. citri (Wu et al., 2019). σ54 and RNA polymerase can form a stable complex to block transcription initiation. To start transcription, the complex must interact with transcriptional activators exhibiting nucleotide hydrolysis activities (Studholme & Dixon, 2003). The sensory modules of these activators are usually located in their N‐terminal regions, typically adopting a response regulator receiver (REC) domain fold (Shingler, 2011). Coincidentally, VemR has only a REC domain. As FleQ is the transcriptional activator of all RpoN2 (σ54)‐dependent flagellar promoters (Hu et al., 2005), VemR may be the regulator of FleQ activity and may thus indirectly regulate RpoN2 function.

Signal transduction via two‐component systems (TCSs) is a common mechanism by which prokaryotes sense and respond to environmental stimuli. A typical TCS consists of two proteins: a histidine kinase (HK) and a response regulator (RR) (Buschiazzo & Trajtenberg, 2019). The HK autophosphorylates its histidine residues via ATP hydrolysis and then transfers the phosphate group to the aspartate residues of the cognate RR to trigger adaptive responses (Stock et al., 2000). In Xanthomonas, five TCSs, namely, RpfC/RpfG (Stock et al., 2000), RavA/RavR (Tao et al., 2014), HpaS/HrpG (Li et al., 2014), HpaS/HpaR2 (Li et al., 2014), and HpaS/VemR (Li, Wang, et al., 2020), have been identified. RpfC/RpfG positively regulates the expression of extracellular enzymes and EPS, and negatively regulates intercellular c‐di‐GMP signal transduction (Stock et al., 2000). RavA phosphorylates RavR, and phosphorylated RavR reduces the level of free c‐di‐GMP in cells and increases EPS production, cell motility, and virulence (Tao et al., 2014). There is a physical interaction between HpaS and HrpG, and hpaS deletion reduces the HrpG phosphorylation level. Deletion of hpaS or hrpG eliminates bacterial virulence and the hypersensitive response (HR) (Li et al., 2014). Similar to HpaS/HrpG, hpaS deletion reduces the phosphorylation level of the orphan RR VemR in vivo, and the HpaS/VemR TCS controls the swimming behaviour of Xcc (Li, Wang, et al., 2020). According to these studies, HpaS phosphorylates at least three RRs (HrpG, VemR, and HpaR2), but how and when HpaS interacts with these RRs needs to be determined.

In addition, Clp is a key regulator of flagellum biosynthesis, EPS production, extracellular enzymes, and the expression of the Hrp system (de Crecy‐Lagard et al., 1990; Hsiao et al., 2005; Hsiao & Tseng, 2002; Lee et al., 2003). Clp functions as a c‐di‐GMP receptor. On binding to c‐di‐GMP, Clp changes its conformation, which eliminates the interaction between Clp and its target gene promoters (Liu et al., 2013). Clp also up‐regulates the expression of the flagellin‐encoding gene fliC and positively regulates Xcc motility (Lee et al., 2003). However, how Clp regulates fliC expression remains unclear.

Here, we studied the roles of VemR, RpoN2, and FleQ in Xcc motility and virulence. We also tested the possible interaction between VemR and RpoN2 or FleQ. In addition, we demonstrated that RavA and VemR form a TCS. We also explored the relationship among RavA, RavR, VemR, and Clp and the mechanism by which RavA/VemR regulates bacterial motility. Our results shed light on the complex regulation of Xcc motility and virulence by TCSs.

2. RESULTS

2.1. RpoN2 and FleQ function downstream of VemR to negatively regulate Xcc virulence

Our previous studies have shown that VemR, but not RpoN2 or FleQ, was necessary for Xcc 8004 virulence (Tao & He, 2010). However, RpoN2 was recently reported to be vital for the virulence of the Xc17 strain (Li, Wang, et al., 2020). To further determine the roles of rpoN2, vemR, and fleQ in the interaction between Xcc and host plants, we constructed single (ΔrpoN2, ΔvemR, ΔfleQ), double (ΔrpoN2/ΔvemR, ΔvemR/ΔfleQ, ΔrpoN2/ΔfleQ), and triple (ΔrpoN2/ΔvemR/ΔfleQ) deletion mutants in Xcc strain Δlac8 (Δlac8, a derivative strain of Xcc 8004, has no β‐galactosidase activity but has the same other phenotypes as Xcc 8004, including virulence, motility, EPS production, biofilm formation, and growth rate; hereafter, we refer to Δlac8 as the wild‐type [WT] strain) (Wang et al., 2018). Then, we compared the virulence of these strains on Brassica oleracea ‘Wenxin'. At 14 days after inoculation, the average lesion length of the WT strain was c.30.7 mm. The virulence of ΔvemR was completely lost but could be restored by complementation with vemR. In contrast, the lesion length of the ΔrpoN2 or ΔfleQ strain was not significantly different from that of the WT strain. Furthermore, the average lesion lengths of the ΔrpoN2/ΔvemR and ΔvemR/ΔfleQ double mutants were approximately 13.7 and 7.7 mm, respectively, demonstrating that their infection abilities were between those of ΔvemR and ΔrpoN2 or ΔfleQ. The virulence of the double mutant ΔrpoN2/ΔfleQ and the triple mutant ΔrpoN2/ΔvemR/ΔfleQ was similar to that of ΔrpoN2 or ΔfleQ (Figure 1a,b). These data indicate that rpoN2 and fleQ are epistatic to vemR and act as negative regulators in the VemR‐mediated signalling pathway.

FIGURE 1.

FIGURE 1

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RpoN2/FleQ and VemR antagonistically regulate virulence and motility in Xanthomonas campestris pv. campestris (Xcc). (a) Virulence phenotypes of the wild type (WT), rpoN2, vemR and fleQ single, double, and triple deletion mutants, and the corresponding complementation strains 14 days postinoculation (dpi) on broccoli cv. Wenxin leaves. (b) Lengths of the lesions on the 14 dpi‐infected leaves caused by the above Xcc strains, as shown in (a). (c) Swarming zones of the above Xcc strains inoculated on swarming plates (NY medium containing 2% glucose and 0.6% agar) at 28°C for 3 days. (d) Average diameters of the swarming swimming zones. (e) Swarming zones of the abovementioned strains on swimming plates (0.03% Bacto peptone, 0.03% yeast extract, and 0.28% agar incubated at 28°C for 5 days). (f) Average diameters of the swimming zones. The values given are the means ± SD from triplicate experiments. C indicates the indicated gene(s) complementation strain of a given mutant (including the indicated gene(s) deletion). As our focus was the relationship between vemR and fleQ/rpoN2, we only analysed the significance of the difference between ΔvemR and the double mutants ΔvemR/ΔfleQ and ΔvemR/ΔrpoN2. **p < 0.01, ***p < 0.001

2.2. RpoN2 and FleQ are epistatic to VemR in the regulation of Xcc motility

RpoN2 and FleQ positively regulate the expression of flagellum biosynthesis genes in the Xc17 strain (Yang et al., 2009), and VemR is indispensable for the regulation of flagellar motility by the Xcc 8004 strain (Tao & He, 2010). In Pseudomonas aeruginosa, FleQ binds to the σ54 factor RpoN to regulate flagellum expression (Dasgupta et al., 2002). In X. citri pv. citri, VemR regulates the transcription of the flagellar gene flgG by interacting with RpoN2 (Wu et al., 2019). Therefore, we wanted to clarify the mechanism by which the rpoN2‐vemR‐fleQ operon regulates Xcc motility.

