Skip to main content
NCBI home page
Molecular Plant Pathology logoLink to Molecular Plant Pathology
. 2026 Oct 19;27(10):e70354. doi: 10.1111/mpp.70354

A Hierarchical OhrR–EmrR Regulatory Circuit Fine‐Tunes the Vfm Quorum‐ Sensing System and Virulence in Dickeya oryzae

Wenqi Huang 1, Ziwei Meng 1, Zhongge Liu 1, Weihan Gu 2, Ming Hu 2, Tao Zhuo 1, Lianhui Zhang 2,✉, Mingfa Lv 1,✉
PMCID: PMC13680859  PMID: 42897884

ABSTRACT

Rice foot rot, caused by Dickeya oryzae, is a destructive bacterial disease. The Vfm quorum‐sensing (QS) system is a key virulence regulator of this pathogen. However, the mechanisms by which it perceives and integrates upstream environmental signals for precise transcriptional control are poorly understood. Here, we identify a cascade regulatory network involving the MarR‐family transcription factors OhrR and EmrR. Deletion of ohrR in D. oryzae EC1 significantly reduced the Vfm signalling activity, diminished the production of virulence factors including phytotoxin zeamines and plant cell wall‐degrading enzymes, and attenuated its pathogenicity. Molecular analyses revealed that OhrR directly bound to the key promoters in the vfm cluster (P vfmA , P vfmK , P vfmE and P vfmF ) and activated their transcription. Intriguingly, OhrR also directly activated the expression of emrR, which encodes another MarR‐family transcriptional regulator. In contrast to the activator role of OhrR, EmrR acted as a feedback inhibitor, directly bound to the same vfm promoter regions but repressed their transcription. OhrR–EmrR thus acts as a two‐tiered ‘activation–inhibition’ regulatory circuit in modulation of the Vfm QS system, in which OhrR functions as a master switch for environmental signal response and initiation of the Vfm system, whereas EmrR acts as a precise brake to prevent overexpression of virulence determinants. The findings from this study unveil the fascinating interaction between OhrR–EmrR and the Vfm QS system, and depict a novel mechanism for precise virulence control in D. oryzae via complex, dynamic transcriptional reprogramming.

Keywords: Dickeya oryzae, OhrR‐EmrR, transcriptional regulator, Vfm quorum sensing, virulence

We report a novel hierarchical regulatory circuit consisting of two MarR‐family transcription factors, OhrR and EmrR, that act antagonistically to dynamically fine‐tune the Vfm QS system and bacterial virulence.

graphic file with name MPP-27-e70354-g004.webp

1. Introduction

Quorum sensing (QS), a process whereby bacterial cells communicate to coordinate gene expression and adapt to environmental changes, is now a well‐characterized phenomenon (Whitehead et al. 2001; Fuqua and Greenberg 2002; Federle and Bassler 2003). QS systems play pivotal roles in diverse bacterial species, regulating processes such as secondary metabolite biosynthesis and virulence‐related functions, including exopolysaccharide (EPS) production, antibiotic/toxin synthesis, biofilm formation, extracellular enzyme secretion and motility (Ohtani et al. 2002; Pemberton et al. 2005; Deng et al. 2010, 2011, 2012; Passos‐da‐Silva et al. 2017). A typical QS system consists of a signal synthase and a signal receptor, such as LuxI and LuxR in Vibrio fischeri and LasI and LasR in Pseudomonas aeruginosa (Fuqua and Greenberg 2002). At low bacterial cell density, LuxI or its homologues produces a basal level of QS signal acylhomoserine lactone (AHL) molecules that is not sufficient to activate the signal receptor LuxR or its homologues. When the bacterial population reaches a threshold level, the accumulated AHL signals interact and activate the receptor LuxR or its homologues, which consequently initiate the transcription of the downstream target genes. By using such a simple and yet elegant population density‐dependent QS mechanism, bacterial pathogens can mount their attack collectively to overwhelm host defence responses and establish infections (Whitehead et al. 2001; Fuqua and Greenberg 2002; Lee and Zhang 2015). However, evidence has emerged that bacterial pathogens can also subtly tap on host stress signals to fine‐tune QS output and modulate virulence gene expression (Lee et al. 2013).

Dickeya oryzae (formerly Erwinia chrysanthemi pv. zeae) is the causal agent of bacterial foot rot, causing significant economic losses in rice and banana production (Samson et al. 2005). It is one of the few Dickeya species with a broad host range, infecting both monocotyledonous and dicotyledonous plants (Sinha and Prasad 1977; Nassar et al. 1994). Similar to the well‐studied model species Dickeya dadantii (Diolez and Coleno 1985; Reverchon and Nasser 2013), genomic and biochemical studies of D. oryzae have identified conserved virulence factors, including plant cell wall‐degrading enzymes (PCWDEs), biofilm formation, EPS, an iron assimilation system, the antioxidant pigment indigoidine and a type III secretion system (T3SS) (Enard et al. 1988; Collmer and Bauer 1994; Reverchon et al. 2002; Yang et al. 2002; Yap et al. 2005; Zhou et al. 2015). Additionally, a family of phytotoxins/antibiotics termed zeamines constitutes a key virulence determinant that appears specific to D. oryzae and some Dickeya solani strains (Wu et al. 2010; Zhou et al. 2011; Cheng et al. 2013; Zhou et al. 2015, 2024). Furthermore, bacterial motility is also known to play a key role in D. oryzae systemic infection (Chen et al. 2020).

Recent research shows that virulence factor production in D. oryzae is controlled by several QS systems. Among them, the AHL‐based QS system influences motility and biofilm formation (Hussain et al. 2008). Putrescine, synthesized by SpeA of either bacterium or plant origin, also acts as a QS signal or a cross‐kingdom signal affecting motility, biofilm and virulence (Shi et al. 2019). The Vfm QS system, which is independent of the AHL‐based QS system, is crucial for the production of zeamine and PCWDEs, and essential for bacterial virulence (Lv et al. 2019; Zhou et al. 2024). Collectively, these findings underscore the important role of QS in D. oryzae EC1 virulence. Furthermore, the Vfm QS plays a dominant role in virulence regulation (Lv et al. 2019). However, the upstream regulatory mechanisms and components integrating environmental cues into these QS networks remain largely unknown.

The MarR‐family transcriptional regulator OhrR (organic hydroperoxide reductase regulator) is critical for several virulence associated traits in D. oryzae , including zeamine production, cellulase secretion, biofilm formation and motility (Lv et al. 2022). Another MarR‐family member, EmrR (multidrug efflux transporter repressor), similarly influences cellulase secretion, biofilm formation and motility (Lv et al. 2026). Both OhrR and EmrR belong to the MarR family, and their respective mutants show significant virulence attenuation. These observations suggest potential interconnections between OhrR, EmrR and the Vfm QS system, but the nature of these relationships is unclear. In this study, we demonstrated that in D. oryzae , OhrR directly and positively regulates the biosynthesis of Vfm signals, while EmrR directly and negatively regulates Vfm biosynthesis. We also showed that OhrR directly activates emrR transcription. These findings unveil a novel hierarchical regulatory circuit modulating QS and virulence in D. oryzae .

