Tyrosine Nitration of Flagellins: a Response of Sinorhizobium meliloti to Nitrosative Stress
ABSTRACT
Rhizobia are bacteria which can either live as free organisms in the soil or interact with plants of the legume family with, as a result, the formation of root organs called nodules in which differentiated endosymbiotic bacteria fix atmospheric nitrogen to the plant’s benefit. In both lifestyles, rhizobia are exposed to nitric oxide (NO) which can be perceived as a signaling or toxic molecule. NO can act at the transcriptional level but can also modify proteins by S-nitrosylation of cysteine or nitration of tyrosine residues. However, only a few molecular targets of NO have been described in bacteria and none of them have been characterized in rhizobia. Here, we examined tyrosine nitration of Sinorhizobium meliloti proteins induced by NO. We found three tyrosine-nitrated proteins in S. meliloti grown under free-living conditions, in response to an NO donor. Two nitroproteins were identified by mass spectrometry and correspond to flagellins A and B. We showed that one of the nitratable tyrosines is essential to flagellin function in motility.
IMPORTANCE Rhizobia are found as free-living bacteria in the soil or in interaction with plants and are exposed to nitric oxide (NO) in both environments. NO is known to have many effects on animals, plants, and bacteria where only a few molecular targets of NO have been described so far. We identified flagellin A and B by mass spectrometry as tyrosine-nitrated proteins in Sinorhizobium meliloti in vivo. We also showed that one of the nitratable tyrosines is essential to flagellin function in motility. The results enhanced our understanding of NO effects on rhizobia. Identification of bacterial flagellin nitration opens a new possible role of NO in plant-microbe interactions.
INTRODUCTION
Soil bacteria that interact with plants are of major importance in agriculture. They include the alphaproteobacteria such as Bradyrhizobium japonicum, Rhizobium etli, and Sinorhizobium meliloti. These bacteria can adopt two different lifestyles. On the one hand, they are found as free-living organisms in the soil, but on the other hand, they are capable of establishing a symbiosis with leguminous plants, such as soybean, common bean, and alfalfa (1, 2). The interaction is initiated when rhizobia move toward plant roots and then express early nodulation genes leading to the production of nod factors (3). Rhizobia induce the formation of new plant root organs (nodules) where they differentiate into bacteroids which fix atmospheric nitrogen to the benefit of the plant. In exchange, they receive carbon sources in the form of dicarboxylic acids and are accommodated in an ecological niche. The model rhizobium S. meliloti performs a symbiosis with plants from the genera Medicago, Melilotus, and Trigonella.
In both situations, whether in the soil or inside nodules, S. meliloti is exposed to nitric oxide (NO). NO is a gas molecule that can be produced in the soil from nitrates in part due to denitrification (4, 5). Denitrification is a form of respiration for bacteria living under oxygen-limited environments in which the sequential transformation of nitrate (NO3−) into nitrite (NO2−) and then into NO, nitrous oxide (N2O), and finally dinitrogen (N2) is coupled to bioenergy production. NO, an intermediate product of denitrification, is a diffusible radical that has been recognized as an integral signaling molecule in mammals and more recently in bacteria and plant cells. In mammals, NO plays pivotal roles in vasodilation, neurotransmission, and immune system function, while in plants, NO controls physiological functions in plant growth, immunity, and development, including seed germination, flowering, fruit ripening, and senescence (6). In bacteria, NO is involved in several biological processes, such as biofilm formation and quorum sensing (7). When present at higher concentrations, NO can be a toxic molecule which limits bacterial growth. As such, it is an important part of the defense arsenal of mammals and plants during host-pathogen interactions (8). Nevertheless, NO has been shown to be involved in all steps of legume symbiosis (9–11).
All of the direct cellular effects of NO are far from being known. Various biomolecules are targeted by NO. Indeed, NO can act on gene transcription via specific regulators, such as NnrR and FixLJ in S. meliloti (12, 13) and DosS/DosT in Mycobacterium tuberculosis (14). When exposed to NO, S. meliloti develops a transcriptional response involving about 100 genes (13), among which are genes implicated in NO degradation (hmp and nor). NO can also act directly on proteins either by iron nitrosylation of heme cofactors, by S-nitrosylation of cysteine residues, or by nitration of tyrosine residues within proteins (15). In contrast to the high number of genes potentially affected by the presence of NO, few proteins are modified posttranslationally by NO (16). Tyrosine nitration has been understudied because this modification has long been considered an irreversible mechanism which might have deleterious consequences instead of a regulatory role. However, there are indications that denitration may occur in vivo (17). Tyrosine nitration is restricted to specific target tyrosine residues and can lead to the activation or inhibition of the target proteins. These modified proteins are challenging to analyze, as this modification is supposed to affect a small number of proteins and only a few tyrosines within each protein (18). Few nitrated tyrosines have been identified in bacteria (19–22). Interestingly ArgR, a transcriptional regulator for arginine biosynthesis in Escherichia coli has been shown to be nitrated by peroxynitrite in vitro on one or more of the three tyrosine residues present on the protein (23). This nitration causes the degradation of the hexameric or trimeric form of the protein which results in the inability to bind the operator sequences in the promoter of arginine biosynthesis genes even in the presence of arginine. The consequence of this is the derepression of transcription. Hence, tyrosine nitration might have a role to play in the regulation of bacterial metabolism.
