S-Nitrosylation of the virulence regulator AphB promotes Vibrio cholerae pathogenesis

Abstract

Vibrio cholerae is the etiologic agent of the severe human diarrheal disease cholera. To colonize mammalian hosts, this pathogen must defend against host-derived toxic compounds, such as nitric oxide (NO) and NO-derived reactive nitrogen species (RNS). RNS can covalently add an NO group to a reactive cysteine thiol on target proteins, a process called protein S-nitrosylation, which may affect bacterial stress responses. To better understand how V. cholerae regulates nitrosative stress responses, we profiled V. cholerae protein S-nitrosylation during RNS exposure. We identified an S-nitrosylation of cysteine 235 of AphB, a LysR-family transcription regulator that activates the expression of tcpP, which activates downstream virulence genes. Previous studies show that AphB C235 is sensitive to O2 and reactive oxygen species (ROS). Under microaerobic conditions, AphB formed dimer and directly repressed transcription of hmpA, encoding a flavohemoglobin that is important for NO resistance of V. cholerae. We found that tight regulation of hmpA by AphB under low nitrosative stress was important for V. cholerae optimal growth. In the presence of NO, S-nitrosylation of AphB abolished AphB activity, therefore relieved hmpA expression. Indeed, non-modifiable aphBC235S mutants were sensitive to RNS in vitro and drastically reduced colonization of the RNS-rich mouse small intestine. Finally, AphB S-nitrosylation also decreased virulence gene expression via debilitation of tcpP activation, and this regulation was also important for V. cholerae RNS resistance in vitro and in the gut. These results suggest that the modulation of the activity of virulence gene activator AphB via NO-dependent protein S-nitrosylation is critical for V. cholerae RNS resistance and colonization.

Introduction

Vibrio cholerae is the causative agent of cholera, a severe watery diarrhea disease. There are millions of cases of cholera and thousands of deaths caused by V. cholerae each year [1]. To colonize and cause disease, V. cholerae uses sophisticated signal transduction pathways to activate a set of virulence factors [2,3]. The master virulence regulator ToxT controls expression of an array of virulence genes, including genes involved in the synthesis of cholera toxin and toxin-coregulated pilus (TCP). The expression of toxT requires ToxR and TcpP, and TcpP is regulated by AphA and AphB. Under anaerobic conditions, tcpP expression increases, which is directly regulated by AphB [4,5], a LysR-family protein that is widely conserved in prokaryotes. Under aerobic conditions, one key cysteine residue (Cys235) of AphB is reversibly oxidized. While under low oxygen conditions, AphB Cys235 is reduced, which promotes oligomerization and subsequently enhances AphB activity, thus activating tcpP.

In addition to activating virulence genes, V. cholerae must also coordinate the sequential expression of a series of factors required for defense against host-derived toxic compounds. One of these compounds elevated during infection is nitric oxide (NO), a toxic radical that disrupts the function of proteins containing cysteine residues, enzymes catalyzing iron-dependent reactions, and members of the electron transport chain [6]. Furthermore, NO reacts with other small molecules produced by the immune system to form other toxic reactive nitrogen species (RNS). In the host, NO is generated by acidified nitrite in the stomach and by enzymes of the nitric oxide synthase (NOS) family, such as inducible NOS (iNOS), which is capable of generating large quantities of NO in an inflammatory setting [7]. It has been reported that patients with cholera have an increase in the expression of iNOS in their small intestines [8–10], suggesting that V. cholerae encounters NO during human infections.

To cope with NO produced during infection, many pathogenic bacteria have evolved mechanisms to convert NO into other, less toxic, nitrogen oxides [6]. In V. cholerae, HmpA, a member of the flavohemoglobin family of enzymes, plays a key role in resistance of NO by catalyzing the conversion of NO to nitrous oxide or nitrate [11]. In the presence of NO, hmpA expression is activated by the NO sensor NorR, a predicted σ54-dependent transcriptional regulator [6]. NorR also activates the expression of nnrS, which encodes a novel protein that is important for RNS resistance [6]. NnrS does not directly remove NO but instead it protects the cellular iron pool from the formation of dinitrosyl iron complexes (DNICs) [12]. NO also impacts cellular signaling through S-nitrosylation of protein cysteine residues [13]. To investigate how NO affects V. cholerae protein S-nitrosylation, we performed a proteomic study of V. cholerae exposed to NO using a modified iodoacetyl isobaric tandem mass tags (iodoTMT) approach [14]. We found that the cysteine 235 residue in the key virulence activator AphB was nitrosylated during RNS exposure. We showed that in the absence of nitrosative stress AphB repressed hmpA expression and that nitrosative stress inactivated AphB via protein S-nitrosylation. We also showed that AphB nitrosylation was important for V. cholerae RNS resistance and pathogenesis in vitro and in vivo.

Results

Proteomic profiling of V. cholerae protein S-nitrosylation upon RNS exposure

To investigate how NO affects V. cholerae protein S-nitrosylation, we applied a modified iodoTMT approach [14] to identify V. cholerae protein S-nitrosylation in the presence of NO (Fig 1A). First, we grew V. cholerae under a virulence-inducing condition (AKI medium) [15] and treated the cells with the NO donor DEA NONOate. The reduced thiols in the extracts were blocked with methyl methanethiosulfonate (MMTS). Nitrosylated thiols were then selectively reduced by sodium ascorbate, and reduced cysteines in proteins were specifically labeled with iodoTMTzero. After digestion by trypsin, peptides were captured by an anti-TMT resin. The eluted labeled peptides were then analyzed by LC-MS/MS. Using this method, we identified 67 proteins containing cysteine S-nitrosylation upon exposure to NO (Fig 1B and S1 Dataset). Several of these identified proteins are important for cellular process, protein synthesis, and energy metabolism implying that S-nitrosylation may have a profound impact on bacterial physiology. A few reactive oxygen species (ROS) resistance proteins, such as thioredoxin peroxidase AhpC, thioredoxin TagD, and glutathione synthetase GssA were also detected, suggesting that there are overlapping roles between ROS and RNS resistance. In mammalian cells, thioredoxins can be nitrosylated and transnitrosylate target proteins [16]. We also detected S-nitrosylation in important virulence factors: toxin co-regulated pilin TcpA and toxin coregulated pilus biosynthesis protein TcpH. Interestingly, we identified S-nitrosylation of the key virulence activator AphB [4,5]. AphB contains three cysteine residues, Cys76, Cys94, and Cys235, and AphB peptides containing Cys235-TMT were detected (Fig 1C). Cys76-TMT was also detected but at a lower abundance than Cys235-TMT (S1 Dataset). In this study, we further investigated the relationship between S-nitrosylation and AphB function.

[Figure omitted. See PDF.]

Fig 1. Profiling cysteine S-nitrosylation in the V. cholerae proteome.

