Mutant construction was performed according to the procedure developed by Arnaud et al. (2004). The msrAB ORF was interrupted by insertion of a non-polar spectinomycin resistance expression cassette, spc (Murphy, 1985) as follows. A DNA fragment of 1,413 bp encompassing the msrAB ORF was amplified from B. cereus genomic DNA by PCR with primers 5′-gaattcTCATGCCTTGAAAGTTACGG-3′ and 5′-agatctTTGGCGTAACGGTAATTGGT-3′, which contained EcoRI and BglII restriction sites, respectively. The amplified DNA fragment was cloned into pCRXL-TOPO (Invitrogen). The resulting pCRXLmsrAB plasmid was digested with StuI. A 1.5 kb SmaI fragment containing spc was purified from pDIA (Laouami et al., 2011) and ligated into StuI-digested pCRXLmsrAB. The resulting plasmid, pCRXLmsrABΔspc, was digested with EcoRI plus BglII. The msrABΔspc fragment was then subcloned into EcoRI/BglII sites of pMAD (Arnaud et al., 2004). This construct was used for B. cereus transformation (Omer et al., 2015). For complementation of the ΔmsrAB mutant with wild-type msrAB gene, the 1,413 bp EcoRI-BglII fragment was cloned into pHT304 (Arantes and Lereclus, 1991). MsrAB is under the control of its own promoter into pHT304-msrAB.

Wild-type B. cereus ATCC 14579 without its pBClin15 plasmid (Madeira et al., 2016a,b), its ΔmsrAB mutant and ΔmsrAB/pHT304msrAB complemented strains were grown in MOD medium supplemented with 30 mM glucose as the carbon source, as previously described (Madeira et al., 2016b). The inoculum was a sample of exponential subculture harvested by centrifugation, washed and diluted in fresh medium to obtain an initial optical density at 600 nm of 0.02. Three independent batch cultures (biological replicates) were carried out at 37°C for each strain.

B. cereus growth was monitored spectrophotometrically at 600 nm. The specific growth rate (μ) was determined using the modified Gompertz equation (Zwietering et al., 1990). Cells and filtered culture supernatants were harvested at the indicated growth stage as previously described (Madeira et al., 2015, 2016b). Exoproteins were immediately precipitated from the culture supernatant using trichloroacetic acid (TCA), as previously described, and stored at 4°C until analysis. The concentrations of substrate, and by-products in the filtered culture supernatants were determined with Enzytec Fluid kits purchased from R-Biofarm, as described by the manufacturer. Exoprotein concentration was determined by the Bradford protein assay (Pierce).

Protein extraction and subsequent digestion were performed as previously described (Madeira et al., 2015). Extracellular and intracellular proteins from the 27 samples (biological triplicates from the three time conditions for the wild-type, ΔmsrAB and ΔmsrAB/pHT304-msrAB strains) were resolved on NuPAGE® 4–12% Bis-Tris gels (Invitrogen) that were run for a short (about 3 mm) electrophoretic migration using NuPAGE MES supplemented with NPAGE antioxidant as the running buffer (Hartmann and Armengaud, 2014). This avoids artefactual protein oxidation. For each of the 54 protein samples, the whole protein content was extracted as a single polyacrylamide band. The bands were subjected to proteolysis with sequencing grade trypsin (Roche) following the ProteaseMAX protocol (Promega), as previously described (De Groot et al., 2009; Clair et al., 2010). NanoLC-MS/MS experiments were performed using an LTQ-Orbitrap XL hybrid mass spectrometer (ThermoFisher) coupled to an Ultimate 3000 nRSLC system (Dionex, ThermoFisher; Dedieu et al., 2011; Madeira et al., 2015).

MS/MS spectra were searched against an in-house polypeptide sequence database corresponding to an improved annotation of the B. cereus ATCC 14,579 genome (Madeira et al., 2016a). The MASCOT Daemon search engine (version 2.3.02; Matrix Science) was used to search tryptic peptides as previously described (Dupierris et al., 2009; Madeira et al., 2016a). The mass spectrometry proteomics data have been deposited in the ProteomeXchange Consortium (http://proteomecentral.proteomexchange.org) via the PRIDE partner repository (http://www.ebi.ac.uk/pride) with the dataset identifiers, PXD006169 and 10.6019/PXD006169 (exoproteome) and, PXD006205 and 10.6019/PXD006205 (cellular proteome).