2. Results

2.1. OhrR Is Required for the Full Vfm QS Signalling Activity

Previous work showed that ohrR mutation results in reduced zeamine production and cellulase activity (Lv et al. 2022), and Vfm QS system mutants are impaired in zeamine and PCWDE production (Lv et al. 2019). To determine if OhrR affects Vfm QS signalling activity, we constructed the reporter strain ∆vfmE(uidA) to monitor the Vfm signal production. The reporter strain was co‐cultured separately with wild‐type EC1 or the strain ∆ohrR at a 1:1 ratio, and the bacterial growth (OD600) and fluorescence (β‐glucuronidase reporter [GUS] using 4‐methylumbelliferyl β‐D‐glucuronide as the substrate) were monitored simultaneously. The growth kinetics of strain EC1 and the strain ∆ohrR in co‐culture were comparable. Interestingly, the GUS fluorescence (normalized to cell density) was significantly lower in the co‐cultures containing the ∆ohrR compared to those with wild‐type EC1 (Figure 1), indicating that OhrR is required for the full Vfm QS signal production.

FIGURE 1.

FIGURE 1

Open in a new tab

OhrR is required for the full Vfm quorum‐sensing (QS) signalling activity. Dynamics of Vfm QS signalling activity in co‐culture. The reporter strain ∆vfmE(uidA) was co‐cultured at a 1:1 ratio with wild‐type EC1 (red lines) or the ∆ohrR mutant (blue lines). Growth (OD600, circles) and normalized GUS fluorescence (fluorescence/OD600, squares) were monitored. Data are the representative of three independent experiments.

2.2. OhrR Positively Regulates Expression of the vfm Cluster

Because Vfm signal production was reduced in the strain ∆ohrR, we examined its effect on the expression of key vfm cluster operons. The promoter regions of the signal synthesis genes (vfmA and vfmK), the AraC‐family regulator gene (vfmE) and a signal transport gene (vfmF) (Figure 2a) were fused to the GFP gene in plasmid pROBE‐NT, generating transcriptional fusion reporter constructs PvfmA‐gfp, PvfmK‐gfp, PvfmE‐gfp and PvfmF‐gfp, and introduced into wild‐type EC1 and the ∆ohrR. GFP fluorescence measured at OD600 = 1.0 was significantly lower in the ∆ohrR background for all four promoters compared to the wild‐type background (Figure 2b). Consistent with this, reverse transcription‐quantitative PCR (RT‐qPCR) analysis confirmed that transcript levels of vfmA, vfmE, vfmF and vfmK were significantly reduced in the mutant ∆ohrR compared to wild‐type EC1 (Figure 2c). These results indicate that OhrR positively regulates the transcription of key operons within the vfm cluster.

FIGURE 2.

FIGURE 2

Open in a new tab

OhrR positively regulates expression of the vfm cluster. (a) Gene organization of the vfm gene cluster. (b) Promoter activity of key vfm operons. GFP fluorescence from transcriptional fusions (PvfmA‐gfp, PvfmK‐gfp, PvfmE‐gfp and PvfmF‐gfp) was measured in wild‐type EC1 and the ∆ohrR mutant at OD600 = 1.0. (c) Reverse transcription‐quantitative PCR analysis of vfmA, vfmE, vfmF and vfmK transcript levels in wild‐type EC1 and the ∆ohrR mutant. Expression was normalized to 16S rRNA and is shown relative to wild‐type EC1 (set to 1). Experiments were repeated at least three times in triplicate and error bars indicate stand deviation. ****p < 0.0001, Student's t‐test.

2.3. OhrR Directly Binds to the Promoters of vfm Genes

Given the reduced Vfm signal and gene expression in the strain ∆ohrR, we investigated whether OhrR directly regulates the vfm cluster. Recombinant OhrR protein was expressed in Escherichia coli and purified (Figure 3a). Electrophoretic mobility shift assays (EMSAs) showed that purified OhrR specifically bound to the promoter fragments of vfmA, vfmE, vfmF and vfmK, causing a mobility shift in a concentration‐dependent manner (Figure 3b and Figure S1). Bioinformatic analysis identified a conserved 20‐bp putative OhrR‐binding motif in these promoter regions (Figure 3c and Figure S2). EMSAs with synthesized oligonucleotides containing these motifs (B vfmA , B vfmE , B vfmF and B vfmK ) confirmed that OhrR directly and specifically bound to this conserved sequence (Figure 3d). These data demonstrate that OhrR directly activates the vfm gene expression by binding to a specific promoter motif.

FIGURE 3.

FIGURE 3

Open in a new tab

OhrR directly binds to the promoters of the vfm genes. (a) Purification of recombinant His‐tagged OhrR protein analysed by SDS‐PAGE. (b) Electrophoretic mobility shift assay (EMSA) showing binding of purified OhrR to the promoter fragments of vfmA, vfmE, vfmF and vfmK, with increasing concentrations of OhrR protein (0, 1, 2 μM) being used. (c) Sequence of the conserved OhrR‐binding motif identified in the promoters of vfmA, vfmE, vfmF and vfmK. (d) EMSA validation using synthesized oligonucleotides containing the predicted OhrR‐binding motifs (B vfmA , B vfmE , B vfmF and B vfmK ).

2.4. OhrR Directly Activates the Expression of emrR

As both OhrR and EmrR are MarR‐family regulators affecting virulence traits, we explored their potential regulatory relationship. The emrR mutant is defective in motility and biofilm formation, and EmrR autoregulates its own expression (Lv et al. 2026). We measured the promoter activity of ohrR and emrR using lacZ transcriptional fusions. The activity of the ohrR promoter was unaffected in a ∆emrR background (Figure 4a). In contrast, the activity of the emrR promoter was significantly lower in the strain ∆ohrR than wild‐type EC1 (Figure 4b). RT‐qPCR confirmed that emrR transcript level was reduced approximately 2.4‐fold in the mutant strain ∆ohrR (Figure 4c). Similarly, we also assessed ohrR promoter activity in the ΔohrR and wild‐type strain EC1, and observed that ohrR promoter activity was markedly decreased in the ΔohrR background (Figure S3). Bioinformatic analysis identified a conserved OhrR‐binding motif in the emrR promoter region (Figure 4d). EMSA confirmed that purified OhrR protein directly bound to this motif (Figure 4e). These results demonstrate that OhrR directly and positively regulates the transcription of emrR.

FIGURE 4.

FIGURE 4

Open in a new tab

OhrR directly activates the transcription of emrR. (a, b) β‐Galactosidase activity assays. Promoter activity of (a) ohrR and (b) emrR was measured in the indicated genetic backgrounds using lacZ transcriptional fusions. (c) Reverse transcription‐quantitative PCR analysis of emrR expression in wild‐type EC1 and the ∆ohrR mutant. (d) Sequence of the predicted OhrR‐binding motif in the emrR promoter region. (e) Electrophoretic mobility shift assay confirming the direct binding of purified OhrR protein to the biotin‐labelled oligonucleotide containing the predicted emrR promoter motif. In (a–c), data are the means ± SD from three biological replicates. ****p < 0.0001, Student's t‐test.