Two types of motilities, swarming and swimming, are key for bacterial movement. Swarming, which involves pilus‐dependent population migration, is an important factor for Xanthomonas virulence (Ryan et al., 2007). Compared with the WT strain, ΔvemR exhibited significantly reduced swarming (approximately 60%). In contrast, the swarming of the ΔrpoN2 or ΔfleQ strain was not different from that of the WT strain. Compared with the WT strain, the swarming abilities of ΔrpoN2/ΔvemR and ΔfleQ/ΔvemR decreased by 33% and 16%, respectively, but those of ΔrpoN2/ΔfleQ and ΔrpoN2/ΔvemR/ΔfleQ increased by 11% and 9%, respectively. These phenotypes could be restored by gene complementation (Figure 1c,d). The above results revealed that VemR positively regulated pilus‐dependent population motility, while RpoN2 and FleQ had antagonistic effects. These data are consistent with the virulence phenotypes, suggesting that Xcc virulence could be affected by this colony movement.

For swimming, the flagellum‐dependent motility of ΔvemR was 224% higher than that of the WT strain. In contrast, this type of motility was markedly reduced after rpoN2 or fleQ deletion. When rpoN2/vemR, vemR/fleQ, rpoN2/fleQ, and rpoN2/vemR/fleQ were deleted, their swimming motility was consistent with the phenotype of the rpoN2 or fleQ mutant (i.e., the migration was greatly decreased). All changes in swimming behaviour could be recovered by complementation (Figure 1e,f). These results suggest that VemR is a negative regulator of flagellum‐based swimming, but RpoN2 and FleQ act as positive regulators, which is contrary to what was observed in the swarming and virulence results. In summary, RpoN2 and FleQ are epistatic to VemR in regulating Xcc motility and virulence.

2.3. VemR interacts with FleQ and regulates its own expression

Our data above show that rpoN2 and fleQ are epistatic to vemR in regulating bacterial virulence and motility, indicating that the expression of VemR could be regulated by RpoN2 and FleQ or that VemR could interact with FleQ and/or RpoN2 to affect their functions. Therefore, we first analysed rpoN2‐vemR‐fleQ operon expression levels in the WT, ΔrpoN2, ΔvemR, and ΔfleQ strains. The rpoN2 promoter was fused to the β‐glucuronidase (gusA) gene to construct the rpoN2p‐gusA plasmid. After transforming this construct into the WT, ΔrpoN2, ΔvemR, and ΔfleQ strains, the GUS activity of the transformants was analysed. The intensity of the blue GUS staining of ΔvemR was stronger than that of WT (Figure S2), and the quantification of GUS activity showed that the rpoN2 promoter activity increased by 799% in ΔvemR. In contrast, the blue colour intensity of ΔrpoN2 and ΔfleQ became weaker, and the quantitative activity of the rpoN2 promoter was reduced by 36.9%–44.9% compared with that in the WT strain (Figure 2a). These results show that VemR down‐regulates rpoN2‐vemR‐fleQ expression, while FleQ and RpoN2 activate the expression of this operon.

FIGURE 2.

FIGURE 2

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VemR directly interacts with FleQ. (a) The rpoN2 promoter activities in the wild type (WT), ΔravA, ΔrpoN2, ΔvemR, and ΔfleQ strains were determined by measuring the β‐glucuronidase (GUS) activity of rpoN2p‐gusA. The values given are the means ± SD from triplicate experiments. Significant differences between a given mutant and the WT strain are shown. **p < 0.01, ***p < 0.001. (b) Bacterial two‐hybrid (B2H) assay for analysis of the interactions of FleQ and VemR. All strains were inoculated on LB and M63 supplemented with X‐gal (LX and M63X, respectively) for 24 h at 30°C. Each spot was inoculated with 2 µl of each of a 10‐fold dilution series (i.e., 10−1, 10−2, 10−3, 10−4, 10−5, and 10−6‐fold, from left to right) of cells in logarithmic growth phase (OD at 600 nm = 0.8). D11K, D56A, and D11K/D56A are VemR phosphorylation‐related site (D11 and D56) mutants. The plasmid pairs T18C/T25 and T18CZIP/T25ZIP were used as negative and positive controls, respectively. T18C‐X and T25‐Y indicate the vectors T18C and T25 harbouring the X and Y genes, respectively. (c) β‐Galactosidase activity of each reporter B2H strain. The values given are the means ± SD from triplicate experiments. Significant differences between the given bacteria and the negative control strain (T18C/T25) are shown. **p < 0.01. (d) Pull‐down assays for analysis of the interactions of FleQ and VemR. Cell lysates containing His‐FleQ (55 kDa) and MBP (maltose‐binding protein)‐VemR (58 kDa) or MBP (44 kDa) were incubated overnight at 4°C with amylose resin. After elution, the protein samples were boiled and separated by SDS‐PAGE and immunoblotted with anti‐MBP and anti‐His antibodies

Then, we analysed whether VemR could interact with FleQ and/or RpoN2. To this end, we first used bacterial two‐hybrid (B2H) assays to detect the interaction between RpoN2/FleQ and VemR. Only the reporter strains expressing both VemR and FleQ grew on the selection plates and produced blue colonies. The strains expressing either VemR or FleQ did not grow on the selection plates and did not produce blue colonies (Figure 2b,c). These data suggest that VemR and FleQ interact in Eschericia coli cells. However, no interaction was found between RpoN2 and VemR in the B2H experiments (Figure S3).

Previous studies have predicted that aspartic acid residues at positions 11 (D11) and 56 (D56) in VemR are key for its phosphorylation. D56 might be the phosphorylation site, and D11 coordinates the Mg2+ cation, allowing proper D56 phosphorylation (Appleby & Bourret, 1999; Delgado et al., 1993). After substituting these aspartic acids with lysine and alanine, the reporter strains expressing the VemR mutants (D11K, D56A, and D11KD56A) and FleQ still grew and turned blue on the selection plates (Figure 2b,c), illustrating that these residue substitutions, which may affect the phosphorylation of VemR, do not influence the interaction between FleQ and VemR.

To further verify the interaction between VemR and FleQ, pull‐down assays were carried out. His‐FleQ, a c.55 kDa protein, could be pulled down by maltose‐binding protein (MBP)‐VemR (58 kDa) but not by MBP (44 kDa) (Figure 2d). Consistent with the results of B2H experiments, protein pull‐down analysis also showed that there is a physical interaction between VemR and FleQ.

2.4. RpoN2‐VemR‐FleQ is epistatic to RavA in the regulation of Xcc virulence and motility

Our previous study showed that RavA was involved in Xcc virulence, EPS production, biofilm formation, and motility (Tao et al., 2014). The phenotypes of the ravA mutant were highly similar to those of the vemR mutant (Tao & He, 2010; Tao et al., 2014). RavA acts as an HK, and the cognate HK of VemR has not been found, suggesting that RavA may interact directly or indirectly with VemR. To determine whether RavA plays a role in the regulation of bacterial motility via the RpoN2‐VemR‐FleQ signalling pathway, we constructed double mutants (ΔrpoN2/ΔravA, ΔvemR/ΔravA, and ΔfleQ/ΔravA), triple mutants (ΔrpoN2/ΔvemR/ΔravA, ΔvemR/ΔfleQ/ΔravA, and ΔrpoN2/ΔfleQ/ΔravA), and quadruple mutants (ΔrpoN2/ΔvemR/ΔfleQ/ΔravA).

