Gram-negative bacteria resist antimicrobial agents by a DzrR-mediated envelope stress response
BMC Biology volume 21, Article number: 62 (2023)
Envelope stress responses (ESRs) are critical for adaptive resistance of Gram-negative bacteria to envelope-targeting antimicrobial agents. However, ESRs are poorly defined in a large number of well-known plant and human pathogens. Dickeya oryzae can withstand a high level of self-produced envelope-targeting antimicrobial agents zeamines through a zeamine-stimulated RND efflux pump DesABC. Here, we unraveled the mechanism of D. oryzae response to zeamines and determined the distribution and function of this novel ESR in a variety of important plant and human pathogens.
In this study, we documented that a two-component system regulator DzrR of D. oryzae EC1 mediates ESR in the presence of envelope-targeting antimicrobial agents. DzrR was found modulating bacterial response and resistance to zeamines through inducing the expression of RND efflux pump DesABC, which is likely independent on DzrR phosphorylation. In addition, DzrR could also mediate bacterial responses to structurally divergent envelope-targeting antimicrobial agents, including chlorhexidine and chlorpromazine. Significantly, the DzrR-mediated response was independent on the five canonical ESRs. We further presented evidence that the DzrR-mediated response is conserved in the bacterial species of Dickeya, Ralstonia, and Burkholderia, showing that a distantly located DzrR homolog is the previously undetermined regulator of RND-8 efflux pump for chlorhexidine resistance in B. cenocepacia.
Taken together, the findings from this study depict a new widely distributed Gram-negative ESR mechanism and present a valid target and useful clues to combat antimicrobial resistance.
Bacterial cells frequently encounter divergent toxic agents in hazardous environments. Cell envelope provides the first line of defense for protecting bacterial cells from damages caused by envelope-targeting antimicrobial agents, oxidative reagents, and other extracellular stresses. Envelope stress response (ESR) mechanisms are crucial for maintenance of bacterial envelope homeostasis by activating protective mechanisms against envelope-damaging stresses . Five widespread ESRs, i.e., Bae, Cpx, RpoE (σE), Rcs, and Psp, have been identified in Escherichia coli and other bacterial species upon exposure to envelope-targeting antimicrobial agents [1, 2], such as vancomycin , β-lactam antibiotics , chlorhexidine [5,6,7,8], chlorpromazine [6, 7], and polymyxin B [9, 10]. These reported ESR mechanisms are mainly through maintaining the compositions of cell envelope [2, 11], and activating the expression of RND efflux pumps [12,13,14].
Among these five widely conserved ESRs, Bae, Cpx, and Rcs use phosphorelay proteins to regulate the expression of target genes. Upon exposure to antimicrobial agents, both Bae and Cpx responses are commonly implicated in regulation of RND efflux pumps , while Rcs response is often associated with altering cell surface structure . Bae response constitutes a two-component system (TCS) sensor kinase BaeS and a response regulator BaeR . In E. coli, the phosphorylated BaeR is required for the direct regulation of RND efflux pump gene acrD, conferring bacterial resistance against SDS upon exposure to indole . In Erwinia amylovora, the phosphorylation and inducible expression of BaeR increase the expression level of MdtABC for bacterial resistance against tannin . Cpx response contains a TCS senor kinase CpxA and a response regulator CpxR together with a lipoprotein NlpE and a periplasmic repressor CpxP . In E. coli, CpxR mediates the indole regulation on RND efflux pump genes acrAB, tolC, acrD, and mdtA, which is required for bacterial resistance against protamine, SDS, and rhodamine 6G [17, 19]. In Vibrio cholerae, CpxR was proposed to directly control the expression of RND efflux pump genes vexAB and vexGH with putative CpxR binding site in their promoter regions in the presence of KCl . Rcs response is a complex regulatory system with phosphorelay proteins, in which RcsC and RcsD are senor kinases, and RcsB serves as a response regulator for modulating gene expression . Unlike the TCS response regulators in ESRs, RcsB can adopt both phosphorylated and unphosphorylated states in gene regulation. The unphosphorylated RcsB can interact with FixJ/NarL family transcriptional regulators, including BglJ, GadE, DctR, and MatA, to regulate bacterial physiology and metabolism, including bacterial motility and cell membrane compositions .
Zeamines and structurally related chemical compounds, i.e., fabclavines, are polyamine antimicrobial agents produced by Dickeya, Serratia, and Xenorhabdus strains [21,22,23,24,25]. In Dickeya and S. plymuthica, zeamines are synthesized by zms (zmn) cluster genes, in which zmsA (zmn10) is one of the key genes required for the biosynthesis of all zeamine molecules [22, 25]. Previous studies unveiled the potent inhibitory activities of zeamines against the growth of nematodes, fungi, oomycetes, and multidrug-resistant bacteria [21, 26, 27] and demonstrated that zeamines act by damaging the cell envelope of Gram-negative bacteria in a way similar to cationic antimicrobial peptide polymyxin B , implicating that zeamines may trigger ESRs in a way similar to polymyxin B [10, 29]. However, our recent study showed that responses to zeamines and polymyxin B in D. oryzae EC1, formerly known as D. zeae EC1 , are different . Exposure to a low level of zeamines (5 μg/ml) could significantly increase the expression level of efflux pump genes desAB, which encode a RND efflux pump DesABC, whereas polymyxin B was unable to trigger desAB expression even at a much higher concentration (100 μg/ml) . This suggests that D. oryzae EC1 may evolve a new ESR mechanism in response to zeamines.
In this study, we found that the dzrR gene, which is located next to the desAB genes in D. oryzae EC1, encodes a key regulator modulating bacterial resistance to zeamines. DzrR induced the transcriptional expression of desAB upon exposure to zeamines likely in a protein phosphorylation-independent manner. In addition, we showed that a range of envelope-targeting antimicrobial agents, including chlorhexidine and chlorpromazine, could also activate desAB expression through DzrR, and this activation was not dependent on the previously characterized canonical ESRs. Furthermore, we found that the DzrR-dependent ESR is widely conserved in Dickeya, Ralstonia, and Burkholderia species. Interestingly, we showed that in B. cenocepacia, although the genomic location of dzrR homolog is far from the desAB homologs, its protein product could still directly activate the efflux pump gene expression in the presence of chlorhexidine. Chlorhexidine is a widely used antimicrobial agent for skin disinfection before surgery and for sterilization of surgical instruments. Identification of this novel and widespread DzrR-mediated ESR mechanism would be of significant implications in our fight against the emergence of antimicrobial resistance.
