Redox controls RecA protein activity via reversible oxidation of its methionine residues

An E. coli strain derivative lacking all peroxide-scavenging activities (catalases KatE and KatG, and peroxidase AhpC), referred to as Hpx^-, accumulates micromolar amounts of H₂O₂ (Seaver and Imlay, 2001). We found the Hpx^- msrA msrB strain to have significantly reduced viability upon UV stress compared to the Hpx^- parental strain with around 1.4 log difference at 50 and 60 J/m² (Figure 1A). UV defect is a hallmark of DNA recombination and/or repair defect. E. coli fights UV-induced DNA damages by both a RecA-dependent homologous recombination (HR) pathway and a RecA-independent UvrABCD-dependent nucleotide excision repair (NER) pathway. Therefore, we deleted either the recA or the uvrA gene in the Hpx^- background. In fact, the recA mutation proved to be lethal in the Hpx^- background in aerobic conditions as previously reported (Park et al., 2005). In contrast, the Hpx^- uvrA strain exhibited no phenotype, presumably because the HR pathway was sufficient to afford UV resistance. We then used Hpx^- uvrA strain to delete both msrA and msrB genes. The resulting Hpx^- msrA msrB uvrA strain showed hypersensitivity to O₂, similar to the Hpx^- recA strain (Figure 1B). Altogether, these observations suggested that in the presence of a higher-than-normal level of H₂O₂ RecA protein might be damaged by oxidation. MsrA/B thus could be essential for maintaining a level of functional reduced RecA above a threshold value. Consistently, we observed that high recA gene dosage suppressed UV sensitivity of the Hpx msrA msrB mutant (Figure 1C) and the O₂ sensitivity of the Hpx^- msrA msrB uvrA (Figure 1D).

Figure 1

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RecA is a multifunctional protein serving in HR and SOS response regulation (Clark and Margulies, 1965; Slilaty and Little, 1987). We used P1 phage-mediated transduction (Miller, 1972) to assess RecA-dependent HR efficiency under oxidative stress. When using the Hpx^- msrA msrB strain as a recipient, the transductant frequency dropped tenfold compared with the parental MsrA/B-proficient strain (Figure 2A). Transductant frequency was restored to the parental strain level by bringing in either plasmid-borne msrA or msrB genes (Figure 2A). Next, we investigated the influence of MsrA/B on the RecA-dependent SOS response. We used a recA-gfp gene fusion to monitor the SOS response. The Hpx^- strain exhibited enhanced SOS induction compared with the wild-type (WT) strain (Figure 2B). This was consistent with a previous work reporting increased DNA damage level in the Hpx^- background (Courcelle et al., 2001; Mancini and Imlay, 2015). When msrA msrB deletions were introduced into the Hpx^- background, the level of SOS induction decreased and was similar to that observed in the WT strain (Figure 2B). The well-known SOS-inducing compound mitomycin C (MMC) was then added. The Hpx^- strain exhibited an enhanced level of recA-gfp expression compared with the WT (Figure 2B). In contrast, deleting msrA msrB genes in the Hpx^- strain impaired MMC induction of recA-gfp (Figure 2B). Altogether, these observations showed that under oxidative stress MsrA/B maintains a level of reduced functional RecA necessary to carry out both efficient recombination and SOS regulation.

Figure 2

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We then characterized the effect of oxidative agents on the RecA protein. First, purified RecA protein was treated with hydrogen peroxide (H₂O₂) (50 mM) for 2 hr and analyzed by mass spectrometry. On the nine Met residues present in RecA, four could be identified within peptides detectable by mass spectrometry after peptidase digestion. In contrast, peptides containing Met28 and Met197 were too small to be detected. Likewise, Met35, Met58, Met164, and Met202 were found to have around 20% of oxidation (Figure 3A). This basal level of oxidation could be explained by the use of in-gel digestion, a technique known to cause Met oxidation (Klont et al., 2018), prior to mass spectrometry analysis. After H₂O₂ treatment, the level of oxidation increased to approximately 90% at Met164- and Met202-containing positions (Figure 3A). Met35 and Met58, being part of the same peptide, we were unable to differentiate the oxidation level of each residue separately. Importantly, no Cys residue was found to be modified. Second, the RecA oxidized above, referred to as RecAox, was treated with MsrA/B enzymes in the presence of 1,4-dithiothreitol (DTT) to yield to RecArep. Mass spectrometry analysis of RecArep revealed a drastic decrease in its content of Met-O residues (Figure 3A). As explained above, gel electrophoresis per se might have induced oxidation of Met residues, and therefore, one must be cautious when considering the quantitation of Met-O as their source might be multiple. However, besides allowing us to identify those Met residues that are sensitive to oxidation and eventually can be reduced by the MsrA/B enzymes, these in vitro characterizations allowed us to guide our in vivo studies presented later in this article. Third, we used a gel shift assay that detects Met-O residues in a polypeptide as Met-O-containing polypeptides run slower than their reduced counterpart on SDS-polyacrylamide gel electrophoresis (SDS-PAGE) (Gennaris et al., 2015). By western blot, we observed that RecA produced in a msrA msrB mutant exposed to HOCl treatment run slower than RecA produced in a WT strain or in a msrA msrB mutant that has not been exposed to HOCl (Figure 3B). These results showed that RecA is subject to oxidation both in vitro and in vivo, and that in both contexts MsrA/B allows reduction of Met-O residues present in RecAox.