For bacterial virulence, the average lesion length of ΔravA was 10 mm, while those of ΔrpoN2/ΔravA and ΔfleQ/ΔravA double mutants increased to 20 and 20.7 mm, respectively, 100% and 107% higher than that of ΔravA. When vemR and ravA were both deleted, the virulence was completely lost, similar to the virulence phenotype of ΔvemR (Figure 3a,b). The lesions of the triple mutants ΔrpoN2/ΔvemR/ΔravA, ΔvemR/ΔfleQ/ΔravA, and ΔrpoN2/ΔfleQ/ΔravA were similar to those of the double mutants ΔrpoN2/ΔvemR, ΔvemR/ΔfleQ, and ΔrpoN2/ΔfleQ, respectively. Similarly, the virulence of the quadruple mutant ΔrpoN2/ΔvemR/ΔfleQ/ΔravA was consistent with that of ΔrpoN2/ΔvemR/ΔfleQ (Figure 3a,b).

FIGURE 3.

FIGURE 3

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The rpoN2‐vemR‐fleQ operon functions downstream of ravA in the regulatory pathway of bacterial virulence and motility. (a) Virulence phenotypes of the wild type (WT), ΔravA, double mutants (ΔrpoN2/ΔravA, ΔvemR/ΔravA, ΔfleQ/ΔravA), triple mutants (ΔrpoN2/ΔvemR/ΔravA, ΔvemR/ΔfleQ/ΔravA, ΔrpoN2/ΔfleQ/ΔravA), quadruple mutant (ΔrpoN2/ΔvemR/ΔfleQ/ΔravA), and corresponding complementation strains 14 days postinoculation (dpi) on broccoli cv. Wenxin leaves. C indicates that the indicated gene (rpoN2, vemR, and fleQ) complements a given mutant. (b) Lengths of the lesions on the 14 dpi‐infected broccoli leaves caused by the above strains, as shown in (a). (c, d) The swarming colony phenotypes (c) and average zone diameters (d) of the above strains, as shown in (a). (e, f) The swimming colony phenotypes (e) and average zone diameters (f) of the above strains, as shown in (a). The values given are the means ± SD from triplicate experiments. As our focus was the relationship of ravA and rpoN2, vemR or fleQ, we only compared the significant difference between ΔravA and the double mutants ΔravA/ΔrpoN2, ΔravA/ΔvemR, and ΔravA/ΔfleQ. **p < 0.01

The diameter of the swarming zone of ΔravA was 10.3 mm, while those of the ΔrpoN2/ΔravA and ΔfleQ/ΔravA mutants increased to 20.7 and 23 mm, respectively (Figure 3c,d). For swimming, the swimming zone diameter of ΔravA was 23 mm, and that of ΔvemR/ΔravA increased to 35 mm. In contrast, the swimming ability was lost in ΔrpoN2/ΔravA and ΔfleQ/ΔravA, similar to the result for ΔrpoN2 or ΔfleQ (Figure 3e,f). The motilities (including swimming and swarming) of the triple mutants (ΔrpoN2/ΔvemR/ΔravA, ΔvemR/ΔfleQ/ΔravA, and ΔrpoN2/ΔfleQ/ΔravA) and the quadruple mutant (ΔrpoN2/ΔvemR/ΔfleQ/ΔravA) were also similar to those of ΔrpoN2 (Figure 3c–f). The above results show that VemR, RpoN2, and FleQ are epistatic to RavA in the regulation of Xcc virulence and motility.

2.5. RavA can interact with and phosphorylate VemR

The function of RRs is usually determined by their phosphorylation state, which is controlled by cognate HKs (Galperin, 2006). Although RavA is the cognate HK of RavR, ravA and ravR deletion did not result in the same phenotypes (Figure 3), implying that RavA might regulate other RR activities. VemR, which contains only a REC domain and lacks any output domain (Qian et al., 2008), was reported as the cognate RR of HpaS (Li, Wang, et al., 2020). However, deletion of hpaS and vemR did not result in the same phenotypes (Li, Wang, et al., 2020), indicating that VemR activity may be regulated by other HKs.

As our above genetic data showed that vemR is epistatic to ravA in the regulation of Xcc motility and virulence, and that ΔvemR and ΔravA have highly similar phenotypes, we hypothesized that RavA is also a cognate HK of VemR. We used B2H assays to analyse whether RavA interacts with VemR. The strains expressing both VemR and RavA grew and turned blue on the detection plates, which was similar to the result for the positive control strain (T18CZIP/T25ZIP), demonstrating that VemR can interact with RavA in E. coli cells. When the histidine residue at position 164 of the RavA phosphorylation site was substituted with alanine (H164A), the interaction decreased (Figure 4a,b). Similarly, the D11K and D11K/D56A substitutions in VemR also weakened its interaction with RavA, but the D56A substitution had little effect, showing that the phosphorylatable H164 of RavA and D11 of VemR might be required, but not essential, for their interaction, while the D56 phosphorylation site of VemR does not significantly influence its interaction with RavA. Nevertheless, these data demonstrated that RavA interacts with VemR in E. coli. We also confirmed the interaction of RavA and VemR by pull‐down tests because MBP‐RavA, not sole MBP, pulled down His‐VemR in vitro (Figure 4c). Taken together, our data demonstrate that RavA and VemR interact with each other and might form a TCS.

FIGURE 4.

FIGURE 4

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RavA interacts with and phosphorylates VemR. (a) Bacterial two‐hybrid (B2H) assays for analysis of the interaction between RavA and VemR in Escherichia coli. All strains were inoculated on LX and M63X at 30°C for 24 h, as shown in Figure 2. D11K, D56A, and D11K/D56A are VemR phosphorylation‐related site (D11 and D56) mutants, while H164A is a RavA phosphorylation site mutant. T18C‐X and T25‐Y indicate the vectors T18C and T25 harbouring the X and Y genes, respectively. (b) β‐Galactosidase activities of each reporter B2H strain. The plasmid pairs T18C/T25 and T18CZIP/T25ZIP were used as negative and positive controls, respectively. The values given are the means ± SD from triplicate experiments. Significant differences between the given bacteria and the negative control strain (T18C/T25) are shown. **p < 0.01. (c) Analysis of the interactions of RavA and VemR by maltose‐binding protein (MBP) pull‐down assays. The experiments were performed as shown in Figure 2d, but His‐VemR (15 kDa) and MBP‐RavA (89 kDa) were used instead of His‐FleQ and MBP‐VemR, respectively. (d) In vivo and in vitro assays for detecting phosphotransfer from RavA to VemR. In vivo: Xanthomonas campestris pv. campestris (Xcc) strains and E. coli strains expressing VemR‐His were cultured in M4M and LB media, respectively. VemR‐His protein was purified with Ni‐NTA resin, and elution samples were separated on Phos‐tag and standard SDS‐PAGE and then transferred to PVDF membranes for western blotting with anti‐His antibody. In vitro: 50 ng of purified RavA was autophosphorylated for 30 min, then 50 ng of purified His‐VemR protein was added and incubated at 28°C. At the indicated time, SDS (1× final concentration) sample buffer was added to stop the reaction. SDS‐PAGE and western blotting were performed as in vivo assays. −, without the RavA protein

To confirm our hypothesis that RavA/VemR is a TCS, we performed phosphotransfer assays. When VemR‐His was expressed in ΔvemR and ΔravA/ΔvemR, the phosphorylation of VemR‐His could be detected in both strains, but the levels were much higher in ΔvemR than in ΔravA/ΔvemR (Figure 4d), suggesting that RavA phosphorylates VemR in Xcc. In E. coli, Flag‐RavA and VemR‐His coexpression resulted in detectable phosphorylation states of VemR, but expression of only VemR did not have this effect, indicating that RavA phosphorylates VemR in E. coli (Figure 4d). Moreover, we performed in vitro phosphorylation assays and found that phosphorylated glutathione S‐transferase (GST)‐RavA transferred the phosphate group to His‐VemR (Figure 4d). These data demonstrate that RavA interacts with and phosphorylates VemR.