Identification of DzrR associated with zeamine resistance in D. oryzae EC1
In D. oryzae EC1, expression of the efflux pump genes desAB was found induced by the self-produced antimicrobial agents zeamines, which suggests a regulatory mechanism for sensing and responding to these toxic agents. To identify this putative regulatory mechanism, a reporter strain ∆zmsK(pPdesAB-Gfp) was generated by introducing a previously constructed reporter plasmid pPdesAB-Gfp (pDesABgfp) , in which the gfp was placed under the control of the desAB promoter, into the ∆zmsK mutant producing only zeamine II  and with a significantly higher expression level of desAB than the wild-type strain EC1 (Additional file 1: Fig. S1). The reporter ∆zmsK(pPdesAB-Gfp) was selected as the parental strain for random transposon mutagenesis with mariner-based transposon carried by pBT20 . The relative fluorescence of transposon mutants cultured in the LS5 medium, which was optimized for zeamine production , was determined to screen for the genes affecting the expression level of desAB. After screening about 9400 mutants, a transposon insertion mutant Dz974, which produced about 16-fold less amount of relative fluorescence than its parental strain ∆zmsK(pPdesAB-Gfp), was obtained (Fig. 1A). Unlike the negative control strain ∆zmsA(pPdesAB-Gfp), which did not produce zeamines , the relative fluorescence of Dz974 did not increase after exogenous supplement of zeamines (20 μg/ml) (Fig. 1A), suggesting that bacterial response to zeamines was abolished in Dz974. To identify the mutated gene in Dz974, the flanking regions of the transposon insertion were determined by FPNI-PCR . The results showed that Dz974 contains a transposon insertion at the 125th base pair of a 738-bp coding sequence of a gene (NCBI accession no. W909_RS06570) next to desAB (Fig. 1B). This gene encodes a putative TCS response regulator with REC and Trans_reg_C domains (Fig. 1C) and was herewith designated as dzrR (Dickeya zeamine resistance regulator).
To validate the role of DzrR in regulation of the desAB expression and zeamine resistance in D. oryzae EC1, we generated an in-frame deletion mutant ∆dzrR and a complementation strain ∆dzrR(pBB-dzrR). Reverse transcription-quantitative PCR (RT-qPCR) analysis showed that deletion of dzrR caused significantly decreased transcript level of desB when ∆dzrR was cultured in LS5 medium (fold change > 2, P < 0.05; Fig. 1C) and in trans expression of dzrR in ∆dzrR could restore desB expression (Fig. 1D). In addition, we found that DzrR was not involved in regulation of zeamine production (Additional file 1: Fig. S2A) and expression of dzrR could not be stimulated by zeamines (Additional file 1: Fig. S2B and S2C). To elucidate whether DzrR contributes to zeamine resistance in D. oryzae EC1, we constructed dzrR deletion mutant using the wild-type EC1 as a parent strain. In addition, to avoid potential interference of the self-produced zeamines in minimal inhibitory concentration (MIC) assay, a dzrR deletion mutant was also generated at the background of the zeamine-minus mutant ∆zmsA. MICs of zeamines for the parent strains, i.e., ∆zmsA and the wild-type strain EC1, dzrR mutants ∆zmsA∆dzrR and ∆dzrR, and the complementation strains ∆zmsA∆dzrR(pBB-dzrR) and ∆dzrR(pBB-dzrR) were determined by broth microdilution method in 96-well plates. The results indicated that similar to desAB , null mutation of dzrR compromised bacterial resistance against zeamines. Deletion of dzrR resulted in about 2–fourfold decrease in the MICs of zeamines compared to the parental strains, which was restored by in trans expression of dzrR (Table 1). These findings suggest that DzrR plays a key role in regulation of the D. oryzae resistance against zeamines by activating the transcriptional expression of the RND efflux pump genes desAB upon exposure to zeamines.
DzrR specifically binds to the promoter region of desAB
To elucidate how DzrR could modulate the expression of desAB, the potential interaction between DzrR and the established 222-bp promoter region of desAB (PdesAB)  was investigated by electrophoretic mobility shift assay (EMSA). The results showed that a final concentration of 0.5 μM or 1 μM of tag-free DzrR could cause mobility shift of biotin-labeled PdesAB (Bound probe, Fig. 2A), suggesting a direct interaction between DzrR and PdesAB. The specific binding of DzrR to PdesAB was validated as addition of excessive unlabeled PdesAB could inhibit the DNA mobility shift, and the same amount of DzrR could not cause mobility shift of the biotin-labeled control DNA fragment provided by the EMSA kit (Fig. 2A). Together with the regulatory role of DzrR on desAB expression in the presence of zeamines (Fig. 1A and D), the above findings demonstrated that DzrR controls the transcriptional expression of desAB likely via directly binding to PdesAB.
To identify the DzrR binding site in PdesAB, DNase I footprinting assay was performed on PdesAB labeled with FAM (5′) and HEX (3′). The results unveiled that DzrR might specifically bind to a 25-bp region (5′-CGGTTCTCCATCATATCTCCATATT-3′) in PdesAB (Fig. 2B and C). For validation, a 37-bp DNA fragment PdesAB(37), containing the above identified putative DzrR binding site in the middle, was generated and labeled with biotin by annealing the primer pairs Biotin-PdesAB(37)-F/PdesAB(37)-R together (nucleotide acid sequences refer to Additional file 2: Table S2) for EMSA analysis. The results showed that addition of DzrR could cause DNA band mobility shift of Biotin-PdesAB(37) (Bound probe, Fig. 2D), validating the DzrR binding site identified in DNase I footprinting assay (Fig. 2B–C).