2.5. EmrR Directly Represses the Vfm QS System

Because OhrR activates both the Vfm system and emrR expression, we next asked if EmrR could modulate the Vfm signal production. Using the ∆vfmE(uidA) reporter strain in a co‐culture assay, we found that Vfm signalling activity was significantly higher in the presence of the mutant strain ∆emrR compared to wild‐type EC1, despite similar growth curves (Figure 5a). Promoter‐GFP fusion assays showed that the activities of P vfmA , P vfmK , P vfmE and P vfmF were all elevated in the ∆emrR background compared to the wild type (Figure 5b). RT‐qPCR confirmed the increased transcript levels of these vfm genes in the mutant strain ∆emrR (Figure 5c). Recombinant EmrR protein was purified (Figure 5d). A conserved 21‐bp EmrR‐binding motif was identified in the promoters of vfmA, vfmE, vfmF and vfmK (Figure 5e), and EMSA verified that EmrR specifically and directly bound to this motif (Figure 5f and Figure S4). These findings establish EmrR as a direct negative regulator of the vfm gene cluster.

FIGURE 5.

FIGURE 5

Open in a new tab

EmrR directly represses the Vfm quorum‐sensing (QS) system. (a) Vfm signal activity in co‐culture of the ∆vfmE(uidA) reporter with wild‐type EC1 (red lines) or the ∆emrR mutant (blue lines). (b) Promoter activity of the vfm genes (PvfmA‐gfp, PvfmK‐gfp, PvfmE‐gfp and PvfmF‐gfp) in wild‐type EC1 and the ∆emrR mutant. (c) Reverse transcription‐quantitative PCR (RT‐qPCR) analysis of vfm gene expression in the ∆emrR mutant relative to wild‐type EC1. (d) Purification of recombinant His‐tagged EmrR protein (SDS‐PAGE). (e) Candidate EmrR‐binding motif identified in the promoters of vfmA, vfmF, vfmE and vfmK.  (f) Electrophoretic mobility shift assay (EMSA) showing binding of purified EmrR to biotin‐labelled oligonucleotides containing the predicted binding motif. All experiments were conducted in triplicate and independently replicated at least three times. In (a–c), data are the means ± SD from three biological replicates. *p < 0.1, ***p < 0.0001, ****p < 0.0001, Student's t‐test.

2.6. Phenotypic Analysis of the ohrR and emrR Double Mutant

Given the opposing effects of OhrR (activator) and EmrR (repressor) on the Vfm system, we constructed a double‐deletion strain ∆ohrRemrR to assess their combined impact. The double mutant ∆ohrRemrR showed a more severe reduction in extracellular cellulase activity than either single‐mutant strain (∆ohrR and ∆emrR) (Figure 6a). Biofilm formation was significantly impaired in all three mutants (∆ohrR, ∆emrR and ∆ohrRemrR) to a similar degree (Figure 6b). Swimming motility was increased in ∆ohrR, decreased in ∆emrR, and restored to near wild‐type levels in the double‐mutant strain ∆ohrRemrR (Figure 6c). The single‐mutant strain ∆emrR maintained wild‐type strain EC1 zeamine levels. Zeamine production was significantly decreased in the mutant strain ∆ohrR, and the double‐deletion mutant strain ∆ohrRemrR displayed zeamine levels comparable to the single‐mutant strain ∆ohrR (Figure 6d). The potato slice inoculation test demonstrated that, in comparison with the single‐mutant strain ∆ohrR and ∆emrR, the double‐deletion mutant strain ∆ohrRemrR exhibited a further diminished capacity to cause maceration on potato slices (Figure 6e,f). These complex phenotypic variations suggest that OhrR and EmrR form a regulatory circuit that finely tunes different virulence factors outputs, with EmrR providing feedback inhibition on the OhrR–Vfm axis.

FIGURE 6.

FIGURE 6

Open in a new tab

Phenotypic characterization of single and double ohrR emrR mutants. (a) Extracellular cellulase activity. (b) Biofilm formation (crystal violet staining). (c) Swimming motility. (d) Zeamine production. (e) Maceration of potato tuber slices by wild‐type EC1, ∆ohrR, ∆emrR and ∆ohrRemrR strains. Data are the means ± SD from three independent experiments. Different letters above bars indicate statistically significant differences (p < 0.05, one‐way ANOVA with post hoc multiple comparisons. Pairwise comparisons included wild‐type EC1 versus each mutant, and comparisons among ΔohrR, ΔemrR and ΔohrRemrR mutants. Different letters represent significant differences).

3. Discussion

This study reveals a precise transcriptional cascade in which the MarR‐family regulators OhrR and EmrR hierarchically and antagonistically modulate the Vfm QS system to control virulence in D. oryzae EC1. We demonstrate that OhrR directly activates the vfm gene cluster (Figures 1, 2, 3) and simultaneously induces the expression of emrR (Figure 4). EmrR then acts as a direct negative feedback regulator of the same vfm cluster (Figure 6). For the regulation of cellulase activity, OhrR and EmrR act congruently as negative regulators, with the double mutant exhibiting an additive effect (Figures 6 and 7). This ‘activation–inhibition’ dual‐layer regulatory circuit (Figure 7) provides a framework for understanding how plant pathogens achieve precise spatiotemporal control of virulence through complex transcriptional reprogramming.

FIGURE 7.

FIGURE 7

Open in a new tab

Schematic model of the OhrR–EmrR regulatory circuit modulating the Vfm quorum‐sensing (QS) system in Dickeya oryzae. OhrR, potentially activated by environmental signals (e.g., oxidative stress), directly binds to and activates the transcription of key operons within the vfm cluster (vfmAZBCD, vfmKLMNOPQRSTUVW, vfmE and vfmFG), promoting Vfm signal synthesis, regulation and transport. Simultaneously, OhrR directly activates the expression of emrR. EmrR protein then acts as a feedback inhibitor, directly binding to the same vfm promoter regions to repress their transcription, preventing over‐activation. This ‘activation–inhibition’ cascade allows for precise, dynamic control of the Vfm QS system and downstream virulence factor production. Dashed lines indicate the putative regulatory patterns that they have not yet been experimentally verified. PWDEs, plant cell wall‐degrading enzymes; T1SS, type I secretion system; T2SS, type II secretion system.

Notably, the predicted OhrR and EmrR binding motifs within the vfm promoters display relatively low sequence conservation across different target sites (Figures 3c and 5e). Poorly conserved or degenerate binding sequences are frequently observed among MarR family transcriptional regulators, and sequence conservation alone is insufficient to confirm authentic protein–DNA interactions (Nazaret et al. 2023). Our cross‐EMSA and competitive EMSA assays nevertheless support the specific binding of OhrR and EmrR to these vfm promoter fragments (Figures [Link], [Link] and S5), reinforcing that biochemical validation is critical to verify predicted binding sites when motif conservation is limited. Similar observations have been reported for other MarR type regulators, where functional binding sites deviate substantially from consensus motifs obtained from different bacterial species (Deochand and Grove 2017; Nazaret et al. 2023).