2.6. RavA/VemR regulates the expression of the flagellar gene fliC by regulating the expression and activity of RpoN2/FleQ

To test whether RavA regulates rpoN2‐vemR‐fleQ operon expression, rpoN2 p‐gusA was also transformed into the ΔravA and WT strains to analyse rpoN2 promoter activity. The GUS staining assays showed that the colonies of the ΔravA mutant containing rpoN2 p‐gusA were bluer than those of the WT strain containing the same plasmid (Figure S2). Compared with WT, ΔravA showed a 668% increase in rpoN2 promoter activity, similar to ΔvemR (Figures 2a and S2). These results suggest that RavA down‐regulates the transcription of rpoN2‐vemR‐fleQ, similar to VemR.

The expression of the flagellin‐encoding gene fliC is regulated by multiple genes, including rpoN2 and fleQ (Figure S1) (Yang et al., 2009). To understand whether VemR and RavA play roles in the regulation of fliC expression, we used promoter‐reporter constructs to detect fliC expression levels. We constructed a fliC p‐lacZ fusion strain based on Δlac8 (Wang et al., 2018) that could directly reflect the expression levels of fliC on plates containing X‐gal (5‐bromo‐4‐chloro‐3‐indolyl β‐d‐galactoside). When rpoN2 or fleQ was deleted, the colonies did not exhibit a blue colour (Figure S4). After deletion of vemR, the colonies turned blue and showed detectable β‐galactosidase activity. β‐Galactosidase activity could not be detected in the double mutants (ΔrpoN2/ΔvemR, ΔvemR/ΔfleQ, and ΔrpoN2/ΔfleQ) or the triple mutant (ΔrpoN2/ΔvemR/ΔfleQ) (Figures 5a,b and S4). These data imply that RpoN2 and FleQ positively regulate fliC transcription, while VemR inhibits the transcription of this gene via RpoN2 and FleQ. Similar to ΔvemR, the fliC promoter activity increased 253% and 284% in ΔravA and ΔvemR/ΔravA, respectively (Figure 5a,c). In addition, ΔrpoN2/ΔravA and ΔfleQ/ΔravA did not exhibit fliC promoter activity (Figure 5a,c). This suggests that RavA/VemR regulates the expression of the flagellin‐encoding gene fliC via RpoN2/FleQ.

FIGURE 5.

FIGURE 5

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RavA regulates fliC expression via RpoN2‐VemR‐FleQ. (a) Plate‐based detection of the effects of rpoN2, vemR, fleQ, and ravA deletion mutations on fliC promoter activity, as determined by measuring β‐galactosidase activity. The XC1214 (Xanthomonas campestris pv. campestris lacZ) gene was integrated downstream of the fliC promoter, and lacZ‐reporter strains were constructed. (b, c) Quantitative analysis of the effects of rpoN2, vemR, fleQ, and ravA single, double, triple, and quadruple mutations on fliC promoter activity. The values given are the means ± SD from triplicate experiments. In rpoN2‐vemR‐fleQ‐related mutants (b), only the vemR single mutation had a significant effect on fliC promoter activity. In ravA‐related mutants (c), ravA mutation significantly increased fliC expression, but fleQ or rpoN2 mutation in the ravA mutant abolished this induction. **p < 0.01, ***p < 0.001

2.7. RavA/VemR regulates fliC expression via FliA

FliA is essential for the transcription of fliC (Figure S1) (Kan et al., 2018). The expression of the flhF‐fleN‐fliA operon is dependent on RpoN2/FleQ and is down‐regulated by FliA (Yang et al., 2009). As RavA and VemR play a crucial role in the transcription of fliC (Figure 5), we asked whether this regulation requires FliA. First, we transformed flhF p ‐gusA into the WT, ΔravA, ΔrpoN2, ΔvemR, and ΔfleQ strains to analyse flhF promoter activity. Only ΔravA and ΔvemR containing flhF p ‐gusA produced a dark blue product (Figure S2), and the activity of the flhF promoter increased by approximately 75% in these strains compared with that in the WT strain (Figure 6a), demonstrating that RavA/VemR down‐regulates the transcription of the flhF‐fleN‐fliA operon. Then, we constructed five fliA‐related mutants (ΔfliA, ΔravR/ΔfliA, ΔravA/ΔfliA, ΔvemR/ΔfliA, and ΔfleQ/ΔfliA) and analysed their virulence and motility. Deletion of fliA did not affect the infection ability of the WT, ΔravR, and ΔfleQ strains but significantly increased the virulence of the ΔravA and ΔvemR strains (Figures 6b and S5), indicating that FliA acts as a negative regulator of virulence in the RavA/VemR‐mediated signalling pathway rather than the RavA/RavR‐mediated signalling pathway.

FIGURE 6.

FIGURE 6

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Effects of fliA mutation on bacterial virulence and motility. (a) The flhF‐fleN‐fliA operon promoter activity in the wild type (WT), ΔravA, ΔrpoN2, ΔvemR, and ΔfleQ strains. The flhFp‐gusA construct (flhF promoter‐gusA fusion) was transformed into the mentioned strains, and β‐glucuronidase (GUS) activity was detected. (b) Lesion lengths (14 days postinoculation) on broccoli leaves infected by the indicated strains (WT, ΔfliA, ΔravR, ΔravA, ΔvemR, ΔfleQ, ΔfliA/ΔravR, ΔfliA/ΔravA, ΔfliA/ΔvemR, and ΔfliA/ΔfleQ). (c) Swarming zone diameters of the above strains. (d) Swimming zone diameters of the above strains. (e) The fliC promoter activity of the above strains in NYG medium. The values given are the means ± SD from triplicate experiments. Significant differences between a single‐gene mutant and the corresponding double‐gene mutant (ΔravR vs. ΔfliA/ΔravR, ΔravA vs. ΔfliA/ΔravA, ΔvemR vs. ΔfliA/ΔvemR, and ΔfleQ vs. ΔfliA/ΔfleQ) are shown. A significant difference between the fliA mutant and the WT strain (ΔfliA vs. WT) is also shown. **p < 0.01, ***p < 0.001

Mutation of fliA in the WT, ΔravR, ΔravA, ΔvemR, or ΔfleQ stains did not change the swarming motility of these strains (Figures 6c and S5). In contrast, the swimming ability of ΔfliA, ΔravR/ΔfliA, ΔravA/ΔfliA, and ΔvemR/ΔfliA decreased 75%–91%, similar to that of ΔfleQ (Figures 6d and S5), whose swimming ability was almost completely lost. The activity of the fliC promoter decreased significantly in the ΔravR/ΔfliA, ΔravA/ΔfliA, and ΔvemR/ΔfliA double mutants (Figures 6e and S5). The above data suggest that fliC expression regulated by RavA/VemR is dependent on RpoN2/FleQ and FliA.