A. Schematic of the nitrosylated thiol labeling approach. V. cholerae was grown under the virulence-inducing conditions and subsequently challenged with 100 μM Diethylammonium (Z)-1-(N,N-diethylamino)diazen-1-ium-1,2-diolate (DEA NONOate) for 1 hr. Free thiols in the extracts were blocked. Upon reduction of S-nitrosylated thiols, proteins were labeled with iodoTMTzero. After digestion with trypsin, peptides were captured by an anti-TMT resin. The eluted labeled peptides were then analyzed by LC-MS/MS. B. Pie chart showing the distribution of SNO-modified cysteine-containing proteins in V. cholerae by protein functional categories. C. LC-MS/MS spectrum of AphB peptides containing Cys235 TMT modification. D. Confirmation of AphB cysteine S-nitrosylation. Cultures of V. cholerae ΔaphB with PBAD-aphBWT and cysteine→serine mutant derivatives with or without DEA NONOate exposure were subjected to the biotin switch assay [17]. Western blot analysis on avidin agarose-pulled down samples was then performed using the anti-AphB antibody.

https://doi.org/10.1371/journal.ppat.1010581.g001

To confirm S-nitrosylation of AphB, we adapted the biotin-switch assay [17] to detect nitrosylated AphB in cells exposed to NO. The nitrosylated proteins were selectively reduced and labeled with biotin. Biotinylated proteins were subsequently enriched by avidin agarose, and AphB was detected via western blot using anti-AphB antibodies (the flowchart in Fig 1D). We also included cysteine-to-serine mutants of AphB as these mutant proteins could not be modified [5]. Without addition of NO, no AphB was nitrosylated (therefore biotinylated) (Fig 1D). When NO was added, S-nitrosylation occurred in both wildtype AphB and AphBC76S/C94S mutants, but not in AphBC235S mutants (Fig 1D). These data suggest that AphB is nitrosylated primarily at Cys235 upon cell exposure to NO.

Reduced AphB directly represses hmpA expression, and S-nitrosylation leads to inactive AphB

AphB is a LysR-family transcription regulator that activates V. cholerae virulence gene expression. To study how S-nitrosylation of AphB may facilitate V. cholerae fitness under nitrosative stress, we first scanned for the consensus sequence motif of AphB binding (Fig 2A) across the V. cholerae genome. We identified an AphB box in the promoter region of hmpA (VCA0183), which encodes a key NO detoxifying enzyme [11]. To determine if AphB directly regulates the hmpA promoter, interaction was assessed by electrophoretic mobility shift assays (EMSA) using purified AphB and biotinylated hmpA promoter DNA fragments which contains the putative AphB binding site (Fig 2A). We found that the reduced form of AphB (in the presence of DTT) could bind the hmpA promoter (Fig 2B), and this binding was specific as inclusion of excess amount of unlabeled hmpA promoter DNA abolished the binding (S1A Fig). The AphB binding affinity to the hmpA promoter was ~50% lower than that of the tcpP promoter, a known target of AphB (S1B Fig). These data suggest that AphB may regulate hmpA expression directly.

[Figure omitted. See PDF.]

Fig 2. AphB S-nitrosylation and hmpA expression.

A. AphB consensus binding site (curated in the CollecTF database: http://collectf.umbc.edu/browse/home/) analysis (generated by WebLogo: https://weblogo.berkeley.edu/logo.cgi) analysis revealed a putative binding site in the hmpA promoter region. Arrows: translational start. B. Gel shift assays using purified AphB-His6 and biotinylated hmpA promoter DNA. AphB-His6 proteins were added (from left to right) at the concentrations of 0, 0.125, 0.25, 0.5, 1, and 2 μM, respectively. 20 nM 50mer dsDNA containing the hmpA promoter region were used in each lane. C-E. The effects of AphB S-nitrosylation on the expression of NO-activated genes. ΔaphB with vector pBAD24, PBAD-aphBWT, or PBAD-aphBC235S harboring hmpA-lacZ (C), norR-lacZ (D), or nnrS-lacZ (E) reporters were grown in the minimal medium containing 0.1% arabinose for 4 hrs microaerobically. When indicated, 100 μM DEA NONOate were then added, and all the cultures were incubated for an additional 2 hrs. β-galactosidase activity was then measured. Data are the means ± SD from three independent experiments. ***: P<0.0005; ****: P<0.0001 (two-way ANOVA).

https://doi.org/10.1371/journal.ppat.1010581.g002

Next, we examined how AphB regulates hmpA. We measured hmpA-lacZ expression in ΔaphB mutants containing an empty vector control, PBAD-aphBWT, and PBAD-aphBC235S. In the absence of NO, hmpA was not expressed, consistent with the previous report [11]. In the presence of NO, hmpA expression was highly induced in ΔaphB and aphBWT strains but was low in the non-modifiable aphBC235S mutant (Fig 2C). These data suggested reduced AphB represses hmpA expression. As norR, which encodes a transcriptional activator that turns on hmpA in the presence of NO, is divergently transcribed from hmpA (Fig 2A) [11], we examined whether AphB also regulates norR expression. By measuring the norR-lacZ activity in different aphB strain backgrounds, we found that norR expression was not controlled by either NO or AphB (Fig 2D). Moreover, since NorR-NO activates nnrS expression [11,12], we also examined whether AphB affects nnrS expression. Fig 2E shows that nnrS was induced by NO independent of AphB. Taken together, these data suggest that AphB represses hmpA expression directly and S-nitrosylation negatively affects AphB activity, as non-modifiable AphBC235S mutants retained AphB repression of hmpA even in the presence of NO.

To further examine the effects of S-nitrosylation on AphB activity, we nitrosylated purified AphBWT and AphBC235S in vitro and tested their ability to bind to the hmpA promoter sequence. AphBC235S proteins had similar binding affinity to the hmpA promoter as that of AphBWT (Compare S2A and S2B Fig). When AphBWT was treated with S-Nitroso-L-glutathione (GSNO), the AphB binding of hmpA was abolished (Fig 3A). Addition of DTT in the reaction restored AphB binding capacity. On the other hand, GSNO did not affect AphBC235S binding of hmpA (Fig 3A, right panel). It has been shown that aerobic growth and oxidative stress lead to reduction of AphB dimerization, thus diminishing activity [5, 18]. To test whether nitrosative stress affects AphB dimerization as well, we conducted in vivo crosslink assays. Microaerobically cultured V. cholerae expressing AphBWT and AphBC235S with or without NO exposure was treated with the crosslinker DSP (dithiobis(succinimidyl propionate)) and western blot analyses were then performed. In the presence of NO, wildtype AphB dimer formation was significantly reduced, whereas similar numbers of dimers of AphBC235S mutant proteins were formed without or with NO (Fig 3B). These data indicate that S-nitrosylation through the Cys235 residue reduces AphB multimerization.

[Figure omitted. See PDF.]

Fig 3. The effects of S-nitrosylation on AphB activity.

A. S-nitrosylation in vitro. 1 μM of purified AphBWT and AphBC235S were incubated with or without 1 mM S-Nitroso-L-glutathione (GSNO) for 15 mins, and gel shift assays were performed. When indicated, extra 100 mM DTT was included in the binding reactions. B. The effect of NO on AphB dimerization. ΔaphB containing PBAD-aphBWT or PBAD-aphBC235S were grown in AKI medium for 4 hrs. 0.1% arabinose was then added. When indicated, 50 μM DEA NONOate was then added, and all the cultures were incubated for an additional 2 hrs. Cell pellets were then incubated with the crosslinking reagent DSP. The samples were then subjected to SDS-PAGE and western blots using anti-AphB antibody.

https://doi.org/10.1371/journal.ppat.1010581.g003

AphB S-nitrosylation is important for V. cholerae RNS resistance in vitro and in vivo

Since AphB regulates the expression of hmpA, which encodes a key RNS resistance enzyme, next we investigated the relationship between AphB S-nitrosylation and RNS resistance. V. cholerae strains were grown in minimal media containing different concentrations of NO donors, and OD600 was then monitored. We found that compared to the wildtype, aphBC235S mutants were more sensitive to NO, similar to that of hmpA deletion mutants, whereas ΔaphB mutants was slightly more resistant to NO (Fig 4A). Inclusion of a constitutively expressed hmpA in aphBC235S mutants could relieve its sensitivity to NO (Fig 4A purple triangles), suggesting that AphBC235S effects act through regulation of hmpA.