Tyrosine-nitrated proteins have also been identified in plants and in particular in legume root nodules. Indeed, two important proteins have been shown to be tyrosine nitrated, i.e., the leghemoglobin, an abundant hemeprotein of legume nodules that plays an essential role as an O2 transporter, and the glutamine synthetase which is responsible for the assimilation of the ammonia released by nitrogen fixation (24, 25). Defining one set of specific targets of NO may help us understand the importance of protein posttranslational modifications in growth inhibition as well as in bacterial adaptation to the soil or plant environment. In this work, we describe the detection of S. meliloti tyrosine-nitrated proteins.
RESULTS
NO-modified protein profiling in S. meliloti.
During the symbiotic interaction between legumes and rhizobia, only two important plant proteins have been shown to be tyrosine nitrated, while no modified rhizobial proteins have been found yet. Rhizobium extraction from nodules can be a limiting step to study protein posttranslational modifications. Hence, we addressed the question of whether there are S. meliloti proteins which can be tyrosine nitrated by using free-living cells. S. meliloti cells were grown in liquid culture for 1 hour in the presence of the following concentrations of the NO donor spermine NONOate (SpNO): 25, 75, and 250 μM. A transcriptomic analysis of the S. meliloti response to 25 μM SpNO has been performed previously (13), and it was shown in that work that cell growth was not significantly affected at this concentration. At the different concentrations mentioned above, bacteria underwent a lag phase but finally resumed growth (see Fig. S1 in the supplemental material). In other biological systems, NO donors can be used up to mM-range concentrations corresponding to a μM range of available NO (7). NO content in the soil is generally in the nM range, but local spikes at the μM range can be found (L. Bakken, personal communication). Soluble proteins were extracted, and the protein nitration profile was analyzed by Western blotting with antibodies directed against nitrotyrosines (Fig. 1). We noticed that a nitrated protein of 38 kDa was readily detected when cells were exposed to 25 μM SpNO or more. Increasing the concentration of SpNO up to 250 μM allowed us to detect two additional proteins of 46 and 49 kDa (Fig. 1). In order to verify whether these bands correspond to tyrosine-nitrated proteins induced by NO, different controls were performed. First, because SpNO generates NO and spermine in the culture, a control experiment where spermine was added alone was performed, and it showed that proteins were not tyrosine nitrated under this condition (Fig. 1). Second, to test the specificity of the antibody, the Western blot membrane was pretreated with dithionite to reduce nitrotyrosines. Under those conditions, tyrosine-nitrated proteins at 38, 46, and 49 kDa (see Fig. S2 in the supplemental material) were no longer detected, confirming that these proteins are tyrosine nitrated.
FIG 1
It has been described that NO can act at the transcriptional level (13, 23). Hence, the expression of genes encoding proteins of 38, 46, and 49 kDa could be induced by NO. To test this hypothesis, soluble proteins from untreated S. meliloti cells were extracted and subsequently treated with SpNO. The nitration of a 38-kDa protein was observed at all SpNO concentrations, albeit with lower intensity than that observed after treatment of the cell culture with SpNO (Fig. 1). Protein nitration was not observed when only spermine was added to the extract. The nitration of 46- and 49-kDa proteins was not visible, suggesting inappropriate in vitro conditions for nitration. Tyrosine nitration can also be mediated by peroxynitrite (ONOO−) which is produced from NO and O2− (18). Therefore, we tested whether the in vitro tyrosine nitration of proteins was detectable when peroxynitrite was added instead of SpNO to protein extracts from untreated S. meliloti cells. A faint overall increase of the nitrated proteins was detected (Fig. 1). The nitrations observed with SpNO seem more specific than the nitrations detected in the presence of peroxynitrite.
Identification of two flagellin nitropeptides.
In order to identify the proteins corresponding to the 38-, 46-, and 49-kDa bands detected by Western blotting, we prepared protein extracts from S. meliloti cells treated or not with 250 μM SpNO. To enrich these protein extracts in nitroproteins, an immunoprecipitation was performed with antinitrotyrosine antibodies coupled to protein A agarose. Only the 46- and 49-kDa proteins were recovered after immunoprecipitation, as assessed by Western blotting (see Fig. S3 in the supplemental material). During immunoprecipitation, the 38-kDa protein nitrotyrosine residues may not have been accessible to the nitrotyrosine antibody. The tyrosine residues may have been reachable for the nitration when cells were intact but not after extraction of the soluble fraction. The corresponding 46- to 49-kDa band was excised from the Imperial-stained gel and analyzed by mass spectrometry using liquid chromatography-tandem mass spectrometry (LC-MS/MS). A total of 3,010 peptides representing 406 proteins were detected from three biologically independent experiments. Among them, only two were found to be tyrosine nitrated (Fig. 2A, B), namely, AVDVAATGQEVVY78DDGTTK and VDTAY263SGMESAIEVVK. The comparison of these peptide sequences to the S. meliloti 2011 genomic database indicates that they belong to flagellin A (FlaA; SMc03037) or flagellin B (FlaB; SMc03038). These proteins display 93% identity (see Fig. S4 in the supplemental material). Despite their similarities, specific peptides from FlaA and FlaB could be detected in our samples. S. meliloti encodes four flagellins, i.e., FlaA, FlaB, FlaD, and FlaC. Specific peptides belonging to FlaD were also detected, indicating that this protein is present in the sample even though nitrated or nitratable peptides were not detected.