A central finding places OhrR and EmrR upstream of the previously characterized VfmIH–VfmE signal transduction axis (Nasser et al. 2013; Lv et al. 2019). OhrR acts as a master switch by directly binding to the promoters of genes for signal synthesis (vfmA and vfmK), regulation (vfmE) and transport (vfmF) to initiate and amplify the Vfm‐QS activity (Figure 3). The absence of OhrR results in a substantial collapse of Vfm signalling and bacterial virulence (Lv et al. 2022). A key feature of this model is the cascading regulatory relationship between OhrR and EmrR (Figure 7). Direct activation of emrR by OhrR establishes an intrinsic feedback loop. Accumulated EmrR subsequently dampens OhrR‐dependent activation by repressing the same set of vfm promoters. This regulatory architecture has two major functional consequences. First, it supports dynamic balance and pulsed virulence responses. Instead of a simple on–off switch, the circuit enables dynamically balanced, potentially pulsed virulence outputs. OhrR can respond to environmental oxidative cues (Duarte and Latour 2010; Garnica et al. 2017; Meireles et al. 2022) and rapidly trigger virulence gene expression. Concurrent induction of EmrR restrains metabolically costly virulence factor production, which may facilitate immune evasion or optimize resource allocation during later infection stages. Second, this module constitutes a signal integration mode. As typical MarR‐family sensors, OhrR likely perceives host‐derived stress signals such as reactive oxygen species (ROS) and integrates these inputs into the QS regulatory network. The OhrR–EmrR hierarchical cascade (Figure 7) therefore allows D. oryzae to fine‐tune virulence according to both population density and prevailing environmental conditions.

Our results, together with previous studies, highlight the complexity of QS regulation within Dickeya. First, there is considerable specificity and strain‐level variation. The Vfm signal molecules differ between D. oryzae and D. dadantii (Lv et al. 2019), and polymorphisms in vfmO/P generate strain‐specific signal analogues (Hugouvieux‐Cotte‐Pattat et al. 2022). Differences in ohrR/emrR expression, protein activity or binding site sequences among strains may further shape these strain‐specific regulatory patterns. Second, cross‐system interactions vary across species. In D. solani, the Vfm and AHL‐based Exp systems do not synergistically control PCWDEs or pathogenicity, and Vfm exerts a more dominant role than in other Dickeya species (Potrykus et al. 2018). Our findings on the internal fine‐tuning of the Vfm system emphasize its central importance, with dedicated regulatory modules such as OhrR–EmrR enabling precise output control. Third, Vfm is embedded within larger global regulatory networks. The Vfm system receives inputs from multiple global regulators including Fis, IHF and PecS (Nasser et al. 2013; Lv et al. 2018; Chen, Hu, et al. 2022; Chen, Li, et al. 2022). The addition of the redox sensing OhrR and its downstream EmrR expands our understanding of how Dickeya integrates diverse environmental signals to optimize virulence.

Beyond elucidating this novel regulatory mechanism, the OhrR–EmrR–Vfm cascade raises questions regarding its evolutionary advantage and its potential as a target for disease control strategies. The ‘activation–inhibition’ circuit likely provides fitness benefits within the dynamic host apoplast, where pathogens encounter fluctuating stressors such as ROS bursts associated with pathogen‐associated molecular pattern (PAMP)‐triggered immunity (Torres et al. 2006; Mittler et al. 2011). The OhrR–EmrR regulatory circuit may equip D. oryzae with a ‘measured aggression’ infection strategy. OhrR, which can sense oxidative stress (Fuangthong and Helmann 2002), triggers rapid virulence gene activation to establish infection and counteract host defences. OhrR‐dependent induction of EmrR then acts as an intrinsic dosage controller, preventing energetically wasteful overproduction of PCWDEs and toxins. Unrestricted production of these factors could trigger severe host tissue damage and activate systemic host defences (Boller and He 2009). Such fine‐tuning may facilitate transition from initial invasion to sustained colonization by balancing virulence investment with immune evasion. This layered regulatory logic is reminiscent of virulence circuits in animal bacterial pathogens, pointing toward convergent evolution of transcription‐based resilience for host adaptation (Galán 2021).

In summary, we have identified and characterized a cascade of transcriptional hierarchy involving OhrR and EmrR that might precisely and dynamically govern the central Vfm QS pathway in D. oryzae . OhrR serves as an environmental cue detector and system initiator, while EmrR functions as an internal feedback modulator. This discovery not only expands the functional repertoire of MarR regulators into the realm of bacterial cell–cell communication but, more importantly, reveals that plant pathogens employ multilayered, logic‐gated regulatory circuits far beyond simple density sensing to achieve optimal virulence expression.

4. Experimental Procedures

4.1. Bacterial Strains and Growth Conditions

Bacterial strains and plasmids used in this study are listed in Table 1. E. coli strains were cultured at 37°C, and D. oryzae EC1 and its derivatives were cultured at 30°C. Lysogeny broth (LB), minimal medium (MM) or LS5 medium (Cheng et al. 2013; Liao et al. 2014) were used as indicated. Antibiotics were used at the following concentrations: polymyxin B sulphate (PB, 25 μg/mL), streptomycin (Str, 50 μg/mL), kanamycin (Kan, 50 μg/mL) and ampicillin (Amp, 100 μg/mL).

TABLE 1.

Strains and plasmids used in this study.