[Figure omitted. See PDF.]

Fig 4. The effects of AphB S-nitrosylation on V. cholerae growth and colonization.

A. NO resistance. Wildtype and ΔhmpA containing pBAD24 or ΔaphB with vector pBAD24, PBAD-aphBWT, or PBAD-aphBC235S were grown in the M9 minimal medium containing 0.1% casein amino acids, 0.1% arabinose, 0.1 mM IPTG, and the different concentrations of DETA-NONOate indicated. When indicated, the strains harbored either pSRK-Gm or Ptac-hmpA plasmids. The cultures were incubated at 37°C without shaking, and the OD600 was measured after 16 hrs. Data are the means ± SD from three independent experiments. *: P<0.05, **: P<0.01; ****: P<0.0001 (two-way ANOVA). B & C. Colonization. 108 cells of wildtype and ΔhmpA mutants (B) or wildtype and aphBC235S mutants (C) were mixed 1:1 and intragastrically administered to mice without aminoguanidine (AG) treatment (-AG) and mice with AG (+AG). At 7-day post-inoculation, V. cholerae colonization in the small intestine was determined. The competitive index (CI) was calculated as the ratio of mutant to wildtype colonies normalized to the input ratio. Horizontal line: mean CI of 5 mice. **: p <0.005 (Mann-Whitney U test).

https://doi.org/10.1371/journal.ppat.1010581.g004

To examine the effect of S-nitrosylation on V. cholerae colonization, we used a streptomycin-treated adult mouse model, in which bacteria experience host-generated oxidative and nitrosative stress [11,19,20]. To mitigate host-derived RNS, we orally administrated aminoguanidine (AG), an iNOS inhibitor that is routinely used to reduce NO production in mice [21,22]. When the ΔhmpA mutant was co-inoculated with wildtype, ΔhmpA mutants displayed a significant colonization defect in both the small intestine (Fig 4B) and in feces (S3A Fig), consistent with the previous report [11]. When mice were treated with AG, however, hmpA mutant colonization was restored (Figs 4B and S3A) to a level similar to that of iNOS-/- mice [11], confirming that V. cholerae uses HmpA to mitigate RNS and that AG treatment is sufficient to remove RNS stress in vivo. When the non-modifiable aphBC235S mutants were tested in this mouse model, we found that in the -AG group, aphBC235S mutants had an approximately 1000-fold reduction in colonization compared to the wildtype. Whereas in +AG group, aphBC235S mutant colonization was only reduced by about 10-fold (Figs 4C and S3B). These data suggest that S-nitrosylation of AphB is important for V. cholerae to resist RNS during colonization.

AphB repression of hmpA prevents intracellular ROS accumulation

We next examined why AphB repression of hmpA is necessary, given that hmpA expression in the presence of NO is regulated by the transcriptional activator NorR (Fig 2C and [11]). We wondered if AphB repression of hmpA is important for V. cholerae fitness. To answer this question, we first compared hmpA transcription in wildtype and ΔaphB under different NO concentrations. In the wildtype, hmpA was induced by 100 μM NO, whereas in ΔaphB mutants, 10 μM NO could induce hmpA (Fig 5A). As a negative control, hmpA was not induced by NO in the aphBC235S mutant (Fig 5A). These data suggest that AphB S-nitrosylation may be less sensitive to NO than that of NorR activation; therefore, AphB can still repress hmpA expression at low concentrations of NO.

[Figure omitted. See PDF.]

Fig 5. The physiological roles of AphB regulation of hmpA.

A. AphB repression of hmpA at low NO concentrations. ΔaphB with vector pBAD24, PBAD-aphBWT, or PBAD-aphBC235S were grown in the minimal medium containing 0.1% arabinose at 37°C microaerobically for 4 hrs. Various concentrations of DEA NONOate were added and incubated for an additional 1 hr. RNA was harvested and hmpA transcripts were quantified by qPCR and normalized against 16S rRNA. Data are the means ± SD from three independent experiments. **: p <0.001; ****: p <0.0001 (two-way ANOVA). B. Hypersusceptibility of HmpA-overexpressing cells to oxidative stress. 0.1% NaN3 and when indicated, 0.2 mM 2,2 -dipyridyl (DP) were added to the log-phase LB cultures for 30 mins. Cultures were then challenged with 2 mM H2O2 at 37°C for 30’. Viable counts were measured by serial dilution and plating onto LB agar. Data are the means ± SD from three independent experiments. *: p < 0.05 (Ordinary one-way ANOVA). C&D. HmpA overproduction on ROS accumulation (C) and growth (D). V. cholerae harboring an empty vector control or Ptac-hmpA plasmid was grown in LB to stationary phase, and DCFDA staining was performed. Representative histograms of DCFDA-stained cells analyzed by using a CytoFLEX flow cytometer were shown in (C) and CFU counts were shown in (D). ***: p <0.0005 (Student t-test). E. Working model. Reduced AphB represses hmpA at low NO concentrations that may be enough to activate NorR. S-nitrosylation of AphB at high NO concentration leads to full induction of hmpA.

https://doi.org/10.1371/journal.ppat.1010581.g005

Previous studies show that although HmpA flavohemoglobin plays an essential role in nitrosative stress responses in E. coli and Salmonella, dysregulation of hmpA may lead to the exacerbation of oxidative stress through Fenton reactions [23, 24]. Thus, we tested whether overexpression of HmpA affects V. cholerae susceptibility to hydrogen peroxide. Log-phase cultures were treated with sodium azide to inhibit respiration and subsequently challenged with H2O2. We found that the survival of HmpA-overexpressing cells was reduced more than 20-fold compared with cells harboring a vector (Fig 5B). Treatment with the iron chelators 2,2 -dipyridyl (DP) rescued the cells (Fig 5B), indicating that the HmpA-induced potentiation of oxidative stress susceptibility is iron-dependent. Furthermore, we measured intracellular reactive oxygen species (ROS) accumulation in stationary-phase cultures using the redox-sensitive, cell-permeable dye 2’,7’-dichlorodihydrofluorescein diacetate (DCFDA). We found that significant more ROS accumulated in HmpA-overexpressed cells (Fig 5C). The viability of HmpA-overexpressed cells was also reduced (Fig 5D). Taken together, these results suggest that AphB-mediated hmpA repression is important for V. cholerae fitness. We propose that reduced AphB plays an important role in the control of hmpA expression when NorR is already activated by NO at low concentrations. At high NO concentrations AphB is nitrosylated and therefore inactivated, which leads to NorR-activated hmpA expression to detoxify RNS (Fig 5E). In addition, when NO is removed, AphB may also play a role to prevent HmpA-mediated oxidative stress by faster restoration of repression of hmpA that is no longer needed. This hypothesis will be tested in a future study.