2.8. clp is epistatic to ravA/vemR in Xcc virulence, not motility, regulation

Clp is a global transcriptional regulator that regulates extracellular enzymes, EPS, motility, and virulence in Xcc (He et al., 2007). As the virulence of the ravA/vemR deletion mutant was similar to that of the clp mutant, we studied the relationship between these genes. First, we constructed five clp‐related mutants: Δclp, ΔravR/Δclp, ΔravA/Δclp, ΔvemR/Δclp, and ΔfleQ/Δclp. None of these strains could infect broccoli leaves (Figures 7a and S5), indicating that Clp is located downstream of RavA/VemR and RavA/RavR, and regulates the virulence of Xcc. Deletion of clp led to a slight improvement in the swarming ability of the WT and ΔfleQ strains but had no effects on the ΔravR, ΔravA, and ΔvemR strains (Figures 7b and S5). Except for ΔfleQ/Δclp, the swimming ability of all the clp‐related mutants increased by approximately 26%–61% (Figures 7c and S5). These changes were not achieved by regulating fliC transcription because clp deletion had no effects on fliC promoter activity (Figures 7d and S5). These data show that Clp and RavA/VemR are involved in different pathways for swimming regulation in Xcc, but both depend on FleQ. Taken together, the results show that Clp is located downstream of RavA/VemR and regulates Xcc virulence, and that these proteins participate in cross‐talk to regulate Xcc motility.

FIGURE 7.

FIGURE 7

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clp is epistatic to ravA/vemR in the regulation of Xanthomonas campestris pv. campestris (Xcc) virulence but not motility. (a) Lesion lengths (14 days postinoculation) of broccoli leaves infected with wild type (WT), Δclp, ΔravR, ΔravA, ΔvemR, ΔfleQ, Δclp/ΔravR, Δclp/ΔravA, Δclp/ΔvemR, and Δclp/ΔfleQ. (b, c) Swarming and swimming zone diameters of the above strains, respectively. (d) The fliC promoter activity was analysed by measuring the β‐galactosidase activity of the above strains. The values given are the means ± SD from triplicate experiments. Significant differences between a single‐gene mutant and the corresponding double‐gene mutant (ΔravR vs. Δclp/ΔravR, ΔravA vs. Δclp/ΔravA, ΔvemR vs. Δclp/ΔvemR, and ΔfleQ vs. Δclp/ΔfleQ) are shown. A significant difference between the clp mutant and the WT strain (Δclp vs. WT) is also shown. *p < 0.05, **p < 0.01, ***p < 0.001

3. DISCUSSION

In the genome of Xcc, rpoN2, vemR, and fleQ are expressed as a transcriptional operon (Tao & He, 2010; Wu et al., 2019). vemR deletion resulted in a significant decrease in the virulence and adaptation of Xcc, but single or double deletion of fleQ and rpoN2 did not affect these behaviours (Figure 1). Interestingly, the phenotypes of ∆rpoN2/∆vemR and ∆vemR/∆fleQ were at an intermediate level between those of ∆vemR and ΔrpoN2 or ΔfleQ, while those of ΔrpoN2/ΔfleQ and ΔrpoN2/ΔvemR/ΔfleQ were similar to those of ΔrpoN2 or ΔfleQ (Figure 1). Notably, our complementation assays showed that rpoN2, vemR, and fleQ rescued the phenotypes of the corresponding mutants (Figure 1), demonstrating that there were no alternative promoters in the rpoN2 and vemR coding regions, and that deletion of rpoN2 or vemR had no polar effect. Therefore, we suggest that fleQ and rpoN2 are epistatic to vemR and that VemR may directly or indirectly regulate FleQ and RpoN2 activities.

VemR is an atypical RR that contains a CheY‐like receiver but lacks an output domain (Qian et al., 2008), suggesting that its function may require its interaction with other proteins. Thus, we tested whether VemR can interact with FleQ and RpoN2, and found that VemR physically binds FleQ but not RpoN2 (Figure 2). As FleQ acts as a transcriptional activator and may interact with RpoN2 to control gene expression (Dasgupta et al., 2002), the regulatory function of VemR in Xcc virulence and motility might depend on its interaction with FleQ. Notably, VemR was recently reported to interact with RpoN2 in X. citri subsp. citri and regulate bacterial virulence and swimming (Wu et al., 2019), but we did not find this interaction in Xcc (Figure S3). Because ∆rpoN2/∆vemR and ∆vemR/∆fleQ have similar phenotypes, VemR may also have the ability to interact with RpoN2 in Xcc, but alternative analysis methods are required to explore this further, as our B2H assays could not verify this hypothesis. Moreover, a recent study showed that VemR directly interacts with the flagellum protein FliM to regulate swimming in Xcc, and this interaction depends on the VemR phosphorylation status (Li, Wang, et al., 2020). Although we could not detect the VemR–RpoN2 interaction via B2H assays and did not perform a VemR–FliM interaction assay, previous reports and our findings illustrate that the regulatory function of VemR is dependent on its interaction with other proteins (Li, Wang, et al., 2020; Wu et al., 2019), although VemR interacts with different proteins (RpoN2, FleQ, FliM, or other unknown partners). The question of how VemR is able to interact with different proteins, as shown by various research groups, needs to be answered in the future. The mechanisms by which VemR regulates the activities of FleQ, RpoN2, or FliM are also unclear and need further study (Figure 8).

FIGURE 8.

FIGURE 8

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Model for the cross‐talk among HpaS‐, RavA‐, and RavS‐mediated signalling pathways in Xanthomonas campestris pv. campestris (Xcc). RavA/VemR regulates flagellin fliC expression via FleQ/RpoN2 and FliA to influence Xcc swimming and regulates FleQ/RpoN2 and Clp activities to affect bacterial swarming. RavA/RavR and RavS/RavR control cellular c‐di‐GMP levels and thus regulate bacterial motility via the c‐di‐GMP receptor Clp. HpaS/VemR regulates bacterial swimming via direct interaction with the flagellum protein FliM. HpaS phosphorylates HrpG and regulates the expression of the type III secretion system (T3SS) and Xcc virulence. Inline graphic positive regulation; Inline graphic negative regulation; solid lines, having experimental evidence; dotted lines, supposed relationship. Components of a given pathway are indicated in the same colour

The function of an RR is typically controlled by its cognate HK, but bioinformatics analysis could not predict the cognate HK of VemR because no HK has been annotated in the genomic context of vemR (Qian et al., 2005). When HpaS was used as bait to screen for its interacting proteins, VemR was identified. Further yeast two‐hybrid, pull‐down, and phosphorylation assays showed that HpaS directly interacts with and phosphorylates VemR, indicating that HpaS and VemR form a TCS (Li, Wang, et al., 2020). Interestingly, mutation of hpaS did not affect the swarming ability of Xcc but significantly reduced its swimming ability. Additionally, double mutation of hpaS and vemR increased the swimming ability to the WT levels but caused the swarming phenotype to be similar to that of the vemR mutant (Li, Wang, et al., 2020). These data indicate that vemR is epistatic to hpaS in Xcc swarming regulation, but vemR and hpaS have antagonistic effects on swimming regulation. Therefore, VemR is not the downstream partner of HpaS in the regulation of Xcc swimming. Further study is needed to determine how HpaS and VemR coregulate Xcc motility.