Strains or plasmids Relevant phenotypes and characteristics a Source or references
Strains
EC1 Wild type of Dickeya oryzae, PBr Hussain et al. (2008)
EC1ΔlacZ lacZ in‐frame deletion mutant derived from EC1, PBr This research
ΔemrR emrR in‐frame deletion mutant derived from EC1, Pbr Lv et al. (2026)
ΔemrR ΔlacZ emrR in‐frame deletion mutant derived from EC1ΔlacZ , PBr Lv et al. (2026)
ΔohrR ohrR in‐frame deletion mutant derived from EC1, PBr Lv et al. (2022)
ΔohrR ΔlacZ ohrR in‐frame deletion mutant derived from EC1ΔlacZ , PBr This research
ΔohrRΔemrR ohrR and emrR double in‐frame deletion mutant from EC1, PBr This research
EC1ΔlacZ (pLME‐emrR) The EC1ΔlacZ was transformed with plasmid pLAME‐emrR, PBr, Ampr This research
ΔemrR ΔlacZ (pLME‐ohrR) The ΔemrR ΔlacZ was transformed with plasmid pLAME‐ohrR, PBr, Ampr This research
EC1ΔlacZ (pLME‐ohrR) The EC1ΔlacZ was transformed with plasmid pLAME‐ohrR, PBr, Ampr This research
ΔohrR ΔlacZ (pLME‐emrR) The ΔohrR ΔlacZ was transformed with plasmid pLAME‐emrR, PBr, Ampr This research
EC1 (pROBE‐vfmA) The EC1 was transformed with plasmid pROBE‐vfmA, PBr, Kanr This research
EC1 (pROBE‐vfmE) The EC1 was transformed with plasmid pROBE‐vfmE, PBr, Kanr This research
EC1 (pROBE‐vfmF) The EC1 was transformed with plasmid pROBE‐vfmF, PBr, Kanr This research
EC1 (pROBE‐vfmK) The EC1 was transformed with plasmid pROBE‐vfmK, PBr, Kanr This research
ΔohrR (pROBE‐vfmA) The ΔohrR was transformed with plasmid pROBE‐vfmA, PBr, Kanr This research
ΔohrR (pROBE‐vfmE) The ΔohrR was transformed with plasmid pROBE‐vfmE, PBr, Kanr This research
ΔohrR (pROBE‐vfmF) The ΔohrR was transformed with plasmid pROBE‐vfmF, PBr, Kanr This research
ΔohrR (pROBE‐vfmK) The ΔohrR was transformed with plasmid pROBE‐vfmK, PBr, Kanr This research
ΔemrR (pROBE‐vfmA) The ΔemrR was transformed with plasmid pROBE‐vfmA, PBr, Kanr This research
ΔemrR (pROBE‐vfmE) The ΔemrR was transformed with plasmid pROBE‐vfmE, PBr, Kanr This research
ΔemrR (pROBE‐vfmF) The ΔemrR was transformed with plasmid pROBE‐vfmF, PBr, Kanr This research
ΔemrR (pROBE‐vfmK) The ΔemrR was transformed with plasmid pROBE‐vfmK, PBr, Kanr This research
EC1 (pROBE‐ohrR) The EC1 was transformed with plasmid pROBE‐ohrR, PBr, Kanr This research
ΔohrR (pROBE‐ohrR) The ΔohrR was transformed with plasmid pROBE‐ohrR, PBr, Kanr This research
ΔvfmE (uidA) The reporter strain of VFM signal, PBr This research
DH5α Escherichia coli strain as host for plasmid constructs derived from pBBR1‐MCS4 Zhou et al. (2011)
HB101 (pRK2013) E. coli, thr leu thi recA hsdR hsdM pro, Kanr Zhou et al. (2011)
CC118λ E. coli strain as host for plasmid constructs derived from pKNG101 Zhou et al. (2011)
BL21 (DE) E. coli strain as host for protein prokaryotic expression plasmids Zhou et al. (2011)
Plasmids
pKNG101 Knockout vector, Strr Zhou et al. (2011)
pLME A promote probe vector that contains lacZ, Ampr Lv et al. (2026)
pLME‐emrR The promoter probe vector pLME was fused the promoter region of emrR, Ampr This research
pLME‐ohrR The promoter probe vector pLME was fused the promoter region of ohrR, Ampr This research
pROBE‐NT A promoter probe vector that contains gfp, Kanr This research
pROBE‐vfmA The promoter probe vector pROBE‐NT was fused the promoter region of vfmA, Kanr This research
pROBE‐vfmE The promoter probe vector pROBE‐NT was fused the promoter region of vfmE, Kanr This research
pROBE‐vfmF The promoter probe vector pROBE‐NT was fused the promoter region of vfmF, Kanr This research
pROBE‐vfmK The promoter probe vector pROBE‐NT was fused the promoter region of vfmK Kanr This research
pROBE‐ohrR The promoter probe vector pROBE‐NT was fused the promoter region of ohrR Kanr This research
pCold‐SUMO‐M Protein prokaryotic expression vector, Ampr Liu et al. (2017)
pCold‐ohrR pCold‐SUMO‐M carries the ohrR coding region, Ampr This research
pET30a Protein prokaryotic expression vector, Kanr Lv et al. (2026)
pET30a‐emrR pET30a carries the emrR coding region, Kanr This research
Open in a new tab
a

PBr, Ampr, Kanr, Strr = resistance to polymyxin B sulphate, ampicillin, kanamycin, and streptomycin, respectively.

4.2. Construction of Deletion Mutants

In‐frame deletion mutants were constructed using the suicide vector pKNG101 via triparental mating and sacB‐based counterselection as previously described (Lv et al. 2018, 2019). Flanking regions (approximately 600 bp) of the target genes were amplified with specific primers (Table S1) and sequentially cloned into pKNG101.

4.3. Construction and Measurement of GFP Reporter Strains

Promoter regions (approximately 300–500 bp upstream of the start codon) were amplified and cloned into the BamHI/HindIII sites of the GFP reporter vector pROBE‐NT. Recombinant plasmids were transferred into D. oryzae strains via triparental mating. Overnight cultures were diluted 1:100 in fresh LB and grown with shaking at 28°C. GFP fluorescence (excitation 488 nm, emission 509 nm) and OD600 were measured periodically using a microplate reader (BioTek). Data from at least three independent biological replicates are presented.

4.4. GUS Activity Assay for Vfm Signal Detection

A Vfm signal reporter strain, ∆vfmE (uidA), was constructed by inserting a promoterless uidA (GUS) gene into the vfmE locus (Lv et al. 2019). To measure signal activity in trans, culture supernatants (20% v/v final) from test strains were added to the reporter culture. For co‐culture assays, the reporter strain was mixed 1:1 with the test strain. Cultures containing 100 μg/mL 4‐methylumbelliferyl β‐D‐glucuronide (MUG) were incubated overnight. Fluorescence (excitation 365 nm, emission 455 nm) and OD600 were measured at the time points as indicated.

4.5. Detection of Extracellular Enzyme Activity

Qualitative assays of extracellular cellulase (Cel) activity were conducted on LB agar plates supplemented with 0.5% carboxymethyl cellulose sodium, as previously described (Chatterjee et al. 1995; Lv et al. 2018). Plates were incubated at 28°C for 16 h prior to staining with Congo red. Excess dye was washed off using 1 M NaCl solution, and the radius of each cellulolytic degradation halo was recorded.

4.6. Bioassay for Zeamine Production

Zeamine production was assessed using an E. coli DH5α growth inhibition assay on LS5 agar plates, following established protocols (Lv et al. 2018, 2019).

4.7. Motility and Biofilm Assays

Swimming motility was tested on 0.2% agar plates containing 1% tryptone and 0.5% NaCl. Plates were point‐inoculated and incubated at 28°C for 15 h before measuring migration diameter. Biofilm formation was quantified in static SOBG medium using the crystal violet staining method as described (Chen et al. 2020). All assays were performed with at least three biological replicates.

4.8. β‐Galactosidase Activity Assay

The promoter regions of ohrR and emrR were fused separately to lacZ in plasmid pLME (Lv et al. 2026). The resulting plasmids were introduced into the relevant D. oryzae ∆lacZ strain. β‐Galactosidase activity was measured in the cells grown to mid‐exponential phase (OD600 = 1.0) using o‐nitrophenyl‐β‐D‐galactopyranoside (ONPG) as a substrate as described (Meng et al. 2020; Lv et al. 2026). Activity is expressed in Miller units.

4.9. RNA Extraction and RT‐qPCR

Total RNA was extracted from bacterial cultures at OD600 = 1.0 using the SV Total RNA Isolation System (Promega). cDNA was synthesized using the FastQuant RT Kit (Tiangen). qPCR was performed with SuperReal PreMix Colour SYBR Green (Tiangen) on a QuantStudio 6 Flex system. The 16S rRNA gene was used as an internal reference. Gene expression changes were calculated using the 2−∆∆Ct method (Livak and Schmittgen 2001). Data represent at least two independent runs with four biological replicates each.