AphB S-nitrosylation reduces virulence gene expression

Since AphB is the key virulence activator that is required for tcpP expression, we next tested how AphB S-nitrosylation affects virulence gene expression. We first assessed the ability of AphB to bind to the tcpP promoter DNA by EMSA. We found that like the hmpA promoter (Fig 3A), S-nitrosylation abolished AphBWT, but not AphBC235S, binding of tcpP promoter (Fig 6A). Of note, AphBC235S proteins had similar binding affinity to the tcpP promoter as that of AphBWT (S2B Fig).

[Figure omitted. See PDF.]

Fig 6. The impact of AphB S-nitrosylation on virulence.

A. Interaction with the tcpP promoter. 1 μM of purified AphBWT or AphBC235S were incubated with or without 1 mM GSNO for 15 mins, and gel shift assays using a 50mer biotin-labeled dsDNA of tcpP were performed. When indicated, extra 100 mM DTT was included in the binding reactions. B. tcpP expression. ΔaphB with vector pBAD24, PBAD-aphBWT, or PBAD-aphBC235S were grown in the minimal medium microaerobically for 3 hrs. Various concentrations of DEA NONOate were added and incubated for an additional 1 hr. RNA was harvested, and tcpP transcripts were quantified by qPCR and normalized against 16S rRNA. Data are the means ± SD from three independent experiments. ****: p <0.0001 (two-way ANOVA). C. TcpA production. V. cholerae strains were grown in the minimal medium microaerobically until mid-log phase. 0.1% arabinose (to induce aphB), 0.1 mM taurocholate (to induce virulence genes), and when indicated, 100 μM DEA NONOate were added and incubated for an additional 3 hrs. Cell lysates were then subjected to SDS-PAGE and western blot analysis using an anti-TcpA antibody. D. In vitro competition of a tcpP constitutive mutant. Wildtype and chromosomal lacZ::Ptac-tcpPH mutants were cocultured in the AKI medium microaerobically for 8 hrs in the absence and in the presence of 100 μM DETA NONOate. CFU was then determined. Horizontal line: Mean of 20 repeats from three independent experiments. **: p <0.01 (Student’s t-test). E. In vivo competition. 108 cells of wildtype and lacZ::Ptac-tcpPH mutants were mixed 1:1 and intragastrically administered to mice without aminoguanidine (AG) treatment (-AG) and mice with AG (+AG). At 7-day post-inoculation, V. cholerae colonization in the small intestine was determined. The competitive index (CI) was calculated as the ratio of mutant to wildtype colonies normalized to the input ratio. Horizontal line: mean CI of 5 mice. **: p <0.005 (Mann-Whitney U test).

https://doi.org/10.1371/journal.ppat.1010581.g006

We then measured S-nitrosylation of AphB on tcpP transcription. We found that high concentrations of NO (100 μM) inhibited tcpP expression when wildtype aphB was present (Fig 6B, circles). In aphBC235S mutants, however, tcpP was still expressed in the presence of NO (Fig 6B, triangles). As controls, tcpP was not induced in ΔaphB mutants (Fig 6B, squares). We then tested NO effects on downstream major virulence factor TcpA production by western blots. Fig 6C shows that little, if any, TcpA was produced in ΔaphB mutants. Addition of NO to wildtype significantly reduced TcpA production. However, NO did not affect TcpA when aphBC235S was expressed or when a constitutively expressed tcpP was inserted on the chromosome (lacZ::Ptac-tcpPH) to bypass the requirement of AphB activation. These data suggest that inactivation of AphB by S-nitrosylation leads to a reduction of virulence gene expression.

We then investigated whether the reduction of virulence via AphB S-nitrosylation could have a physiological role. We first performed growth competition assays in AKI medium with or without NO stresses using wildtype and the Ptac-tcpPH strains. We found that the mutants displayed a slight growth defect in the absence of NO, but the viability of Ptac-tcpPH strains was significantly reduced (Figs 6D and S3C). We then tested the Ptac-tcpPH strain competitiveness in vivo. The mutant was defective in colonization of the gut of RNS-rich mice (-AG), but not in that of RNS-mitigated mice (+AG) (Figs 6E and S3C). These data suggest that it may be important for V. cholerae to reduce virulence gene expression when high RNS is encountered.

Discussion

In this study, we found that when V. cholerae cells were exposed to nitric oxide, the key virulence regulator AphB was nitrosylated at Cys235, which inactivated AphB and allowed expression of the AphB-repressed hmpA. HmpA contains an iron-heme moiety that directly catalyzes the NO-decomposition reaction as well as a flavoreductase domain that mediates the transfer of the electrons to and from NO [25]. HmpA plays a critical role in V. cholerae RNS resistance in vitro and in vivo (Fig 4B and [11]). HmpA homologs are important for detoxification of NO during infection with other pathogenic bacteria, such as E. coli, Yersinia pestis, Staphylococcus aureus, and Salmonella enterica [23,26–28]. HmpA is also needed for V. fischeri initiation of symbiosis with squid hosts [29]. Previous studies show that in other pathogenic bacteria, several virulence regulatory proteins are nitrosylated under RNS exposure. For example, S-nitrosylation of SsrB Cys203, a two-component response regulator in Salmonella that activates Salmonella pathogenicity island-2 (SPI-2) type III secretion system, inhibits the DNA-binding capacity of SsrB, and exhibits increased fitness when exposed to RNS in vivo [30]. In S. aureus, S-nitrosylation of the quorum sensing (QS) regulator AgrA leads to decreased transcription of the agr operon and quorum sensing-activated toxin genes. In addition, in E. coli, nitrosylated OxyR activates genes that encode enzymes controlling S-nitrosylation and protecting against nitrosative stress under anaerobic conditions [31]. Interestingly, similar to that of AphB, OxyR also uses the reactive thiols to sense oxygen as well as ROS [32,33].

It is intriguing that V. cholerae deploys AphB to repress hmpA, which is only activated by NorR in the presence of NO [11]. In other pathogenic bacteria, such as Salmonella, HmpA production needs to be tightly regulated [23,24,34] because under aerobic conditions without nitrosative stress, elevated expression of hmpA increases bacterial susceptibility to ROS. We showed here that constitutive expression of hmpA in V. cholerae was detrimental to bacterial survival against ROS attacks or when bacteria were grown to stationary phase (Fig 5), suggesting the importance of AphB repression of hmpA. Interestingly, in almost all bacteria that harbor a hmpA gene, a NO-responsive repressor NsrR is involved in (or predicted to be) regulating hmpA [35]. These include Gram-positive bacilli and actinobacteria; β-proteobacteria such as Ralstonia, Burkholderia, Bordetella; γ-proteobacteria such as all members in Enterobacteria, Shewanella, all other Vibrionales except V. cholerae. The only exception is that in Pseudomonas, hmpA is predicted to be activated by NorR, as in the case of V. cholerae. NsrR belongs to the Rrf2 family of transcriptional regulators. It acts as a strong repressor of hmpA transcription and uses the iron-sulfur cluster to sense NO and relieve the repression of hmpA [36]. In general, repressors can reduce the basal level expression of a promoter and activators elicit high degree of induction [37], leading us to speculate that V. cholerae recruits AphB to serve as a repressor to perform the function of NsrR. It would be interesting to test if a similar repressor is present in other bacteria lacking NsrR, such as Pseudomonas, as well.