Regulatory capacity of DzrR is likely independent on protein phosphorylation
DzrR is a proposed TCS response regulator with REC and Trans_reg_C domains (Fig. 1C). Bioinformatics analysis showed that its best-characterized homolog is AdeR in Acinetobacter baumannii (NCBI accession no. WP_000459547.1), which has a relatively high level of amino acid sequence identity (52%) and similarity (68%) compared to DzrR, and adopts a phosphorylated manner to regulate the expression of the genes encoding a RND efflux pump AdeABC [34, 35]. Sequence alignment performed among the REC domains of DzrR, AdeR, and three well-studied canonical TCS response regulators in E. coli, i.e., PhoB, OmpR, and CheY, illustrated the potential conserved five αβ folds and amino acid residues required for protein phosphorylation (E22, D23, and D66) and signal transduction (T93 and K115)  (Additional file 1: Fig. S3A). To test whether the DzrR function is dependent on phosphorylation, DzrR variants with E22, D23, and D66 being replaced by alanine respectively were generated by site-directed mutagenesis. MIC assay showed that in trans expression of the E22, D23, and D66 variants in ∆zmsA∆dzrR and ∆dzrR mutants respectively could restore zeamine resistance (Table 1). The results suggest that these conserved amino acid residues associated with protein phosphorylation were not responsible for the DzrR function in zeamine resistance. RT-qPCR assay further confirmed that substitution of D66, which is the key amino acid residue required for phosphoryl group acceptance , did not affect the DzrR-dependent expression of desB in the presence of zeamines (Additional file 1: Fig. S3B).
To further assess the potential involvement of phosphorylation in modulation of DzrR activity, we conducted the alanine scanning mutagenesis by substituting each of the 111 amino acid in the REC domain of DzrR (with the exception of 10 alanines, E22, D23, and D66). These variants as well as their wild-type dzrR were cloned in the vector pBBR1-MCS4 and introduced into the ∆zmsA∆dzrR and ∆dzrR mutants, respectively, and assayed for their ability to restore the mutant growth in the presence of lethal concentration of zeamines. The results showed that all the DzrR variants, including those of the conserved residues T93 and K115 involved in signal transduction  (Additional file 1: Fig. S3A), could rescue the mutant growth in the LB medium supplemented with zeamines at a final concentration of 900 μg/ml (Additional file 1: Fig. S4 and S5). These findings suggest that DzrR might adopt an unphosphorylated form to mediate zeamine response and resistance in D. oryzae EC1.
Zeamines induce desAB expression through DzrR but independent on canonical ESRs
A previous study indicated that zeamines target bacterial cell envelope , which together with our results suggest that DzrR might be a key regulator associated with ESR in D. oryzae. To test this possibility, the relative fluorescence of ∆zmsA(pPdesAB-Gfp) was determined upon exposure to a few known envelope-targeting antimicrobial agents and signaling compounds by Gfp transcriptional fusion assay following the established protocol . The results indicated that antimicrobial agents that target cell envelope, i.e., chlorhexidine, chlorpromazine, and xinjunan [5, 37], significantly induced desAB expression at a sub-inhibitory concentration whereas the signaling compounds, i.e., spermidine , putrescine , and indole [17, 40], did not have a comparable regulatory effect at the same concentration (Fig. 3A, structures of chemical compounds refer to Additional file 1: Fig. S6). In Pseudomonas aeruginosa, chlorhexidine stimulates the expression of RND efflux pump MexCD-OprJ through RpoE ESR . However, in D. oryzae, deletion of rpoE (NCBI accession no. W909_RS14550) did not affect the stimulated expression of desAB upon exposure to chlorhexidine or other three envelope-targeting antimicrobial agents, i.e., chlorpromazine, xinjunan, and zeamines (Fig. 3B). Similarly, inactivation of the other four canonical ESR regulatory genes, i.e., baeR (NCBI accession no. W909_RS13980), cpxR (NCBI accession no. W909_RS18830), rcsB (NCBI accession no. W909_RS05280), or pspF (NCBI accession no. W909_RS11125) did not compromise the stimulated expression of desAB in the presence of zeamines (Fig. 3B). These findings suggest DzrR represents a new ESR mechanism against zeamines in D. oryzae.
DzrR homologs found in Dickeya, Ralstonia, and Burkholderia species have comparable ESR activity
DesB encodes a RND efflux pump inner membrane protein, which is crucial for determining the substrate profile and specificity of the RND efflux pump DesABC . Our previous study indicated that homologs of desB are conserved in various bacterial species . It is interesting to explore whether DzrR is also widely conserved. We therefore conducted a BLASTp search using the amino acid sequence of DzrR as the bait. The results indicated that DzrR homologs sharing a high level of sequence identity (above 50%) and similarity (above 67%) were detected not only in Dickeya, but also in a range of bacterial species, including Ralstonia and Burkholderia (Additional file 2: Table S3). Sequence alignment of DzrR and its homologs from D. dadantii 3937 , R. solanacearum GMI1000 , B. glumae BGR1 , B. cepacia ATCC 25416 , and B. cenocepacia J2315 and H111 [7, 45,46,47] revealed that the REC and Trans_reg_C domains of DzrR homologs had a high level of sequence identity (above 72%) and similarity (above 85%) with their counterparts in DzrR, respectively (Fig. 4A and C). The high level of similarity suggests that DzrR homologs found in Dickeya, Ralstonia, and Burkholderia species may have a similar ESR activity as DzrR. To verify this possibility, zeamine resistance assay was performed on the ∆zmsA∆dzrR and ∆dzrR mutants expressing the dzrR homologs from D. dadantii 3937 (dzrR3937), R. solanacearum GMI1000 (dzrRGMI1000), and B. cepacia ATCC 25416 (dzrR25416), respectively. The results showed that heterologous expression of the dzrR homologs could fully restore the zeamine resistance (Table 1).
For further validation, we conducted RT-qPCR analysis, DNase I footprinting assay, and EMSA. The RT-qPCR results showed that heterologous expression of the dzrR homologs resulted in a more than twofold of increase in transcript level of desB compared to the vector control (Fig. 4D). Furthermore, DNase I footprinting assay and EMSA confirmed that the DzrR homolog found in B. cepacia ATCC 25416, i.e., DzrR25416, could specifically bind to the same region in PdesAB as DzrR did (Figs. 2B, 4B, and E). These findings suggest that DzrR homologs found in Dickeya, Ralstonia, and Burkholderia species have a similar activity in modulating desAB expression against zeamines.
