A Novel Small RNA, DsrO, in Deinococcus radiodurans Promotes Methionine Sulfoxide Reductase (msrA) Expression for Oxidative Stress Adaptation

Yun Chen; Mingming Zhao; Mengli Lv; Min Lin; Jin Wang; Kaijing Zuo

doi:10.1128/aem.00038-22

. 2022 May 16;88(11):e00038-22. doi: 10.1128/aem.00038-22

A Novel Small RNA, DsrO, in Deinococcus radiodurans Promotes Methionine Sulfoxide Reductase (msrA) Expression for Oxidative Stress Adaptation

Yun Chen ^a, Mingming Zhao ^b, Mengli Lv ^a, Min Lin ^b, Jin Wang ^b,^✉, Kaijing Zuo ^a,^✉

Editor: Isaac Cann^c

PMCID: PMC9195949 PMID: 35575549

ABSTRACT

Reactive oxygen species (ROS) can cause destructive damage to biological macromolecules and protein dysfunction in bacteria. Methionine sulfoxide reductase (Msr) with redox-active Cys and/or seleno-cysteine (Sec) residues can restore physiological functions of the proteome, which is essential for oxidative stress tolerance of the extremophile Deinococcus radiodurans. However, the underlying mechanism regulating MsrA enzyme activity in D. radiodurans under oxidative stress has remained elusive. Here, we identified the function of MsrA in response to oxidative stress. msrA expression in D. radiodurans was significantly upregulated under oxidative stress. The msrA mutant showed a deficiency in antioxidative capacity and an increased level of dabsyl-Met-S-SO, indicating increased sensitivity to oxidative stress. Moreover, msrA mRNA was posttranscriptionally regulated by a small RNA, DsrO. Analysis of the molecular interaction between DsrO and msrA mRNA demonstrated that DsrO increased the half-life of msrA mRNA and then upregulated MsrA enzyme activity under oxidative stress compared to the wild type. msrA expression was also transcriptionally regulated by the DNA-repairing regulator DrRRA, providing a connection for further analysis of protein restoration during DNA repair. Overall, our results provide direct evidence that DsrO and DrRRA regulate msrA expression at two levels to stabilize msrA mRNA and increase MsrA protein levels, revealing the protective roles of DsrO signaling in D. radiodurans against oxidative stress.

IMPORTANCE The repair of oxidized proteins is an indispensable function allowing the extremophile D. radiodurans to grow in adverse environments. Msr proteins and various oxidoreductases can reduce oxidized Cys and Met amino acid residues of damaged proteins to recover protein function. Consequently, it is important to investigate the molecular mechanism maintaining the high reducing activity of MsrA protein in D. radiodurans during stresses. Here, we showed the protective roles of an sRNA, DsrO, in D. radiodurans against oxidative stress. DsrO interacts with msrA mRNA to improve msrA mRNA stability, and this increases the amount of MsrA protein. In addition, we also showed that DrRRA transcriptionally regulated msrA gene expression. Due to the importance of DrRRA in regulating DNA repair, this study provides a clue for further analysis of MsrA activity during DNA repair. This study indicates that protecting proteins from oxidation is an effective strategy for extremophiles to adapt to stress conditions.

KEYWORDS: Deinococcus radiodurans, oxidative stress, MsrA, DsrO, small RNA, posttranscriptional regulation

INTRODUCTION

Reactive oxygen species (ROS) accumulation is the indirect consequence of failed or inefficient electron-transfer processes (1). Nearly all organisms produce ROS when they are exposed to UV light, gamma and X-rays, and abiotic and biotic stresses (2). ROS such as H₂O₂ can produce hydroxyl radicals that oxidize both bases (adenine and guanine) and ribose moieties of DNA. Damaged guanine allows its electrons to enter into nearby oxidized bases, producing a wide variety of DNA lesions (3, 4). The polyunsaturated fatty acids of cell membranes are easily peroxidated in some bacteria, producing the cytotoxic compound 4-hydroxy-2-nonenal (5). In addition, ROS have high oxidation reactivity toward sulfur-containing amino acids and metal-containing cofactor sites in proteins, causing irreversible inactivation of many different proteins (6). The sulfur atom in the thioether side chain of Met can be oxidized to sulfoxide, converting Met into methionine sulfoxide (Met-SO). Met-SO residues can be irreversibly further oxidized to methionine sulfone (Met-O) (7). Overall, oxidative damage caused by ROS has devastating effects on DNA, lipid structure, and protein activity in most organisms.

Deinococcus radiodurans is an extremophile with an extremely high tolerance to gamma radiation, UV irradiation, desiccation, and other oxidative stresses (8). To cope with these stressors, D. radiodurans has evolved efficient antioxidative ROS scavenging strategies. D. radiodurans has a relatively low number of respiratory chain enzymes and induces the glyoxylate bypass of the tricarboxylic acid cycle to reduce endogenous ROS production (9). The antioxidant compound deinoxanthin provides D. radiodurans with a distinctive pale brown color and endows it with high ROS-elimination capacity together with manganese (10). With the help of the DNA recombination repair system (RecA, PprA, and other recombination regulators), D. radiodurans displays a remarkable ability to reverse DNA damage and reconstitute the genome (11). D. radiodurans is also well equipped with additional ROS scavenging enzymes (e.g., catalase and superoxide dismutase) (12). These different strategies work effectively to restore damaged D. radiodurans cells to a normal physiological state. Although the molecular mechanism of DNA repair has been described in detail (13 –15), the details of the regulatory mechanism involved in recovering ROS-oxidized protein function in D. radiodurans are unclear.

By comparing key enzymes related to amino acid biosynthesis among different bacterial species using Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis, enzymes in the methionine biosynthesis pathway were found to be incomplete in D. radiodurans (16). Culture experiments in media without methionine confirmed that D. radiodurans CGMC1.3828T requires exogenous methionine for growth (16). Accumulating evidence indicates that the level of protein oxidation is negatively correlated with the survival of irradiated bacteria (17, 18). Protein oxidation is likely an important cause rather than a consequence of radiation toxicity; thus, the ability to protect proteins against oxidation to a large extent distinguishes radiation-resistant from radiation-sensitive Deinococcus species (18 –20). Because Met residues are particularly sensitive to oxidative damage, the limited Met resources in D. radiodurans likely contribute to a stronger ability to rescue oxidized proteins compared with sensitive bacteria (21).

Methionine sulfoxide reductase (Msr) is a key enzyme in reducing oxidized Met residues in proteins to restore their function (22). Bacterial mutants for msrA and/or msrB display ROS-sensitive phenotypes. The two forms of Msr that reduce l-Met-S-(O) are MsrA and MsrB; MsrA functions on both peptide-bound l-Met-S-(O) and free l-Met-S-(O) (23, 24). Msr proteins are evolutionarily conserved and function similarly in response to ROS exposure. Msr expression is controlled by alternative sigma transcription factors in different bacterial species (25, 26). D. radiodurans contains two types of Msr proteins, MsrA and MsrB. MsrB is less efficient due to its higher km-MetSO values and slower turnover in Trx/TrxR-based assays (27, 28). It is therefore likely that MsrA proteins in D. radiodurans are the primary protein reducer under oxidative stress, although the regulatory mechanism of msrA is unknown. In this study, we found that a small RNA (sRNA), DsrO, probably acting in concert with the transcription factor DrRRA, regulates msrA expression for protein translation in D. radiodurans exposed to oxidative stress.

RESULTS

msrA expression in D. radiodurans is strongly induced by oxidative stress.

Methionine sulfoxide reductases (Msr proteins) repair oxidized methionine residues of damaged proteins and are found in nearly all organisms (29). Bacterial mutants lacking Msr proteins are often sensitive to oxidative stress; in pathogenic bacteria, the mutants exhibit decreased virulence via reduced survival probability in host cells (29). To analyze the function of MsrA in D. radiodurans during oxidative stress, we cultured D. radiodurans R1 cells in growth medium with different concentrations of hydrogen peroxide (H₂O₂) (Fig. 1). The real-time quantitative PCR (qRT-PCR) results showed that msrA mRNA expression was significantly upregulated by H₂O₂ exposure, with the highest expression (13-fold higher than that of the untreated control) in medium containing 60 mM H₂O₂ (Fig. 1A). However, when H₂O₂ concentrations were higher than 80 mM, msrA transcript levels were unchanged compared to untreated cells. Further analysis of msrA temporal expression in D. radiodurans R1 treated with 80 mM H₂O₂ indicated that the expression of msrA reached the maximum level after 10 min (Fig. 1B). msrA expression was also induced by exogenous treatments with UV light (for 11 min), heat (48°C for 5 h), or cold (4°C for 5 h) stress, suggesting that msrA responds to a variety of different stresses (see Fig. S1 in the supplemental material).

FIG 1 — *msrA* expression is strongly induced by oxidative stress in *D. radiodurans*. (A and C) *msrA* transcriptional level (A) and protein level (C) in *D. radiodurans* treated with different concentrations of H₂O₂ for 30 min. (B and D) *msrA* transcriptional level (B) and protein level (D) in *D. radiodurans* treated with 80 mM H₂O₂ for different times. The *msrA* transcriptional level was analyzed by qRT-PCR with 16S rRNA as the reference gene. GroEL was used as the internal control for Western blotting. Asterisks indicate statistically significant differences of the value compared to that of untreated cells (one-way analysis of variance [ANOVA], Dunnett’s multiple-comparison test; ***, P ≤ 0.001; *, P ≤ 0.05). The experiments were performed at least three times, and the data are presented as means ± standard error of the mean (SEM).

Maintenance of increased MsrA protein synthesis levels over time is critical for the functions of antioxidative enzyme activity. We thus investigated whether MsrA protein levels in D. radiodurans R1 correspond to changes in msrA gene expression levels under H₂O₂ treatment. Western blot results indicated that the highest level of MsrA occurred during exposure to 80 mM H₂O₂. Consistent with the gene expression results, there were no significant changes compared to the control group when the H₂O₂ concentration was higher than 80 mM (Fig. 1C). MsrA protein synthesis in D. radiodurans treated with 80 mM H₂O₂ continuously increased between 0 and 30 min compared with the control group and then obviously declined (Fig. 1D). These results indicate that the levels of both msrA mRNA and the MsrA protein can be quickly upregulated in response to oxidative treatments. In bacteria, mRNA levels are often tightly correlated with the levels of the corresponding protein translation over time (30). Given that the highest level of MsrA protein occurred after the peak of msrA mRNA levels, we hypothesized that there is a molecular mechanism of maintaining msrA mRNA levels in D. radiodurans to increase protein translation under oxidative stress.

Deletion of the msrA gene reduces the total antioxidant capacity and tolerance to H₂O₂ of D. radiodurans.

To characterize the physiological function of MsrA in D. radiodurans, we first analyzed the growth phenotypes of the wild-type strain (WT), mock strain (ΔmsrA-pRADZ3), msrA knockout strain (ΔmsrA), and complementation strain under oxidative stress (Fig. 2). Under normal growth conditions, there was no significant difference in growth between any of the tested strains. However, under oxidative stress conditions (80 mM H₂O₂ for 30 min), the viability of ΔmsrA decreased significantly compared to WT, whereas the complementation strain had a similar viability to the WT (Fig. 2A). There were no significant differences in growth among WT, ΔmsrA, and complementary strains in response to UV light, heat, or cold stress (Fig. S2), indicating that msrA may not function under these stress treatments.