FIG 2
FlaA and FlaB sequences contain 9 tyrosines but only 3 peptides with a tyrosine could be detected in mass spectrometry after trypsic digestion, among which two were detected as nitrated while the third one (VGSASDNAAYWSIATTMR) was recovered only in its nonnitrated form. Hence, either this peptide is indeed not nitrated or is nitrated but hardly detected in its nitrated form. It has to be noticed that FlaA and FlaB are 41-kDa proteins. One hypothesis to explain this discrepancy with the observed size of the proteins could be the presence of varied posttranslational modifications (nitration, phosphorylation, and glycosylation).
In an attempt to quantify the proportion of nitrated Fla proteins and to identify the 38-kDa protein which we failed to enrich by immunoprecipitation (Fig. S3), total protein extracts were separated on an SDS-PAGE gel, and a band corresponding to 35- to 55-kDa proteins was cut and analyzed by mass spectrometry. We failed to detect nitrated peptides reproducibly (data not shown), probably due to a very small amount of nitrotyrosine protein. FlaA and FlaB nitropeptides could not be detected in these experiments; therefore, we could not determine the proportion of nitrated FlaA and FlaB compared with the total FlaA and FlaB peptides recovered.
FlaA and FlaB proteins are both tyrosine nitrated.
To confirm the results obtained with mass spectrometry, we tested whether mutating flaA and/or flaB genes impaired the immunodetection of the 46- and 49-kDa nitroproteins. Deletion mutants of flaA or flaB or both were constructed. The mutant strains were exposed to SpNO as described before. The nitrated proteins of 46 and 49 kDa were still detected in flaA or flaB single-deletion mutants, whereas they were absent in the flaA flaB double mutant (Fig. 2C). This result confirmed that the detected bands correspond to FlaA and FlaB and indicated that both flagellins are nitrated.
Complementation of the double mutant with the plasmid carrying flaA alone (Fig. 2D) restored 46- and 49-kDa nitration which is consistent with the identification of FlaA among the nitroproteins. Nevertheless, nitrations were not recovered when the double mutant was complemented by expressing flaB alone. To determine whether this lack of nitration complementation could be due to a defect in flaB expression, we analyzed the expression of flaA and flaB in the different strains by reverse transcriptase quantitative PCR (RT-qPCR) (see Fig. S5 in the supplemental material). The results showed that transcription of flaB in the flaA flaB double mutant complemented with the plasmid carrying flaB with its native promoter was comparable to flaB expression in the wild-type (WT) strain, while complementation with flaA with its native promoter showed an overexpression of flaA. These results suggest that to get nitration complementation, restoring the WT level of expression of flaB is not sufficient. We could also determine by RT-qPCR that there was no significant transcription induction of flaA and flaB in response to SpNO (Fig. S5).
To confirm the identification of the two nitrotyrosines Y78 and Y263, we mutated both residues by site-directed mutagenesis in flaA. The mutated flagellin was expressed on a plasmid introduced in the double mutant. Nitration at 46 and 49 kDa was still detectable, suggesting that additional tyrosines are nitrated in FlaA (Fig. 2D).
In order to further confirm the identification of FlaA and FlaB, we analyzed the nitration profile of flagellar structure mutants after SpNO treatment. FlaC and FlaD constitute the filament together with FlaA and FlaB. Expression levels of flaA and flaB were not impaired in these mutants, and nitration of FlaA and FlaB was not affected (see Fig. S6 in the supplemental material). Similarly, expression of flaA and flaB was not affected after mutating motC encoding a periplasmic motility protein, and nitration of FlaA and FlaB still occurred. In contrast, expression of flaA and flaB was impaired in mutants of the genes encoding the flagellum hook (FlgE) or the basal body (FlgI, FliG, and FliM). Expression of flagellins was affected in fliM mutants, as described previously (26). Coherently, nitration of FlaA and FlaB was not detectable in flgE, flgI, fliG, and fliM mutants. These data are in agreement with the identification of FlaA and FlaB as nitrated proteins after SpNO treatment.
Analysis of NO effect on flagellar structure and bacterial motility.
We wanted to analyze the effect of NO on functions linked to flagella, as these structures are built up by the polymerization of flagellins. Modification of these proteins could have an impact on flagellar assembly and function. First, we evaluated the effect of NO on flagella by transmission electron microscopy. Flagella were observed after 1 hour of treatment with SpNO or with spermine as a control. Representative images are presented in Fig. 3 (A, B, C). Under all conditions tested, there were cells without flagella and cells with one or multiple flagella (up to five) as well as some broken flagella. It was not possible to accurately assess the proportion of unflagellated cells or the number of flagella per cell, so we could not see an effect of SpNO on these parameters. The overall length of the flagellum was not statistically different between untreated, spermine-, or SpNO-treated samples (Fig. 3D). In consequence, it was not possible to further analyze an effect of tyrosine nitration of the flagellins on flagellar structure.