DzrR-regulating expression of the desABC homologs is conserved in B. cenocepacia
Considering the wide distribution of dzrR and desB homologs in Gram-negative bacteria, it is intriguing whether they are genetically located adjacent to each other as dzrR and desB in D. oryzae EC1. Genome sequence analysis revealed that desB and dzrR homologs are located next to each other in Dickeya and Ralstonia strains (Fig. 5A; Additional file 2: Table S4) but varied in genetic arrangement in Burkholderia species, showing three types of genetic arrangement: (I) dzrR and desB homologs are located adjacent to each other but transcribed divergently (B. glumae, B. stagnalis, and B. ubonensis); (II) dzrR and desB homologs are neighboring with the same transcriptional orientation (B. pseudomultivorans and B. multivoran); (III) dzrR and desB homologs are distantly located in the same chromosome (B. pyrrocinia, B. cenocepacia, and B. seminalis) or different chromosomes (B. stabilis, B. cepacia, B. metallica, B. lata, and B. contaminans) (Additional file 2: Table S4). In addition, the genetic organization of the desABC homologs in Dickeya, Ralstonia, and Burkholderia strains are different. Unlike Dickeya strains, in which desC encoding the outer membrane protein of the DesABC efflux pump is not at the same location as desAB in the bacterial genome, the desAB homologs in Ralstonia and Burkholderia strains are located together with desC homologs as a single operon (Fig. 5A; Additional file 2: Table S4). Interestingly, although the genetic organization and arrangement of dzrR and desABC homologs are varied in Dickeya, Ralstonia, and Burkholderia strains, a conserved box (5′-N5-CTCCATC-N2-A-N-CTCCAT-N2-T-3′), which harbors two direct 5′-CTCCAT-3′ sequence repeats, was found in the promoter region of all the desAB homologs in Dickeya, Ralstonia, and Burkholderia strains (Fig. 5B). These findings suggest that in Burkholderia strains, the DzrR homologs may directly regulate the expression of distantly located desAB homologs. To validate this hypothesis, EMSA and DNase I footprinting assay were performed between DzrR25416 and the promoter fragment of desAB homologs found in B. cepacia ATCC 25,416 (PdesB25416). The results showed that DzrR25416 directly interacted with the proposed dzrR box in PdesB25416 (Fig. 5C and D). In B. cenocepacia J2315, the desAB homologs, i.e., BCAM0927 and BCAM0926, which encode the RND-8 efflux pump , share about 77% and 86% similarity with their counterparts in D. oryzae EC1, respectively (Fig. 5A). Previous studies demonstrated that their expression is stimulated by exposure to chlorhexidine and chlorpromazine [6, 7]. However, the direct regulator of the RND-8 efflux pump has not yet been characterized [6, 7]. Given the presence of the conserved dzrR box in the promoter of the RND-8 efflux operon in B. cenocepacia strains (Fig. 5B), and the fact that chlorhexidine and chlorpromazine also induced the expression of desAB in D. oryzae EC1 (Fig. 3A), we speculated that the DzrR homolog, although its coding gene is located at a distant location, might play a role in modulation of the RND-8 efflux operon expression upon exposure to chlorhexidine and chlorpromazine. The potential regulatory linkage between DzrR homolog and RND-8 efflux operon was thus investigated in a well-studied strain B. cenocepacia H111 . Expression of RND-8H111 efflux operon in the wild-type strain H111 and the dzrRH111 mutant, i.e., ∆dzrRH111, were compared by determining the relative fluorescence of gfp reporter strains H111(pPRND-8-Gfp) and ∆dzrRH111(pPRND-8-Gfp) upon exposure to chlorhexidine and chlorpromazine, respectively. The results showed that inactivation of dzrRH111 basically abolished the chlorhexidine or chlorpromazine stimulated expression of RND-8H111 efflux operon (Fig. 5E). In addition, null mutation of dzrRH111 increased the chlorhexidine susceptibility in B. cenocepacia H111, which was fully restored by in trans expression of the dzrRH111 in the mutant (Fig. 5F). These findings thus demonstrated unequivocally that DzrRH111 modulates chlorhexidine resistance through regulating the expression of RND-8H111 efflux operon in B. cenocepacia H111.
ESRs are crucial to maintain bacterial envelope homeostasis upon exposure to envelope-targeting antimicrobial agents and other environmental stresses . In this study, we documented a new ESR mediated by DzrR, which could respond to a range of structurally divergent envelope-targeting antimicrobial agents and directly induced the transcriptional expression of the efflux pump genes desAB in D. oryzae EC1. We previously reported that D. oryzae EC1 produces at least two potent antibiotics and phytotoxins, i.e., zeamine and zeamine II [21, 22]. The subsequent study unveiled that a zeamine-inducible RND efflux pump designated as DesABC plays a key role in protection of D. oryzae EC1 against zeamines . Identification of DzrR thus depicts a molecular mechanism with which the pathogen modulates the expression of DesABC efflux pump for self-protection and hence enables the pathogen to produce high level of zeamines against its host plants and microbial competitors. DzrR is a response regulator containing a typical REC-Trans_reg_C domain structure (Fig. 1C). Significantly, dzrR and desABC are widely conserved in Dickeya, Ralstonia, and Burkholderia strains, underlining the biological importance and ecological significance of this newly identified ESR in protection of bacterial cells against various toxic chemicals that can damage bacterial cell envelope.
Phosphorelay systems are critical for bacterial response and resistance to envelope-targeting antimicrobial agents and other stress environmental cues. Among the five well characterized ESRs in E. coli, three are ESRs with phosphorelay proteins, including Bae, Cpx, and Rcs . The Bae and Cpx ESRs commonly rely on the stimulated expression and sensor-dependent phosphorylation of response regulators to activate resistance mechanisms against envelope-targeting antimicrobial agents [13, 17], whereas in the Rcs ESR, the response regulator RcsB could regulate different sets of the target genes with phosphorylated or unphosphorylated form . Compared to the TCS response regulators in Bae and Cpx ESRs, DzrR seem to evolve a different mechanism of regulation. Our previous study showed that a TCS sensor kinase gene located adjacent to dzrR, i.e., S1 (NCBI accession no. W909_RS06575) (Fig. 1B), was not involved in regulation of zeamine resistance . Unlike CpxR , BaeR [13, 17], and AdeR [34, 35], our results indicated that DzrR activated the zeamine resistance mechanism by regulating the expression of desAB likely in a protein phosphorylation-independent manner. Mutations on the proposed key amino acids required for protein phosphorylation of DzrR did not alter zeamine resistance of D. oryzae (Table 1) or DzrR regulation on desB expression (Additional file 1: Fig. S3). And alanine scanning mutagenesis of the REC domain of DzrR did not identify a single key amino acid residue critical to the function of DzrR (Additional file 1: Fig. S4 and S5). In trans expression of the Trans_reg_C domain of DzrR in ∆zmsA∆dzrR and ∆dzrR mutants could not restore zeamine resistance (Table 1). Alternatively, DzrR is orphaned, or may function as a transcriptional regulator in a way similar to the unphosphorylated RcsB, regulating the target gene expression by cooperating with other family proteins other than TCS proteins. On the other hand, unlike BaeR and CpxR whose expression could be stimulated by envelope-targeting antimicrobial agents , we found that dzrR was constitutively expressed along with bacterial growth disregarding the presence or absence of zeamines (Additional file 1: Fig. S2), and DzrR is required for the zeamine-induced expression of desAB (Figs. 1A and 3B). Further study is surely needed to elucidate how DzrR could modulate the expression of these ESR genes.