4.10. Protein Purification and EMSA

The coding sequences of ohrR and emrR were cloned into pCold‐SUMO‐M or pET‐30a vectors (Liu et al. 2017; Xu et al. 2023; Lv et al. 2026), respectively. Recombinant His‐tagged proteins were expressed in E. coli BL21 (DE3) and purified using Ni‐NTA resin (Clontech). Promoter DNA fragments or biotin‐labelled oligonucleotides were incubated with purified proteins in binding buffer for 30 min at room temperature. Protein–DNA complexes were resolved on 6% non‐denaturing polyacrylamide gels in 0.5× TBE buffer and detected using the Chemiluminescent Nucleic Acid Detection Module (Thermo Fisher Scientific).

4.11. Prediction of OhrR and EmrR Binding Motifs

Putative OhrR and EmrR binding sites in the promoter regions of vfmA, vfmE, vfmF and vfmK were predicted by sequence alignment using known motifs (implemented via Clustal Omega v. 6.0). The OhrR motif was the published Bacillus subtilis OhrR binding site (5′‐TTAATTGTGTACAATTTA‐3′; Fuangthong et al. 2001), and the EmrR seed was the E. coli EmrR consensus (5′‐CTGTCGTTACTATATCGGCTG‐3′; Xiong et al. 2000). Approximately 500 bp sequences upstream of each gene start codon were aligned against the seed sequences to retrieve candidate binding fragments. The obtained candidate sites were subsequently verified via EMSA. Sequence logos of our identified OhrR and EmrR motifs were generated using WebLogo v. 3.

4.12. Statistical Analysis

All experiments were performed with at least three independent biological replicates. Data are presented as mean ± SD. Statistical significance was determined using Student's t‐test or one‐way ANOVA with appropriate post hoc tests, as indicated in the figure legends. Differences were considered significant at p < 0.05. All analyses were performed using GraphPad Prism v. 8.0 software.

Author Contributions

Wenqi Huang: investigation. Ziwei Meng: investigation. Zhongge Liu: investigation. Weihan Gu: methodology. Ming Hu: methodology. Tao Zhuo: formal analysis, methodology. Lianhui Zhang: writing – review and editing, methodology, funding acquisition. Mingfa Lv: writing – review and editing, writing – original draft, supervision, funding acquisition, formal analysis, project administration.

Funding

This work was supported by grants from the National Natural Science Foundation of China (No. 32570212, U22A20480) and the Science Foundation of Fujian Provincial Science & Technology Department (2023J01440, 2022J01128).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Specific competitive electrophoretic mobility shift assays (EMSAs) for OhrR and EmrR binding to the vfmK promoter, with the pelE promoter fragment serving as a negative control.

MPP-27-e70354-s002.docx (168KB, docx)

Figure S2: Localization of OhrR and EmrR binding motifs within the vfmA, vfmE, vfmF and vfmK promoter regions. Schematic representation showing the positions of identified OhrR and EmrR binding motifs upstream of the translation start sites of vfmA, vfmE, vfmF and vfmK.

MPP-27-e70354-s005.docx (241.6KB, docx)

Figure S3: ohrR promoter activity assay in wild‐type EC1 and ΔohrR. p < 0.0001 (t‐test).

MPP-27-e70354-s006.docx (29.1KB, docx)

Figure S4: Cross‐electrophoretic mobility shift assay (EMSA) validation of binding specificity for OhrR and EmrR toward their respective sites in the vfmA promoter.

MPP-27-e70354-s004.docx (235.3KB, docx)

Figure S5: Electrophoretic mobility shift assay (EMSAs) using the vfmA promoter probe with OhrR and EmrR at different molar ratios.

MPP-27-e70354-s003.docx (73.7KB, docx)

Table S1: Primers used in this study.

MPP-27-e70354-s001.docx (21.8KB, docx)

Contributor Information

Lianhui Zhang, Email: lhzhang01@scau.edu.cn.

Mingfa Lv, Email: mingfalv2010@126.com.