FIG 3
Next, we choose to analyze bacterial swimming and swarming motilities which are dependent on flagella (27–30). Different concentrations of spermine or SpNO were added to swimming or swarming plates containing 0.3% Bacto agar or 0.6% Noble agar, respectively (see Fig. S7 in the supplemental material). Motility was observed after 24 and 48 h. As expected, the flaA flaB mutant displayed neither swimming nor swarming motility. SpNO negatively affected motility compared with that of the spermine control. This negative effect on swimming motility was significant at 250 μM SpNO at 24 h and at 48 h. At 48 h, it was also possible to see a significant effect for 75 μM SpNO. SpNO impaired swarming motility at all concentrations and time points tested. The motility decrease could in part be due to an effect of NO on cell growth in particular at 24 h, as SpNO increased lag-phase duration in a dose-dependent manner (Fig. S1). Swimming in the presence of SpNO was comparable to that of untreated samples at 48 h, indicating that the SpNO effect on swarming motility is not a mere toxic effect. The stronger impact of NO on swarming motility is consistent with a higher number of flagella under swarming conditions (30). Thus, reduced motility correlated with conditions where higher flagellin tyrosine nitration is observed. However, we cannot discard the possibility that NO could impair bacterial motility through other mechanisms.
Importance of the nitratable tyrosines on flagellin function.
In order to try to link NO effect on motility and flagellin tyrosine nitration, the structure and function of flagellins from flaA flaB mutants complemented with flaA or flaA Y78F Y263F were tested. Mutating flaA and flaB had a major impact on bacterial flagellar structure (Fig. 4A, D). The motility was completely lost in the flaA flaB double mutant (Fig. 5A) as reported (28). Motility was also lost in the flaA single mutant but only partially reduced after mutating flaB (Fig. 5A), suggesting that FlaB is less important than FlaA for the flagellar structure, as already described (31). It was possible to restore flagellar structures (Fig. 4B, D) and some motility of flaA flaB (Fig. 5A) by expressing flaA. To determine the importance of the two identified nitratable tyrosines, we tried to complement flaA flaB by expressing FlaA-bearing mutations on both tyrosines Y78 and Y263 (AFF). We found that it was possible to get short flagellum-like structures (Fig. 4C, D) but it was not possible to restore motility in the complemented mutants (Fig. 5A), and it was not due to a problem of gene expression (Fig. S5). Interestingly, mutating the nitratable tyrosines of flagellins seemed to produce an effect similar to SpNO treatment on bacterial motility.
FIG 4
FIG 5
Expressing FlaA-bearing mutations on both tyrosines Y78 and Y263 (AFF) in the double mutant flaA flaB could overly impair the structuring of the flagella. Moreover, we also do not expect all tyrosines to be affected by nitration upon NO treatment. We therefore decided to test FlaAFF expression in flaA mutants. Complementing the flaA mutant with FlaA restores motility, but complementing by FlaAFF did not restore motility (Fig. 5A). With the goal to identify which of the tyrosines, namely, Y78 or Y263, is essential for flagellar assembly and movement, we complemented the flaA mutant with FlaA mutated either in Y78 or Y263 (Fig. 5B). Expression of flaA Y263F restored motility to the same level as the expression of flaA, but expression of flaA Y78F was not as efficient, indicating that Y78 is essential to flagellar structure. Expression of flaA- and flaA-bearing mutations Y78F or Y263F were equivalent in the complemented mutants (see Fig. S8 in the supplemental material). The importance of Y78 in flagellar function in motility suggests that modification of this tyrosine by nitration could have a major effect on flagellar structure and function.
DISCUSSION
To understand how rhizobia respond to nitrosative stress in soil or in the plant environment during symbiosis, we searched for S. meliloti proteins targeted by NO and particularly tyrosine-nitrated proteins. Here, we identified tyrosine-nitrated proteins in S. meliloti in vivo. We found three tyrosine-nitrated proteins in S. meliloti in response to the NO donor SpNO by Western blotting with an antibody directed against nitrotyrosines. Two nitropeptides belonging to flagellin A and B were identified by mass spectrometry. Mutant analysis and complementation experiments confirmed the identification. We used a non-a priori approach to evidence tyrosine-nitrated proteins. Instead, in previous studies in bacteria, only candidate proteins have been analyzed for their in vitro nitration after purification (19–21) or to investigate posttranslational modifications by two-dimensional (2D) gel and mass spectrometry to characterize the nitrotyrosine residue (22). Detection of the nitrated flagellins was possible only after enrichment by immunoprecipitation. Despite that enrichment, many nonnitrated peptides were detected under our conditions in mass spectrometry, suggesting a low specificity of the immunoprecipitation even though it proved efficient to recover nitropeptides from bacterial cells without an a priori approach. Identification of tyrosine nitration remains challenging to date due to the low yield of nitration in vivo, low numbers of proteins undergoing nitration, and low numbers of nitratable tyrosines (15–17). Indeed, we could detect only three nitroproteins in Western blot analysis. Technical difficulties like enriching the nitrated proteins or peptides and detecting the nitrated peptides in mass spectrometry exacerbate the problem. Finding nitrated flagellins could be due to the fact that flagellins are abundant bacterial proteins, and it does not exclude that other proteins are nitrated, like the 38 kDa that could not be identified. Similarly, enzymes from the tricarboxylic acid cycle have been identified while studying S-nitrosylation in Medicago truncatula and S. meliloti (32); probably, they are among the most abundant proteins in the cells.