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By using electron microscopy (Lusetti et al., 2003b), we observed that, in the presence of single-strand binding protein (SSB), RecA formed extended filaments on circular single strand (css) DNA (Figure 3—figure supplement 1). RecAox was unable to do so (Figure 3C), whereas RecArep recovered part of its ability to form nucleoprotein filaments in the presence of SSB (Figure 3—figure supplement 1). Using enzymatic assays, we showed that, as previously reported (Kowalczykowski and Krupp, 1987), RecA exhibited DNA-dependent ATPase activity in the presence of SSB with an apparent kcat of 25.22 µM⋅min⁻¹. In contrast, RecAox was unable to hydrolyze ATP (SSB) with an apparent kcat of 0.26 µM⋅min⁻¹, whereas RecArep recovered a significant level of activity with an apparent kcat of 9.66 µM⋅min⁻¹ (Figure 3D). RecA is also known to promote DNA strand exchange (Cox and Lehman, 1982; Lindsley and Cox, 1990). We compared the capacities of RecA, RecAox, and RecArep to promote strand exchange between cssDNA and linear double strand (lds) DNA in the presence of SSB. Over time, formation of nicked circular product (N.P.) was observed in the RecA-containing sample but not in the RecAox-containing reaction (Figure 3—figure supplement 1). RecArep was also able to promote DNA strand exchange, though with reduced efficiency compared to RecA (Figure 3—figure supplement 1). Together, our results show that RecAox protein cannot catalyze HR, which MsrA/B is partially able to restore.

Met35, which is targeted by oxidation, is located at the monomer–monomer interface in the RecA proto-filament (Figure 4A; McGrew and Knight, 2003; Chen et al., 2008). We replaced Met35 by either Gln (Q), a mimetic of Met-O, to test the consequences of oxidizing position 35, or Leu (L), to test the physicochemical constraints prevailing at that position (Figure 4B; Drazic et al., 2013). We reasoned that the variant containing Leu would inform us on the physicochemical constraints prevailing at position 35, and the variant containing Gln would inform us specifically on the effect of oxidation of Met35. To test activities of RecA variants, we used sensitivities to MMC and O₂ as well as induction of the SOS sfiA gene as readouts. Expression of RecA^M35L and RecA^M35Q variants was unable to rescue hypersensitivities of ∆recA and Hpx^- ∆recA strains to MMC and O₂, respectively (Figure 4C). Moreover, the SOS response was abrogated in the ∆recA strain harboring RecA^M35L and RecA^M35Q (Figure 4D–F). We performed biochemical analyses of the RecA^M35L variant and found that it had lost ATPase activity, nucleofilament formation, and DNA strand exchange capacities (Figure 5; Figure 5—figure supplement 1). Altogether, these studies indicated that position 35 is functionally important, with little tolerance for substitution at this position. Surprisingly, such an apparently essential role of Met35 was overlooked in the previous multiple mutagenesis studies (McGrew and Knight, 2003). Overall, these results did not allow us to evidence specific effect of oxidation of Met35 residue, besides the fact that it would be, as any other modification of that position, highly detrimental for both DNA repair and SOS regulation.

Figure 4

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Met164, identified above as a target of oxidation (Figure 3A), is located in the L1 region that interacts with DNA (Figure 4A; McGrew and Knight, 2003; Chen et al., 2008). Applying the same logic as for the study of position 35, we carried out phenotypic analyses to investigate the functionality of Gln- or Leu-containing variants at position 164. Expression of the RecA^M164Q variant failed to complement hypersensitivity to MMC of ∆recA strain (Figure 4C). In contrast, expression of the RecA^M164L variant allowed full complementation (Figure 4C). Similarly, the Hpx^- ∆recA strain was hypersensitive to O₂ when expressing RecA^M164Q but fully viable when expressing RecA^M164L (Figure 4C). The SOS response was followed by monitoring sfiA::lacZ expression, RecA steady-state level and LexA cleavage by immunoblotting, and cellular filamentation by phase contrast microscopy. A strain synthesizing RecA^M164L exhibited WT-like features of MMC-induced SOS response within all tests listed above (Figure 4D–F). Surprisingly, a strain synthesizing RecA^M164Q showed constitutive high levels of sfiA::lacZ expression, a high steady-state level of RecA, decreased levels of LexA, and strong cellular filamentation even in the absence of MMC (Figure 4D–F). This phenotype was surprising and somehow echoed the long-studied recA730 allele (RecA^E38K) as both alleles turn on the SOS response constitutively (Witkin et al., 1982). In contrast, recA730 bears no phenotype regarding MMC sensitivity (Figure 6A). In order to better compare both alleles, we introduced the recA-M164Q allele in the chromosome. The resulting strain recapitulated all the phenotypes observed with a plasmid encoded RecA^M164Q version (Figure 6A, B). First, it showed the expression of the sfiA::lacZ fusion even in the absence of MMC, that is, it was constitutive for SOS activation. Second, it proved to be sensitive to MMC treatment, indicating that RecA^M164Q protein is deficient in recombination-mediated repair. Then, we asked whether making that Met164 position refractory to oxidation would be beneficial or detrimental. For this, we introduced the recA-M164L allele in the chromosome and ran a growth competition assay between the WT and recA-M164L-containing strains. Under aerobic condition, the recA-M164L containing strain showed a clear advantage over the WT strain as, after a 120 hr growth, the population of the mutant counted for 70% of the total (Figure 7A, B). In contrast, under anaerobic condition, both strains grew at the same rate and a steady 50% ratio of both population was kept throughout the growth (Figure 7A, B). This experiment showed that preventing oxidation of Met164 position provides E. coli with a better fitness.