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

References

  1. Boller, T. , and He S. Y.. 2009. “Innate Immunity in Plants: An Arms Race Between Pattern Recognition Receptors in Plants and Effectors in Microbial Pathogens.” Science 324: 742–744. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Chatterjee, A. , Cui Y., Liu Y., Dumenyo C. K., and Chatterjee A. K.. 1995. “Inactivation of rsmA Leads to Overproduction of Extracellular Pectinases, Cellulases, and Proteases in Erwinia carotovora subsp. carotovora in the Absence of the Starvation/Cell Density‐Sensing Signal, N‐(3‐Oxohexanoyl)‐L‐Homoserine Lactone.” Applied and Environmental Microbiology 61: 1959–1967. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Chen, S. , Hu M., Hu A., et al. 2022. “The Integration Host Factor Regulates Multiple Virulence Pathways in Bacterial Pathogen Dickeya zeae MS2.” Molecular Plant Pathology 23: 1487–1507. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Chen, Y. , Li Y., Zhu M., et al. 2022. “The GacA‐GacS Type Two‐Component System Modulates the Pathogenicity of Dickeya oryzae EC1 Mainly by Regulating the Production of Zeamines.” Molecular Plant–Microbe Interactions 35: 369–379. [DOI] [PubMed] [Google Scholar]
  5. Chen, Y. , Zhou J., Lv M., Liang Z., Parsek M. R., and Zhang L. H.. 2020. “Systematic Analysis of c‐di‐GMP Signaling Mechanisms and Biological Functions in Dickeya zeae EC1.” mBio 11: e02993‐20. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Cheng, Y. , Liu X., An S., et al. 2013. “A Nonribosomal Peptide Synthase Containing a Stand‐Alone Condensation Domain Is Essential for Phytotoxin Zeamine Bosynthesis.” Molecular Plant–Microbe Interactions 26: 1294–1301. [DOI] [PubMed] [Google Scholar]
  7. Collmer, A. , and Bauer D. W.. 1994. “ Erwinia chrysanthemi and Pseudomonas syringae : Plant Pathogens Trafficking in Extracellular Virulence Proteins.” Current Topics in Microbiology and Immunology 192: 43–78. [DOI] [PubMed] [Google Scholar]
  8. Deng, Y. , Schmid N., Wang C., et al. 2012. “Cis‐2‐Dodecenoic Acid Receptor RpfR Links Quorum‐Sensing Signal Perception With Regulation of Virulence Through Cyclic Dimeric Guanosine Monophosphate Turnover.” Proceedings of the National Academy of Sciences of the United States of America 109: 15479–15484. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Deng, Y. , Wu J., Eberl L., and Zhang L. H.. 2010. “Structural and Functional Characterization of Diffusible Signal Factor Family Quorum‐Sensing Signals Produced by Members of the Burkholderia cepacia Complex.” Applied and Environmental Microbiology 76: 4675–4683. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Deng, Y. , Wu J., Tao F., and Zhang L. H.. 2011. “Listening to a New Language: DSF‐Based Quorum Sensing in Gram‐Negative Bacteria.” Chemical Reviews 111: 160–173. [DOI] [PubMed] [Google Scholar]
  11. Deochand, D. K. , and Grove A.. 2017. “MarR Family Transcription Factors: Dynamic Variations on a Common Scaffold.” Critical Reviews in Biochemistry and Molecular Biology 52: 595–613. [DOI] [PubMed] [Google Scholar]
  12. Diolez, A. , and Coleno A.. 1985. “Mu‐lac Insertion‐Directed Mutagenesis in a Pectate Lyase Gene of Erwinia chrysanthemi .” Journal of Bacteriology 163: 913–917. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Duarte, V. , and Latour J. M.. 2010. “PerR vs OhrR: Selective Peroxide Sensing in Bacillus subtilis .” Molecular BioSystems 6: 316–323. [DOI] [PubMed] [Google Scholar]
  14. Enard, C. , Diolez A., and Expert D.. 1988. “Systemic Virulence of Erwinia chrysanthemi 3937 Requires a Functional Iron Assimilation System.” Journal of Bacteriology 170: 2419–2426. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Federle, M. J. , and Bassler B. L.. 2003. “Interspecies Communication in Bacteria.” Journal of Clinical Investigation 112: 1291–1299. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Fuangthong, M. , Atichartpongkul S., Mongkolsuk S., and Helmann J. D.. 2001. “OhrR Is a Repressor of ohrA, a Key Organic Hydroperoxide Resistance Determinant in Bacillus subtilis .” Journal of Bacteriology 183: 4134–4141. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Fuangthong, M. , and Helmann J. D.. 2002. “The OhrR Repressor Senses Organic Hydroperoxides by Reversible Formation of a Cysteine‐Sulfenic Acid Derivative.” Proceedings of the National Academy of Sciences of the United States of America 99: 6690–6695. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Fuqua, C. , and Greenberg E. P.. 2002. “Listening in on Bacteria: Acyl‐Homoserine Lactone Signaling.” Nature Reviews Molecular Cell Biology 3: 685–695. [DOI] [PubMed] [Google Scholar]
  19. Galán, J. E. 2021. “ Salmonella typhimurium and Inflammation: A Pathogen‐Centric Affair.” Nature Reviews Microbiology 19: 716–725. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Garnica, O. A. , Das K., and Dhandayuthapani S.. 2017. “OhrR of Mycobacterium smegmatis Senses and Responds to Intracellular Organic Hydroperoxide Stress.” Scientific Reports 7: 3922. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Hugouvieux‐Cotte‐Pattat, N. , Royer M., Gueguen E., et al. 2022. “Specificity and Genetic Polymorphism in the Vfm Quorum Sensing System of Plant Pathogenic Bacteria of the Genus Dickeya .” Environmental Microbiology 24: 1467–1483. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Hussain, M. B. , Zhang H. B., Xu J. L., Liu Q., Jiang Z., and Zhang L. H.. 2008. “The Acyl‐Homoserine Lactone‐Type Quorum‐Sensing System Modulates Cell Motility and Virulence of Erwinia chrysanthemi pv. zeae .” Journal of Bacteriology 190: 1045–1053. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Lee, J. , Wu J., Deng Y., et al. 2013. “A Cell‐Cell Communication Signal Integrates Quorum Sensing and Stress Response.” Nature Chemical Biology 9: 339–343. [DOI] [PubMed] [Google Scholar]
  24. Lee, J. , and Zhang L.. 2015. “The Hierarchy Quorum Sensing Network in Pseudomonas aeruginosa .” Protein & Cell 6: 26–41. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Liao, L. , Cheng Y., Liu S., et al. 2014. “Production of Novel Antibiotics Zeamines Through Optimizing Dickeya zeae Fermentation Conditions.” PLoS One 9: e116047. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Liu, G. D. , Zhou S. F., Ding X. C., et al. 2017. “Soluble Expression of Recombinant cMyc, Klf4, Oct4, and Sox2 Proteins in Bacteria and Transduction Into Living Cells.” International Journal of Ophthalmology 10: 560–566. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Livak, K. J. , and Schmittgen T. D.. 2001. “Analysis of Relative Gene Expression Data Using Real‐Time Quantitative PCR and the 2−∆∆CT Method.” Methods 25: 402–408. [DOI] [PubMed] [Google Scholar]
  28. Lv, M. , Chen Y., Hu M., et al. 2022. “OhrR Is a Central Transcriptional Regulator of Virulence in Dickeya zeae .” Molecular Plant Pathology 23: 45–59. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Lv, M. , Chen Y., Liao L., et al. 2018. “Fis Is a Global Regulator Critical for Modulation of Virulence Factor Production and Pathogenicity of Dickeya zeae .” Scientific Reports 8: 341. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Lv, M. , Hu M., Li P., Jiang Z., Zhang L. H., and Zhou J.. 2019. “A Two‐Component Regulatory System VfmIH Modulates Multiple Virulence Traits in Dickeya zeae .” Molecular Microbiology 112: 1493–1509. [DOI] [PubMed] [Google Scholar]
  31. Lv, M. , Huang W., Meng Z., et al. 2026. “The MarR Family Transcriptional Regulator EmrR Negatively Regulates the Type III Secretion System (T3SS) and Positively Modulates Pathogenicity in Dickeya oryzae .” Molecular Plant Pathology 27: e70255. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Meireles, D. A. , da Silva Neto J. F., Domingos R. M., Alegria T. G. P., Santos L. C. M., and Netto L. E. S.. 2022. “Ohr–OhrR, a Neglected and Highly Efficient Antioxidant System: Structure, Catalysis, Phylogeny, Regulation, and Physiological Roles.” Free Radical Biology and Medicine 185: 6–24. [DOI] [PubMed] [Google Scholar]
  33. Meng, X. , Ahator S. D., and Zhang L. H.. 2020. “Molecular Mechanisms of Phosphate Stress Activation of Pseudomonas aeruginosa Quorum Sensing Systems.” mSphere 5: e00119‐20. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Mittler, R. , Vanderauwera S., Suzuki N., et al. 2011. “ROS Signaling: The New Wave?” Trends in Plant Science 16: 300–309. [DOI] [PubMed] [Google Scholar]
  35. Nassar, A. , Bertheau Y., Dervin C., Narcy J. P., and Lemattre M.. 1994. “Ribotyping of Erwinia chrysanthemi Strains in Relation to Their Pathogenic and Geographic Distribution.” Applied and Environmental Microbiology 60: 3781–3789. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Nasser, W. , Dorel C., Wawrzyniak J., et al. 2013. “Vfm a New Quorum Sensing System Controls the Virulence of Dickeya dadantii .” Environmental Microbiology 15: 865–880. [DOI] [PubMed] [Google Scholar]
  37. Nazaret, F. , Alloing G., Mandon K., and Frendo P.. 2023. “MarR Family Transcriptional Regulators and Their Roles in Plant‐Interacting Bacteria.” Microorganisms 11: 1936. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Ohtani, K. , Hayashi H., and Shimizu T.. 2002. “The luxS Gene Is Involved in Cell‐Cell Signalling for Toxin Production in Clostridium perfringens .” Molecular Microbiology 44: 171–179. [DOI] [PubMed] [Google Scholar]
  39. Passos‐da‐Silva, D. , Schofield M. C., Parsek M. R., and Tseng B. S.. 2017. “An Update on the Sociomicrobiology of Quorum Sensing in Gram‐Negative Biofilm Development.” Pathogens 6: 51. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Pemberton, C. L. , Whitehead N. A., Sebaihia M., et al. 2005. “Novel Quorum‐Sensing‐Controlled Genes in Erwinia carotovora subsp. carotovora: Identification of a Fungal Elicitor Homologue in a Soft‐Rotting Bacterium.” Molecular Plant–Microbe Interactions 18: 343–353. [DOI] [PubMed] [Google Scholar]
  41. Potrykus, M. , Hugouvieux‐Cotte‐Pattat N., and Lojkowska E.. 2018. “Interplay of Classic Exp and Specific Vfm Quorum Sensing Systems on the Phenotypic Features of Dickeya solani Strains Exhibiting Different Virulence Levels.” Molecular Plant Pathology 19: 1238–1251. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Reverchon, S. , and Nasser W.. 2013. “ Dickeya Ecology, Environment Sensing and Regulation of Virulence Programme.” Environmental Microbiology Reports 5: 622–636. [DOI] [PubMed] [Google Scholar]
  43. Reverchon, S. , Rouanet C., Expert D., and Nasser W.. 2002. “Characterization of Indigoidine Biosynthetic Genes in Erwinia chrysanthemi and Role of This Blue Pigment in Pathogenicity.” Journal of Bacteriology 184: 654–665. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Samson, R. , Legendre J. B., Christen R., Saux M. F., Achouak W., and Gardan L.. 2005. “Transfer of Pectobacterium chrysanthemi (Burkholder et al. 1953) Brenner et al. 1973 and Brenneria paradisi Aca to the Genus Dickeya gen. nov. as Dickeya chrysanthemi comb. nov. and Dickeya paradisiaca comb. nov. and Delineation of Four Novel Species, Dickeya dadantii sp. nov., Dickeya dianthicola sp. nov., Dickeya dieffenbachiae sp. nov. and Dickeya zeae sp. nov.” International Journal of Systematic and Evolutionary Microbiology 55: 1415–1427. [DOI] [PubMed] [Google Scholar]
  45. Shi, Z. , Wang Q., Li Y., et al. 2019. “Putrescine Is an Intraspecies and Interkingdom Cell–Cell Communication Signal Modulating the Virulence of Dickeya zeae .” Frontiers in Microbiology 10: 1950. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Sinha, S. K. , and Prasad M.. 1977. “Bacterial Stalk Rot of Maize, Its Symptoms and Host‐Range.” Zentralblatt für Bakteriologie, Parasitenkunde, Infektionskrankheiten und Hygiene 132: 81–88. [DOI] [PubMed] [Google Scholar]
  47. Torres, M. A. , Jones J. D., and Dangl J. L.. 2006. “Reactive Oxygen Species Signaling in Response to Pathogens.” Plant Physiology 141: 373–378. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Whitehead, N. A. , Barnard A. M., Slater H., Simpson N. J., and Salmond G. P.. 2001. “Quorum‐Sensing in Gram‐Negative Bacteria.” FEMS Microbiology Reviews 25: 365–404. [DOI] [PubMed] [Google Scholar]
  49. Wu, J. , Zhang H. B., Xu J. L., Cox R. J., Simpson T. J., and Zhang L. H.. 2010. “ 13C Labeling Reveals Multiple Amination Reactions in the Biosynthesis of a Novel Polyketide Polyamine Antibiotic Zeamine From Dickeya zeae .” Chemical Communications 46: 333–335. [DOI] [PubMed] [Google Scholar]
  50. Xiong, A. , Gottman A., Park C., Baetens M., Pandza S., and Matin A.. 2000. “The EmrR Protein Represses the Escherichia coli emrRAB Multidrug Resistance Operon by Directly Binding to Its Promoter Region.” Antimicrobial Agents and Chemotherapy 44: 2905–2907. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Xu, Y. , Dong M., Wang Q., et al. 2023. “Soluble Expression of Antimicrobial Peptide BSN‐37 From Escherichia coli by SUMO Fusion Technology.” Protein Journal 42: 563–574. [DOI] [PubMed] [Google Scholar]
  52. Yang, C. H. , Gavilanes‐Ruiz M., Okinaka Y., et al. 2002. “ hrp Genes of Erwinia chrysanthemi 3937 Are Important Virulence Factors.” Molecular Plant–Microbe Interactions 15: 472–480. [DOI] [PubMed] [Google Scholar]
  53. Yap, M. N. , Yang C. H., Barak J. D., Jahn C. E., and Charkowski A. O.. 2005. “The Erwinia chrysanthemi Type III Secretion System Is Required for Multicellular Behavior.” Journal of Bacteriology 187: 639–648. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Zhou, J. , Cheng Y., Lv M., et al. 2015. “The Complete Genome Sequence of Dickeya zeae EC1 Reveals Substantial Divergence From Other Dickeya Strains and Species.” BMC Genomics 16: 571. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Zhou, J. , Hu M., and Zhang L.. 2024. “ Dickeya Diversity and Pathogenic Mechanisms.” Annual Review of Microbiology 78: 621–642. [DOI] [PubMed] [Google Scholar]
  56. Zhou, J. , Zhang H., Wu J., et al. 2011. “A Novel Multidomain Polyketide Synthase Is Essential for Zeamine Production and the Virulence of Dickeya zeae .” Molecular Plant–Microbe Interactions 24: 1156–1164. [DOI] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