Interestingly, although DzrR homologs have been found in a range of bacterial pathogens (Additional file 2: Table S3), their biological functions and mode of actions are hardly characterized. To our knowledge, AdeR is the only characterized response regulator among the DzrR homologs (Additional file 2: Table S3), which shares about 52% identity and 68% similarity with DzrR at amino acid level. AdeR and its cognate sensor kinase AdeS constitute a functional TCS, which confers antibiotic resistance by regulating expression of the RND efflux pump genes adeABC [34, 35]. However, DzrR and AdeR differ in at least two aspects. Firstly, the functionality of AdeR depends on phosphorylation , whereas the DzrR activity does not seem to rely on phosphorylation (Additional file 1: Fig. S3, S4, and S5; Table 1). Secondly, the binding site of AdeR contains a 10-bp direct repeat DNA sequence (5′-AAGTGTGGAGNAAGTGTGGAG-3′) , which is not found in the target promoter of DzrR. Similarly, the AdeR binding site is also not present in the promoter region of desAB in D. oryzae EC1. Instead, our data demonstrated that DzrR recognizes the promoter sequence with two direct 5′-CTCCAT-3′ repeats (Fig. 5B). Given that the DzrR25416 from B. cepacia ATCC 25416, which shares about 70% identity and 80% similarity with DzrR at amino acid level (Fig. 5A), could also cause mobility shift of its target PdesB25416 (Fig. 5C), our phylogenic analysis in DzrR homologs supports the notion that the DzrR homologs could be classed into at least two groups according to their DNA binding sites (Additional file 1: Fig. S7) [48, 49]. In addition, we noticed that overexpression of dzrR and its homologs in dzrR mutant totally restores the zeamine resistance (Table 1), but only partly rescue desB expression (Fig. 4D), suggesting that DzrR could activate other genes involved in zeamine resistance than DesABC efflux pump encoding genes, which requires a further elucidation.
DzrR-dependent expression of desAB in D. oryzae EC1 was not only induced by zeamines but also by other structurally divergent envelope-targeting antimicrobial agents, i.e., chlorhexidine, chlorpromazine, and xinjunan (Fig. 3A, Additional file 1: Fig. S6). In particular, xinjunan, which is structurally more similar to zeamines than other two (Additional file 1: Fig. S6), could activate higher level of desAB expression than zeamines did (Fig. 3B). However, the polyamine signaling compounds, i.e., spermidine and putrescine, which are structurally similar to zeamines than chlorhexidine and chlorpromazine, did not activate desAB expression at the same concentration (Fig. 3A, Additional file 1: Fig. S6). The fact that all the tested membrane-targeting antibiotics used in this study, including zeamines, chlorhexidine, chlorpromazine, and xinjunan, could induce desAB expression in a DzrR-dependent manner (Fig. 3B) indicates that DzrR is a key component of ESR, which typically responds to and induces resistance mechanisms against various envelope-targeting toxic agents. On the other hand, the data from this study also imply that different bacteria may respond to the same envelope-targeting antimicrobial agent with different ESR mechanisms. In P. aeruginosa, the stimulated expression of MexCD-OprM in the presence of chlorhexidine and chlorpromazine is RpoE-dependent , while in Salmonella enterica serovar enteritidis ATCC 13,076, it is Cpx ESR that responds to chlorhexidine and triggers the expression of a range of membrane-associated proteins . In view of these findings, we tested whether RpoE, Cpx, and other three well established ESRs were associated with the zeamine-induced desAB expression. However, the results precluded the involvement of these canonical ESRs in response to zeamines in D. oryzae (Fig. 3B). In addition, we found that the DzrR homologs in Burkholderia are functionally conserved with DzrR in D. oryzae as they could rescue zeamine resistance of dzrR mutants (Table 1), and could also regulate the expression of RND-8 efflux operon upon exposure to chlorhexidine (Fig. 5E). Apparently, much remains to be done to decipher the molecular mechanisms that bridge sensing and responding to various envelope stress conditions.
In this study, we found three types of genetic arrangement of dzrR and desABC homologs in Burkholderia (Additional file 2: Table S4). Chromosomal rearrangements including inversion and translocation often happen during evolution of Burkholderia . The result obtained from the phylogenomic study suggested that variation in genetic arrangement of dzrR and desABC homologs in different Burkholderia species was established during Burkholderia speciation (Additional file 1: Fig. S8) [51,52,53,54,55]. In type I arrangement, dzrR and desAB homologs are located adjacent to each other with different transcriptional orientation as the case of dzrR and desAB in D. oryzae EC1 (Fig. 5A), which may represent the ancestor form (Additional file 1: Fig. S8). The dzrR homologs may experience inversion and translocation during Burkholderia evolution, resulting in the establishment of type II and type III genetic arrangements (Additional file 1: Fig. S8). Although desC homologs in the desABC operons of Ralstonia and Burkholderia species show a relatively low level of amino acid similarity (Fig. 5A), we proposed that the desABC operons with different genetic organization and arrangement in Dickeya, Ralstonia, and Burkholderia species may share a more recent ancestor. This is evident as a high level of sequence similarity is detected in the 5′-noncoding region of desAB in Dickeya, Ralstonia, and Burkholderia species (Additional file 1: Fig. S9), especially the dzrR box (Fig. 5B) and the fragments I–III of an 84 (82)-bp noncoding region (Additional file 1: Fig. S9). The variation in genetic organization of desABC operons in Dickeya, Ralstonia, and Burkholderia may be established during the evolutionary divergence of Pectobacteriaceae. Similar to the well-known Gammaproteobacterium E. coli but dissimilar to the Betaproteobacteria Burkholderia and Ralstonia, the RND efflux pumps in Dickeya seem to share with only one outer membrane protein encoded by desC .