Figure 6

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Figure 7

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Taken together, these results strongly show that oxidation of Met164 modifies both repair and regulatory properties of RecA. Unexpectedly, oxidation of Met164 constitutively activates the SOS system, yet the consequences appear to be physiologically deleterious for E. coli.

We conducted biochemistry-based studies to investigate the mechanistic basis of the SOS-constitutive and repair-deficient phenotype conferred by the RecA^M164Q variant. We purified RecA^M164Q and added it along with SSB protein to cssDNA (Lusetti et al., 2003b; Weinstock et al., 1981; Cox and Lehman, 1982; Kowalczykowski and Krupp, 1987; Lindsley and Cox, 1990). We observed the formation of extended nucleoprotein filaments of similar or identical size as those obtained with WT RecA (Figure 8A). Surprisingly, SSB was not required. In the absence of SSB, RecA^M164Q produced normal-length nucleoprotein filaments, whereas RecA^wt formed only short filaments (Figure 8A). Next, enzymatic assays revealed that SSB was dispensable to facilitate ATPase activity of the RecA^M164Q variant in the presence of cssDNA from M13 phage, and the apparent kcat of RecA^M164Q was 21.75 µM⋅min⁻¹ compared with the 10.78 µM⋅min⁻¹ observed for the RecA^wt (Figure 8B). Because at such concentration SSB is known to stimulate RecA-promoted nucleoprotein filament formation by melting DNA secondary structures (Kowalczykowski and Krupp, 1987), these observations indicated that the RecA^M164Q variant displayed an enhanced filament propagation capacity, pointing to a potential hyper-reactivity of RecA toward ssDNA. Electrophoresis mobility shift assay (EMSA) techniques showed that the RecA^M164Q variant binds cssDNA with the same affinity as RecA but fails to bind circular double strand (cds) nicked DNA (Figure 8—figure supplement 1). These observations provided a molecular explanation as to why changing Met164 for Q, by inference oxidizing Met164 in Met-O, enabled RecA to turn on the SOS response even in the absence of MMC-induced DNA damage. We observed that the RecA^M164Q variant was unable to initiate the DNA strand exchange, suggesting a defect in homologous pairing (Figure 8—figure supplement 1). The failure of the variant to promote DNA strand exchange in vitro accounts for the in vivo hypersensitivity to MMC exposure of a strain synthesizing the RecA^M164Q variant. Altogether, these biochemical investigations provided a detailed analysis of features that RecA lost upon oxidation of Met164 and provided us with a molecular explanation for both in vivo constitutive SOS activation and failure to repair MMC-inflicted damages.

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The results above revealed a high level of heterogeneity between sensitivity of the different Met residues to oxidative agents. Besides the fact that the consequences of oxidation varied depending upon which Met residue had been modified, Met residues also showed various intrinsic sensitivities to HOCl depending upon their position in the polypeptide. To complete our study, we investigated whether we could further classify the Met residue as a function of their sensitivity to increasing concentration of HOCl. We focused our attention on the four Met residues studied above, that is, Met35, 58, 164, and 202. Met58 proved to be totally resistant to oxidation (Figure 8D). This is consistent with the fact it is not solvent-exposed. Met202 was the more sensitive. Interestingly, mimetic oxidation variant RecA^M202Q has been previously studied (McGrew and Knight, 2003; Hörtnagel et al., 1999) and shown to exhibit properties close to WT, strongly supporting the notion that oxidation at that position would bear no effect on RecA activity. Met164 was extremely sensitive as well, although slightly less reactive than Met202, but far more sensitive than Met35 (Figure 8D). These results showed that Met residues are not equivalent in the face of a potent oxidative agent such as HOCl and that M164 is highly prone to oxidation.

All strains are listed in the Key resources table. All strains used for in vivo experiments are E. coli MG1655 derivatives. Strains were constructed by transduction from P1 grown on strains from Keio or LCB collections (Baba et al., 2006), or they were constructed as described by Datsenko and Wanner, 2000 protocol using lambda Red recombination. pCP20 was used to remove the cassette when it was required. All constructions were confirmed by PCR. For strains carrying spectinomycin (spec) resistance cassette, a derivative Datsenko–Wanner protocol was used. Briefly, an FRT-flanked spectinomycin resistance (spc) cassette was amplified using chromosomal DNA from LL401 (ara_p::erpA) strain as a template, 5-K7wanner-spc forward primer, and 3-K7wanner-spc reverse primer (Key Resources Table). The amplified spc cassette fragment was inserted into pGEMTeasy vector by TA cloning, yielding pGEMT-spc. recA::spc strain was then constructed in a one‐step inactivation, and a DNA fragment containing spc gene flanked with a 5′ and 3′ region bordering recA gene was amplified by PCR using pGEMT-spc cassette as a template and oligonucleotides 5wanner-recA and 3wanner-recA (Key Resources Table). Strain BW25113, carrying the pKD46 plasmid, was transformed by electroporation with the amplified fragment, and Spc-resistant colonies were selected. The replacement of chromosomal recA gene by spc gene was checked by PCR amplification. Phage P1 was used to transduce the mutation into MG1655 and Hpx^- strains, yielding recA::spc and Hpx^- recA::spc strains, respectively.