As the vemR/r avA double mutant has similar phenotypes, including swimming, swarming, and virulence, as the vemR single mutant, vemR has an epistatic relationship with ravA (Figure 3), leading us to ask whether RavA and VemR interact directly or indirectly. Our B2H, pull‐down, and phosphorylation assays showed that RavA interacted with and phosphorylated VemR (Figure 4). We also found that changes in the key residues for the phosphorylation of RavA and VemR might influence their interaction in the B2H assays (Figure 4). Together, these genetic and biochemical analyses indicate that RavA and VemR also form a TCS that regulates Xcc motility and virulence.

Previous studies have shown that the expression of flagellar genes is regulated by cascade signal transduction pathways (Figure S1). RpoN2/FleQ regulates the expression of the class II gene operon flhF‐fleN‐fliA. FliA can regulate the expression of its own operon and the fliC gene (Yang et al., 2009). Through analysis of fliC promoter activity, RpoN2 and FleQ were found to activate fliC transcription, and RavA and VemR inhibited fliC transcription via RpoN2/FleQ (Figures 5 and S4). Analysis of the transcriptional expression of flhF‐fleN‐fliA also revealed that RavA/VemR can down‐regulate the transcription of this operon and that RavA/VemR regulates fliC expression via a mechanism dependent on FliA (Figures 6 and S5). Based on the above results, we have discovered a new signalling pathway that regulates the expression of the flagellar gene fliC (Figure 8). RavA senses an unknown signal (which might be an intracellular cue) and undergoes autophosphorylation, followed by transfer of the phosphate group to VemR. VemR interacts with the transcription activator FleQ and regulates FleQ activity and its own transcription. RpoN2 and FleQ regulate the expression of flhF‐fleN‐fliA and influence fliC expression, thus affecting the swimming ability of Xcc (Figure 8). Simultaneously, FliA regulates the expression of genes involved in c‐di‐GMP turnover and thus indirectly influences Xcc swimming ability (Yang et al., 2009) (Figure S1). As RavR is important for c‐di‐GMP metabolism and Xcc swimming (Tao et al., 2014), we also analysed the role of FliA in RavR‐mediated fliC expression and found that FliA was also vital for RavR signalling (Figure 6). Therefore, FliA is required for the effects of both VemR and RavR in the regulation of Xcc swimming (Figure 8).

In addition to VemR, RavA can also form a TCS with RavR to regulate bacterial motility and virulence (Tao et al., 2014). Moreover, RavS/RavR is also an important TCS for Xcc adaptation and virulence (He et al., 2007). Although we uncovered the functions of these TCSs in Xcc, the relationship among RavA/RavR, RavA/VemR, and RavS/RavR remains unclear. Because RavS is a transmembrane HK while RavA is a cytoplasmic HK, they might sense extra‐ and intracellular signals, respectively (Figure 8), and promote different signal transduction pathways. Similarly, HpaS is also a transmembrane HK that may also transduce extracellular signals to regulate VemR, HrpG, or HrpR2 activity. However, how and when HpaS phosphorylates these proteins is unknown. Uncovering the complex regulatory network by these newly identified branched TCSs (RavA/RavR, RavA/VemR, RavS/RavR, HpaS/VemR, HpaS/HrpG, and HpaS/HpaR2) in Xcc will deepen our understanding of bacterial adaptation to environments (Figure 8). To best understand these complex cross‐talk networks, two aspects should be considered in future studies: (a) the affinities and phosphoryl‐transfer efficiencies, and (b) the subcellular locations and concentrations of an HK (RavS, RavA, HpaS) and its cognate RR (RavR, VemR, HrpG, HpaR2) in different environmental and cellular conditions.

RavA/RavR and RavS/RavR are involved in c‐di‐GMP signalling, which is important for bacterial motility, biofilm formation, EPS production, and virulence (He et al., 2007; Tao et al., 2014). These behaviours are also regulated by Clp (He et al., 2007), RavA, and VemR (Tao & He, 2010; Tao et al., 2014). Because Clp acts as a c‐di‐GMP receptor (He et al., 2007), we also analysed the relationship between RavA/VemR and Clp. Notably, the regulation of swimming by Clp is independent of VemR and flagellar regulators, including RpoN2 and FliA, but requires FleQ (Figure 7). However, virulence regulation by VemR is partly dependent on Clp. The results indicate that Clp and VemR are involved in different regulatory pathways of Xcc swimming but have an epistatic relationship in regulating the virulence of Xcc. They also suggest that the Clp‐mediated c‐di‐GMP signalling pathway participates in cross‐talk with RavA/VemR signalling. Because RavA/RavR is a crucial regulatory module in c‐di‐GMP signalling that regulates Xcc motility (Tao et al., 2014), RavA might regulate RavR activity to change cellular c‐di‐GMP levels and thus affect Clp activity. Although mutation of clp in the ravR mutant significantly altered bacterial virulence and swimming, swarming and flic promoter activity were not regulated by Clp in the ΔravR strains. Thus, RavA influences VemR and RavR activities, thereby regulating Clp function in an unknown manner (Figure 8).

In conclusion, our results indicate that VemR controls bacterial motility and virulence by interacting with the transcriptional activator FleQ. This interaction may affect the activity and expression of RpoN2/FleQ to regulate the transcription of flagellum‐related genes (Figure 8). Overall, our findings illustrate that the RavA/VemR TCS plays a key regulatory role in Xcc motility and virulence.

4. EXPERIMENTAL PROCEDURES

4.1. Bacterial strains, plasmids, media, and growth conditions

The bacterial strains and plasmids used in this study are listed in Table S1. The Xcc strains were grown in nutrient‐yeast‐glycerol (NYG) medium (Daniels & Barber, 1984), NY medium (NYG medium without glycerol), or minimal medium 4 (M4) modified (M4M) medium (each litre of which contained 10.5 g Na2HPO4, 4.5 g NaH2PO4, 1 g (NH4)2SO4, 0.5 g C6H5Na3O7, 0.2 g MgSO4, 8 g casein enzymatic hydrolysate, and 1 g d‐glucose) at 28°C. E. coli strains were incubated in Luria Bertani (LB) medium (Miller, 1972) or minimal medium 63 (M63) (each litre of which contained 100 mM KH2PO4, 75 mM KOH, 15 mM (NH4)2SO4, 1 mM MgSO4, 3.9 μM FeSO4, and 22 mM d‐glucose). Antibiotics at the following final concentrations were used: 25 μg/ml kanamycin (Kan), 50 μg/ml rifampicin (Rif), 100 μg/ml ampicillin (Amp), 50 μg/ml spectinomycin (Spc), 5 μg/ml gentamicin (Gm), and 25 μg/ml chloramphenicol (Cm).