FIG 2 — The deletion of the *msrA* gene in *D. radiodurans* decreases its tolerance to oxidative stress. (A) H₂O₂ sensitivity assays of different *D. radiodurans* strains. Spotted agar plates after H₂O₂ treatment and serial dilution. (Left) 0 mM H₂O₂; (right) 80 mM H₂O₂ for 30 min; WT, wild-type strain, Δ*msrA*, *msrA*-deleted mutant; *msrA-pRADZ3*, *msrA* mutant transformed with *pRADZ3* empty plasmid; *msrA-com*, *msrA* mutant supplemented with the *msrA* gene. (B to D) Total antioxidant capacity (B), ROS level (C), and intracellular carbonylation level (D) of different strains after H₂O₂ treatment. The total level of various antioxidant macromolecules, antioxidant small molecules, and enzymes in a system reflects the total antioxidant capacity of the bacterial cells. Rosup is a reactive oxygen positive-control reagent. Asterisks indicate a statistically significant difference in the value compared to that of untreated cells (one-way ANOVA, Dunnett’s multiple-comparison test; ***, P ≤ 0.001; *, P ≤ 0.05). Experiments were performed at least three times, and the data are presented as the means ± SEM.

ROS accumulation has been suggested to be the main cause of oxidative damage, leading to decreased bacterial growth. Carbonylated protein levels can serve as an indicator of an oxidative cellular status; dysfunctional carbonylated proteins are more prone to irreversible oxidation of other amino acid residues or to the formation of protein aggregates. We therefore compared the total antioxidant capacity, ROS content, and carbonylation level in the WT, msrA-pRADZ3, ΔmsrA, and complementation strains under oxidative stress. As expected, deletion of the msrA gene significantly reduced intracellular antioxidant capacity (Fig. 2B). In the medium with 80 mM H₂O_2, ROS levels rose sharply in ΔmsrA and ΔmsrA-pRADZ3 compared to the WT, supporting that MsrA acts as an oxidant scavenger under oxidative stress (31) (Fig. 2C), and in the 80 mM H₂O₂ treatment, msrA mutants had twice the level of carbonylated proteins as the WT (Fig. 2D). These data further support that MsrA is necessary for normal antioxidant capacity in D. radiodurans.

DsrO is associated with oxidative stress, and its mutation reduces the tolerance of D. radiodurans to H₂O₂.

Given that the peak levels of msrA mRNA and MsrA protein were staggered, we speculated that there is a factor such as a small RNA (sRNA) in D. radiodurans that posttranscriptionally regulates msrA. We thus used an sRNA target prediction program to screen sRNAs potentially binding to msrA mRNA using D. radiodurans genomic and transcriptomic data (RNAalifold, http://rna.tbi.univie.ac.at/cgi-bin/RNAWebSuite/RNAalifold.cgi) (32). Among the 23 sRNAs associated with the oxidative response, Dsr19 was predicted to be most likely to bind to the stem-loop region of msrA mRNA through a 12-nucleotide (nt) complementary fragment (5′-UUUGUUUGCUGG-3′) (Fig. 3A and B). To clearly show the characteristics of Dsr19 during oxidative stress, we renamed Dsr19 DsrO (Deinococcus radiodurans small RNA related to oxidation).

FIG 3 — A noncoding RNA, DsrO, directly binds to the stem-loop structure of *msrA* mRNA. (A) Predicted binding pattern between *msrA* mRNA and DsrO based on the bioinformatics analysis. (B) Affinity coefficient between DsrO and *msrA* mRNA detected by the microscale thermophoresis (MST) method. (C to F) The transcriptional level of DsrO in response to the treatments of 80 mM H₂O₂ (C), UV radiation (D), heat (48°C) (E), and cold (4°C) (F) in *D. radiodurans*. Asterisks indicate statistically significant differences compared to untreated cells (one-way ANOVA, Dunnett’s multiple-comparison test; ***, P ≤ 0.001; *, P ≤ 0.05). Experiments were performed at least three times, and data are presented as means ± SEM.

To determine whether DsrO and msrA may interact, we characterized the expression patterns of both genes under stress treatments using qRT-PCR. DsrO expression levels were quickly and significantly enhanced with increasing H₂O₂ concentration in the medium (≤80 mM). In the WT, DsrO was upregulated for up to 50 min in the 80 mM H₂O₂ treatment group (Fig. 3C). In contrast to msrA, DsrO could quickly respond to UV light (Fig. 3D), heat (Fig. 3E), and cold (Fig. 3F), indicating that DsrO participates in the regulation of several abiotic stress responses in D. radiodurans.

To characterize the physiological function of DsrO, we investigated the growth phenotypes of the WT, the mock strain (ΔDsrO-pRADZ3), the DsrO knockout strain (ΔDsrO), and the complementation strain under oxidative stress (Fig. 4). Under normal conditions (CK), there was no significant difference in growth between the tested strains. However, the DsrO mutant strain showed strong sensitivity to the 80 mM H₂O₂ treatment, whereas the complementation strain was nearly identical to the WT (Fig. 4A). Consistent with the experiments of msrA function analysis, significantly reduced total antioxidant capacity and greatly increased intracellular ROS levels were observed in the DsrO knockout compared to the WT (Fig. 4B and C). Contrary to our expectations, exposure to UV light, heat, or cold stress did not affect bacterial growth in any of the tested strains (Fig. S3). These observations showed that only oxidative stress influenced growth in the DsrO mutant strain, even though DsrO expression was significantly induced during several abiotic stress conditions (Fig. 3C to F). In conclusion, based on the above-described results, we propose that DsrO regulates the oxidative stress response of D. radiodurans by directly regulating msrA mRNA levels.

FIG 4 — The DsrO knockout mutant is sensitive to oxidative stress. (A) Phenotypes of different strains under treatment with 0 mM H₂O₂ and 80 mM H₂O₂ for 30 min. The bacterial cells were serially diluted 10 times. (Left) untreated control; (right) 80 mM H₂O₂ treatment. WT, wild-type strain; ΔDsrO, DsrO-deleted mutant; DsrO-*pRADZ3*, the DsrO mutant with *pRADZ3* empty plasmid; DsrO-com, DsrO mutant supplemented with the DsrO gene. (B and C) Total antioxidant capacity and ROS level of WT, ΔDsrO, DsrO-pRADZ3, and DsrO-com. Rosup is a reactive oxygen species positive-control reagent. One-way ANOVA and Dunnett’s multiple-comparison test; ***, P ≤ 0.001; *, P ≤ 0.05. Experiments were performed at least three times, and the data are presented as the means ± SEM.

DsrO directly stabilizes msrA expression by base pairing within the coding sequence.

To determine how DsrO interacts with its target, msrA, we performed 5′ RACE (rapid amplification 5′ end of full-length cDNA) analysis and confirmed that DsrO transcription started from the nucleotide of adenine and that the full length of the DsrO gene was 83 bp (Fig. S4). We further applied microscale thermophoresis (MST) to identify the binding strength between DsrO and msrA mRNA. The binding force curve indicated that DsrO could directly bind to msrA mRNA with a dissociation constant (K_d) of 3.80 ± 2.77 μM (Fig. 3B). When all of the bases in the complementation sequence of DsrO or msrA mRNA were mutated, the binding force between DsrO and msrA mRNA disappeared (Fig. S5C), supporting that the interaction occurs between bases 66 and 77 in DsrO and 54 and 65 in msrA. To determine which nucleotide is critical for their interactions, we mutated each base in these regions of DsrO and msrA and tested the affinity strength (Fig. S5A). All of the binding capacity was lost when the first two bases (66 and 67) in DsrO were mutated (Fig. S6A and B); other point mutations changed the dissociation constant, and these changes were significant compared to the WT (Fig. S6C to F). For msrA, all binding capacity was lost when base 64 or 65 (corresponding to base 66 or 67 in DsrO) was mutated (Fig. S7A and B). As with DsrO, the effects of other point mutations on the dissociation constant were significant in comparison to the WT (Fig. S7C to F). In conclusion, the first two bases at the 5′ end of the DsrO ring are the key bases for the interaction between DsrO and msrA mRNA in D. radiodurans.

To test the effects of these two bases on the physiological functions of D. radiodurans under oxidative stress, we investigated the growth phenotypes of the WT, mock strain (DsrO-pRADZ3), knockout mutant strain (DsrO-m3 or DsrO-m4), and complementation strain under oxidative stress (Fig. 5). Under normal growth conditions, there was no significant difference in growth between any of the tested strains; however, the DsrO mutant strains (DsrO-m3/m4) were significantly more sensitive to treatment with 80 mM H₂O₂, whereas the complementation strain was nearly identical to the WT (Fig. 5A). Significantly reduced total antioxidant capacity and strongly increased intracellular ROS levels were also observed in the DsrO-m3/m4 mutant strains in comparison to the WT (Fig. 5B). The effect of msrA point mutation on the oxidative tolerance of D. radiodurans was almost the same as that of DsrO; two mutations in msrA (at base 64 or 65, corresponding to base 66 and base 67 in DsrO) led to high oxidative stress sensitivity (Fig. 5C). These msrA-m3 and msrA-m4 mutants were sensitive to oxidative stress, with significantly decreased physiological indices (intracellular ROS levels, total antioxidant capacity) compared with the WT (Fig. 5D). We thus conclude that the first two bases located on the stem-loop structure of DsrO not only affect the binding strength between DsrO and msrA but also determine the level of tolerance to oxidative stress in D. radiodurans.

FIG 5 — The key base mutation of the binding region between DsrO and *msrA* confers a phenotype sensitive to oxidative stress. (A and C) Survival of DsrO (A) or *msrA* (C) point-mutation strains upon exposure to 80 mM H₂O₂. (B and D) Total antioxidant capacity and ROS level of different DsrO (B) or *msrA* (D) point-mutant strains upon exposure to 80 mM H₂O₂. WT, wild-type strain; ΔDsrO/*msrA*, DsrO/*msrA*-deleted mutant; DsrO/*msrA-pRADZ3*, the DsrO/*msrA* mutant with *pRADZ3* empty plasmid; DsrO/*msrA*-com, DsrO mutant supplemented with DsrO/*msrA* gene; DsrO/*msrA*-m3, DsrO/*msrA*_com strain with the 1st mutated base, DsrO/*msrA*_m4, DsrO_com strain with the 2nd mutated base; Rosup is a reagent that acts as a positive control. Asterisks indicate statistically significant differences compared to untreated cells. One-way ANOVA and Dunnett’s multiple-comparison test; ***, P ≤ 0.001; *, P ≤ 0.05. Experiments were performed at least three times, and the data are presented as the means ± SEM.

msrA targeted by DsrO forms a stem-loop structure that is required for msrA mRNA stability and protein translation.

The sRNAs can have a direct effect on the stability or translation levels of target mRNAs (33, 34). We therefore analyzed the half-life of msrA mRNA in D. radiodurans R1 and the mutant strains (msrA-m3 and msrA-m4). Analysis of mRNA stability showed that the msrA mRNA half-life was 5 min in the WT and only 2 min in the ΔDsrO and DsrO-m3/m4 mutant strains (Fig. 6A). Western blot and level of carbonylation results showed that the protein synthesis of MsrA was reduced in DsrO knockout and DsrO-m3/m4 point mutation strains in comparison with the WT, indicating that DsrO-m3/m4 point mutation likely affects protein translation (Fig. 6B and C). This indicated that DsrO and its key bases are essential for maintaining the stability of msrA mRNA.

FIG 6 — DsrO promotes the stability of *msrA* mRNA and enhances MsrA protein translation. (A) Time of *msrA* mRNA reaching its half-life in different strains. (B) Western blot of MsrA in different strains; bands of target protein (MsrA) and reference protein (GroEL) are indicated by arrows. WT, wild-type strain; ΔDsrO, DsrO deleted mutant; DsrO-pRADZ3, the DsrO mutant transformed with *pRADZ3* empty plasmid; DsrO-com, DsrO mutant supplemented with DsrO gene. (C) Intracellular carbonylation level and the percentage reduction of dabsyl-MetSO peak area in different samples. (D) The contents of intracellular Met-SO in DsrO mutants. One-way ANOVA and Dunnett’s multiple-comparison test; ***, P ≤ 0.001; *, P ≤ 0.05.