The ‘recA–gfp’ fusion gfp gene was introduced just after the recA gene without removing the stop codon of recA in the chromosome. The gfp gene has its own ribosome binding site, and the recA–gfp fusion is transcriptional. For construction of the single-copy recA-gfp+ fusion, a fragment containing the promoterless gfp+ gene and the chloramphenicol resistance cassette was amplified from the PprgH plasmid with primers 5-recA-GFP and 3-recA-K7Cat (Key Resources Table). All gfp+ constructs were first integrated into the chromosome of E. coli strain BW25113 by using the Lambda Red system. Phage P1 was then used to transduce the recA-gfp+ fusion into MG1655, Hpx^-, ΔmsrA ΔmsrB, and Hpx ΔmsrA ΔmsrB strains.

All plasmids used were sequenced to confirm their construction and are listed in the Key Resources Table. Plasmids with the recA wt or recA with methionine mutations were constructed by the two-step PCR fragments method using oligonucleotides listed in the Key Resources Table, constructs were sub-cloned in pGEMT-easy vector (Promega), and then plasmids were double digested by SacII/SpeI and cloned in pBBR-MC2LL*. Plasmids used for the suppression and complementation of msrA msrB deletions phenotypes are derivatives of pUC18 vector. Plasmid used for RecA^M35L and RecA^M164Q overexpression was constructed by the direct mutagenesis of the pEAW260 using modified Megaprimer protocol (Kirsch and Joly, 1998). The PCR program was adapted for the Phusion polymerase enzyme. In both cases, T7-pro was used as the upstream primer and the downstream primer was recA*-M35L-rev or recA*-M164Q-rev.

All chemicals used were purchased from Sigma, Euromedex, or Fisher Scientific. Antibiotics were used at the following concentrations: chloramphenicol (25 μg/mL), ampicillin (50 μg/mL), kanamycin (50 μg/mL), and spectinomycin (75 μg/mL).

Cells were grown in Lysogeny Broth (LB) medium; for solid medium, 15 g/L of agar was supplemented. In aerobic condition, cells were grown in flask with a 1/10 vol of the flask capacity at 37°C with shaking at 150 rpm. In anaerobic condition, cells were grown in Coy Chamber; the medium was preincubated at least 16 hr before the experiments.

The M13mp18 RFI and css forms were prepared as described (Haruta et al., 2003) or purchased from Biolabs. The M13mp18 lds was obtained by the digestion of the M13mp18 RFI with the BamHI enzyme or by the double digestion with the BglII and DraIII enzymes (use only for the EM experiment). The cds nicked DNA was obtained by the digestion of pEAW951 (pBR22 carrying 2000 bp of M13mp18) with the DNaseI following the protocol described by Shibata et al., 1981. Digest products were gel purified using Promega kit. All DNA concentrations are reported in terms of μM of nucleotides (μM/nt).

The E. coli MsrA and MsrB (Grimaud et al., 2001), RecA (Cox et al., 1981; Britt et al., 2011), and SSB (Lohman et al., 1986) proteins were purified as described previously. RecA^M35L and RecA^M164Q were purified following the same protocol as RecA wt (Cox et al., 1981; Britt et al., 2011). The concentrations of MsrA and MsrB were determined by the Bradford method using the Bio-Rad Protein assay kit. The concentrations of SSB and RecA proteins were determined using native extinction coefficients: ε = 2.23 × 10⁴ M⁻¹ cm⁻¹ for RecA proteins and ε = 2.38 × 10⁴ M⁻¹ cm⁻¹ for SSB. All proteins were tested for endo- and exonuclease contamination. DNA degradation of cds, lds, and cssDNA by the proteins prep was assayed. No contaminating endo- or exonuclease activity was detected.

WT and ∆msrA ∆msrB mutants were grown aerobically at 37°C with shaking (150 rpm) in 10 mL of LB medium in 100-mL flasks. At A_600nm ≈0.5, cells were subjected to NaOCl treatment (1.75 or 2 mM). Cells were then incubated at 37°C with shaking for 2 hr. Samples were precipitated with trichloroacetic acid (TCA), and the pellets were then washed with ice-cold acetone, suspended in SB buffer, heated at 95°C, and loaded on a 12% SDS-PAGE gel for immunoblot analysis using anti-RecA antibodies (Abcam 63797). The protein amounts loaded were standardized by taking into account the A_600nm values of the cultures.

RecA oxidation was performed in 1× RecA buffer (25 mM Tris-acetate, 10 mM magnesium acetate, 3 mM potassium glutamate and 5% [w/v] glycerol), 50 mM hydrogen peroxide (H₂O₂) was added to 1.05 µM RecA, and the mixture was incubated for 2 hr at 37°C. Oxidation reaction was stopped by adding 400 U/mL catalase bovine, and the mix was incubated for 45 min at room temperature to remove the H₂O₂. The non-oxidized RecA was treated similarly except that nuclease-free water was used instead of H₂O₂. MsrA/B reparation assay was performed using aliquot of RecA oxidized, 2 µM MsrA, 10 µM MsrB, and 50 mM DTT (used for the enzyme recycling activity of Msr) were added, and the reaction was incubated 2 hr at 37°C to allow the reparation. In control, non-oxidized RecA was treated similarly except that RecA conservation buffer was used instead of MsrA and MsrB. Also, 2 µL of each reaction was mixed with 3 µL of cracking buffer heated for 10 min and run on 12% Tris glycine SDS-PAGE. Gels were stained with Coomassie Blue, the protein bands were cut and analyzed in-gel digestion, and mass spectrometric analysis was done at the Mass Spectrometry Facility (Biotechnology Center, University of Wisconsin-Madison).