V. cholerae resides in both aquatic environments and in human small intestines. AphB plays a pivotal role in the life cycle of V. cholerae. During the transition from oxygen-rich environments to the oxygen-poor intestine, AphB Cys235 is reduced and can activate downstream virulence genes. The rate of reduction of AphB is low and another redox-sensing regulator, OhrR, is needed to jump-start virulence gene expression [38]. During ROS insults, AphB Cys235 is oxidized, which leads to the derepession of ohrA, encoding an organic hydroperoxide resistant protein [18]. This pathway of regulation can explain our data that aphBC235S mutant colonization was not fully restored in the +AG mice (Fig 4C), as ohrA expression was repressed in aphBC235S mutants. Similar to that of AphB sensing nitrosative stress we showed in this study, AphB Cys235 is less sensitive to oxidation than that of OhrR oxidation, thus OhrA production is tightly regulated. Evidently, AphB Cys235 is less prone to redox potential changes, and this may be important for V. cholerae physiology.

AphB is a key virulence regulator. S-nitrosylation of AphB leads to the reduction of tcpPH expression and downstream TcpA production (Fig 6). An inability to turn off tcpPH expression in the presence of nitrosative stress resulted in growth disadvantages in vitro and in vivo (Fig 6). The mechanisms behind this are currently unknown. We tested tcpA deletion in Ptac-tcpPH strain and found that it still displayed the growth defect with NO (S4 Fig), suggesting that dysregulated TcpA pilin production is not involved in the RNS resistance. Interestingly, V. cholerae employs QS systems to temporally control virulence during infection. QS represses virulence gene expression and biofilm formation while activating production of extracellular proteases, suggesting the importance of QS in entering and exiting the host [39–42]. Host-derived RNS increases as the infection progresses [8], which increases the rate of detachment from the epithelium and subsequently could permit individual cells to establish new infection foci in the intestine or to exit the host. AphB S-nitrosylation therefore provides another layer of regulation to repress virulence as needed.

Methods and materials

Ethics statement

All animal experiments were performed in strict accordance with the animal protocol (Protocol No. 805287) that was approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Pennsylvania.

Strains, plasmids, and culture conditions

V. cholerae El Tor C6706 [43] was used as the primary wildtype strain in this study. Both V. cholerae and E. coli were propagated in LB (Luria-Bertani) Miller medium with appropriate antibiotics at 37°C, unless otherwise noted. AKI medium was used to induce virulence gene expression [44]. Cultures were grown aerobically (shaking at 250 rpm) or microaerobically (stationary growth). For supplying NO donors in cultures, either diethylammonium (Z)-1-(N,N-diethylamino)diazen-1-ium-1,2-diolate (DEA NONOate)(half-life of 2 minutes at 37°C) or (Z)-1-[N-(2-aminoethyl)-N-(2-ammonioethyl)amino]diazen-1-ium-1,2-diolate (DETA NONOate)(half-life of 20 hours at 37°C) (Cayman Chemical) was used.

In-frame deletion of aphB (VC1049), PBAD-aphB wildtype and cysteine to serine mutant derivatives, and plasmids to make recombinant AphB proteins (AphB-His6) were constructed as described previously [5]. Transcriptional hmpA- (VCA0183), nnrS- (VC2330), and norR- (VCA0182) LacZ reporters, and hmpA in-frame deletion were constructed as described in [11]. Ptac-hmpA plasmid was constructed by cloning hmpA coding sequences into pMal-c2x (New England Biolab) and then subcloned to pSRK-Gm [45]. Chromosomal aphB variants used for mouse experiments were constructed by the Tn7-based genomic integration system [46]. Briefly, DNA sequences of aphB wildtype or C235S mutant with the native promoter (341 bp upstream of ATG) were PCR amplified and cloned into the pGP704-mTn7 vector via NotI restriction cloning. The insertion of the aphB wildtype or C235S mutant into the V. cholerae genome was done by triparental mating among V. cholerae ΔaphB, E. coli S17-1 λpir carrying the helper plasmid pUX-BF13, and E. coli SM10 λpir carrying aphB variants on pGP704-mTn7. Exconjugants were selected on thiosulfate-citrate-bile salts-sucrose (TCBS) agar plates containing gentamicin (20 μg/ml). The insertion of mini Tn7 cassette at the intergenic region of VC0487 and VC0488 was confirmed by PCR and sequencing using flanking primers of insertion site. The chromosomal lacZ::Ptac-tcpPH strain was constructed by the multiplex genome editing by natural transformation (MuGENT) method [47]. Briefly, Ptac promoter and tcpPH sequence were PCR amplified from pMal-c2x (New England Biolab) and V. cholerae C6706 genome respectively, stitched with the lacZ UP and Down 3kb arms, cotransformed into V. cholerae C6706 with VC1807::spec DNA, and selected on LB agar plates containing spectinomycin (100 μg/ml). The insertion of Ptac-tcpPH was confirmed by PCR and sequencing. The primer sequences used for the study and further details about the constructions are available upon request.

Proteomic profiling of thiol S-nitrosylation in V. cholerae

Iodoacetyl isobaric tandem mass tags (iodoTMT) labeling of S-nitrosylated thiols was performed by modifying the method described previously [14,48]. Briefly, V. cholerae were grown in AKI medium without shaking for 4 hrs and then challenged with 100 μM DEA NONOate for 1 hr. The cell lysates were first incubated with 20 mM methyl methanethiosulfonate (MMTS) (Pierce) and then reduced with 20 mM sodium ascorbate. Proteins containing reduced thiols were then labeled with iodoTMTzero and digested with trypsin (8 μg trypsin/100 μg proteins). IodoTMTzero-labeled peptides were enriched by using the Anti-TMT Resin and eluted by TMT elution buffer (Thermo Scientific). The eluted peptides were submitted to Poochon Scientific (Frederick, MD). Liquid chromatography tandem mass spectrometry (LC-MS/MS) analysis was performed using a Q Exactive Orbitrap mass spectrometer (Thermo Scientific) coupled with an UltiMate 3000 nano UPLC system (Thermo Scientific). Peptide sequences were identified using the Proteome Discoverer 2.4 software (Thermo, San Jose, CA) based on the SEQUEST and percolator algorithms. MS/MS spectra were searched against a UniProt V. cholerae protein database and a common contaminants database using full tryptic specificity with up to two missed cleavages. Variable modifications included in the search were an addition mass of TMTzero (324.2 Da) on cysteine. Consensus identification lists were generated with false discovery rates set at 1% for protein, peptide, and site identifications.

Detection of S-nitrosylated AphB using biotin switch assays

Overnight cultures of ΔaphB with PBAD-aphBWT, PBAD-aphBC76S/C94S, and PBAD-aphBC235S were inoculated into the AKI medium and incubated microaerobically at 37°C for 4 hrs. 0.1% arabinose and, when indicated, 100 μM DEA-NONOate were added and further incubated for 1 hr. Bacterial cells were collected and subjected to Cayman Chemical’s S-nitrosylated protein detection assay kit (biotin switch) [17] with modifications. Briefly, total proteins were treated with MMTS to block free thiols, reduced by sodium ascorbate, and labeled with biotin. Biotinylated proteins were then enriched by using avidin agarose (Pierce). Proteins from both flowthrough and eluted samples were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and AphB was detected by western blot using an anti-AphB antibody [5].