MsrA can catalyze the reduction of methionine-S-sulfoxide (Met-S-SO) to methionine in proteins and free amino acids. We therefore used high-performance liquid chromatography (HPLC) to analyze the contents of intracellular Met-SO in DsrO mutants. The metabolites measured were dabsyl-Met-S-SO, dabsyl-Met-R-SO, and dabsyl-Met. In the ΔDsrO mutant lines (DsrO-m3, DsrO-m4; msrA-m3, msrA-m4), dabsyl-Met-S-SO content was reduced by 30% compared to the WT; complementation of DsrO in ΔDsrO recovered the levels to 97% of the WT (Fig. 6D). In summary, these results suggest that the sRNA DsrO directly stabilizes the target mRNA msrA to promote its translation under oxidative stress.

msrA gene expression is transcriptionally regulated by the DrRRA transcription factor.

To identify the protein that transcriptionally regulates msrA in response to oxidative stress, we screened transcription factors coexpressed with msrA based on the transcriptomic data and a previously reported promoter analysis (35). DrRRA (Deinococcus radiodurans radio-resistance regulator) was predicted to bind to the msrA promoter based on previous transcriptomic data and interaction predictions. To confirm the interaction of DrRRA with the msrA promoter, we expressed and purified an N-terminally His-tagged DrRRA fusion protein from Escherichia coli and performed electrophoretic mobility shift assays (EMSAs). We observed a specific shift in mobility of the msrA promoter probe (500 bp) and a concomitant loss of the free msrA promoter probe, and stronger bands were detected with increasing concentrations of the DrRRA protein (Fig. 7A). In the DrRRA mutant cells, msrA expression was reduced, and no change occurred in response to oxidative stress (Fig. 7B). These data together indicate that DrRRA directly and specifically binds to the promoter region of msrA. Further analysis of the drRRA knockout line indicated that the deletion significantly decreased its tolerance to treatment with H₂O₂ (Fig. 7C), supporting that msrA is transcriptionally regulated by DrRRA.

FIG 7 — DrRRA is the transcriptional regulator of the *msrA* gene in *D. radiodurans*. (A) Electrophoretic mobility shift assay of DrRRA binding to the Cy5-labeled *msrA* promoter specifically. (Lane 1 [left] and Lane 2 [right]) Detection result of *dr_0997* promoter; lane 1 is the band of the single labeled *dr_0997* probe; lane 2 is the result of the labeled *dr_0997* probe with 6 μg DrRRA protein. Lanes 3 to 9) Detection result of *msrA* promoter; (lane 3) DrRRA protein; (lanes 4 to 8 reaction system contained 40 ng labeled *msrA* probe and 0 μg, 3 μg, 6 μg, 9 μg, and 12 μg DrRRA protein. (Lane 9) Reaction system containing 40 ng labeled *msrA* probe, 500 ng unlabeled *msrA* probe, and 6 μg DrRRA protein. (B) The deletion of the DrRRA gene in *D. radiodurans* resulted from the induction decrease of *msrA* expression activated by the oxidative stress. (C) H₂O₂ sensitivity assays of different DrRRA strains. Spotted agar plates after H₂O₂ treatment and serial dilution. Different strains were treated with 0 to 100 mM H₂O₂ for 30 min and dotted in the plates. WT, wild-type strain, Δ*drRRA*, *drRRA* deleted mutant. One-way ANOVA, Dunnett’s multiple-comparison test; ***, P ≤ 0.001; *, P ≤ 0.05. Experiments were performed at least three times, and data are presented as means ± SEM.

DISCUSSION

To survive in extreme environments, D. radiodurans has developed efficient antioxidant systems to decrease ROS production, repair DNA damage, and recover the functions of oxidized proteins (36). Compared to the ROS-sensitive E. coli, D. radiodurans has highly active protein repair systems to mitigate ROS damage and decrease oxidized protein and protein carbonylation. Because the Msr protein is a typical enzyme with the ability to repair oxidized Met (37), we investigated the function of MsrA in D. radiodurans and analyzed its regulatory mechanisms. Here, we report the function of MsrA in D. radiodurans during oxidative stress and show that msrA is regulated by both the sRNA DsrO and the transcription factor DrRRA to quickly respond to oxidative stress, improving the antioxidant ability of D. radiodurans compared to ROS-sensitive bacteria (Fig. 8).

FIG 8 — Diagram of DsrO posttranscriptionally regulating *msrA* in *D. radiodurans*. (i) *D. radiodurans* cells produce ROS under the treatment of exogenous H₂O₂ or oxidative stresses. ROS oxidize the methionine residue of proteins into methionine sulfoxide, inactivating the protein or causing cell death. (ii) When exogenous H₂O₂ enters the *D. radiodurans* cell, the sRNA DsrO and *msrA* are rapidly induced. At the same time, the transcription factor DrRRA upregulates the expression of *msrA* at the transcriptional level. (iii) DsrO binds to the stem-ring structure of *msrA* mRNA and thus enhances the stability of *msrA* mRNA and improves MsrA expression at the translational level. (iv) The increased MsrA reduces the methionine sulfoxide on the protein to methionine to restore protein function, thus promoting the oxidation resistance of *D. radiodurans*. The large gray-green circle represents the 50s large subunit of the ribosome, the small circle represents the 30 s small subunit of the ribosome, the blue stem-loop represents sRNA DsrO, the light purple stem-loop represents *msrA* mRNA, the gray fold represents DrRRA protein, and the bright purple fold represents MsrA protein. The dark blue folds represent proteins in the cell, and the rose dots above represent Met residues on the proteins.

MsrA in D. radiodurans R1 primarily functions under oxidative stress.

Different abiotic and biotic stresses can result from ROS accumulation and cell damage in bacteria (38). Methionine, as an initiator amino acid residue, is fundamental for protein translation and ranks as the most sensitive to oxidation (39, 40). To restore the function of the damaged protein, msrA is strongly upregulated in response to high radiation, desiccation, or low temperature (41). In D. radiodurans, we also found that msrA expression was significantly enhanced when the cells were exposed to 80 mM H₂O₂, UV radiation, or temperature stress (Fig. 1, Fig. S1). In D. radiodurans cells exposed to 60 mM H₂O₂ for 30 min, the maximum transcription level of msrA was almost 13-fold higher than that in the control group (Fig. 1A). msrA knockout D. radiodurans cells were susceptible to H₂O₂ damage (Fig. 2), and the msrA mutant had a deficiency in antioxidative capacity and increased levels of dabsyl-Met-S-SO. These results were consistent with the modification function of MsrA protein previously observed in other organisms (7).

Contrary to our expectations, msrA knockouts were not sensitive to UV radiation or heat/cold stress, even though msrA was strongly upregulated in all three stress conditions in D. radiodurans (Fig. S2). These results indicate that MsrA is a key protein in the response to H₂O₂ treatment but does not function during UV radiation or temperature stress. Another possibility is that a reduction in the level of damaged proteins requires the participation of both MsrA and MsrB under UV radiation or hot/cold stress. Similar to most other organisms, the D. radiodurans genome contains one copy each of msrA and msrB (42). Based on their primary sequence and protein structures, MsrA and MsrB have no significant homology. MsrA and MsrB possess the ability to reverse S,R-MetSO forms into Met (43). Molecular analysis indicated that msrA and msrB have different expression patterns and physiological roles when bacterial cells are exposed to various abiotic stresses, such as iron limitation (44). Consequently, future work will aim to functionally analyze MsrB and its connection with MsrA in protein repair under UV radiation or hot/cold stress.

msrA mRNA stability is crucial for the antioxidant capacity of D. radiodurans.

The sRNAs typically bind target mRNAs through complementation, and this binding alters mRNA stability and translation (45, 46). Using sRNA target mRNA analysis and molecular interaction analysis, we demonstrated that the sRNA DsrO binds to the coding sequence region of msrA mRNA in vitro (Fig. 3). Site-directed mutagenesis of the complementary sequence between DsrO and msrA demonstrated that the 1st and 2nd bases of DsrO are crucial for the stability of msrA mRNA (Fig. S6 and S7). As expected, viability was decreased in DsrO and msrA knockouts and point mutants when treated with 80 mM H₂O₂ (Fig. 5). We thus propose that DsrO is a positive regulator of msrA mRNA stability, and this improved mRNA stability benefits MsrA protein translation, ultimately enhancing MsrA enzyme activity (Fig. 6). Bacterial sRNAs often negatively regulate transcription by binding the mRNA of a target gene and enhancing transcription termination (34). Contrary to the regulatory pattern of most investigated bacterial sRNAs, DsrO expression accelerated the translation of msrA, as indicated by Western blotting (Fig. 6). One possible explanation is that the interaction between DsrO and msrA mRNA forms a secondary structure to activate translation. This regulatory mechanism is similar to the activation of translation fepA mRNA by the binding sRNAs OmrA and OmrB in E. coli (47).

D. radiodurans has an unusual capability to repair extensive DNA damage, maintaining genome stability under abiotic stress conditions, including high gamma and UV radiation, hydrogen peroxide exposure, and desiccation (48). The efficient DNA repair system in D. radiodurans requires the participation of RecA, PprA, and DNA glucoamylase (49). Under high H₂O₂ levels, the Mets in E. coli RecA may be oxidized to MetSO, losing function. MsrA/B are required for maintaining the recombinase function of RecA under oxidative stress (50). These findings imply that the regulation of oxidized protein repair involves cross talk with the DNA repair signaling pathway in D. radiodurans. In this study, we demonstrated that DrRRA transcriptionally regulates msrA expression by directly binding to its promoter (−73 bp to −54 bp: CTCCCGGCGCCTGTCCCCTC). DrRRA has a regulatory role in multiple antioxidation and DNA repair pathways for preserving DNA in D. radiodurans (35). The deletion of drRRA results in decreased viability under oxidative stress as well as the delay of genome recovery, indicating that DrRRA and DsrO may jointly regulate DNA and protein modification. Because msrA expression is regulated by DrRRA at the transcriptional level and by DsrO at the posttranscriptional level, future work should focus on determining whether other proteins/factors integrate the regulatory activities of DrRRA and DsrO in D. radiodurans.

MATERIALS AND METHODS

Bacterial strains and plasmids.

The liquid growth media used were TGY broth (1% tryptone, 0.1% glucose, 0.5% yeast extract, pH 7.0) and LB broth (1% tryptone, 1% yeast extract, 0.5% NaCl, pH 7.0). The solid growth media used were TGY and LB agar (TGY or LB broth with 1.5% agar). Deinococcus radiodurans R1 and its derivatives were grown in or on TGY with appropriate antibiotics (Table 1) at 30°C. Escherichia coli was grown in or on LB with appropriate antibiotics (Table 1) at 37°C. All liquid cultures were incubated with shaking at 220 rpm. The plasmids and strains used in this study are listed in Table 1.

TABLE 1.