4.2. Construction and complementation of the deletion mutant strains

The promoter of fliC was fused with the lac Z gene and integrated into the β‐galactosidase‐deficient strain Δlac8 to construct the reporter strain Δlac8‐fliC‐lacZ (Wang et al., 2018). To construct the ravA deletion mutant, c.500 bp fragments flanking ravA were amplified and then cloned together into the HindIII/XbaI sites of pK18mobsacB (Schäfer et al., 1994). The recombinant plasmid pK18mobsacB‐ravA was introduced into Δlac8‐fliC‐lacZ (WT) by triparental conjugation (Li & Wang, 2011). After two cycles of selection, the primer set ravA‐FF/ravA‐RR was used to identify the ΔravA mutant. The same genetic manipulation method was used to obtain other mutants. The primers used in this study are listed in Table S2.

For complementation assays, the promoter and the coding sequence of the given gene were amplified and cloned into pXE64G (Lee et al., 2012) to generate complementation plasmids. The complementation plasmids were introduced into the corresponding mutants by electroporation.

4.3. Pathogenicity assays

Six‐week‐old plants of broccoli cv. Wenxin were used as host plants. The virulence of Xcc strains was determined by leaf clipping, as described elsewhere (Qian et al., 2005). Lesion lengths were measured 2 weeks after inoculation on 10 leaves for each strain.

4.4. Motility assays

Swarming motility was determined on NY plates containing 2% glucose and 0.6% agar. Swimming motility was assessed on 0.28% agar plates containing 0.03% Bacto peptone and 0.03% yeast extract as described previously (Qi et al., 2020).

4.5. β‐Galactosidase activity assays

β‐Galactosidase (LacZ) activity analysis was carried out as previously described (Griffith & Wolf, 2002; Smale, 2010; Wang et al., 2018). For plate assays, 1 µl of diluted cell culture was inoculated on NYG plates containing 40 mg/L X‐gal. After incubating at 28°C for 2 days, the plates were checked for the appearance of blue colonies. For quantitative assays of β‐galactosidase activity, cells (logarithmic growth phase) were collected by centrifugation. Equal amounts (OD600 = 2.0) of bacteria were resuspended in 1 ml of Z buffer (0.4 mg lysozyme, 60 mM Na2HPO4, 40 mM NaH2PO4, 10 mM KCl, 1 mM MgSO4, and 50 mM β‐mercaptoethanol [β‐ME]) and lysed at room temperature for 30 min. Then, 100 μl of supernatant was added to 200 μl of o‐nitrophenyl‐β‐galactoside (ONPG) (4 mg/ml) and incubated at 37°C. When the sample appeared yellow, the reaction was stopped with 0.2 M Na2CO3 and β‐galactosidase activity was measured by determining the A420. The assays were performed in triplicate. The following formula was used for activity calculation: Unit = (A420 × 1000)/(t × v × OD600), where t = elapsed time (in min), v = volume of culture (in ml), and OD600 = optical density of the culture used.

4.6. β‐Glucuronidase activity assays

Bacteria (logarithmic growth phase) were collected by centrifugation, washed twice with double deionized water (ddH2O), and resuspended in ddH2O (OD600 = 1.0). One hundred microlitres of the bacterial suspension was mixed with 200 μl of GUS staining solution (50 mM NaH2PO4, 50 mM Na2HPO4, 10 mM Na2EDTA, 0.1% Triton X‐100, 0.5 mM K3[Fe(CN)6], 0.5 mM K4[Fe(CN)6], 20 mM X‐Gluc), incubated at 37°C for 4 h, and the colour change was observed (Kosugi et al., 1990). The method for quantification of GUS activity was almost the same as that used for measuring β‐galactosidase activity. The differences were as follows: (a) the lysis buffer contained 50 mM NaH2PO4, 50 mM Na2HPO4, 10 mM Na2EDTA, 0.1% Triton X‐100, 0.1% SDS, and 10 mM β‐ME, (b) the reaction substrate was 2 mM 4‐methylumbellifery‐β‐d‐glucuronide (4‐MUG), and (c) the fluorescence excitation wavelength was 365 nm and the emission wavelength detected was 455 nm.

4.7. Bacterial two‐hybrid assays

The Bacterial Adenylate Cyclase (CyaA) Two‐Hybrid System (Karimova et al., 1998) was used to detect protein–protein interactions. VemR, RavA, FleQ, and RpoN2 were fused with the T25 or T18 fragment of CyaA and cotransformed into the reporter strain BTH101. Selection and data analysis were carried out as described in a previous report (Karimova et al., 1998). Briefly, the resulting strains were grown overnight in liquid media containing the corresponding antibiotics and washed twice with ddH2O. Then, the cell suspension was diluted and spotted on LB medium or auxotrophic M63 medium containing 40 mg/ml X‐gal and cultured at 30°C for 24 h.

4.8. Site‐directed mutagenesis

A Q5 Site‐Directed Mutagenesis Kit (NEB) was used for site‐directed mutagenesis. The recombinant plasmid containing the target gene was used as the PCR template. The mutants were obtained by PCR using primer pairs containing the mutant sequences. The amplified product was treated with KLD enzyme mix and transformed into E. coli DH5α. Mutation was confirmed by DNA sequencing.

4.9. Protein pull‐down assays

The PCR‐amplified vemR or ravA full‐length coding region was digested with NdeI and EcoRI, and cloned into the pMAL‐c5× vector (NEB), and the fleQ and vemR PCR fragments were digested with BamHI and HindIII and cloned into the pQE80L vector (Qiagen). The recombinant vectors were transferred into E. coli BL21 (DE3) cells. Protein synthesis was induced by 0.3 mM isopropyl‐β‐d‐thiogalactopyranoside (IPTG). The bacterial cells were collected and lysed by ultrasonication. Both MBP‐VemR/FleQ‐His and MBP‐RavA/VemR‐His were incubated with amylose resin overnight at 4°C, followed by multiple washes with Tri‐buffered saline (50 mM Tris‐HCl, 150 mM NaCl, pH 7.5) to eliminate the influence of contaminants, and proteins were eluted with 10 mM maltose. Electrophoresis was performed by 12% sodium dodecyl sulphate‐polyacrylamide gel electrophoresis (SDS‐PAGE), and immunoblotting was performed with an anti‐MBP antibody and an anti‐His antibody. The MBP protein was used as a negative control (Liu et al., 2011).

4.10. Protein phosphorylation analysis

In vivo VemR phosphorylation states were detected in Xcc and E. coli. In E. coli, the strains expressing VemR‐His and RavA‐Flag/VemR‐His were cultured in LB medium to an OD600 of 0.6. Then, 0.3 mM IPTG was added to induce protein expression, and the cells were cultured for 5 h at 25°C. Then, the bacterial cells were collected and lysed by ultrasonication to prepare total proteins. VemR‐His was purified with Ni‐NTA, and 10‐µl elution samples were subjected to Phos‐tag SDS‐PAGE as described previously (Li et al., 2014) except that Zn2+ was used instead of Mn2+. Protein transfer from the gel to a polyvinylidene difluoride (PVDF) membrane and western blotting were performed as described in a previous report (Li et al., 2014). Anti‐FLAG and anti‐His antibodies were used for protein detection. In Xcc, the plasmids encoding VemR‐His were transformed into ΔvemR and ΔravA/ΔvemR. The resulting strains were cultured in M4M medium to an OD600 of 0.8, and protein expression was induced by 0.3 mM IPTG. Then, the bacterial cells were collected and lysed by ultrasonication to prepare total proteins. VemR‐His protein purification and Phos‐tag SDS‐PAGE were performed as discussed above. For in vitro assays, GST‐RavA and VemR‐His were expressed in E. coli BL21 and then purified. The autophosphorylation of GST‐RavA and phosphotransfer assays were performed as described previously (Tao et al., 2014). The phosphorylation states of VemR‐His were detected in Phos‐tagged gels as described above for in vivo assays.