In summary, this study depicts a new ESR mechanism mediated by DzrR, which can respond to zeamines and other envelope-targeting antimicrobial agents. DzrR plays a key role in positive regulation of the RND efflux pump DesABC, which confers resistance to zeamines. Interestingly, DzrR-DesABC system appears to be widely conserved in the Gram-negative microorganisms including agricultural and medical important bacterial pathogens, but so far, the biological functions and signaling mechanisms are hardly investigated in other microorganisms. The findings from this study not only present a new mechanism with which D. oryzae could protect itself against zeamines, but also provide useful clues for elucidating its role in other bacterial pathogens and a potential target for tackling emergence of antimicrobial resistance.
Bacterial strains, plasmids, primers, and growth conditions
Bacterial strains, plasmids, and primers used in this study are listed in Additional file 2: Table S1 and S2 [31, 32, 46, 47, 56], respectively. D. oryzae strains were routinely grown at 28 °C in Luria–Bertani (LB) medium [per liter contains tryptone 10 g, yeast extract 5 g, NaCl 10 g, pH 7.0], minimal medium (MM) [per liter contains 10.5 g K2HPO4, 4.5 g KH2PO4, 2.0 g (NH4)2SO4, 2.0 g mannitol, 2.0 g glycerol, 0.2 g MgSO4·7H2O, 0.01 g CaCl2, 0.005 g FeSO4·7H2O, and 0.002 g MnCl2·4H2O, pH 7.0], or LS5 medium [per liter contains 5.25 g K2HPO4, 2.25 g KH2PO4, 10.0 g sucrose, 3.6 g NH4NO3, 1.0 g KCl, and 0.25 g MgSO4·7H2O, pH 7.0] . E. coli and B. cenocepacia derivatives were routinely grown at 37 °C in LB medium. Antibiotics were added at following concentrations when necessary: gentamycin, 50 μg/ml; streptomycin, 50 μg/ml; ampicillin, 100 μg/ml; kanamycin, 100 μg/ml; polymyxin B, 50 μg/ml.
Construction of in-frame deletion, complementation, and heterologous expression strains
Construction of in-frame deletion, complementation, and heterologous expression strains of D. oryzae were following the methods described previously . Briefly, for construction of in-frame deletion mutants of D. oryzae EC1, the downstream and upstream regions of the target genes were cloned in the suicide vector pKNG101. The resultant constructs were transformed into the wild-type strain EC1 or a zmsA in-frame deletion mutant respectively by triparental mating. The transformants grown on MM agar plates supplemented with 5% (wt/vol) sucrose were confirmed by PCR and DNA sequencing to identify the desired deletion mutants. For complementation and heterologous expression, the open reading frames (ORFs) of dzrR and its homologs were cloned in pBBR1-MCS4 respectively for constructing expression constructs, which were introduced into dzrR mutants by triparental mating. The transformants were screened on MM agar plates containing ampicillin and confirmed by PCR. In-frame deletion of dzrRH111 in B. cenocepacia H111 was performed following the previously described method . Briefly, the downstream and upstream regions of dzrRH111 were cloned in the suicide vector pK18 for constructing the in-frame deletion construct pK18-dzrRH111, which was transformed into B. cenocepacia H111 by triparental mating. The dzrRH111 mutants were screened on LB agar plates (without NaCl) containing 10% (wt/vol) sucrose, and confirmed by PCR and DNA sequencing. The complementation strain ∆dzrRH111(pBB(K)-dzrRH111) was constructed by transformed the pBB(K)-dzrRH111 plasmid that express dzrRH111 in trans into the dzrRH111 mutant.
Transposon mutagenesis analysis
To identify the potential regulatory genes of desAB expression, the reporter strain ∆zmsK(pPdesAB-Gfp) was randomly mutated with mariner-based transposon carried by pBT20  through biparental mating on YEB agar plates [per liter contains10 g tryptone, 5 g yeast extract, 10 g KCl, 10 g sucrose, 0.5 g MgSO4·7H2O, and 18 g agar, pH 7.0]. The mutant colonies selected on the MM agar medium supplemented with antibiotics were inoculated and cultured in 96-well plates with LS5 medium plus kanamycin at 28 °C for 48 h. The cell culture dilutions were transferred into 96-well black clear-bottom plates for measurement of optical density at 600 nm (OD600) and fluorescence density (excitation at 485 nm and emission at 535 nm)  using BioTek SYNERGY H1 microplate reader (Vermont, USA). The fluorescence of each sample was calculated by dividing the fluorescence density reading with the optical density reading. The relative fluorescence of each mutant was expressed as the fluorescence of each mutant normalized to the fluorescence of ∆zmsK(pPdesAB-Gfp). The mutants showing over twofold changes in relative fluorescence with the parental strain ∆zmsK(pPdesAB-Gfp) were selected as candidates for further analysis. To localize the transposon insertion sites within the mutants, the flanking regions of the transposon insertion sites were analyzed by the FPNI-PCR method  using Green Taq Mix (Vazyme, Nanjing, China). The first step of FPNI-PCR was conducted using the fresh bacterial cultures as the templates with the following conditions: 1 cycle at 95 °C for 90 s, 2 cycles at 94 °C for 10 s, 62 °C for 30 s, and 72 °C for 2 min, 1 cycle at 94 °C for 10 s, 1 cycle at 25 °C for 2 min, 1 cycle at 72 °C for 2 min, 2 cycles at 94 °C for 10 s, 62 °C for 30 s, and 72 °C for 2 min, 1 cycle at 94 °C for 10 s, 1 cycle at 44 °C for 30 s, followed by 1 cycle at 72 °C for 7 min. The second step of FPNI-PCR was conducted using an aliquot of 1 μl of each PCR product from the first step of FPNI-PCR as the temple with the following conditions: 1 cycle at 95 °C for 90 s, 30 cycles at 94 °C for 10 s, 62 °C for 30 s, and 72 °C for 2 min, followed by 1 cycle at 72 °C for 5 min. The third step of FPNI-PCR was conducted using an aliquot of 1 μl of each PCR product with tenfold dilution from the second step of FPNI-PCR as the temple with the following conditions: 1 cycle at 95 °C for 90 s, 30 cycles at 94 °C for 10 s, 62 °C for 30 s, and 72 °C for 2 min, followed by 1 cycle at 72 °C for 5 min.