The list of plasmids and strains

Plasmid/strain	Description	Source
pRADZ3	Shuttle vector for E. coli and D. radiodurans, Chl^r (D. radiodurans), Amp^r (E. coli)	Laboratory stock
pKatAPH3	To amplify the kanamycin resistance gene	Laboratory stock
pKatAAD2	To amplify the spectinomycin resistance gene	Laboratory stock
D. radiodurans R1	Wild-type, served as the strain for generating the mutants	Laboratory stock
E. coli DH5α	The strain for expressing the shuttle plasmid pRADZ3	CW Biotech
ΔmsrA mutant	D. radiodurans with genomic deletion of msrA gene	This study
msrA-pRADZ3	msrA mutant with pRADZ3 shuttle plasmid introduced into its genome	This study
msrA_com	Complementation of the msrA deletion in D. radiodurans, transformation of msrA mutant with pRADZ3 plasmid expressing D. radiodurans msrA gene	This study
msrA_m3	msrA complementary strain with a mutated base which is the first base in the binding sequence	This study
msrA_m4	msrA complementary strain with a mutated base which is the second base in the binding sequence	This study
ΔDsrO mutant	D. radiodurans with genomic deletion of DsrO gene	This study
DsrO_pRADZ3	DsrO mutant with pRADZ3 shuttle plasmid introduced into its genome	This study
DsrO_com	Complementation of the DsrO deletion in D. radiodurans, transformation of DsrO mutant with pRADZ3 plasmid expressing D. radiodurans DsrO gene	This study
DsrO_m3	DsrO complementary strain with a mutated base which is the first base in the binding sequence	This study
DsrO_m4	DsrO complementary strain with a mutated base which is the first base in the binding sequence	This study

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To generate the msrA and DsrO mutants, sequences 1 kb upstream (U) and 1 kb downstream (D) of the gene and the spectinomycin resistance gene (S) were amplified separately. Then, the three fragments were assembled in a molar ratio of 1:1:1 under the catalysis of homologous recombinase. Finally, the assembled product was used as a template, and P1-F/P3-R was used as a primer to amplify the fusion fragment USD. Finally, the fused fragment was introduced into D. radiodurans as previously reported (51). For complementation of the msrA gene in D. radiodurans, msrA and the associated promoter were inserted into the pRADZ3 plasmid between the endonuclease BamHI and SpeI sites, and then the recombinant plasmid was transformed into D. radiodurans R1.

H₂O₂ stress treatment and oxidative tolerance phenotype analysis.

D. radiodurans strain R1 and its derivatives were grown in TGY broth until the beginning of the logarithmic growth phase (optical density at 600 nm [OD₆₀₀], 0.5 to 0.8) and then transferred into fresh TGY broth with or without H₂O₂ (final concentration, 80 mM) for 30 min. For oxidative tolerance analysis of the different strains, 100 μL of bacterial culture was added to 900 μL of phosphate-buffered saline (PBS). Cultures were serially diluted 10-fold to a final concentration of 10⁻⁵. The diluted culture (7 μL) was dropped vertically onto TGY plates in the order of concentration from 10⁻¹ to 10⁻⁵. The plates were incubated at 30°C for 3 days, and growth phenotypes were recorded.

Total RNA extraction and Northern blotting.

Total RNA extraction was performed following the protocol of the PureLink RNA minikit (Invitrogen, California, USA). For Northern blotting, 10 μg of total RNA from different samples was separated on a 6% urea gel (National Diagnostics, Georgia, USA) for 1 to 1.5 h at 120 V. After electrophoresis, the RNA was transferred to a nylon membrane using semidry transfer blotting. The membrane was incubated at 60°C for 1 to 2 h to facilitate cross-linking to the EDC (1-ethyl-3-[3-dimethylaminopropyl]carbodiimide) reagent.

The hybridization probes were labeled following the instructions of the Pierce RNA 3′ end biotinylation kit (Thermo Fisher Scientific), and then the biotin-labeled probes were hybridized with the membrane at 37°C overnight in a hybridization oven. The nylon membranes were washed twice with stringent buffer, and the hybridization signals were detected following the procedure of the chemiluminescent nucleic acid detection module (Thermo Fisher Scientific).

Rapid amplification of the 5′ end (RACE) of the DsrO gene.

The 5′/3′ RACE kit 2nd generation (Roche, Mannheim, Germany) was used to amplify the end of the DsrO gene. The primers were designed according to the genome sequence of the D. radiodurans R1 strain. The full-length cDNA of the gene was amplified after cDNA synthesis (cDNA synthesis buffer, 4 μL; deoxynucleotide mixture, 2 μL; primer SP1, 1 μL; total RNA, 1 μg; transcriptor reverse transcriptase, 1 μL; and double-distilled water _[ddH2O], up to 20 μL) and 2 cycles of nested PCR (dA-tailed cDNA, 5 μL; oligonucleotide dT-anchor primer, 1 μL; primer SP2, 1 μL; deoxynucleotide mixture, 1 μL; expanded high-fidelity buffer, 5 μL; double-distilled water, up to 50 μL).

Real-time quantitative PCR (qRT-PCR).

Gene expression was analyzed with qRT-PCR. First-strand cDNA synthesis was performed with the PrimeScript RT reagent kit (TaKaRa Biomedical Technology, Beijing, China). First, 2 μL 5× genomic DNA (gDNA) eraser buffer, 1 μL gDNA eraser, and 1 μg total RNA were incubated at 42°C for 2 min to remove genomic DNA; then we added 1 μL PrimeScript RT enzyme mix I, 4 μL RT primer mix, 4 μL 5× PrimeScript buffer 2, and 1 μL RNase-free distilled water (dH₂O) into the reaction liquid. qRT-PCR was performed with SYBR qPCR master mix (Vazyme, Nanjing, China) following the manufacturer’s instructions. The reaction system included 10 μL Hiscript III RT supermix, 1 μL RT-F primer, 1 μL RT-R primer, and 8 μL ddH₂O, and the 16S rRNA gene was used as the internal control. The 2^–ΔΔ^CT values were calculated to indicate the expression levels of the target genes (52).

Bioinformatics analysis.

To predict the binding sequence of DsrO in the msrA gene, IntaRNA software was used (http://rna.informatik.uni-freiburg.de/IntaRNA/Input.jsp) (53). The minimum binding nucleotide was set to 7. The secondary structures of msrA mRNA and DsrO were predicted with RNAalifold (http://rna.tbi.univie.ac.at/cgi-bin/RNAWebSuite/RNAalifold.cgi) (54). The fold algorithms and basic options used were minimum free energy (MFE), partition function, and the avoidance of isolated base pairs.

Microscale thermophoresis analysis.

Full-length sRNA was synthesized by in vitro transcription; 5′ fluorescein amidite (FAM)-labeled mRNA was purchased from Shanghai GenePharma (Shanghai, China). The labeled mRNAs were then diluted to a fluorescence intensity of 200 to 1,000 according to the protocol in the user manual of the Monolith NT.115 system (NanoTemper Technologies, Munich, Germany). The detailed procedure was followed as previously reported (55, 56).

Total antioxidant capacity, reactive oxygen (ROS) level, and intracellular carbonylation level analysis.

Bacterial ROS levels were measured with a ROS assay kit (Beyotime, Shanghai, China) containing the 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA) fluorescent probe according to the manufacturer’s protocol (57). Bacteria loaded with the DCFH-DA probe were cultured to the initial logarithmic growth phase. Bacteria were collected by centrifugation and washed three times with TGY medium to remove probe contamination. Bacteria were then treated with H₂O₂ (final concentration, 80 mM) for 30 min. Rosup was used as the positive control as described in the kit brochure. Fluorescence intensity was analyzed at 488 nm excitation and 525 nm emission wavelengths.

Ferric reducing ability of plasma (FRAP) was performed to analyze the antioxidant capacity of bacteria using a total antioxidant capacity assay kit (Beyotime, Shanghai, China) (58). Cells were collected for ultrasonication at the beginning of the logarithmic growth phase. The bacteria were then centrifuged at 1,150 × g for 10 min. The supernatant (200 μL) was transferred to a new Eppendorf tube, and 180 μL of FRAP buffer was added. The mixture was incubated at 37°C for 3 to 5 min, and then the OD was measured at 593 nm.

A protein carbonyl colorimetric assay kit (Cayman Chemical, Ann Arbor, MI, USA) (59) was used to measure the protein carbonyl content of cell lysates in 96-well plates. Bacteria at the beginning of the logarithmic stage were collected for ultrasonication. The bacteria were then centrifuged at 3,000 rpm for 10 min. The supernatant (10 μL) was transferred to new plates and diluted 10 times. The amount of protein was calculated from a bovine serum albumin (BSA) standard curve (0.25 to 1.0 mg/mL). The amount of protein-hydrozone produced was quantified spectrophotometrically at an absorbance between 360 and 380 nm.

Stability measurement of msrA mRNA.

Deinococcus radiodurans R1 and its derivatives were cultured to the initial logarithmic growth phase. Bacteria were treated with H₂O₂ (to a final concentration of 80 mM) for 30 min before rifampin (400 μg/mL) was added to stop gene transcription. At different time points after rifampin treatment (0 min, 2 min, 4 min, 6 min, 8 min, and 10 min), 2 mL of culture was collected from each sample. Total RNA was extracted as described above, and the level of msrA mRNA was detected with a real-time fluorescence quantification kit (Invitrogen).

Western blot analysis.

Total protein was extracted from the H₂O₂-stressed and untreated bacteria as described above. The total protein (10 μg) of each sample was denatured at 100°C for 5 min and separated by 12% SDS-PAGE at 4°C for 1.5 h. After electrophoresis, the separated proteins were transferred from the gel onto a polyvinylidene difluoride (PVDF) membrane with a semidry electroblotter. The membrane was blocked with skimmed milk for 1 h. After that, the membranes were hybridized overnight with the primary anti-rabbit MsrA antibody (1:1,000). The primary polyclonal antibody against MsrA was produced by Genscript Biotechnology Company based on the MsrA protein sequence (Genscript, Nanjing, China). Next, the membrane was washed three times and treated with 1:5,000 goat anti-rabbit horseradish peroxidase (HRP) secondary antibody. Detection of the MsrA band was performed using the SuperSignal West Femto monoclonal kit (Thermo Fisher Scientific). The GroEL protein was used as the reference protein.

Enzymatic activity analysis of MsrA.

MsrA enzymatic activity was analyzed as previously described (60). Dabsyl-Met (50 mM) was oxidized overnight by 500 mM H₂O₂, the supernatant was filtered by a C₁₈ column, and finally, Dabsyl-MetSO was obtained when the column was eluted with acetonitrile. The reaction solution and the termination solution were prepared according to previous reports (60). Total protein extracts from different samples were added to the reaction solution. The reaction was terminated after incubation at 37°C for 1 h, and then the samples were centrifuged at 12,000 × g for 30 min at 4°C. The supernatant was used for HPLC. HPLC parameters and program settings all followed the procedure of the report of Vieira Dos Santos et al. (60).

Electrophoretic mobility shift assay (EMSA).

His-tagged DrRRA was expressed using the pET30a (+) plasmid in E. coli strain BL21 and purified with His-tagged beads (Invitrogen, USA). The purified proteins were used in EMSA. EMSAs were performed according to the manufacturer’s instructions (Odyssey Infrared EMSA kit, LI-COR). DNA fragments used for Cy5-labeled probes were amplified by PCR with the labeled primers (Table S1). The Cy5-labeled DNA probe (40 ng) was incubated with different quantities of DrRRA protein and a positive dr_0997 probe at 30°C for 30 min in binding buffer (20 μL). After incubation for 30 min, the solution, including protein-bound and free DNA, was separated by electrophoresis on 4.0% native polyacrylamide gels with 0.5× TBE buffer (44.5 mM Tris-HCl, 44.5 mM boric acid, and 1 mM EDTA, pH 8.0) at 100 V and 4°C. The gel was detected by an Amersham Imager 600.

Data availability.

The genome sequence of Deinococcus radiodurans R1 has been deposited in NCBI GenBank under accession numbers CP015081 (https://www.ncbi.nlm.nih.gov/genome/?term=CP015081)(new) and AE000513 (https://www.ncbi.nlm.nih.gov/genome/?term=AE000513) (old). The accession numbers of the D. radiodurans genes used in this study are as follows: msrA (A2G07_04355 and DR_1849) and DR_0997 (A2G07_08515). The primers used in this study are listed in Table 2. Other relevant data are available within this article and its associated supplemental material.

TABLE 2.