Figure S1: Specific competitive electrophoretic mobility shift assays (EMSAs) for OhrR and EmrR binding to the vfmK promoter, with the pelE promoter fragment serving as a negative control.

MPP-27-e70354-s002.docx (168KB, docx)

Figure S2: Localization of OhrR and EmrR binding motifs within the vfmA, vfmE, vfmF and vfmK promoter regions. Schematic representation showing the positions of identified OhrR and EmrR binding motifs upstream of the translation start sites of vfmA, vfmE, vfmF and vfmK.

MPP-27-e70354-s005.docx (241.6KB, docx)

Figure S3: ohrR promoter activity assay in wild‐type EC1 and ΔohrR. p < 0.0001 (t‐test).

MPP-27-e70354-s006.docx (29.1KB, docx)

Figure S4: Cross‐electrophoretic mobility shift assay (EMSA) validation of binding specificity for OhrR and EmrR toward their respective sites in the vfmA promoter.

MPP-27-e70354-s004.docx (235.3KB, docx)

Figure S5: Electrophoretic mobility shift assay (EMSAs) using the vfmA promoter probe with OhrR and EmrR at different molar ratios.

MPP-27-e70354-s003.docx (73.7KB, docx)

Table S1: Primers used in this study.

MPP-27-e70354-s001.docx (21.8KB, docx)

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

Articles from Molecular Plant Pathology are provided here courtesy of Wiley

RESOURCES