After identifying tyrosine-nitrated flagellins, we investigated a possible impact of NO on flagellar structure and function. Indeed, the structures of some proteins have been shown to be modified by tyrosine nitration; such is the case, for example, of human lactoferrin which loses its antibacterial properties (33) or mammalian cytochrome c which displays a change of affinity for cardiolipin (34). In addition, in vitro tyrosine nitration of flagellin from Proteus mirabilis created a derivative unable to aggregate, as shown by studying its conformational transitions and structural properties (35). Searching for structural modifications on flagella by transmission electron microscopy, we could not show that flagellum length was impaired by NO. Under our in vivo conditions, the proportion of proteins that were tyrosine nitrated or the extent of tyrosine residues nitrated per protein could be insufficient to detect any modification at the flagellar structure level. Investigating a possible role of NO in flagellar function, we showed that swimming and even more swarming are affected by SpNO. Because a growth defect in the presence of SpNO could partly explain these phenotypes, we could not conclude that these effects are linked to flagellin nitration. Development of single-cell tracking to follow SpNO effects on S. meliloti is worth considering in the future for analyzing the motility in real time. In parallel, we showed that mutating the nitratable Y78 in flagellin impaired bacterial motility, suggesting that modification of that residue, such as nitration, could affect flagellar functionality. Mutating Y263 had less of an impact. Correlatively, Y78 is found to be more conserved among rhizobia than Y263 (B. Gourion, personal communication). Y263 was not even conserved among different S. meliloti strains. Y263 is part of the variable region of flagellin, while Y78 is found in the conserved region of flagellin that is critical to the flagellar assembly (36).
The presence of NO has been evidenced at all steps of various symbiotic interactions and in particular in the symbiosis between S. meliloti and M. truncatula (11). We showed that the bacteria actively contributed to the protection of glutamine synthetase, a key enzyme in nitrogen assimilation, toward its tyrosine nitration which inactivates the enzyme (24, 37). The results presented here show that S. meliloti proteins can be affected by NO-mediated posttranslational modification. Identifying flagellins as nitroproteins presents new questions on the role of NO and flagellins in symbiosis. Some published data show that flagellin-dependent motility is dispensable for nodulation but could be an advantage in efficiency of nodulation and in competition for nodule occupancy (27, 38–41). During symbiosis in determinate or indeterminate nodules, flagellum-related genes are downregulated (42, 43), but unexpectedly, the flagellar regulon is then upregulated in the nitrogen-fixing zone (zone III) in indeterminate nodules (44), suggesting putative different roles of flagella during symbiosis. Similarly in pathogenesis, nonmotile Ralstonia bacteria still express flagellin fliC in the plant host (45).
It was shown that plants respond to flagellin by the production of NO upon bacterial invasion (46) and that protein tyrosine nitrations were linked to this NO production (47). Flagellin from rhizobia are divergent and do not induce defense reactions (48–50), but the NO response to rhizobium flagellin was not tested. It would be interesting to investigate whether rhizobium flagellin actually triggers NO production at the beginning of symbiosis. NO via flagellin tyrosine nitration could help or hinder the nodule initiation and development. Indeed, it has been shown that NO was present at the earliest step of the interaction and especially along the infection thread (9). Even though the amount of NO has not been estimated, we could speculate that NO has a role in flagellin nitration at this stage to inactivate flagella. Finally, posttranslational modifications in bacteria have been correlated with changes in lifestyle (51), and we hypothesize that NO could play a role in the transition from saprophytic life to symbiotic life. In plant-pathogen interactions, other posttranslational modifications of flagellins have been described, and interestingly, they can modify the outcome of the process. Plants produce glycosidases that contribute to the release of flagellin immunopeptides, and Pseudomonas bacteria can evade by using modified glycans (52). Unglycosylated flagella showed an increase in bacterial virulence in Xanthomonas bacteria, while flagellar assembly and bacterial motility were not affected in most mutants (53). Phosphorylation of flagellin FliC in Pseudomonas bacteria did not affect its motility but did affect the type 2 secretion system and biofilm formation (54).
In conclusion, our results provide novel insights into a possible response to NO of rhizobia via modification of flagellins and open new perspectives toward a possible role of NO in plant-microbe interactions through bacterial flagellin nitration.
MATERIALS AND METHODS
Bacterial strains and growth conditions.