Primers used in this study

Primer name	Sequence (5′–3′)^a	Purpose
P1-F	TAGTCCTGCACCCACTTCAGGGCCG	ΔmsrA construct
P1-R	TCGGTCTCCATGCGAGCTCGAATTCGATAGGGACCGGAAAGCGGATTTAA
P2-F	ATCCGCTTTCCGGTCCCTATCGAATTCGAGCTCGCATGGAGACCGA
P2-R	TCGGTCTGCCACAGCACCCGCATTATTTGCCGACTACCTTGGTGAT
P3-F	ACCAAGGTAGTCGGCAAATAATGCGGGTGCTGTGGCAGACCGAGGC
P3-R	GCTCGCACAGCGTGGTGTATCTGGC
P4-F	TCCAGCACACTGGCGGCCGTTACTAGTTCAGGAACCAGCCGTCGTCA	msrA_com construct
P4-R	GCATGCCTGCAGGTCGAATCGGATCCGCTCGGTGAAGACCCGGGGG	msrA_com construct
P5-F	CACCGCTTAACATCACCCCT	ΔDsrO construct
P5-R	TGCTCGATGAGTTTTTCTAAGGATAACTTTGTTTTACGAGCGTT
P6-F	CGTAAAACAAAGTTATCCTTAGAAAAACTCATCGAGCATCAAATG
P6-R	CCTGGTTCAAACAACGAAAACATGGAGACCGAGGGCCCTTGACA
P7-F	AAGGGCCCTCGGTCTCCATGTTTTCGTTGTTTGAACCAGGTGCA
P7-R	GGGGATGAACGGCATTCTCT
P8-F	TCCAGCACACTGGCGGCCGTTACTAGTCCACCCTCCCCGACGCCCTGCACG	DsrO_com construct
P8-R	GCATGCCTGCAGGTCGAATCGGATCCCGCCGTCCAGCAAACAAAAAACG	DsrO_com construct
msrA RT-F	CGCCGCACAATAGCAGCATG	qRT-PCR for msrA expression pattern
msrA RT-R	AGCGGTAATCGGGATTGGGCA	qRT-PCR for msrA expression pattern
DsrO RT-F	ACAGGTCTACGTTTCACTGTGGATA	qRT-PCR for msrA expression pattern
DsrO RT-R	CGCCGTCCAGCAAACAAAAAACGCT	qRT-PCR for msrA expression pattern
16s RT-F	ATTCCTGGTGTAGCGGTG	qRT-PCR for 16s rRNA
16s RT-R	ATTCCTGGTGTAGCGGTG	qRT-PCR for 16s rRNA
msrA SP1	TCAGCGGAGTTTGTCGCCGTAGTACTGGCGCAGCT	5′ RACE for msrA
msrA SP2	CCGAGCTGCTCGATCATTTCGCGGGCCGTCTGTTC
msrA SP3	AGCGGTAATCGGGATTGGGCACCGTACCGCCGATG
DsrO SP1	CGCCGTCCAGCAAACAAAAAACGCTGAACAGTCAT	5′ RACE for DsrO
DsrO SP2	CAGCAAACAAAAAACGCTGAACAGTCATGCTTGAAAATCCGC
DsrO SP3	AAAAACGCTGAACAGTCATGCTTGAAAATCCGCTTCAAAAGTT

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The underlining indicates the restriction enzyme cutting site of Spe1(P4/P8-F) and BamHI(P4/P8-R).

ACKNOWLEDGMENTS

This work was funded by Key Research and Development Projects (no. 2018YFA0901000 and 2018YFA0901003), the Ministry of Agriculture Transgenic Program (no. 2019ZX08010-002), the National Natural Science Foundation of China (no. 31800061, 31570080, and 31500063), and the BIO-Agri project of Shanghai Jiao Tong University (SJTU). We also appreciate support from the Agricultural Science and Technology Innovation Program of Chinese Academy of Agricultural Sciences (CAAS).

We declare no conflicts of interest.

K. Zuo, M. Lin, and J. Wang. designed and conducted the experiments. Y. Chen, M. Lv, and M. Zhao performed the experiments. K. Zuo wrote and revised the paper.

Footnotes

Supplemental material is available online only.

Supplemental file 1

Fig. S1 to S7. Download aem.00038-22-s0001.pdf, PDF file, 0.7 MB^{(710.2KB, pdf)}

Contributor Information

Jin Wang, Email: wangjin@caas.cn.

Kaijing Zuo, Email: kjzuo@sjtu.edu.cn.

Isaac Cann, University of Illinois at Urbana-Champaign.