Gel retardation assays

AphBWT-His6 and AphBC235S-His6 proteins were expressed and purified on nickel columns according to the manufacturer’s instructions (Qiagen). Double-stranded biotin-labeled hmpA (CAATAAATTAGGTTTTGGCATGCAATCTGCTCATTAAGAAGGCTTGTGTAGAGTGGCTCA) and tcpP (GCAATTAAGTTCTCATTATCAACTGCAGAATTAGATTGCAAATAATTATATTAAAAAAAA) probes were prepared by annealing the single-stranded 5’biotin-labeled sense and antisense 50mer oligos after denaturation. Binding reactions contained various amount of AphB proteins and 20 nM of DNA in a buffer consisting of 10 mM Tris-HCl (pH 7.5), 1 mM EDTA, 50 mM KCl, 100 μg/ml BSA, 10 μg of poly dI-dC/ml, and when indicated, 100 mM dithiothreitol (DTT). After 20 minutes of incubation at 25°C, samples were size-fractionated using 5% polyacrylamide gels in 0.5X Tris-acetate-EDTA (TAE) buffer. The biotin-labeled free DNA and protein-DNA complexes were electrophoretic transferred via binding reactions to nylon membrane and visualized using horseradish peroxidase (HRP) conjugated streptavidin (Pierce). To S-nitrosylate purified AphB proteins, DTT was first removed from AphBWT and AphBC235S by dialysis. 1 μM proteins were then incubated with 1 mM S-Nitroso-L-glutathione (GSNO) (Cayman Chemical) for 15 mins at 25°C.

Gene expression studies

Overnight cultures of ΔaphB with vector pBAD24, PBAD-aphBWT, or PBAD-aphBC235S harboring different lacZ reporters were inoculated 1:100 into the minimal medium described in [11] containing 0.1% arabinose. After 4 hrs of growth, various amounts of DEA-NONOate indicated were added to the cultures. Two hours later, β-galactosidase activity was measured. For qPCR quantification of hmpA and tcpP transcripts, the above cultures were treated with DEA-NONOate for one hour before bacterial cells were harvested and total RNA was purified by using RNeasy Kit (Qiagen). RNA reverse transcription was performed by using the SuperScript II Kit (Invitrogen). Quantitative realtime qPCR was performed on a CFX96 Real-Time system (BioRad) using primers specific for hmpA and tcpP. The 16S rRNAs were used as internal controls in all reactions.

Detection of TcpA and AphB

Whole-cell lysates were prepared from bacterial cultures and samples were normalized to the amount of total protein as assayed by the Biorad protein assay (Biorad). The samples were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) on a 12% polyacrylamide gel and transferred to nitrocellulose membrane for western blot analysis using polyclonal rabbit anti-TcpA [40] or AphB antibody[5].

In vivo crosslinking of AphB

ΔaphB containing PBAD-aphBWT or PBAD-aphBC235S plasmids were grown in the AKI medium for 4 hrs. 0.1% arabinose and when indicated, 50 μM DEA NONOate were then added and all the cultures were incubated for an additional 2 hrs. Cells were resuspended in PBS and treated with 0.8 mM crosslinking reagent DSP (dithiobis(succinimidyl propionate))(Thermo Fisher) on ice for 2 hrs. The cross-linking reactions were terminated by adding 50 mM Tris-HCl (pH 7.5). Cell pellets were resuspended in SDS loading buffer without reducing agents and AphB multimers were detected by western blots using AphB antibody.

Intracellular ROS accumulation

V. cholerae containing a vector control or Ptac-hmpA plasmid were grown in LB with 0.5 mM IPTG at 37°C for 8 hrs with aeriation. Cell pellets were washed with PBS and resuspended in PBS with 20 μM of the redox-sensitive, cell-permeable dye 2’,7’-dichlorodihydrofluorescein diacetate (DCFDA) (Abcam). Cells were then stained for 1 hr at 37°C, and DCFDA-stained cells were subjected to flow cytometry (CytoFLEX, BD) for ROS detection using the 488 nm laser for excitation and detected at 535 nm.

V. cholerae mouse colonization

The streptomycin-treated adult mouse model was used to examine V. cholerae RNS resistance in vivo. Briefly, six-week-old CD-1 mice were used. 0.5% (wt/vol) streptomycin and 0.5% aspartame were added to the drinking water throughout the experiment. After 3 days of streptomycin treatment, 1 mg/ml aminoguanidine (AG, Acros Organics) was added to the drinking water of +AG group of mice and throughout remainder of the experiment. Additionally, the equivalent of 1 mg of AG was orally gavaged into each mouse in the +AG group of mice daily throughout the remainder of the experiment. One day after starting the AG treatment, approximately 108 CFU of each of the two differentially labeled strains (wildtype lacZ+ and mutant lacZ-) were mixed at a 1:1 ratio and intragastrically administered to each mouse. Fecal pellets were collected from each mouse at the indicated time points, resuspended in LB, serially diluted, and then plated on plates containing 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside (X-gal) and appropriate antibiotics. At day 7, small intestine tissues were harvested and CFU of small intestine-colonized V. cholerae were enumerated. The competitive index was calculated as the ratio of mutant to wildtype colonies normalized to the input ratio.

Supporting information

S1 Fig. EMSA controls.

A. Specificity of AphB binding of hmpA. AphB-His6 proteins were mixed with 20 nM 50mer biotin-labeled double-stranded hmpAp DNA in the absence or in the presence of 1 μg unlabeled hmpA promoter DNA. B. AphB on tcpP. AphB-His6 proteins added (from left to right) at concentrations of 0, 0.125, 0.25, 0.5, 1, and 2 μM, respectively. 20 nM 50mer dsDNA containing the tcpP promoter region were used in each lane.

https://doi.org/10.1371/journal.ppat.1010581.s001

(PDF)

S2 Fig. EMSA using purified AphBC235S-His6.

AphB-His6 proteins added (from left to right) at concentrations of 0, 0.125, 0.25, 0.5, 1, and 2 μM, respectively. 20 nM 50mer dsDNA containing hmpA (A) or tcpP (B) promoter region were used in each lane.

https://doi.org/10.1371/journal.ppat.1010581.s002

(PDF)

S3 Fig. Streptomycin-treated adult mouse colonization.

108 cells of wildtype and ΔhmpA mutants (A) or wildtype and aphBC235S mutants (B) or wildtype and Ptac-tcpPH mutants (C) were mixed 1:1 and intragastrically administered to mice without aminoguanidine (AG) treatment (-AG) and mice with AG (+AG). V. cholerae CFU from fecal samples was determined daily. The competitive index (CI) was calculated as the ratio of mutant to wildtype colonies normalized to the input ratio. Horizontal line: mean CI of 5 mice. *: p <0.05; **: p <0.005; ***: p <0.001 (Mann-Whitney U test).

https://doi.org/10.1371/journal.ppat.1010581.s003

(PDF)

S4 Fig. TcpA effects in the tcpP constitutive mutant.