S. meliloti cells (Table 1) were grown at 28°C in plates with Luria-Bertani medium supplemented with 2.5 mM MgSO4 and 2.5 mM CaCl2 (LBMC) and then in liquid cultures for one night with the appropriate antibiotics. Cells were then grown to exponential phase (optical density at 600 nm [OD600], 0.2 to 0.3) in Vincent minimal media (VMM) as previously described (13). Cells were then treated with the NO donor spermine NONOate (SpNO; reference no. [ref.] 82150; Cayman) at concentrations of 25, 75, or 250 μM (from a stock solution of 100 mM in 0.01 M NaOH) for 1 h, at 28°C, under agitation. Spermine (25, 75, or 250 μM; from a stock solution of 100 mM in 0.01 M NaOH; Sigma) was also used as a control. For motility tests, the cells were grown overnight at 28°C in tryptone yeast (TY) medium (5 g·liter−1 tryptone, 3 g·liter−1 yeast extract, and 15 g·liter−1 agar) with 6 mM CaCl2. Antibiotics were added at the following concentrations: 100 μg/ml streptomycin (Sm), 100 μg/ml neomycin (Nm), and 10 μg/ml tetracycline (Tc).
TABLE 1
| Strain or plasmid | Description | Reference or source |
|---|---|---|
| Strains | ||
| Sinorhizobium meliloti | ||
| GMI11495 | Wild-type strain (Smr), Rm2011 background | 60 |
| CBT2352 | GMI11495 ΔflaA, Smr | This work |
| CBT2363 | GMI11495 ΔflaA+pFAJ-flaA, Smr, Tcr | This work |
| CBT2365 | GMI11495 ΔflaA+pFAJ-flaA Y78F Y263F, Smr, Tcr | This work |
| CBT2367 | GMI11495 ΔflaA+pFAJ, Smr, Tcr | This work |
| CBT2369 | GMI11495 ΔflaB, Smr | This work |
| CBT2429 | GMI11495 ΔflaA ΔflaB, Smr | This work |
| CBT2443 | GMI11495 ΔflaA ΔflaB+pFAJ, Smr, Tcr | This work |
| CBT2444 | GMI11495 ΔflaA ΔflaB+pFAJ-flaA, Smr, Tcr | This work |
| CBT2445 | GMI11495 ΔflaA ΔflaB+pFAJ-flaB, Smr, Tcr | This work |
| CBT2446 | GMI11495 ΔflaA ΔflaB+pFAJ-flaAY78F Y263F, Smr, Tcr | This work |
| CBT2512 | GMI11495 ΔflaA+pFAJ-flaAY78F, Smr, Tcr | This work |
| CBT2513 | GMI11495 ΔflaA+pFAJ-flaAY263F, Smr, Tcr | This work |
| 2011mTn5STM.4.03.B12 | GMI11495 SMc03019(fliG)::Tn5, Smr, Nmr | 60 |
| 2011mTn5STM.3.11.D06 | GMI11495 SMc03032(flgI)::Tn5, Smr, Nmr | 60 |
| 2011mTn5STM.1.04.C04 | GMI11495 SMc03040(flaC)::Tn5, Smr, Nmr | 60 |
| 2011mTn5STM.3.01.B08 | GMI11495 SMc03043(motC)::Tn5, Smr, Nmr | 60 |
| 2011mTn5STM.4.04.B01 | GMI11495 SMc03047(flgE)::Tn5, Smr, Nmr | 60 |
| 2011mTn5STM.4.22.C08 | GMI11495 SMc03039(flaD)::Tn5, Smr, Nmr | 60 |
| 2011mTn5STM.2.01.H08 | GMI11495 SMc03021(fliM)::Tn5, Smr, Nmr | 60 |
| Escherichia coli | ||
| DH5α | supE44Δlac*U169 F80ΔlacZ ΔM15 hsdR17 rec*A1 endA1 gyr*A96 thi-1 relA1 | Invitrogen |
| Plasmids | ||
| pGEM-T | Cloning vector (Apr) | Promega |
| pJQ200mp19 | Gene replacement vector (Gmr) | 61 |
| pFAJ1702 | Stable RK2-derived cloning vector (Apr Tcr) | 62 |
S. meliloti growth sensitivity to SpNO.
Cells grown at 28°C to exponential phase (OD600, 0.2) in Vincent minimal media (VMM) were distributed in 96-well plates (200 μl). Cells were left untreated or were treated with spermine or SpNO at 25, 75, or 250 μM. OD600 was measured at 28°C every 400 s during 22 h in a multiwell plate reader (Fluostar Omega, BMG Labtech). Five technical repeats of each condition were performed on the same plate. The experiment was performed three times independently.
Deletion mutants and plasmid constructs.
To construct the deletion mutants flaA, flaB, and flaA flaB, DNA fragments upstream (flaA [515 bp] and flaB [435 bp]) and downstream (flaA [506 bp] and flaB [513 bp]) of the region of interest were cloned in pGEM-T with SacI/BamHI and BamHI/SalI restriction sites, respectively (Table 2). The cloned fragments were verified by DNA sequencing. They were assembled as SacI-BamHI and BamHI-SalI fragments into SacI-SalI-cut pJQ200mp19. Plasmids were introduced in S. meliloti by electroporation, and then deletion mutants were isolated as described (37). The deletion was verified by PCR and sequencing. To complement these deleted strains, flaA (1,185 bp) and flaB (1,185 bp) were cloned with their promoter regions (488 bp and 342 bp, respectively) into the pGEM-T vector with HindIII/BamHI restriction sites (Table 2). The cloned fragments were verified by DNA sequencing and then integrated as HindIII-BamHI fragments in HindIII-BamHI-cut pFAJ1702. Plasmids were introduced in S. meliloti by electroporation. For site-directed mutagenesis, overlapping primers were designed (Table 2) to introduce the desired sequences to change tyrosine into phenylalanine at positions 78 and 263. Replacing tyrosine by phenylalanine has already been described by studying tyrosine nitration (24). The pGEM-T plasmids with cloned flaA was used as the template for amplification. The sample was treated with DpnI to remove the methylated template prior to E. coli transformation. The mutated flaA sequences were verified by sequencing. To get flaA with Y78F Y263F mutations, the site-directed mutagenesis was performed on flaA with the Y78F mutation plasmid.