The digestion was performed as outlined on the website: http://www.biotech.wisc.edu/ServicesResearch/MassSpec/ingel.htm. In short, Coomassie Blue R-250 stained gel pieces were de-stained twice for 5 min in MeOH/H₂O/NH₄HCO₃(50%:50%:100 mM), dehydrated for 5 min in ACN/H₂O/NH₄HCO₃(50%:50%:25 mM), then once more for 1 min in 100% ACN, dried in a Speed-Vac for 2 min, reduced in 25 mM DTT (dithiotreitol in 25 mM NH₄HCO₃) for 30 min at 52°C, alkylated with 55 mM IAA (iodoacetamide in 25 mM NH₄HCO₃) in darkness at room temperature for 30 min, washed twice in H₂O for 30 s, equilibrated in 25 mM NH₄HCO₃ for 1 min, dehydrated for 5 min in ACN/H₂O/NH₄HCO₃(50%:50%:25 mM), and then once more for 30 s in 100% ACN, dried again and rehydrated with 20 μL of trypsin solution (10 ng/μL trypsin Gold [Promega] in 25 mM NH₄HCO₃/0.01% ProteaseMAX w/v [Promega]). Additionally, 30 μL of digestion solution (25 mM NH₄HCO₃/0.01% ProteaseMAX w/v [Promega]) was added to facilitate complete rehydration and excess overlay needed for peptide extraction. The digestion was conducted for 3 hr at 42°C. Peptides generated from digestion were transferred to a new tube and acidified with 2.5% trifluoroacetic acid (TFA) to 0.3% final. Degraded ProteaseMAX was removed via centrifugation (max speed, 10 min) and the peptides solid phase extracted (ZipTip C18 pipette tips Millipore, Billerica, MA).

Peptides were eluted off the C18 SPE column with 1 μL of acetonitrile/H₂O/TFA (60%:40%:0.2%) into 1.5 mL Protein LoBind tube (Eppendorf), 0.5 µL was deposited onto the Opti-TOF 384-well plate (Applied Biosystems, Foster City, CA), and re-crystallized with 0.4 µL of matrix (5 mg/mL α-cyano-4hydroxycinnamic acid in acetonitrile/H₂O/TFA [60%:40%:0.1%]). Peptide Mass Fingerprint analysis was performed on a 4800 matrix-assisted laser desorption/ionization-time of flight-time of flight mass spectrometer (Applied Biosystems, Foster City, CA). In short, peptide fingerprint was generated scanning 700–4000 Da mass range using 1000 shots acquired from 20 randomized regions of the sample spot at 3600 intensity of OptiBeam on-axis Nd:YAG laser with 200 Hz firing rate and 3–7 ns pulse width in positive reflectron mode. Fifteen most abundant precursors, excluding trypsin autolysis peptides and sodium/potassium adducts, were selected for subsequent tandem MS analysis, where 1200 total shots were taken with 4200 laser intensity and 1.5 kV collision-induced activation using air. Post-source decay fragments from the precursors of interest were isolated by timed-ion selection and reaccelerated into the reflectron to generate the MS/MS spectrum. Raw data was deconvoluted using GPS Explorer software and submitted for peptide mapping and MS/MS ion search analysis with an in-house licensed Mascot search engine (Matrix Science, London, UK) with cysteine carbamidomethylation as fixed modification plus methionine oxidation and asparagine/glutamine deamidation as variable modifications. The relative Met-O % was calculated using the values of the isotope cluster area of the initial mass, the mass +16 and +32 Da for each peptide. The mean values of the biological triplicates are shown in the histogram plot.

RecA oxidation was performed in buffer Tris-HCl 10 mM pH 7.4, DTT 0.1 mM, EDTA 0.01 mM, and glycerol 5%. RecA was incubated with varying molar excess of HOCl (Acros Organics) for 30 min at 25°C. Following incubation, reaction was stopped with 50 mM of L-methionine. For the identification of peptides, samples were reduced and alkylated before digestion of O/N with trypsin at 30°C in 50 mM NH₄HCO₃(pH 8.0). Peptides were dissolved in solvent A (0.1% TFA in 2% ACN), directly loaded onto reversed-phase pre-column (Acclaim PepMap 100, Thermo Scientific), and eluted in backflush mode. Peptide separation was performed using a reversed-phase analytical column (Acclaim PepMap RSLC, 0.075 × 250 mm, Thermo Scientific) with a linear gradient of 4–36% solvent B (0.1% FA in 98% ACN) for 36 min, 40–99% solvent B for 10 min, and holding at 99% for the last 5 min at a constant flow rate of 300 nL/min on an EASY-nLC 1000 UPLC system. The peptides were analyzed by an Orbitrap Fusion Lumos tribrid mass spectrometer (Thermo Fisher Scientific). The peptides were subjected to NSI source followed by tandem mass spectrometry (MS/MS) in Fusion Lumos coupled online to the UPLC. Intact peptides were detected in the Orbitrap at a resolution of 120,000. Peptides were selected for MS/MS using HCD setting at 30; ion fragments were detected in the Orbitrap at a resolution of 30,000. A targeted mass list was included for the 11 theoretical peptide sequences (see Table 1), considering potential oxidation of Met and charges states +2 or +3. The electrospray voltage applied was 2.1 kV. MS1 spectra were obtained with an AGC target of 4E5 ions and a maximum injection time of 50 ms, and targeted MS2 spectra were acquired with an AGC target of 2E5 ions and a maximum injection time of 60 ms. For MS scans, the m/z scan range was 350–1800. The resulting MS/MS data was processed and quantified using Skyline (version 19.1.0.193). Trypsin was specified as cleavage enzyme allowing up to two missed cleavages, four modifications per peptide, and up to three charges. Mass error was set to 10 ppm for precursor ions and 0.05 Da for fragment ions. Oxidation on Met was considered as variable modification.