Gfp transcriptional fusion assay
The reporter plasmids PdzrR-Gfp and pPRND-8-Gfp were constructed as pPdesAB-Gfp  by amplification of the promoter regions of dzrR and RND-8H111 efflux operon using primer pairs pPdzrR-Gfp-F/pPdzrR-Gfp-R and pPRND-8-Gfp-F/pPRND-8-Gfp-R and fusing them with the coding region of Gfp in the plasmid pPROBE-NT, respectively. The fluorescence of bacterial strains containing the gfp reporter constructs were determined by measuring the average fluorescence of 50,000 cells using a CytoFLEX flow cytometer (Beckman Coulter, Brea, CA, USA) . The relative fluorescence was expressed as the fluorescence of bacterial cells containing a relevant reporter construct normalized to the fluorescence of the corresponding control sample.
Reverse transcription-quantitative PCR (RT-qPCR) assay
RT-qPCR assay was performed as previously described . Briefly, bacterial strains were cultured in LS5 medium and harvested at an OD600 about 1.5. RNA of bacterial strains was extracted from three milliliters of bacterial cultures using the RiboPure RNA purification kit (Thermo Fisher Scientific, MA, USA). RNA purity and integrity were analyzed by gel electrophoresis, and by determination of the ratios of A260/A280 and A260/A230 using a NanoDrop 2000c (Thermo Fisher Scientific, MA, USA). For RT-qPCR analysis, cDNA was synthesized from an aliquot of 100 ng of RNA sample using a FastKing RT kit (with gDNase) (Tiangen Biotech, Beijing, China). RT-qPCR analysis was performed on a Quantstudio 6 Flex system using PowerUp SYBR green master mix (Thermo Fisher Scientific, MA, USA) with the following conditions: 1 cycle at 50 °C for 2 min and 95 °C for 2 min, followed by 40 cycles at 95 °C for 15 s, 57 °C for 15 s, and 72 °C for 30 s. Specific primer pairs 16S-F/16S-R and desB-F/desB-R targeting 16S rRNA gene and desB respectively were used for RT-qPCR. The 16S rRNA gene was used as the endogenous reference. The fold change of desB expression was calculated through 2−∆∆CT method .
MIC (minimal inhibitory concentration) assay
The minimal inhibitory concentration assay of zeamines was performed as previously described  following the recommendations from the Clinical and Laboratory Standards Institute. In brief, fresh bacterial cultures at an OD600 about 0.5 were added to each well of 96-well plates (1%, vol/vol) with 100 μl LB medium containing twofold dilutions of zeamines to reach 2.0 × 105 CFU/ml. The 96-well plates were incubated at 28 °C for 18 h, and MIC of zeamines for bacterial strains was defined as the lowest antibiotic concentration with no visible cell growth.
Envelope-targeting antimicrobial agents and signaling compounds
Zeamines were purified using the absorbent resin XAD7 (Sigma-Aldrich, China) and confirmed by liquid chromatography-mass spectrometry (LC–MS) following the method described previously . Briefly, the supernatants of the wild-type strain EC1 cultured in LS5 medium were passed through a column with absorbent resin XAD7 (Sigma-Aldrich, China). The column was then washed with double-distilled H2O and methanol, and crude zeamine antibiotics were eluted from the absorbent resin by acetone. Confirmation of zeamines was performed by liquid chromatography-mass spectrometry (LC–MS) using an Agilent 1260 infinity system equipped with a Phenomenex Luna column (C18, 250 by 4.6 mm, 5 μm) coupled with a Bruker maxis Q-TOF mass spectrometer to identify three main zeamines, i.e., zeamine, zeamine I, and zeamine II. A gradient of 5% to 95% (CH3CN supplemented with 1% formic acid in H2O) at a flow rate of 1 ml/min was conducted in 20 min to elute and separate zeamines. The mass spectrometer was operated in positive mode for detection of three main zeamines using the following parameters: scan range from 100 to 2,000 m/z, ESI source type, end plate offset at − 500 V, capillary at 4500 V, nebulizer gas (N2) at 0.8 bar, dry gas at 5.0 L/min, dry temperature at 180 °C, and the collision cell RF at 800.0 Vpp. Chlorhexidine (Macklin, Shanghai), chlorpromazine hydrochloride (Macklin, Shanghai), xinjunan (ALTA, China), spermidine trihydrochloride (Sigma-Aldrich, China), and putrescine dihydrochloride (Sigma-Aldrich, China) used in this study were obtained commercially as indicated. The structures of chemical molecules are shown in Additional file 1: Fig. S6.
Purification of DzrR and DzrR25416 was conducted following the established method  with minor modifications. Briefly, the coding sequences of dzrR and dzrR25416 were amplified using the primer pairs GST-DzrR-F/GST-DzrR-R and GST-DzrR25416-F/GST-DzrR25416-R, respectively, and introduced into the pGEX-6p-1 plasmid. The resultant constructs were transformed into E. coli BL21 for protein expression under the treatment of isopropyl-β-D-thiogalactopyranoside IPTG (0.1 mM) at 16 °C for overnight. The bacterial cells were collected and disrupted by French ® Pressure Cell (Homogenising Systems Ltd, UK). The cell-free supernatants collected by centrifugation and filter-sterilization were loaded on the column with ProteinIso ® GST Resin (TransGen Biotech, China). GST-DzrR and GST-DzrR25416 were eluted with the buffer containing 5 mM glutathione reduced. GST-tag cleavage was performed using the PreScission Protease to obtain tag-free DzrR and DzrR25416. Protein purity of DzrR and DzrR25416 was analyzed by SDS-PAGE gel (Additional file 1: Fig. S10C and S10F).