Wildtype and lacZ::Ptac-tcpPH mutants or wildtype and lacZ::Ptac-tcpPH/ΔtcpA mutants were cocultured in the AKI medium microaerobically for 8 hrs in the absence and in the presence of 100 μM DETA NONOate. CFU was then determined. Data are the means ± SD from 6 independent experiments. ***: p <0.005 (Ordinary one-way ANOVA).

https://doi.org/10.1371/journal.ppat.1010581.s004

(PDF)

S1 Dataset. V. cholerae S-nitrosylation proteome.

https://doi.org/10.1371/journal.ppat.1010581.s005

(XLSX)

Acknowledgments

We thank Drs. Andrew Stern, Yunduan Wang, and Mark Goulian for technical support and helpful discussion.

Citation: Chen J, Byun H, She Q, Liu Z, Ruggeberg K-G, Pu Q, et al. (2022) S-Nitrosylation of the virulence regulator AphB promotes Vibrio cholerae pathogenesis. PLoS Pathog 18(6): e1010581. https://doi.org/10.1371/journal.ppat.1010581

References

1. Clemens JD, Nair GB, Ahmed T, Qadri F, Holmgren J. Cholera. Lancet. 2017;390(10101):1539–49. pmid:28302312.

2. Matson JS, Withey JH, DiRita VJ. Regulatory networks controlling Vibrio cholerae virulence gene expression. Infect Immun. 2007;75(12):5542–9. Epub 2007/09/19. pmid:17875629; PubMed Central PMCID: PMC2168339.

3. Hsiao AN, Zhu J. Pathogenicity and virulence regulation of Vibrio cholerae at the interface of host-gut microbiome interactions. Virulence. 2020;11(1):1582–99. WOS:000588140600001. pmid:33172314

4. Kovacikova G, Lin W, Skorupski K. The LysR-type virulence activator AphB regulates the expression of genes in Vibrio cholerae in response to low pH and anaerobiosis. Journal of bacteriology. 2010;192(16):4181–91. Epub 2010/06/22. pmid:20562308; PubMed Central PMCID: PMC2916415.

5. Liu Z, Yang M, Peterfreund GL, Tsou AM, Selamoglu N, Daldal F, et al. Vibrio cholerae anaerobic induction of virulence gene expression is controlled by thiol-based switches of virulence regulator AphB. Proc Natl Acad Sci U S A. 2011;108(2):810–5. Epub 2010/12/29. pmid:21187377; PubMed Central PMCID: PMC3021084.

6. Poole RK. Nitric oxide and nitrosative stress tolerance in bacteria. Biochemical Society transactions. 2005;33(Pt 1):176–80. Epub 2005/01/26. pmid:15667299.

7. Fang FC, Vazquez-Torres A. Reactive nitrogen species in host-bacterial interactions. Current opinion in immunology. 2019;60:96–102. WOS:000493218500014. pmid:31200187

8. Janoff EN, Hayakawa H, Taylor DN, Fasching CE, Kenner JR, Jaimes E, et al. Nitric oxide production during Vibrio cholerae infection. The American journal of physiology. 1997;273(5 Pt 1):G1160–7. Epub 1997/12/31. pmid:9374715.

9. Qadri F, Raqib R, Ahmed F, Rahman T, Wenneras C, Das SK, et al. Increased levels of inflammatory mediators in children and adults infected with Vibrio cholerae O1 and O139. Clinical and diagnostic laboratory immunology. 2002;9(2):221–9. Epub 2002/03/05. pmid:11874856; PubMed Central PMCID: PMC119937.

10. Rabbani GH, Islam S, Chowdhury AK, Mitra AK, Miller MJ, Fuchs G. Increased nitrite and nitrate concentrations in sera and urine of patients with cholera or shigellosis. The American journal of gastroenterology. 2001;96(2):467–72. Epub 2001/03/10. pmid:11232692.

11. Stern AM, Hay AJ, Liu Z, Desland FA, Zhang J, Zhong Z, et al. The NorR regulon is critical for Vibrio cholerae resistance to nitric oxide and sustained colonization of the intestines. mBio. 2012;3(2):e00013–12. Epub 2012/04/19. pmid:22511349; PubMed Central PMCID: PMC3345576.

12. Stern AM, Liu B, Bakken LR, Shapleigh JP, Zhu J. A novel protein protects bacterial iron-dependent metabolism from nitric oxide. Journal of bacteriology. 2013;195(20):4702–8. Epub 2013/08/13. pmid:23935055; PubMed Central PMCID: PMC3807435.

13. Hess DT, Matsumoto A, Kim SO, Marshall HE, Stamler JS. Protein S-nitrosylation: purview and parameters. Nat Rev Mol Cell Biol. 2005;6(2):150–66. pmid:15688001.

14. Qu Z, Meng F, Bomgarden RD, Viner RI, Li J, Rogers JC, et al. Proteomic quantification and site-mapping of S-nitrosylated proteins using isobaric iodoTMT reagents. J Proteome Res. 2014;13(7):3200–11. Epub 2014/06/14. pmid:24926564; PubMed Central PMCID: PMC4084841.

15. Iwanaga M, Yamamoto K. New medium for the production of cholera toxin by Vibrio cholerae O1 biotype El Tor. J Clin Microbiol. 1985;22(3):405–8. Epub 1985/09/01. pmid:2995438; PubMed Central PMCID: PMC268420.

16. Sengupta R, Holmgren A. Thioredoxin and thioredoxin reductase in relation to reversible S-nitrosylation. Antioxidants & redox signaling. 2013;18(3):259–69. Epub 2012/06/19. pmid:22702224.

17. Jaffrey SR, Snyder SH. The biotin switch method for the detection of S-nitrosylated proteins. Sci STKE. 2001;2001(86):pl1. Epub 2001/12/26. pmid:11752655.

18. Liu Z, Wang H.; Zhou Z.; Sheng Y.; Naseer N.; Kan B.; Zhu J. Thiol-based switch mechanism of virulence regulator AphB modulates oxidative stress response in Vibrio cholerae. Molecular microbiology. 2016;102(5):939–49. PubMed Central PMCID: PMC5123930. pmid:27625149

19. Spees AM, Wangdi T, Lopez CA, Kingsbury DD, Xavier MN, Winter SE, et al. Streptomycin-Induced Inflammation Enhances Escherichia coli Gut Colonization Through Nitrate Respiration. mBio. 2013;4(4):e00430–13. pmid:23820397; PubMed Central PMCID: PMC3705454.

20. Wang H, Xing X, Wang J, Pang B, Liu M, Larios-Valencia J, et al. Hypermutation-induced in vivo oxidative stress resistance enhances Vibrio cholerae host adaptation. PLoS Pathog. 2018;14(10):e1007413. pmid:30376582; PubMed Central PMCID: PMC6226196.

21. Cross AH, Misko TP, Lin RF, Hickey WF, Trotter JL, Tilton RG. Aminoguanidine, an inhibitor of inducible nitric oxide synthase, ameliorates experimental autoimmune encephalomyelitis in SJL mice. J Clin Invest. 1994;93(6):2684–90. PubMed Central PMCID: PMC294515. pmid:7515395

22. Winter SE, Winter MG, Xavier MN, Thiennimitr P, Poon V, Keestra AM, et al. Host-derived nitrate boosts growth of E. coli in the inflamed gut. Science. 2013;339(6120):708–11. pmid:23393266; PubMed Central PMCID: PMC4004111.