A modified alcian method was used to visualize RecA filaments. Activated grids were prepared as described previously (Lusetti et al., 2003b). Samples for electron microscopy analysis were prepared as follows. All incubations were carried out at 37°C. Native RecA, RecAox, RecArep, RecAox, RecA^M164Q, or RecA^M35L (6.7 µM) was preincubated with 20 µM M13mp8 circular ssDNA or with ldsDNA in a 1× RecA buffer for 10 min. An ATP regeneration system (10 units/mL creatine phosphokinase and 12 mM phosphocreatine) was also included in the preincubation along with 1 mM DTT. ATP (3 mM) with or without 2 µM SSB protein was added, and the reaction was incubated for an additional 10 min after which ATPγS was added to 3 mM to stabilize the filaments and further incubated for an additional 3 min. In EM experiments involving linear dsDNA (lds), the experiment procedures were done without SSB proteins. In order to make sure that the filaments observed were not formed by RecA joining on its own, the control experiment was done without DNA, and no RecA filament was observed (data not shown).

The reaction mixture was diluted to a final DNA concentration of 0.4 ng/µL in 10 mM HEPES, 200 mM ammonium acetate, 10% glycerol (pH adjusted to 7.5), and adsorbed on alcian grid for 3 min. The grid was then touched to a drop of the above buffer followed by floating on a drop of the same buffer for 1 min. The sample was then stained by touching to a drop of 5% uranyl acetate followed by floating on a fresh drop of the same solution for 30 s. Finally, the grid was washed by touching to a drop of double-distilled water followed by immersion in two 10-mL beakers of double-distilled water. After the sample was dried, it was rotary-shadowed with platinum. This protocol is designed for visualization of complete reaction mixtures, and no attempt was made to remove unreacted material. Although this approach should yield results that give a true insight into reaction components, it does lead to samples with a high background of unreacted proteins. Imaging and photography were carried out with a Hitachi 7600 Transmission Electron Microscope equipped with a Hamamatsu ORCA HR camera. Digital images of the nucleoprotein filaments were taken at ×15,000 magnification except for SSB DNA molecules shown in the inset where the magnification is ×60,000. Imaging was also done using TECNAI G2 12 Twin Electron Microscope (FEI Co.) equipped with a 4k × 4k Gatan Ultrascan CCD camera at ×15,000 magnification.

For the measurement, filaments were randomly selected. For filament measurement on cssDNA, 10 circular filaments each from RecA wt + SSB, RecA wt No SSB, RecA M164Q + SSB, and RecA M164Q No SSB were selected. For filament measurement on ldsDNA, 54 and 53 linear filaments of respectively RecAwt and RecA M164Q were selected. In all experiments, filaments were measured three times using Metamorph analysis software. The average length of each filament was calculated in μm. The scale bar was used as a standard to calculate the number of pixels per μm.

A coupled enzyme spectrophotometric assay was used to measure the DNA-dependent RecA ATPase activity as described (Lindsley and Cox, 1990). All assays were carried out using a Varian Cary 300 dual-beam spectrophotometer equipped with a temperature-controlled 12-cell changer. The cell path length is 1 cm, and the bandwidth was 2 nm. All reactions were carried out at 37°C in 1× RecA buffer, an ATP regeneration system (10 U/mL pyruvate kinase, 3 mM phosphoenolpyruvate), a coupled detection system (10 U/mL lactate dehydrogenase, 3 mM NADH), 3 mM ATP, 3 µM of RecA proteins, no or 0.5 µM of SSB protein, and 5 µM of M13mp18 cssDNA. Graphs represent the mean rate of the ATP hydrolysis, and errors represent the standard deviation from the biological triplicates. The apparent kcat was calculated as described previously (Piechura et al., 2015) by dividing the observed ATP hydrolysis rate by the number of RecA binding sites (i.e. the concentration in nucleotide divided by 3).

Three-strand exchange reactions were carried out in 1× RecA buffer as described (Lusetti et al., 2003a). All reactions contained an ATP regeneration system (10 unit/mL pyruvate kinase and 2.5 mM phosphoenolpyruvate). Reactions were incubated at 37°C. RecA proteins (3.5 μM) were preincubated with 10 μM M13mp18 circular ssDNA for 10 min. Then, SSB protein (1 μM) and 3 mM ATP were added and incubated for an additional 10 min. Reactions were initiated by the addition of M13mp18 linear dsDNA to 10 μM. Aliquot of 10 μL was removed at the indicated time points. The reaction was stopped by the addition of 5 μL of a solution 3:2 Ficoll:SDS. Samples were subjected to electrophoresis on 0.8% on Tris-acetate EDTA (TAE) agarose gel, stained with ethidium bromide, and visualized with Typhoon FLA-9000 (GE Healthcare). Experiments were run in triplicate, and results of a representative experiment are shown.