TABLE 2
| Target | Forward primer sequence (5′–3′) | Reverse primer sequence (5′–3′) |
|---|---|---|
| Deletion mutant construct | ||
| flaA upstream | AGGGAGCTCCCTTCATGATCTCGGAGATCCGC | AATGGATCCGCTCGTCATATTCGTTGTCCCTAC |
| flaA downstream | GAAGGATCCTTCCGCTAAGAAGACATGCAATGG | GTAGTCGACTACGCGGCGTTGTCGGCGGCCTG |
| flaB upstream | AGGGAGCTCCTCGCCATCCAGGCTCTTTCG | AATGGATCCGCTCGTCATGGTTTAGTG |
| flaB downstream | GAAGGATCCTTCCGCTAATCCGACACGGAAAG | ACAGTCGACACCTTGGCCGCACCGAGACC |
| Complementation mutants | ||
| flaA promoter and flaA | AGGAAGCTTCCTTCATGATCTCGGAGATCCGC | TAAGGATCCTTAGCGGAAGAGCGAAAGGACGTTCTGG |
| flaB promoter and flaB | TTCAAGCTTGAAGACATGCAATGGCGGACG | TAAGGATCCTTAGCGGAAGAGCGTAAGGACGTTTTCC |
| Site-directed mutagenesis | ||
| flaA or flaB Y78F mutation | GCCAAGGTCGACACCGCCTTCTCCGGTATGGAATCGGC | GCCGATTCCATACCGGAGAAGGCGGTGTCGACCTTGGC |
| flaA or flaB Y263F mutation | CGGGTCAGGAAGTCGTCTTCGACGACGGCACGACGAAATG | CATTTCGTCGTGCCGTCGTCGAAGACGACTTCCTGACCCG |
| RT-qPCR | ||
| flaA | CAGACTCAGCAGCAGCTC | AGAGCGAAAGGACGTTCTG |
| flaB | CGATCAGCTCAAGGACCAG | CGCTCTTGGTGACGGGAC |
| Smb21134 (reference gene) | GACGGAAGCGGAGGCGATGG | CGCCAGCCGTGCGAGTTTCT |
Protein extraction and immunoprecipitation.
The cells were collected by centrifugation (20,000 × g, 10 min, and 4°C). For protein extraction, Bugbuster (Merck) reconstituted with Lysonase (Merck) and with 1 mM phenylmethylsulfonyl fluoride (PMSF) was added to the cell pellet (75 μl per 2-ml cell pellet). The samples were incubated for at least 30 min under gentle agitation (room temperature [RT]) and then centrifuged (20,000 × g, 20 min, and 4°C). The supernatant was collected, and proteins were quantified by the Bradford assay. To perform nitration on the extracted proteins, SpNO (25, 75 or 250 μM) or peroxynitrite (5 μM; ref. 81565; Cayman) was incubated for 1 h at RT. Spermine (250 μM) was also used as a control. For immunoprecipitation, the Bugbuster supernatant was diluted 1:3 in Tris-buffered salt (TBS; 20 mM Tris [pH 7.5] and 150 mM NaCl) and filtered on an Ultracell-4 30K device. TBS was added once more and filtered. The retentate was brought to a concentration of 1 mg/ml. Nitrotyrosine sorbent (25 μl; ref. 389549; Cayman) was added to a 500-μl sample and agitated for 2 h at RT. The beads were collected by centrifugation (1,000 × g, 30 sec) and washed three times in TBS. The elution was performed with 50 μl Laemmli buffer (55) for 5 min at 95°C, and the supernatant was collected after centrifugation. Experiments were repeated three times.
Western blot analysis.
Proteins (15 to 20 μg) were separated by 10% (wt/vol) SDS-PAGE electrophoresis and then transferred onto 0.2-μm polyvinylidene difluoride (PVDF) membranes using the Transblot Turbo system (Bio-Rad). The membrane was blocked in TBS-0.1% Tween with 3% bovine serum albumin (BSA) for 1 h at RT. The nitrotyrosine monoclonal antibody (ref. 189542; Cayman) was diluted 1:250 in TBS-Tween with 1% BSA and incubated with the membrane overnight at 4°C. After the membrane was washed with TBS-Tween, an anti-mouse horseradish peroxidase (HRP)-conjugated antibody was added (dilution of 1:10,000 in TBS-Tween with 1% BSA) for 1 h at RT. The labeled proteins were detected by chemiluminescence using the Clarity Western ECL substrate (Bio-Rad) and ChemiDoc Touch imaging instrument (Bio-Rad). The 40- to 55-kDa portion of the membrane stained with Ponceau was used as a loading control. Experiments were repeated at least three times.