Supporting information

FIGURE S1 Transcription cascade of the Xanthomonas campestris pv. campestris (Xcc) flagellar genes. Three classes of flagellar genes are regulated via a transcription cascade by different sigma and anti‐sigma factors. RpoD (σ70) regulates the expression of class I genes (fleQ, rpoN2 [σ54] and flgM [anti‐sigma factor]). FleQ and RpoN2 (σ54) regulate the expression of class II genes (F‐T3SS, Body hook, flhF, fleN and fliA [σ28]). FliA (σ28) controls the expression of class III genes. FlgM (anti‐sigma factor) binds FliA and negatively regulates its activity. This model is a modification of a previously reported model diagram (Yang et al., 2009). Inline graphic positive regulation; Inline graphic negative regulation

Click here for additional data file. (36.7KB, docx)

FIGURE S2 Effect of ravA, vemR, rpoN2, and fleQ mutations on rpoN2 and flhF promoter activities. The promoters of rpoN2 and flhF were fused with the gusA gene to form the vectors rpoN2 p‐gusA and flhF p‐gusA. These constructs were transformed into the wild type (WT), ΔravA, ΔrpoN2, ΔvemR, and ΔfleQ strains. The resulting bacterial strains were cultured in NYG broth to OD600 = 0.8, and the cells were collected. Coloured substances produced by the collected bacteria were observed by a standard staining method

Click here for additional data file. (684.5KB, tif)

FIGURE S3 There was no physical interaction between RpoN2 and VemR in the bacterial two‐hybrid (B2H) experiment. The plasmid pairs T18C/T25 and T18CZIP/T25ZIP were used as negative and positive controls, respectively. All strains were inoculated on LB medium supplemented with X‐Gal at 30°C for 24 h, and colour production was observed

Click here for additional data file. (913.6KB, tif)

FIGURE S4 Regulatory roles of RpoN2‐VemR‐FleQ and RavA in fliC promoter activity. Double mutant (ΔrpoN2/ΔvemR, ΔvemR/ΔfleQ, and ΔrpoN2/ΔfleQ), triple mutant (ΔrpoN2/ΔvemR/ΔfleQ, ΔrpoN2/ΔvemR/ΔravA, ΔvemR/ΔfleQ/ΔravA, and ΔrpoN2/ΔfleQ/ΔravA), and quadruple mutant (ΔrpoN2/ΔvemR/ΔfleQ/ΔravA) strains as well as their complementation strains were inoculated on NYG medium containing 40 mg/L X‐gal. Coloured colonies were observed after culturing at 28°C for 3 days

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FIGURE S5 Influence of fliA and clp mutations on Xanthomonas campestris pv. campestris (Xcc) virulence and motility. (a) Virulence phenotypes of fliA‐ and clp‐related mutants. Swarming (b) and swimming (c) phenotypes of fliA‐ and clp‐related mutants. (d) The fliC promoter activities in the fliA and clp mutants were tested on NYG plates containing 40 mg/L X‐gal

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TABLE S1 Bacterial strains and plasmids used in this study

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TABLE S2 Primers used in this study

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ACKNOWLEDGEMENTS

We thank Professor Shu‐Fen Weng at National Chung Hsing University for the gift of the pXE64G plasmid. This work was funded by the Natural Science Foundation of Hainan Province (grant no. 320RC478), the National Natural Science Foundation of China (grant no. 31660507), the Key R&D Program of Hainan Province (grant no. ZDYF2019063), and the Funding for the Construction of World First Class Discipline of Hainan University (grant no. RZZX201903).

Lin, M. , Wu, K. , Zhan, Z. , Mi, D. , Xia, Y. , Niu, X. , et al (2022) The RavA/VemR two‐component system plays vital regulatory roles in the motility and virulence of Xanthomonas campestris . Molecular Plant Pathology, 23, 355–369. 10.1111/mpp.13164

Maojuan Lin and Kejian Wu contributed equally to the study.

DATA AVAILABILITY STATEMENT

The raw data from this study are available from the corresponding author upon reasonable request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

FIGURE S1 Transcription cascade of the Xanthomonas campestris pv. campestris (Xcc) flagellar genes. Three classes of flagellar genes are regulated via a transcription cascade by different sigma and anti‐sigma factors. RpoD (σ70) regulates the expression of class I genes (fleQ, rpoN2 [σ54] and flgM [anti‐sigma factor]). FleQ and RpoN2 (σ54) regulate the expression of class II genes (F‐T3SS, Body hook, flhF, fleN and fliA [σ28]). FliA (σ28) controls the expression of class III genes. FlgM (anti‐sigma factor) binds FliA and negatively regulates its activity. This model is a modification of a previously reported model diagram (Yang et al., 2009). Inline graphic positive regulation; Inline graphic negative regulation

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FIGURE S2 Effect of ravA, vemR, rpoN2, and fleQ mutations on rpoN2 and flhF promoter activities. The promoters of rpoN2 and flhF were fused with the gusA gene to form the vectors rpoN2 p‐gusA and flhF p‐gusA. These constructs were transformed into the wild type (WT), ΔravA, ΔrpoN2, ΔvemR, and ΔfleQ strains. The resulting bacterial strains were cultured in NYG broth to OD600 = 0.8, and the cells were collected. Coloured substances produced by the collected bacteria were observed by a standard staining method

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FIGURE S3 There was no physical interaction between RpoN2 and VemR in the bacterial two‐hybrid (B2H) experiment. The plasmid pairs T18C/T25 and T18CZIP/T25ZIP were used as negative and positive controls, respectively. All strains were inoculated on LB medium supplemented with X‐Gal at 30°C for 24 h, and colour production was observed

Click here for additional data file. (913.6KB, tif)

FIGURE S4 Regulatory roles of RpoN2‐VemR‐FleQ and RavA in fliC promoter activity. Double mutant (ΔrpoN2/ΔvemR, ΔvemR/ΔfleQ, and ΔrpoN2/ΔfleQ), triple mutant (ΔrpoN2/ΔvemR/ΔfleQ, ΔrpoN2/ΔvemR/ΔravA, ΔvemR/ΔfleQ/ΔravA, and ΔrpoN2/ΔfleQ/ΔravA), and quadruple mutant (ΔrpoN2/ΔvemR/ΔfleQ/ΔravA) strains as well as their complementation strains were inoculated on NYG medium containing 40 mg/L X‐gal. Coloured colonies were observed after culturing at 28°C for 3 days

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FIGURE S5 Influence of fliA and clp mutations on Xanthomonas campestris pv. campestris (Xcc) virulence and motility. (a) Virulence phenotypes of fliA‐ and clp‐related mutants. Swarming (b) and swimming (c) phenotypes of fliA‐ and clp‐related mutants. (d) The fliC promoter activities in the fliA and clp mutants were tested on NYG plates containing 40 mg/L X‐gal

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TABLE S1 Bacterial strains and plasmids used in this study

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TABLE S2 Primers used in this study

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Data Availability Statement

The raw data from this study are available from the corresponding author upon reasonable request.

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