Cells were grown in LB in anaerobic condition until OD₆₀₀: 0.3–0.5. Then, they were serial diluted by a factor of 10 in Phosphate-buffered saline (PBS buffer), and 100 μL of the adequate dilution was spread in duplicate on LB plates and incubated at 37°C for 16 hr, one duplicate in anaerobic condition and the other in aerobic condition. Ampicillin was added to the culture and to the plates medium for complementation/suppression assay.

Cells were grown in aerobic condition at 37°C until OD₆₀₀: 0.2–0.3 and serial diluted in PBS. Then, pure culture and dilutions from 10⁻¹ to 10⁻⁶ were spotted on LB plates in several replicates. Replicated plates were exposed to increasing UV doses with a Crosslinker (Startalinker, Stargene). Plates were incubated 16 hr at 37°C, and survival cells were counted to determine the viability at different UV doses.

Transductant frequency assay with P1 phage was used as reporter of HR. P1 used for the assay was prepared as follows: P1 was grown on lacZ::cat MG1655 derivatives in soft agar plate and incubated 16 hr at 30°C. The top of the plate was collected, chloroform was added, and the mixture was vortexed and incubated at room temperature for 10 min. Agar and P1 were separated by centrifugation, 10 min at 4500 rcf at 4°C. The upper layer containing the phage was collected and conserved at 4°C. P1 was serial diluted in PBS, and the titration of the phage was determined by spotting 10 μL of each dilution on MG1655 soft agar plates, the pfu/mL was between 10¹⁰ to 10¹¹. For the assay, strains were grown in aerobic condition in LB until OD₆₀₀≈1.0. Also, 50 mL of cells were centrifuged 10 min at 4500 rcf. Pellets were washed with 50 mL of PMC buffer (1× PBS, 10 mM MgSO₄, and 5 mM CaCl₂) followed by a second centrifugation of 10 min. Pellets were resuspended in 5 mL of PMC. Then, 1 mL of cells (≈ 2 × 10¹⁰) was infected with the P1 phage (≈ 10⁸, leading to a m.o.i. of around 0.005) and incubated at 37°C for 10 min without shaking. Also, 1 mL of non-infected cells was subject to the same protocol and used as negative control. Then, cells were centrifuged 3 min at 5500 rcf. Fresh LB supplemented with 10 mM Na citrate was used to resuspend cells pellets followed by 1 hr of incubation at 37°C with shaking. Then, 100 μL of cells was serial diluted in PBS and spotted on LB plates to determine the cfu/mL, and 100 μL of cells was spread on plates containing the adequate antibiotics. Experiments were at least replicated three times. The transductant frequency was calculated as follows: f – number of clones antibiotic-resistant per mL divided by the pfu per mL used for the infection. The differences and p-values of the transductant frequencies were determined by a Mann–Whitney statistical test. Values are represented in boxplot, and the median is indicated with a bold black line.

LB agar plates containing or not 1 μg/mL MMC were prepared. MG1655 recA::spc strain harboring various versions of recA mutated plasmids was streaked onto each plate followed by overnight incubation at 37°C. The growth of each strain was checked by scoring for colony formation. Hpx^- recA::spc strain was transformed with the same plasmids as cited above and tested for its ability to grow in the presence or absence of oxygen.

Microscopic examination of recA::spc strain cultures harboring various versions of recA mutated plasmids was realized using Motic BA310E 600 Biological Microscope. Light microscopy phase contrast imaging of E. coli cells in stationary phase cultured in LB were realized. E. coli cells carrying different recA mutated plasmids presented filamentous (RecA^M164Q) or normal (RecA, RecA^M35Q, RecA^M35L, RecA^M164L) morphologies. Experiments were run in triplicate, and results of a representative experiment are shown.

Transcriptional fusion sfiA-lacZ was used as reporters of the SOS expression level. This construction was a gift from B. Michel (ΔlacZ::cat sfiA::Mud lacZ cat-amp) (Grompone et al., 2004). It is important to note that strains carrying this construction are mutated for sfiA as lacZ gene cassette disrupted the sfiA gene, as indicated in the Key Resources Table. Strains carrying the sfiA-lacZ reporter were grown in exponential phase OD₆₀₀:0.2–0.4 and exposed or not to 0.25 µg/mL MMC for 3 hr. The sfiA expression level was determined by β-galactosidase assays as described (Miller, 1972).

The chromosomal recA-M164L was generated by lambda red recombination adapted from Blank et al., 2011. The recA-M164L sce-I::kan DNA fragment was amplified by PCR using primers (5for recA M164L SceI K7kan/3rev recA K7Kan) with plasmid pWRG100 as a template. The resulting PCR product was electroporated into MG1655 cells after 1 hr of lambda recombinase expression from pWRG99 plasmid. After 1 hr of phenotypic expression, the cells were plated on LB containing 25 μg/mL kanamycin. Selected colonies contained a recA-M164L::sce-I::kan cassette, and harboring the pWRG99 plasmid has been newly isolated at 30°C. The kanamycin cassette was then removed by counter selection using lambda red recombination to insert a oligonucleotide recA hybrid of M164L-Scarless-f and M164L-Scarless-rv. The recA-M164L hybrid product was then electroporated into MG1655 recA-M164L::sce-I::kan expressing lambda recombinase from pWRG99, and after 1 hr of phenotypic expression serially diluted and plated on NA containing 100 μg/mL ampicillin and 1 μg/mL anhydrotetracycline (aTc). pWRG99 also encoded for the meganuclease I-Sce-I under the control of an aTc-inducible promoter. The proper integration (positive Sce-I-resistant clones) of the recA-M164L mutational change was confirmed by diagnostic PCR and then sequenced.