Electrophoretic mobility shift assay (EMSA)
To generate biotin-labeled DNA probes for EMSA, the primer Biotin-PdesAB(37)-F with biotin at its 5′-end was synthesized by Invitrogen (Guangzhou, China), and the biotin-labeled PdesAB(37) was generated by annealing the biotin labeled Biotin-PdesAB(37)-F with PdesAB(37)-R. DNA fragments of PdesAB and PdesB25416 were amplified by PCR with primer pairs Probe-PdesAB-F/Probe-PdesAB-R and Probe-PdesB25416-F/Probe-PdesB25416-R, respectively, and labeled with biotin following the protocol of Biotin 3′ End DNA Labeling Kit (Thermo Fisher Scientific, MA, USA). EMSA assay was performed on tag-free DzrR or DzrR25416 using LightShift ® Chemiluminescent EMSA Kit (Thermo Fisher Scientific, MA, USA) following the method described previously . Briefly, an aliquot of 20 μl binding reaction solution containing tag-free DzrR or DzrR25416, and the DNA probes at a final concentration of 20 fmol were incubated at 28 °C for 20 min. After incubation, the binding reaction mixtures were loaded and electrophoresed on 6% native polyacrylamide gel. DNA probes were transferred to a nylon membrane and detected by a chemiluminescence system (Tanon, Shanghai, China). A DNA probe complexed with DzrR or DzrR25416 represents a shit in migration compared to the free probe. Full images of EMSA results were presented in Additional file 1: Fig. S10A, S10B, S10D, and S10E.
DNase I footprinting assay
DNase I footprinting assay was performed following the method described previously with minor modifications . Briefly, the promoter regions of desAB in D. oryzae EC1 and B. cenocepacia H111 were amplified using the primer pairs Probe-PdesAB-F/Probe-PdesAB-R and Probe-PdesB25416-F/Probe-PdesB25416-R modified with FAM (Probe-PdesAB-F and Probe-PdesB25416-F) and HEX (Probe-PdesAB-R and Probe-PdesB25416-R) at their 5′-end to generate DNA probes. In the binding process, about 400 ng purified DNA probes were incubated with 1 μM tag-free DzrR or DzrR25416 at 25 °C for 30 min in a 40 μl binding reaction solution as described in EMSA assay. After the incubation, an aliquot of 10 μl solution with 0.03 units of DNase I (Promega, USA) was added. The samples were incubated for 1 min, and the reaction was stopped by adding the DNase I stop solution supplied by the RQ1 RNase-Free DNase kit (Promega, USA). Total DNA was purified by phenol/chloroform extraction and dissolved in 10 μl ddH2O after ethanol precipitation. DNA samples were run on 3730XL system and the data were analyzed by Peak Scanner 2 (Applied Biosystems, USA).
Phylogenic analysis of DzrR homologs
A total of 36 amino acid sequences of DzrR homologs obtained from NCBI (Additional file 2: Table S3) were aligned with DzrR by ClustalW and analyzed in MEGA6  by using the Maximum Likelihood method based on the best-fit model (“LG + F”)  with 1000 bootstrap support.
Whole-genome phylogenic analysis of Burkholderia strains
A total of 14 whole genomic sequences of Burkholderia strains were downloaded from NCBI with the accessions described in Additional file 2: Table S5. The single-copy core gene analysis of all genomic sequences was performed through GET_HOMOLOGUES software . The amino acid sequences of core genes were aligned by MAFF v7.490 , and multiple-sequence alignment (MSA) was filtered for columns with high proportions of missing data by trimAl v1.4 . The filtered sequences were concatenated to construct the Maximum Likelihood phylogenetic tree by IQ-TREE v1.6 . The phylogenic tree was adjusted and viewed in iTOL v6.4 .
All experiments were individually performed for at least twice, each with three replicates. Differences of the relative fluorescence between bacterial strains were evaluated using a two-tailed unpaired Student’s t test performed with the GraphPad Prism 7.0 software (GraphPad, La Jolla, CA). A P value of less than 0.05 was considered as significant. As the data of fold change of desB expression do not satisfy Student’s t test assumptions, a permutation test was performed with the R software (ver. 4.2.2) as recommended . A P value of less than 0.05 and the fold change of desB expression above twofold were considered as significant.
Availability of data and materials
The genomic sequence of D. oryzae EC1 in NCBI is accessible with no. NZ_CP006929.1 .
Dickeya zeamine resistance regulator
Electrophoretic mobility shift assay
Envelope stress response
Minimal inhibitory concentration
Reverse transcription-quantitative PCR
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We thank Prof. Deng Yinyue and Dr. Song Shihao for their kindly help in providing the B. cenocepacia H111 strain and pK18 plasmid.
This study is supported by the National Natural Science Foundation of China (32000085, U22A20480, 32072450, and 31972230), KeyRealm R&D Program of Guangdong Province (2018B020205003, 2020B020209001), Guangdong Basic and Applied Basic Research Foundation (2020A1515110465, 2020A1515011534), Guangzhou Science and Technology Plan (201804020066, 202102080488), Guangdong Forestry Science and Technology Innovation Project (2020KJCX009), and Natural Science Foundation of Anhui Province (2108085QC118).
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Fig. S1. Determination of the expression of desAB in the wild-type strain EC1, ∆zmsA, and ∆zmsK. Fig. S2. DzrR was not involved in zeamine production and could not be stimulated by zeamines. Fig. S3. Regulatory role of DzrR is likely independent on protein phosphorylation. Fig. S4. Alanine scanning mutagenesis analysis for detection of the key amino acid residues required for the DzrR function at the background of ∆zmsA∆dzrR. Fig. S5. Alanine scanning mutagenesis analysis for detection of the key amino acid residues required for the DzrR function at the background of ∆dzrR. Fig. S6. Structures of the chemical compounds used in this study to test the signaling activity of DzrR in D. oryzae. Fig. S7. Phylogenic relationship of DzrR and its homologs. Fig. S8. Genetic arrangement of dzrR homologs and desABC operons in Burkholderia strains. Fig. S9. Characteristics of the 5′-noncoding regions of desAB in Dickeya, Ralstonia, and Burkholderia strains. Fig. S10. Full images for the cropped regions displayed in main figures and the confirmation of protein purity.
Table S1. Bacterial strains and plasmids used in this study. Table S2. Primers used in this study. Table S3. Characteristics of DzrR homologs in Gram-negative bacteria. Table S4. Homologs of DzrR and DesB are genetically associated in Dickeya, Ralstonia, and Burkholderia. Table S5. Characteristics of Burkholderia genomes for phylogenomic analysis.
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Liang, Z., Lin, Q., Wang, Q. et al. Gram-negative bacteria resist antimicrobial agents by a DzrR-mediated envelope stress response. BMC Biol 21, 62 (2023). https://doi.org/10.1186/s12915-023-01565-7
- Envelope stress response
- RND efflux pump