23. Bang IS, Liu L, Vazquez-Torres A, Crouch ML, Stamler JS, Fang FC. Maintenance of nitric oxide and redox homeostasis by the salmonella flavohemoglobin hmp. J Biol Chem. 2006;281(38):28039–47. pmid:16873371.

24. Wu G, Corker H, Orii Y, Poole RK. Escherichia coli Hmp, an "oxygen-binding flavohaemoprotein", produces superoxide anion and self-destructs. Archives of microbiology. 2004;182(2–3):193–203. Epub 2004/09/02. pmid:15340787.

25. Poole RK, Hughes MN. New functions for the ancient globin family: bacterial responses to nitric oxide and nitrosative stress. Mol Microbiol. 2000;36(4):775–83. Epub 2000/06/09. pmid:10844666.

26. Richardson AR, Dunman PM, Fang FC. The nitrosative stress response of Staphylococcus aureus is required for resistance to innate immunity. Mol Microbiol. 2006;61(4):927–39. WOS:000239336100008. pmid:16859493

27. Stevanin TM, Read RC, Poole RK. The hmp gene encoding the NO-inducible flavohaemoglobin in Escherichia coli confers a protective advantage in but not in vitro: Links resisting killing within macrophages, with swarming motility. Gene. 2007;398(1–2):62–8. WOS:000248784200008. pmid:17611046

28. Sebbane F, Lemaitre N, Sturdevant DE, Rebeil R, Virtaneva K, Porcella SF, et al. Adaptive response of Yersinia pestis to extracellular effectors of innate immunity during bubonic plague. P Natl Acad Sci USA. 2006;103(31):11766–71. WOS:000239616400061. pmid:16864791

29. Wang Y, Dunn AK, Wilneff J, McFall-Ngai MJ, Spiro S, Ruby EG. Vibrio fischeri flavohaemoglobin protects against nitric oxide during initiation of the squid-Vibrio symbiosis. Mol Microbiol. 2010;78(4):903–15. Epub 2010/09/08. pmid:20815823; PubMed Central PMCID: PMC2978254.

30. Husain M, Jones-Carson J, Song M, McCollister BD, Bourret TJ, Vazquez-Torres A. Redox sensor SsrB Cys203 enhances Salmonella fitness against nitric oxide generated in the host immune response to oral infection. Proc Natl Acad Sci U S A. 2010;107(32):14396–401. pmid:20660761; PubMed Central PMCID: PMC2922535.

31. Seth D, Hausladen A, Wang YJ, Stamler JS. Endogenous protein S-Nitrosylation in E. coli: regulation by OxyR. Science. 2012;336(6080):470–3. Epub 2012/04/28. pmid:22539721.

32. Zheng M, Aslund F, Storz G. Activation of the OxyR transcription factor by reversible disulfide bond formation. Science. 1998;279(5357):1718–21. Epub 1998/03/28. pmid:9497290.

33. Kim SO, Merchant K, Nudelman R, Beyer WF Jr., Keng T, DeAngelo J, et al. OxyR: a molecular code for redox-related signaling. Cell. 2002;109(3):383–96. Epub 2002/05/23. pmid:12015987.

34. Choi G, Kim D, Im H, Choi SH. A Nitric Oxide-Responsive Transcriptional Regulator NsrR Cooperates With Lrp and CRP to Tightly Control the hmpA Gene in Vibrio vulnificus. Front Microbiol. 2021;12:681196. Epub 2021/06/08. pmid:34093504; PubMed Central PMCID: PMC8175989.

35. Rodionov DA, Dubchak IL, Arkin AP, Alm EJ, Gelfand MS. Dissimilatory metabolism of nitrogen oxides in bacteria: comparative reconstruction of transcriptional networks. PLoS Comput Biol. 2005;1(5):e55. Epub 2005/11/02. pmid:16261196; PubMed Central PMCID: PMC1274295.

36. Crack JC, Le Brun NE. Redox-Sensing Iron-Sulfur Cluster Regulators. Antioxid Redox Signal. 2018;29(18):1809–29. Epub 2017/10/03. pmid:28967283.

37. Ptashne M. Repressors. Curr Biol. 2007;17(17):R740–1. Epub 2007/09/07. pmid:17803917.

38. Liu Z, Wang H, Zhou Z, Naseer N, Xiang F, Kan B, et al. Differential Thiol-Based Switches Jump-Start Vibrio cholerae Pathogenesis. Cell reports. 2016;14(2):347–54. pmid:26748713; PubMed Central PMCID: PMC4715633.

39. Zhu J, Mekalanos JJ. Quorum sensing-dependent biofilms enhance colonization in Vibrio cholerae. Dev Cell. 2003;5(4):647–56. Epub 2003/10/11. pmid:14536065.

40. Zhu J, Miller MB, Vance RE, Dziejman M, Bassler BL, Mekalanos JJ. Quorum-sensing regulators control virulence gene expression in Vibrio cholerae. Proceedings of the National Academy of Sciences of the United States of America. 2002;99(5):3129–34. pmid:11854465.

41. Hammer BK, Bassler BL. Quorum sensing controls biofilm formation in Vibrio cholerae. Molecular microbiology. 2003;50(1):101–4. Epub 2003/09/26. pmid:14507367.

42. Vaitkevicius K, Lindmark B, Ou G, Song T, Toma C, Iwanaga M, et al. A Vibrio cholerae protease needed for killing of Caenorhabditis elegans has a role in protection from natural predator grazing. Proc Natl Acad Sci U S A. 2006;103(24):9280–5. Epub 2006/06/07. pmid:16754867; PubMed Central PMCID: PMC1482601.

43. Joelsson A, Liu Z, Zhu J. Genetic and phenotypic diversity of quorum-sensing systems in clinical and environmental isolates of Vibrio cholerae. Infect Immun. 2006;74(2):1141–7. pmid:16428762; PubMed Central PMCID: PMC1360356.

44. Iwanaga M, Yamamoto K, Higa N, Ichinose Y, Nakasone N, Tanabe M. Culture conditions for stimulating cholera toxin production by Vibrio cholerae O1 El Tor. Microbiol Immunol. 1986;30(11):1075–83. Epub 1986/01/01. pmid:3543624

45. Khan SR, Gaines J, Roop RM, Farrand SK. Broad-host-range expression vectors with tightly regulated promoters and their use to examine the influence of TraR and TraM expression on Ti plasmid quorum sensing. Appl Environ Microb. 2008;74(16):5053–62. WOS:000258602100009. pmid:18606801

46. Beyhan S, Tischler AD, Camilli A, Yildiz FH. Transcriptome and phenotypic responses of Vibrio cholerae to increased cyclic di-GMP level. Journal of bacteriology. 2006;188(10):3600–13. Epub 2006/05/05. pmid:16672614; PubMed Central PMCID: PMC1482859.

47. Dalia AB, McDonough E, Camilli A. Multiplex genome editing by natural transformation. Proc Natl Acad Sci U S A. 2014;111(24):8937–42. pmid:24889608; PubMed Central PMCID: PMC4066482.

48. Zhou Y, Pu Q, Chen J, Hao G, Gao R, Ali A, et al. Thiol-based functional mimicry of phosphorylation of the two-component system response regulator ArcA promotes pathogenesis in enteric pathogens. Cell reports. 2021;37(12):110147. Epub 2021/12/23. pmid:34936880; PubMed Central PMCID: PMC8728512.