Cell fitness was determined for each strain using a growth competition assay described by Henrikus et al., 2019. In general, this two-color colony assay is based on the color difference of Ara+ and Ara− colonies on tetrazolium arabinose indicator plates (TA plates). Ara– colonies typically are red colored, while Ara+ colonies are white. Ara+ and Ara– cells can be counted, and thus fitness in a mixed population of two strains can be assessed. In preparation for the assay, individual overnight cultures of Ara– and Ara+ cells were grown in 3 mL of LB at 37°C in a 50-mL conical tube with an inclination of 90°. The next day a mixed culture of Ara– and Ara+ cells was set up at a 1:1 ratio by volume. To start the experiment, 3 mL of medium was inoculated with 30 μL of the mixed culture and grown at 37°C in a 50-mL conical tubes with an inclination of 90°. Fitness was assessed over the period of 120 hr; cells were serial diluted in PBS at 0, 24, 48, 72, 96, and 120 hr. The dilutions were spread on plates containing TA plates and incubated at 37°C for 16 hr before counting. We performed this assay-competing Ara+ cells of recA-M164L allele with Ara– cells of the recAWT and vice versa.

To study the expression of recA in WT, Hpx^-, and Hpx^- ∆msrA ∆msrB backgrounds, overnight bacterial cultures were diluted 1:100 in LB and incubated at 37°C for 5 hr until early stationary phase. For SOS response induction, overnight cultures were diluted 1:100 in LB and incubated at 37°C for 1 hr (early exponential phase). MMC (Sigma) was then added to a final concentration of 0.25 μg/mL, and the cultures were incubated for four additional hours (early stationary phase) at 37°C. Cells were then diluted in PBS. Data acquisition and analysis were performed using a Cytomics FC500-MPL cytometer (Beckman Coulter, Brea, CA). Data were collected for 100,000 events per sample and analyzed with CXP and FlowJo 8.7 software.

E. coli ΔrecA sfiA::lacZ strains transformed with plasmid carrying either WT RecA or RecA mutants were grown overnight. The next day subcultures were made and grown at 37°C until the OD₆₀₀ reached 0.2. Cultures were split, and 0.25 µg/mL MMC was added or not for each. The cultures were grown further at 37°C, and 1 mL of culture from each strain was aliquoted at 3 hr. The culture aliquots were washed once in cold PBS. The RecA protein levels were normalized to bacterial growth and resolved in 4–20% SDS-PAGE. The resolved bands were blotted to nitrocellulose membranes and probed with anti-LexA (1∶5000) and anti-RecA (1∶20,000) antibodies. Goat anti-rabbit IgG-Alk Phos (Chemicon International) and anti-rabbit HRP (Promega) were used as the secondary antibody at 1∶10,000 dilutions to detect RecA and LexA proteins, respectively. Chemiluminescence detection was done using Amersham ECL western blotting kit and staining solution by using NBT/BCIP (Sigma). All the experiments were repeated at least three times for each RecA mutant, and representative result experiments are shown.

The ability of RecA proteins to bind DNA was assayed by EMSA. All reactions were realized in 1× RecA buffer supplemented with 1 mM DTT. Then, 20 μM of DNA, cssM13mp18 DNA, or pEAW951 nicked plasmid (cdsDNA) were incubated with RecA proteins at the indicated concentrations for 5 min at 37°C. Reactions were initiated by adding 3 mM of ATPγS followed by 30 min of incubation at 37°C. Reactions were stopped by adding 1:5 volume of glycerol EDTA dye (GED). Samples were submitted to electrophoresis in 0.8% TAE agarose gel, stained with ethidium bromide, and visualized with Typhoon FLA-9000 (GE Healthcare).

Mann–Whitney U tests were performed using the QI-Macros software (KnowWare International, Inc, Denver, CO).

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank the members of the Barras, Ezraty, Cox, and Casadesus groups for comments on the manuscript, advice, and discussions throughout the work. CH was supported by the Fondation pour la Recherche Médicale (FRM-FDT20150532554) and by AMidex. This work was supported by grants from Agence Nationale Recherche (ANR) (# ANR-METOXIC), CNRS (#PICS-PROTOX), Aix‐Marseille Université, Institut Pasteur, and the ANR-10-LABX-62-IBEID. Work from the Cox Lab was supported by grant GM32335 from the National Institute of General Medical Sciences (NIH). We thank the Wisconsin Veterinary Diagnostic Laboratory of the University of Wisconsin-Madison for allowing us to use their Hitachi 7600 TEM. We acknowledge the use of instrumentation supported by the UW MRSEC (DMR-1121288) and UW NSEC (DMR-0832760). We also thank M Modesti, B Michel, PL Moreau, C Jourlin-Castelli, L Espinosa, and JF Collet for helpful suggestions, reagents, and comments on the manuscript.

© 2021, Henry et al.

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