To test the specificity of the primary antibody, the membrane was treated after protein transfer with a dithionite solution (10 mM dithionite sodium and 50 mM pyridine acetate [pH 5] for 1 h) to reduce nitrotyrosines. Membranes were washed with H2O and TBS-Tween and then processed as described above.
Mass spectrometry.
Immunoprecipitated proteins were separated on 10% SDS-PAGE in a TGX gel (Bio-Rad). The gel was stained with Imperial stain (Thermo Scientific). A band was cut at around 46 to 49 kDa, digested by trypsin, and analyzed by mass spectrometry (conditions are detailed in supplemental material). Experiments were repeated three times. Mass spectrometry was performed by coupling an Orbitrap Fusion Lumos Tribrid system (Thermo Fisher Scientific) with an UltiMate 3000 RSLCnano system (Thermo Fisher Scientific) at the Plateforme d'Analyse Protomique de Paris Sud Ouest (PAPPSO) platform, Jouy-en-Josas, France (http://pappso.inra.fr/). The S. meliloti 2011 database (GMI11495-Rm2011G; https://bbric.toulouse.inra.fr/reference/ws/reference/docid/6288a62ddf483b63eb630ebaf4854a9b.html) was searched by the X!TandemPipeline (open source software developed by PAPPSO; version 3.3.5) (56).
RNA extraction and RT-qPCR.
S. meliloti cells (5 ml) were grown in VMM to exponential phase (OD600, 0.2 to 0.3) and treated or not with SpNO for 1 h at 28°C. Cells were collected by centrifugation (10 min, 3,000 × g, and 4°C). RNA extraction and RT-qPCR proceeded as described previously (57). Expression of flaA and flaB was measured compared to a genomic DNA scale and normalized using Smb21134 as a reference (58). The primers used are presented in Table 2. Experiments were repeated at least three times.
Transmission electron microscopy.
S. meliloti cells (5 ml) were grown in VMM to exponential phase (OD600, 0.2 to 0.3) and then were untreated or treated with 250 μM spermine or SpNO for 1 h at 28°C. Cells were collected by centrifugation for 5 min at 3,000 × g and 4°C. The supernatant was removed, and 1 ml of paraformaldehyde (4% in TBS) was added to the cell pellet. The samples were adsorbed on 400 mesh copper-coated grids and negatively stained with 1% (wt/vol) aqueous uranyl acetate. The grids were observed with a transmission electron microscope (HT-7700 120 kV; Hitachi). Pictures were taken when cells presenting flagella were noticed. Results from 15 images of flagellated cells from each condition per experiment and from five independent experiments were collected. Flagellum length was measured with ImageJ software.
Bacterial motility assay.
The bacterial motility assays were adapted from Peláez-Vico et al. (59). Cells were grown overnight at 28°C in TY medium with CaCl2 to reach an OD600 between 1 and 1.2 the next morning. For the swimming motility assay, 3 μl of cells at an OD600 of 1 were spotted in triplicate on swimming plates (VMM and 0.3% Bacto agar). For the swarming motility assay, the cells were washed twice in VMM and then resuspended in 1/10th VMM and adjusted to an OD600 of 10. Cells (4 μl) were spotted in triplicate on swarming plates (VMM and 0.6% Noble agar). Spermine or SpNO at 25, 75, or 250 μM was added when the agar plates were poured. Plates were incubated for 48 h at 28°C. The plates were scanned at 24 and 48 h. The growth area was measured by ImageJ software. Experiments were repeated at least three times.
ACKNOWLEDGMENTS
P.B. was the recipient of a Contrat Jeune Scientifique INRA. This work was supported by the French Laboratory of Excellence project “TULIP” (ANR-10-LABX-41) and the “Agence Nationale de la Recherche” (STAYPINK-ANR-15-CE20-0005). The funders had no role in the study design, data collection and interpretation, or the decision to submit the work for publication.
We thank C. Pichereaux (FR3450, IPBS, Université de Toulouse, France) for initial MS analyses. We thank B. Happel and A. Becker (University of Marburg, Germany) for providing S. meliloti Tn5 mutants. We thank the CMEAB team (Université de Toulouse, France) for assistance in transmission electron microscopy, particularly D. Goudounèche. We thank M. F. Jardinaud for critical statistical analysis of the data.
A.-C.C., C.H., and C.P. designed and performed the experiments. P.B. performed initial experiments. A.-C.C., C.B., and E.M. analyzed the corresponding results and wrote the paper.
We declare that we have no conflicts of interest with the contents of this article.
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Information & Contributors
Information
Published In
Applied and Environmental Microbiology
Volume 87 • Number 1 • 17 December 2020
eLocator: e02210-20
Editor: Gladys Alexandre, University of Tennessee at Knoxville
PubMed: 33067191
Copyright
© 2020 American Society for Microbiology. All Rights Reserved.
History
Received: 8 September 2020
Accepted: 13 October 2020
Published online: 17 December 2020
Keywords
Contributors
Editor
Gladys Alexandre
Editor
University of Tennessee at Knoxville
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2020.
Tyrosine Nitration of Flagellins: a Response of Sinorhizobium meliloti to Nitrosative Stress. Appl Environ Microbiol
87:e02210-20.
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