REFERENCES

1.Kaludercic N, Deshwal S, Lisa F. 2014. Reactive oxygen species and redox compartmentalization. Front Physiol 5:285–300. 10.3389/fphys.2014.00285. [DOI] [PMC free article] [PubMed] [Google Scholar]
2.Yang L, Mih N, Anand A, Park JH, Tan J, Yurkovich JT, Monk JM, Lloyd CJ, Sandberg TE, Seo SW, Kim D, Sastry AV, Phaneuf P, Gao Y, Broddrick JT, Chen K, Heckmann D, Szubin R, Hefner Y, Feist AM, Palsson BO. 2019. Cellular responses to reactive oxygen species are predicted from molecular mechanisms. Proc Natl Acad Sci USA 116:14368–14373. 10.1073/pnas.1905039116. [DOI] [PMC free article] [PubMed] [Google Scholar]
3.Cadet J, Davies KJA. 2017. Oxidative DNA damage & repair: an introduction. Free Radic Biol Med 106:100–111. 10.1016/j.freeradbiomed.2017.03.030. [DOI] [PMC free article] [PubMed] [Google Scholar]
4.Poetsch AR. 2020. The genomics of oxidative DNA damage, repair, and resulting mutagenesis. Comput Struct Biotechnol J 18:207–219. 10.1016/j.csbj.2019.12.013. [DOI] [PMC free article] [PubMed] [Google Scholar]
5.Yalcinkaya T, Uzilday B, Ozgur R, Turkan I, Mano J. 2019. Lipid peroxidation-derived reactive carbonyl species (RCS): their interaction with ROS and cellular redox during environmental stresses. Environ Exp Bot 165:139–149. 10.1016/j.envexpbot.2019.06.004. [DOI] [Google Scholar]
6.Shari F, Ibrahim AY, Hassan JK, Al-Salman HNK. 2020. Study effect of sulfur containing amino acid supplementation on oxidative stress, inflammatory markers and serum lipid in relation to obese subjects. Int J Pharm Sci 12:362–370. 10.31838/ijpr/2020.12.01.062. [DOI] [Google Scholar]
7.Ezraty B, Gennaris A, Barras F, Collet JF. 2017. Oxidative stress, protein damage and repair in bacteria. Nat Rev Microbiol 15:385–396. 10.1038/nrmicro.2017.26. [DOI] [PubMed] [Google Scholar]
8.Liu Y, Zhang C, Wang Z, Lin M, Wang J, Wu M. 2021. Pleiotropic roles of late embryogenesis abundant proteins of Deinococcus radiodurans against oxidation and desiccation. Comput Struct Biotechnol J 19:3407–3415. 10.1016/j.csbj.2021.05.051. [DOI] [PMC free article] [PubMed] [Google Scholar]
9.Ranawat P, Rawat S. 2017. Radiation resistance in thermophiles: mechanisms and applications. World J Microb Biot 33:112–134. 10.1007/s11274-017-2279-5. [DOI] [PubMed] [Google Scholar]
10.Adamec F, Farci D, Bína D, Litvín R, Khan T, Fuciman M, Piano D, Polívka T. 2020. Photophysics of deinoxanthin, the keto-carotenoid bound to the main S-layer unit of Deinococcus radiodurans. Photochem Photobiol Sci 19:495–503. 10.1039/D0PP00031K. [DOI] [PubMed] [Google Scholar]
11.Jeong SW, Kim MK, Zhao L, Yang SK, Jung JH, Lim HM, Lim S. 2021. Effects of conserved wedge domain residues on DNA binding activity of Deinococcus radiodurans RecG helicase. Front Genet 12:634615–634626. 10.3389/fgene.2021.634615. [DOI] [PMC free article] [PubMed] [Google Scholar]
12.Lim S, Jung JH, Blanchard L, de Groot A. 2019. Conservation and diversity of radiation and oxidative stress resistance mechanisms in Deinococcus species. FEMS Microbiol Rev 43:19–52. 10.1093/femsre/fuy037. [DOI] [PMC free article] [PubMed] [Google Scholar]
13.Agapov A, Esyunina D, Kulbachinskiy A. 2019. Gre-family factors modulate DNA damage sensing by Deinococcus radiodurans RNA polymerase. RNA Biol 16:1711–1720. 10.1080/15476286.2019.1656027. [DOI] [PMC free article] [PubMed] [Google Scholar]
14.Cox MM, Keck JL, Battista JR. 2010. Rising from the ashes: DNA Repair in Deinococcus radiodurans. PLoS Genet 6:e1000815. 10.1371/journal.pgen.1000815. [DOI] [PMC free article] [PubMed] [Google Scholar]
15.Daly MJ. 2009. A new perspective on radiation resistance based on Deinococcus radiodurans. Nat Rev Microbiol 7:237–245. 10.1038/nrmicro2073. [DOI] [PubMed] [Google Scholar]
16.Zhou Y, Shen P, Lan Q, Deng C, Zhang Y, Li Y, Wei W, Wang Y, Su N, He F, Xie Q, Lyu Z, Yang D, Xu P. 2017. High-coverage proteomics reveals methionine auxotrophy in Deinococcus radiodurans. Proteomics 17:1700072–1700074. 10.1002/pmic.201700072. [DOI] [PubMed] [Google Scholar]
17.Ahmad S, Khan H, Shahab U, Rehman S, Rafi Z, Khan MY, Ansari A, Siddiqui Z, Ashraf JM, Abdullah SM, Habib S, Uddin M. 2017. Protein oxidation: an overview of metabolism of sulphur containing amino acid, cysteine. Front Biosci (Schol Ed) 9:71–87. 10.2741/s474. [DOI] [PubMed] [Google Scholar]
18.Krisko A, Radman M. 2010. Protein damage and death by radiation in Escherichia coli and Deinococcus radiodurans. Proc Natl Acad Sci USA 107:14373–14377. 10.1073/pnas.1009312107. [DOI] [PMC free article] [PubMed] [Google Scholar]
19.Radman M. 2016. Protein damage, radiation sensitivity and aging. DNA Repair (Amst) 44:186–192. 10.1016/j.dnarep.2016.05.025. [DOI] [PubMed] [Google Scholar]
20.Daly MJ, Gaidamakova EK, Matrosova VY, Vasilenko A, Zhai M, Leapman RD, Lai B, Ravel B, Li SM, Kemner KM, Fredrickson JK. 2007. Protein oxidation implicated as the primary determinant of bacterial radioresistance. PLoS Biol 5:e92. 10.1371/journal.pbio.0050092. [DOI] [PMC free article] [PubMed] [Google Scholar]
21.Chang RL, Stanley JA, Robinson MC, Sher JW, Li Z, Chan YA, Omdahl AR, Wattiez R, Godzik A, Matallana-Surget S. 2020. Protein structure, amino acid composition and sequence determine proteome vulnerability to oxidation-induced damage. EMBO J 39:e104523. 10.15252/embj.2020104523. [DOI] [PMC free article] [PubMed] [Google Scholar]
22.Achilli C, Ciana A, Minetti G. 2015. The discovery of methionine sulfoxide reductase enzymes: an historical account and future perspectives. Biofactors 41:135–152. 10.1002/biof.1214. [DOI] [PubMed] [Google Scholar]
23.Fu X, Adams Z, Liu R, Hepowit NL, Wu Y, Bowmann CF, Moskovitz J, Maupin-Furlow JA. 2017. Methionine sulfoxide reductase A (MsrA) and its function in ubiquitin-like protein modification in Archaea. mBio 8:e01169-17. 10.1128/mBio.01169-17. [DOI] [PMC free article] [PubMed] [Google Scholar]
24.Boschi-Muller S. 2018. Molecular mechanisms of the methionine sulfoxide reductase system from Neisseria meningitidis. Antioxidants 7:131–142. 10.3390/antiox7100131. [DOI] [PMC free article] [PubMed] [Google Scholar]
25.Hopman CT, Speijer D, van der Ende A, Pannekoek Y. 2010. Identification of a novel anti-σE factor in Neisseria meningitidis. BMC Microbiol 10:164–176. 10.1186/1471-2180-10-164. [DOI] [PMC free article] [PubMed] [Google Scholar]
26.You C, Sekowska A, Francetic O, Martin-Verstraete I, Wang Y, Danchin A. 2008. Spx mediates oxidative stress regulation of the methionine sulfoxide reductases operon in Bacillus subtilis. BMC Microbiol 8:128–141. 10.1186/1471-2180-8-128. [DOI] [PMC free article] [PubMed] [Google Scholar]
27.Grimaud R, Ezraty B, Mitchell JK, Lafitte D, Briand C, Derrick PJ, Barras F. 2001. Repair of oxidized proteins. Identification of a new methionine sulfoxide reductase. J Biol Chem 276:48915–48920. 10.1074/jbc.M105509200. [DOI] [PubMed] [Google Scholar]
28.Kim YK, Shin YJ, Lee WH, Kim HY, Hwang KY. 2009. Structural and kinetic analysis of an MsrA-MsrB fusion protein from Streptococcus pneumoniae. Mol Microbiol 72:699–709. 10.1111/j.1365-2958.2009.06680.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
29.Kappler U, Nasreen M, McEwan A. 2019. New insights into the molecular physiology of sulfoxide reduction in bacteria. Adv Microb Physiol 75:1–51. 10.1016/bs.ampbs.2019.05.001. [DOI] [PubMed] [Google Scholar]
30.Oertlin C, Lorent J, Murie C, Furic L, Topisirovic I, Larsson O. 2019. Generally applicable transcriptome-wide analysis of translation using anota2seq. Nucleic Acids Res 47:e70. 10.1093/nar/gkz223. [DOI] [PMC free article] [PubMed] [Google Scholar]
31.Levine RL, Berlett BS, Moskovitz J, Mosoni L, Stadtman ER. 1999. Methionine residues may protect proteins from critical oxidative damage. Mech Ageing Dev 107:323–332. 10.1016/s0047-6374(98)00152-3. [DOI] [PubMed] [Google Scholar]
32.Bernhart SH, Hofacker IL, Will S, Gruber AR, Stadler PF. 2008. RNAalifold: improved consensus structure prediction for RNA alignments. BMC Bioinformatics 9:474. 10.1186/1471-2105-9-474. [DOI] [PMC free article] [PubMed] [Google Scholar]
33.Storz G, Opdyke JA, Zhang A. 2004. Controlling mRNA stability and translation with small, noncoding RNAs. Curr Opin Microbiol 7:140–144. 10.1016/j.mib.2004.02.015. [DOI] [PubMed] [Google Scholar]
34.Buddeweg A, Sharma K, Urlaub H, Schmitz RA. 2018. sRNA₄₁ affects ribosome binding sites within polycistronic mRNAs in Methanosarcina mazei Gö1. Mol Microbiol 107:595–609. 10.1111/mmi.13900. [DOI] [PubMed] [Google Scholar]
35.Wang L, Xu G, Chen H, Zhao Y, Xu N, Tian B, Hua Y. 2008. DrRRA: a novel response regulator essential for the extreme radioresistance of Deinococcus radiodurans. Mol Microbiol 67:1211–1222. 10.1111/j.1365-2958.2008.06113.x. [DOI] [PubMed] [Google Scholar]
36.Qi HZ, Wang WZ, He JY, Ma Y, Xiao FZ, He SY. 2020. Antioxidative system of Deinococcus radiodurans. Res Microbiol 171:45–54. 10.1016/j.resmic.2019.11.002. [DOI] [PubMed] [Google Scholar]
37.Shome A, Sarkhel R, Apoorva S, Nair SS, Chauhan TKS, Bhure SK, Mahawar M. 2020. Role of protein repair enzymes in oxidative stress survival and virulence of Salmonella. Ann Microbiol 70:55–65. 10.1186/s13213-020-01597-2. [DOI] [Google Scholar]
38.Wang Y, Branicky R, Noë A, Hekimi S. 2018. Superoxide dismutases: dual roles in controlling ROS damage and regulating ROS signaling. J Cell Biol 217:1915–1928. 10.1083/jcb.201708007. [DOI] [PMC free article] [PubMed] [Google Scholar]
39.Brosnan JT, Brosnan ME, Bertolo RFP, Brunton JA. 2007. Methionine: a metabolically unique amino acid. Livestock Sci 112:2–7. 10.1016/j.livsci.2007.07.005. [DOI] [Google Scholar]
40.Martínez Y, Li X, Liu G, Bin P, Yan W, Más D, Valdivié M, Hu CA, Ren W, Yin Y. 2017. The role of methionine on metabolism, oxidative stress, and diseases. Amino Acids 49:2091–2098. 10.1007/s00726-017-2494-2. [DOI] [PubMed] [Google Scholar]
41.Pelle E, Maes D, Huang X, Frenkel K, Pernodet N, Yarosh DB, Zhang Q. 2012. Protection against UVB-induced oxidative stress in human skin cells and skin models by methionine sulfoxide reductase A. J Cosmet Sci 63:359–364. [PubMed] [Google Scholar]
42.Ezraty B, Aussel L, Barras F. 2005. Methionine sulfoxide reductases in prokaryotes. Biochim Biophys Acta 1703:221–229. 10.1016/j.bbapap.2004.08.017. [DOI] [PubMed] [Google Scholar]
43.Sreekumar PG, Hinton DR, Kannan R. 2011. Methionine sulfoxide reductase A: structure, function and role in ocular pathology. World J Biol Chem 2:184–192. 10.4331/wjbc.v2.i8.184. [DOI] [PMC free article] [PubMed] [Google Scholar]
44.Romsang A, Atichartpongkul S, Trinachartvanit W, Vattanaviboon P, Mongkolsuk S. 2013. Gene expression and physiological role of Pseudomonas aeruginosa methionine sulfoxide reductases during oxidative stress. J Bacteriol 195:3299–3308. 10.1128/JB.00167-13. [DOI] [PMC free article] [PubMed] [Google Scholar]
45.Quendera AP, Seixas AF, Dos Santos RF, Santos I, Silva JPN, Arraiano CM, Andrade JM. 2020. RNA-binding proteins driving the regulatory activity of small non-coding RNAs in bacteria. Front Mol Biosci 7:78–87. 10.3389/fmolb.2020.00078. [DOI] [PMC free article] [PubMed] [Google Scholar]
46.Shokeen S, Patel S, Greenfield TJ, Brinkman C, Weaver KE. 2008. Translational regulation by an intramolecular stem-loop is required for intermolecular RNA regulation of the par addiction module. J Bacteriol 190:6076–6083. 10.1128/JB.00660-08. [DOI] [PMC free article] [PubMed] [Google Scholar]
47.Jagodnik J, Chiaruttini C, Guillier M. 2017. Stem-loop structures within mRNA coding sequences activate translation initiation and mediate control by small regulatory RNAs. Mol Cell 68:158–170. 10.1016/j.molcel.2017.08.015. [DOI] [PubMed] [Google Scholar]
48.Cox MM, Battista JR. 2005. Deinococcus radiodurans: the consummate survivor. Nat Rev Microbiol 3:882–892. 10.1038/nrmicro1264. [DOI] [PubMed] [Google Scholar]
49.Slade D, Radman M. 2011. Oxidative stress resistance in Deinococcus radiodurans. Microbiol Mol Biol Rev 75:133–191. 10.1128/MMBR.00015-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
50.Henry C, Loiseau L, Vergnes A, Vertommen D, Mérida-Floriano A, Chitteni-Pattu S, Wood EA, Casadesús J, Cox MM, Barras F, Ezraty B. 2021. Redox controls RecA protein activity via reversible oxidation of its methionine residues. Elife 10:e63747. 10.7554/eLife.63747. [DOI] [PMC free article] [PubMed] [Google Scholar]
51.Xu G, Wang L, Chen H, Lu H, Ying N, Tian B, Hua Y. 2008. RecO is essential for DNA damage repair in Deinococcus radiodurans. J Bacteriol 190:2624–2628. 10.1128/JB.01851-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
52.Livak KJ, Schmittgen TD. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) method. Methods 25:402–408. 10.1006/meth.2001.1262. [DOI] [PubMed] [Google Scholar]
53.Mann M, Wright PR, Backofen R. 2017. IntaRNA 2.0: enhanced and customizable prediction of RNA-RNA interactions. Nucleic Acids Res 45(W1):W435–W439. 10.1093/nar/gkx279. [DOI] [PMC free article] [PubMed] [Google Scholar]
54.Ge P, Zhang S. 2013. Incorporating phylogenetic-based covarying mutations into RNAalifold for RNA consensus structure prediction. BMC Bioinformatics 14:142–110. 10.1186/1471-2105-14-142. [DOI] [PMC free article] [PubMed] [Google Scholar]
55.Chen Y, Xue D, Sun W, Han J, Li J, Gao R, Zhou Z, Zhang W, Chen M, Lin M, Wang J, Zuo K. 2019. sRNA OsiA stabilizes catalase mRNA during oxidative stress response of Deincoccus radiodurans R1. Microorganisms 7:422–434. 10.3390/microorganisms7100422. [DOI] [PMC free article] [PubMed] [Google Scholar]
56.Zhang H, Zhan Y, Yan Y, Liu Y, Hu G, Wang S, Yang H, Qiu X, Liu Y, Li J, Lu W, Elmerich C, Lin M. 2019. The Pseudomonas stutzeri-specific regulatory noncoding RNA, NfiS, targets katB mRNA encoding a catalase essential for optimal oxidative resistance and nitrogenase activity. J Bacteriol 201:e00334-19. 10.1128/JB.00334-19. [DOI] [PMC free article] [PubMed] [Google Scholar]
57.Huang C, Gan D, Luo F, Wan S, Chen J, Wang A, Li B, Zhu X. 2019. Interaction mechanisms between the NOX4/ROS and RhoA/ROCK1 signaling pathways as new anti-fibrosis targets of ursolic acid in hepatic stellate cells. Front Pharmacol 10:431–447. 10.3389/fphar.2019.00431. [DOI] [PMC free article] [PubMed] [Google Scholar]
58.Zamani B, Daneshzad E, Azadbakht L. 2019. Dietary total antioxidant capacity and risk of gastrointestinal cancers: a systematic review and meta-analysis of observational studies. Arch Iran Med 22:328–335. [PubMed] [Google Scholar]
59.Marottoli FM, Trevino TN, Geng X, Arbieva Z, Kanabar P, Maienschein-Cline M, Lee JC, Lutz SE, Tai LM. 2021. Autocrine effects of brain endothelial cell-produced human apolipoprotein E on metabolism and inflammation in vitro. Front Cell Dev Biol 9:668296–668306. 10.3389/fcell.2021.668296. [DOI] [PMC free article] [PubMed] [Google Scholar]
60.Vieira Dos Santos C, Cuiné S, Rouhier N, Rey P. 2005. The Arabidopsis plastidic methionine sulfoxide reductase B proteins. Sequence and activity characteristics, comparison of the expression with plastidic methionine sulfoxide reductase A, and induction by photooxidative stress. Plant Physiol 138:909–922. 10.1104/pp.105.062430. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

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

Supplementary Materials

Supplemental file 1

Fig. S1 to S7. Download aem.00038-22-s0001.pdf, PDF file, 0.7 MB^{(710.2KB, pdf)}

Data Availability Statement

TABLE 2.

Primers used in this study

Primer name	Sequence (5′–3′)^a	Purpose
P1-F	TAGTCCTGCACCCACTTCAGGGCCG	ΔmsrA construct
P1-R	TCGGTCTCCATGCGAGCTCGAATTCGATAGGGACCGGAAAGCGGATTTAA
P2-F	ATCCGCTTTCCGGTCCCTATCGAATTCGAGCTCGCATGGAGACCGA
P2-R	TCGGTCTGCCACAGCACCCGCATTATTTGCCGACTACCTTGGTGAT
P3-F	ACCAAGGTAGTCGGCAAATAATGCGGGTGCTGTGGCAGACCGAGGC
P3-R	GCTCGCACAGCGTGGTGTATCTGGC
P4-F	TCCAGCACACTGGCGGCCGTTACTAGTTCAGGAACCAGCCGTCGTCA	msrA_com construct
P4-R	GCATGCCTGCAGGTCGAATCGGATCCGCTCGGTGAAGACCCGGGGG	msrA_com construct
P5-F	CACCGCTTAACATCACCCCT	ΔDsrO construct
P5-R	TGCTCGATGAGTTTTTCTAAGGATAACTTTGTTTTACGAGCGTT
P6-F	CGTAAAACAAAGTTATCCTTAGAAAAACTCATCGAGCATCAAATG
P6-R	CCTGGTTCAAACAACGAAAACATGGAGACCGAGGGCCCTTGACA
P7-F	AAGGGCCCTCGGTCTCCATGTTTTCGTTGTTTGAACCAGGTGCA
P7-R	GGGGATGAACGGCATTCTCT
P8-F	TCCAGCACACTGGCGGCCGTTACTAGTCCACCCTCCCCGACGCCCTGCACG	DsrO_com construct
P8-R	GCATGCCTGCAGGTCGAATCGGATCCCGCCGTCCAGCAAACAAAAAACG	DsrO_com construct
msrA RT-F	CGCCGCACAATAGCAGCATG	qRT-PCR for msrA expression pattern
msrA RT-R	AGCGGTAATCGGGATTGGGCA	qRT-PCR for msrA expression pattern
DsrO RT-F	ACAGGTCTACGTTTCACTGTGGATA	qRT-PCR for msrA expression pattern
DsrO RT-R	CGCCGTCCAGCAAACAAAAAACGCT	qRT-PCR for msrA expression pattern
16s RT-F	ATTCCTGGTGTAGCGGTG	qRT-PCR for 16s rRNA
16s RT-R	ATTCCTGGTGTAGCGGTG	qRT-PCR for 16s rRNA
msrA SP1	TCAGCGGAGTTTGTCGCCGTAGTACTGGCGCAGCT	5′ RACE for msrA
msrA SP2	CCGAGCTGCTCGATCATTTCGCGGGCCGTCTGTTC
msrA SP3	AGCGGTAATCGGGATTGGGCACCGTACCGCCGATG
DsrO SP1	CGCCGTCCAGCAAACAAAAAACGCTGAACAGTCAT	5′ RACE for DsrO
DsrO SP2	CAGCAAACAAAAAACGCTGAACAGTCATGCTTGAAAATCCGC
DsrO SP3	AAAAACGCTGAACAGTCATGCTTGAAAATCCGCTTCAAAAGTT

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The underlining indicates the restriction enzyme cutting site of Spe1(P4/P8-F) and BamHI(P4/P8-R).

[B1] 1.Kaludercic N, Deshwal S, Lisa F. 2014. Reactive oxygen species and redox compartmentalization. Front Physiol 5:285–300. 10.3389/fphys.2014.00285. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B2] 2.Yang L, Mih N, Anand A, Park JH, Tan J, Yurkovich JT, Monk JM, Lloyd CJ, Sandberg TE, Seo SW, Kim D, Sastry AV, Phaneuf P, Gao Y, Broddrick JT, Chen K, Heckmann D, Szubin R, Hefner Y, Feist AM, Palsson BO. 2019. Cellular responses to reactive oxygen species are predicted from molecular mechanisms. Proc Natl Acad Sci USA 116:14368–14373. 10.1073/pnas.1905039116. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B3] 3.Cadet J, Davies KJA. 2017. Oxidative DNA damage & repair: an introduction. Free Radic Biol Med 106:100–111. 10.1016/j.freeradbiomed.2017.03.030. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B4] 4.Poetsch AR. 2020. The genomics of oxidative DNA damage, repair, and resulting mutagenesis. Comput Struct Biotechnol J 18:207–219. 10.1016/j.csbj.2019.12.013. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B5] 5.Yalcinkaya T, Uzilday B, Ozgur R, Turkan I, Mano J. 2019. Lipid peroxidation-derived reactive carbonyl species (RCS): their interaction with ROS and cellular redox during environmental stresses. Environ Exp Bot 165:139–149. 10.1016/j.envexpbot.2019.06.004. [DOI] [Google Scholar]

[B6] 6.Shari F, Ibrahim AY, Hassan JK, Al-Salman HNK. 2020. Study effect of sulfur containing amino acid supplementation on oxidative stress, inflammatory markers and serum lipid in relation to obese subjects. Int J Pharm Sci 12:362–370. 10.31838/ijpr/2020.12.01.062. [DOI] [Google Scholar]

[B7] 7.Ezraty B, Gennaris A, Barras F, Collet JF. 2017. Oxidative stress, protein damage and repair in bacteria. Nat Rev Microbiol 15:385–396. 10.1038/nrmicro.2017.26. [DOI] [PubMed] [Google Scholar]

[B8] 8.Liu Y, Zhang C, Wang Z, Lin M, Wang J, Wu M. 2021. Pleiotropic roles of late embryogenesis abundant proteins of Deinococcus radiodurans against oxidation and desiccation. Comput Struct Biotechnol J 19:3407–3415. 10.1016/j.csbj.2021.05.051. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B9] 9.Ranawat P, Rawat S. 2017. Radiation resistance in thermophiles: mechanisms and applications. World J Microb Biot 33:112–134. 10.1007/s11274-017-2279-5. [DOI] [PubMed] [Google Scholar]

[B10] 10.Adamec F, Farci D, Bína D, Litvín R, Khan T, Fuciman M, Piano D, Polívka T. 2020. Photophysics of deinoxanthin, the keto-carotenoid bound to the main S-layer unit of Deinococcus radiodurans. Photochem Photobiol Sci 19:495–503. 10.1039/D0PP00031K. [DOI] [PubMed] [Google Scholar]

[B11] 11.Jeong SW, Kim MK, Zhao L, Yang SK, Jung JH, Lim HM, Lim S. 2021. Effects of conserved wedge domain residues on DNA binding activity of Deinococcus radiodurans RecG helicase. Front Genet 12:634615–634626. 10.3389/fgene.2021.634615. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B12] 12.Lim S, Jung JH, Blanchard L, de Groot A. 2019. Conservation and diversity of radiation and oxidative stress resistance mechanisms in Deinococcus species. FEMS Microbiol Rev 43:19–52. 10.1093/femsre/fuy037. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B13] 13.Agapov A, Esyunina D, Kulbachinskiy A. 2019. Gre-family factors modulate DNA damage sensing by Deinococcus radiodurans RNA polymerase. RNA Biol 16:1711–1720. 10.1080/15476286.2019.1656027. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B14] 14.Cox MM, Keck JL, Battista JR. 2010. Rising from the ashes: DNA Repair in Deinococcus radiodurans. PLoS Genet 6:e1000815. 10.1371/journal.pgen.1000815. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B15] 15.Daly MJ. 2009. A new perspective on radiation resistance based on Deinococcus radiodurans. Nat Rev Microbiol 7:237–245. 10.1038/nrmicro2073. [DOI] [PubMed] [Google Scholar]

[B16] 16.Zhou Y, Shen P, Lan Q, Deng C, Zhang Y, Li Y, Wei W, Wang Y, Su N, He F, Xie Q, Lyu Z, Yang D, Xu P. 2017. High-coverage proteomics reveals methionine auxotrophy in Deinococcus radiodurans. Proteomics 17:1700072–1700074. 10.1002/pmic.201700072. [DOI] [PubMed] [Google Scholar]

[B17] 17.Ahmad S, Khan H, Shahab U, Rehman S, Rafi Z, Khan MY, Ansari A, Siddiqui Z, Ashraf JM, Abdullah SM, Habib S, Uddin M. 2017. Protein oxidation: an overview of metabolism of sulphur containing amino acid, cysteine. Front Biosci (Schol Ed) 9:71–87. 10.2741/s474. [DOI] [PubMed] [Google Scholar]

[B18] 18.Krisko A, Radman M. 2010. Protein damage and death by radiation in Escherichia coli and Deinococcus radiodurans. Proc Natl Acad Sci USA 107:14373–14377. 10.1073/pnas.1009312107. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B19] 19.Radman M. 2016. Protein damage, radiation sensitivity and aging. DNA Repair (Amst) 44:186–192. 10.1016/j.dnarep.2016.05.025. [DOI] [PubMed] [Google Scholar]

[B20] 20.Daly MJ, Gaidamakova EK, Matrosova VY, Vasilenko A, Zhai M, Leapman RD, Lai B, Ravel B, Li SM, Kemner KM, Fredrickson JK. 2007. Protein oxidation implicated as the primary determinant of bacterial radioresistance. PLoS Biol 5:e92. 10.1371/journal.pbio.0050092. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B21] 21.Chang RL, Stanley JA, Robinson MC, Sher JW, Li Z, Chan YA, Omdahl AR, Wattiez R, Godzik A, Matallana-Surget S. 2020. Protein structure, amino acid composition and sequence determine proteome vulnerability to oxidation-induced damage. EMBO J 39:e104523. 10.15252/embj.2020104523. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B22] 22.Achilli C, Ciana A, Minetti G. 2015. The discovery of methionine sulfoxide reductase enzymes: an historical account and future perspectives. Biofactors 41:135–152. 10.1002/biof.1214. [DOI] [PubMed] [Google Scholar]

[B23] 23.Fu X, Adams Z, Liu R, Hepowit NL, Wu Y, Bowmann CF, Moskovitz J, Maupin-Furlow JA. 2017. Methionine sulfoxide reductase A (MsrA) and its function in ubiquitin-like protein modification in Archaea. mBio 8:e01169-17. 10.1128/mBio.01169-17. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B24] 24.Boschi-Muller S. 2018. Molecular mechanisms of the methionine sulfoxide reductase system from Neisseria meningitidis. Antioxidants 7:131–142. 10.3390/antiox7100131. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B25] 25.Hopman CT, Speijer D, van der Ende A, Pannekoek Y. 2010. Identification of a novel anti-σE factor in Neisseria meningitidis. BMC Microbiol 10:164–176. 10.1186/1471-2180-10-164. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B26] 26.You C, Sekowska A, Francetic O, Martin-Verstraete I, Wang Y, Danchin A. 2008. Spx mediates oxidative stress regulation of the methionine sulfoxide reductases operon in Bacillus subtilis. BMC Microbiol 8:128–141. 10.1186/1471-2180-8-128. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B27] 27.Grimaud R, Ezraty B, Mitchell JK, Lafitte D, Briand C, Derrick PJ, Barras F. 2001. Repair of oxidized proteins. Identification of a new methionine sulfoxide reductase. J Biol Chem 276:48915–48920. 10.1074/jbc.M105509200. [DOI] [PubMed] [Google Scholar]

[B28] 28.Kim YK, Shin YJ, Lee WH, Kim HY, Hwang KY. 2009. Structural and kinetic analysis of an MsrA-MsrB fusion protein from Streptococcus pneumoniae. Mol Microbiol 72:699–709. 10.1111/j.1365-2958.2009.06680.x. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B29] 29.Kappler U, Nasreen M, McEwan A. 2019. New insights into the molecular physiology of sulfoxide reduction in bacteria. Adv Microb Physiol 75:1–51. 10.1016/bs.ampbs.2019.05.001. [DOI] [PubMed] [Google Scholar]

[B30] 30.Oertlin C, Lorent J, Murie C, Furic L, Topisirovic I, Larsson O. 2019. Generally applicable transcriptome-wide analysis of translation using anota2seq. Nucleic Acids Res 47:e70. 10.1093/nar/gkz223. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B31] 31.Levine RL, Berlett BS, Moskovitz J, Mosoni L, Stadtman ER. 1999. Methionine residues may protect proteins from critical oxidative damage. Mech Ageing Dev 107:323–332. 10.1016/s0047-6374(98)00152-3. [DOI] [PubMed] [Google Scholar]

[B32] 32.Bernhart SH, Hofacker IL, Will S, Gruber AR, Stadler PF. 2008. RNAalifold: improved consensus structure prediction for RNA alignments. BMC Bioinformatics 9:474. 10.1186/1471-2105-9-474. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B33] 33.Storz G, Opdyke JA, Zhang A. 2004. Controlling mRNA stability and translation with small, noncoding RNAs. Curr Opin Microbiol 7:140–144. 10.1016/j.mib.2004.02.015. [DOI] [PubMed] [Google Scholar]

[B34] 34.Buddeweg A, Sharma K, Urlaub H, Schmitz RA. 2018. sRNA₄₁ affects ribosome binding sites within polycistronic mRNAs in Methanosarcina mazei Gö1. Mol Microbiol 107:595–609. 10.1111/mmi.13900. [DOI] [PubMed] [Google Scholar]

[B35] 35.Wang L, Xu G, Chen H, Zhao Y, Xu N, Tian B, Hua Y. 2008. DrRRA: a novel response regulator essential for the extreme radioresistance of Deinococcus radiodurans. Mol Microbiol 67:1211–1222. 10.1111/j.1365-2958.2008.06113.x. [DOI] [PubMed] [Google Scholar]

[B36] 36.Qi HZ, Wang WZ, He JY, Ma Y, Xiao FZ, He SY. 2020. Antioxidative system of Deinococcus radiodurans. Res Microbiol 171:45–54. 10.1016/j.resmic.2019.11.002. [DOI] [PubMed] [Google Scholar]

[B37] 37.Shome A, Sarkhel R, Apoorva S, Nair SS, Chauhan TKS, Bhure SK, Mahawar M. 2020. Role of protein repair enzymes in oxidative stress survival and virulence of Salmonella. Ann Microbiol 70:55–65. 10.1186/s13213-020-01597-2. [DOI] [Google Scholar]

[B38] 38.Wang Y, Branicky R, Noë A, Hekimi S. 2018. Superoxide dismutases: dual roles in controlling ROS damage and regulating ROS signaling. J Cell Biol 217:1915–1928. 10.1083/jcb.201708007. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B39] 39.Brosnan JT, Brosnan ME, Bertolo RFP, Brunton JA. 2007. Methionine: a metabolically unique amino acid. Livestock Sci 112:2–7. 10.1016/j.livsci.2007.07.005. [DOI] [Google Scholar]

[B40] 40.Martínez Y, Li X, Liu G, Bin P, Yan W, Más D, Valdivié M, Hu CA, Ren W, Yin Y. 2017. The role of methionine on metabolism, oxidative stress, and diseases. Amino Acids 49:2091–2098. 10.1007/s00726-017-2494-2. [DOI] [PubMed] [Google Scholar]

[B41] 41.Pelle E, Maes D, Huang X, Frenkel K, Pernodet N, Yarosh DB, Zhang Q. 2012. Protection against UVB-induced oxidative stress in human skin cells and skin models by methionine sulfoxide reductase A. J Cosmet Sci 63:359–364. [PubMed] [Google Scholar]

[B42] 42.Ezraty B, Aussel L, Barras F. 2005. Methionine sulfoxide reductases in prokaryotes. Biochim Biophys Acta 1703:221–229. 10.1016/j.bbapap.2004.08.017. [DOI] [PubMed] [Google Scholar]

[B43] 43.Sreekumar PG, Hinton DR, Kannan R. 2011. Methionine sulfoxide reductase A: structure, function and role in ocular pathology. World J Biol Chem 2:184–192. 10.4331/wjbc.v2.i8.184. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B44] 44.Romsang A, Atichartpongkul S, Trinachartvanit W, Vattanaviboon P, Mongkolsuk S. 2013. Gene expression and physiological role of Pseudomonas aeruginosa methionine sulfoxide reductases during oxidative stress. J Bacteriol 195:3299–3308. 10.1128/JB.00167-13. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B45] 45.Quendera AP, Seixas AF, Dos Santos RF, Santos I, Silva JPN, Arraiano CM, Andrade JM. 2020. RNA-binding proteins driving the regulatory activity of small non-coding RNAs in bacteria. Front Mol Biosci 7:78–87. 10.3389/fmolb.2020.00078. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B46] 46.Shokeen S, Patel S, Greenfield TJ, Brinkman C, Weaver KE. 2008. Translational regulation by an intramolecular stem-loop is required for intermolecular RNA regulation of the par addiction module. J Bacteriol 190:6076–6083. 10.1128/JB.00660-08. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B47] 47.Jagodnik J, Chiaruttini C, Guillier M. 2017. Stem-loop structures within mRNA coding sequences activate translation initiation and mediate control by small regulatory RNAs. Mol Cell 68:158–170. 10.1016/j.molcel.2017.08.015. [DOI] [PubMed] [Google Scholar]

[B48] 48.Cox MM, Battista JR. 2005. Deinococcus radiodurans: the consummate survivor. Nat Rev Microbiol 3:882–892. 10.1038/nrmicro1264. [DOI] [PubMed] [Google Scholar]

[B49] 49.Slade D, Radman M. 2011. Oxidative stress resistance in Deinococcus radiodurans. Microbiol Mol Biol Rev 75:133–191. 10.1128/MMBR.00015-10. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B50] 50.Henry C, Loiseau L, Vergnes A, Vertommen D, Mérida-Floriano A, Chitteni-Pattu S, Wood EA, Casadesús J, Cox MM, Barras F, Ezraty B. 2021. Redox controls RecA protein activity via reversible oxidation of its methionine residues. Elife 10:e63747. 10.7554/eLife.63747. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B51] 51.Xu G, Wang L, Chen H, Lu H, Ying N, Tian B, Hua Y. 2008. RecO is essential for DNA damage repair in Deinococcus radiodurans. J Bacteriol 190:2624–2628. 10.1128/JB.01851-07. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B52] 52.Livak KJ, Schmittgen TD. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) method. Methods 25:402–408. 10.1006/meth.2001.1262. [DOI] [PubMed] [Google Scholar]

[B53] 53.Mann M, Wright PR, Backofen R. 2017. IntaRNA 2.0: enhanced and customizable prediction of RNA-RNA interactions. Nucleic Acids Res 45(W1):W435–W439. 10.1093/nar/gkx279. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B54] 54.Ge P, Zhang S. 2013. Incorporating phylogenetic-based covarying mutations into RNAalifold for RNA consensus structure prediction. BMC Bioinformatics 14:142–110. 10.1186/1471-2105-14-142. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B55] 55.Chen Y, Xue D, Sun W, Han J, Li J, Gao R, Zhou Z, Zhang W, Chen M, Lin M, Wang J, Zuo K. 2019. sRNA OsiA stabilizes catalase mRNA during oxidative stress response of Deincoccus radiodurans R1. Microorganisms 7:422–434. 10.3390/microorganisms7100422. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B56] 56.Zhang H, Zhan Y, Yan Y, Liu Y, Hu G, Wang S, Yang H, Qiu X, Liu Y, Li J, Lu W, Elmerich C, Lin M. 2019. The Pseudomonas stutzeri-specific regulatory noncoding RNA, NfiS, targets katB mRNA encoding a catalase essential for optimal oxidative resistance and nitrogenase activity. J Bacteriol 201:e00334-19. 10.1128/JB.00334-19. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B57] 57.Huang C, Gan D, Luo F, Wan S, Chen J, Wang A, Li B, Zhu X. 2019. Interaction mechanisms between the NOX4/ROS and RhoA/ROCK1 signaling pathways as new anti-fibrosis targets of ursolic acid in hepatic stellate cells. Front Pharmacol 10:431–447. 10.3389/fphar.2019.00431. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B58] 58.Zamani B, Daneshzad E, Azadbakht L. 2019. Dietary total antioxidant capacity and risk of gastrointestinal cancers: a systematic review and meta-analysis of observational studies. Arch Iran Med 22:328–335. [PubMed] [Google Scholar]

[B59] 59.Marottoli FM, Trevino TN, Geng X, Arbieva Z, Kanabar P, Maienschein-Cline M, Lee JC, Lutz SE, Tai LM. 2021. Autocrine effects of brain endothelial cell-produced human apolipoprotein E on metabolism and inflammation in vitro. Front Cell Dev Biol 9:668296–668306. 10.3389/fcell.2021.668296. [DOI] [PMC free article] [PubMed] [Google Scholar]

[B60] 60.Vieira Dos Santos C, Cuiné S, Rouhier N, Rey P. 2005. The Arabidopsis plastidic methionine sulfoxide reductase B proteins. Sequence and activity characteristics, comparison of the expression with plastidic methionine sulfoxide reductase A, and induction by photooxidative stress. Plant Physiol 138:909–922. 10.1104/pp.105.062430. [DOI] [PMC free article] [PubMed] [Google Scholar]

PERMALINK

A Novel Small RNA, DsrO, in Deinococcus radiodurans Promotes Methionine Sulfoxide Reductase (msrA) Expression for Oxidative Stress Adaptation

Yun Chen

Mingming Zhao

Mengli Lv

Min Lin

Jin Wang

Kaijing Zuo

Roles

ABSTRACT

INTRODUCTION

RESULTS

msrA expression in D. radiodurans is strongly induced by oxidative stress.

FIG 1.

Deletion of the msrA gene reduces the total antioxidant capacity and tolerance to H2O2 of D. radiodurans.

FIG 2.

DsrO is associated with oxidative stress, and its mutation reduces the tolerance of D. radiodurans to H2O2.

FIG 3.

FIG 4.

DsrO directly stabilizes msrA expression by base pairing within the coding sequence.

FIG 5.

msrA targeted by DsrO forms a stem-loop structure that is required for msrA mRNA stability and protein translation.

FIG 6.

msrA gene expression is transcriptionally regulated by the DrRRA transcription factor.

FIG 7.

DISCUSSION

FIG 8.

MsrA in D. radiodurans R1 primarily functions under oxidative stress.

msrA mRNA stability is crucial for the antioxidant capacity of D. radiodurans.

MATERIALS AND METHODS

Bacterial strains and plasmids.

TABLE 1.

H2O2 stress treatment and oxidative tolerance phenotype analysis.

Total RNA extraction and Northern blotting.

Rapid amplification of the 5′ end (RACE) of the DsrO gene.

Real-time quantitative PCR (qRT-PCR).

Bioinformatics analysis.

Microscale thermophoresis analysis.

Total antioxidant capacity, reactive oxygen (ROS) level, and intracellular carbonylation level analysis.

Stability measurement of msrA mRNA.

Western blot analysis.

Enzymatic activity analysis of MsrA.

Electrophoretic mobility shift assay (EMSA).

Data availability.

TABLE 2.

ACKNOWLEDGMENTS

Footnotes

Contributor Information

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

TABLE 2.

ACTIONS

PERMALINK

RESOURCES

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Links to NCBI Databases

Deletion of the msrA gene reduces the total antioxidant capacity and tolerance to H₂O₂ of D. radiodurans.

DsrO is associated with oxidative stress, and its mutation reduces the tolerance of D. radiodurans to H₂O₂.

H₂O₂ stress treatment and oxidative tolerance phenotype analysis.