Analyses of changes of peptides and proteins in terms of abundance were achieved by comparing the spectral counts of proteins after voom transformation of abundance values using the R package LIMMA (Ritchie et al., 2015), as previously described (Madeira et al., 2016b). Data were normalized using the trimmed mean of M-values (TMM), implemented in the R package edgeR (Robinson et al., 2010). For quantitative comparisons, data were filtered to have two valid values in at least two biological replicates. Since we were specifically interested in the comparison between wild-type, ΔmsrAB mutant and the complemented strain ΔmsrAB/pHT-msrAB, we conducted differential analysis between WT and ΔmsrAB, as well as ΔmsrAB and ΔmsrAB/pHT-msrAB, and WT and ΔmsrAB/pHT-msrAB, individually. Differential protein and peptide abundances between WT and ΔmsrAB, between ΔmsrAB and ΔmsrAB/pHT-msrAB, and between WT and ΔmsrAB/pHT-msrAB were considered significant at stringent p-values (≤0.01). The results are presented as log₂ fold-changes.

Total RNA was prepared as described previously (Omer et al., 2015). Real-time RT-PCR was performed using the iScript™ One-Step RT-PCR kit with SYBR® Green following the manufacturer's protocol (Biorad). The msrAB-specific primer pair used in this study was: 5′-TTCTGGTACACAGGTGGTC-3′ and 5′-AAAGCGTCCACTCTGCTCAA-3′. Gene expression was normalized by the ΔΔCT analysis. The 16s rDNA was used as the reference gene in the calculations. The 16S rDNA-specific primer pair was 5′-TCCAACTGATGGCGGAC-3′ and 5′-TCACGCCCAGATTCTTTTTGC-3′. Rapid amplification of 5′ complementary cDNA ends (5′RACE) was performed using the 5′/3′ RACE kit (Sigma). The msrAB specific primers SP1, SP2 and SP3 were: 5′-ATGTCCCGTCGTTTCTGAAC-3, 5′-TCAAATGGCGAAACCATACA-3′ and 5′-CCATACACCAGAAGCACCCT-3′, repectively.

Sigma's non-specific protease activity assay was used to determine the protease activity of filtered culture supernatant. In this assay, casein acts as a substrate. Tyrosine, which is released on hydrolysis of casein by proteases, is able to react with Folin-Ciocalteu's reagent to produce a blue chromophore. The quantity of this chromophore was measured by means of its absorbance value by spectrophotometry. Absorbance values generated by the activity of the protease were compared to a standard curve, which was generated on the basis of known quantities of tyrosine. From the standard curve, the activity of protease samples was determined in units, corresponding to the amount in micromoles of tyrosine equivalents released from casein per minute. Experiments were performed twice for each of the 27 filtered culture supernatants. Statistical differences were evaluated by the Student's t-test.

The survival of WT, ΔmsrAB mutant, and complemented ΔmsrAB mutant were determined as follows. After 24 h incubation at 37°C on glucose containing MOD medium, cultures were transferred to 4°C. An aliquot of each culture was collected before and after 1, 2, 3, 4, and 5 days of exposure to 4°C. Viable cells were determined by serial dilution of cultures in PBS, plating on LB agar, and incubation overnight (37°C). Experiments were performed in triplicate. Statistical differences were evaluated by the Student's t-test.

Genome analyses of B. cereus ATCC 14579 identified an ORF (BC_5436) encoding a cytoplasmic protein annotated as MsrAB (NP_835097). This predicted cytoplasmic protein is composed of 321 amino acids and has a molecular weight of 36,938 Da. MsrAB and its gene msrAB are strongly conserved in members of the B. cereus group (data not shown). We mapped the transcriptional start site of msrAB by 5′RACE. The transcriptional start site (G) was located 23 nt upstream of the translational start codon and was preceded by a region similar to σE consensus-35 (TAATATG) and -10 (CATACTG) boxes separated by 13 nt. Furthermore, msrAB appeared to be followed by an inverted repeat (ΔG° = 23.6 kcal/mol) that may a transcriptional terminator (Figure S1). This indicates that msrAB may be transcribed as a single unit. To determine whether there is any regulation of msrAB, mRNA levels were measured at early exponential (EE), late exponential (LE) and stationary (S) growth phases. Figure 1 shows that there was about a 30-fold increase in msrAB expression for cells harvested at the S growth phase compared with the EE growth phase. B. cereus msrAB expression was thus maximal over the stationary phase. Similar stationary phase-induced expression of msr genes has been documented in several bacteria (Moskovitz et al., 1995; Vattanaviboon et al., 2005; Alamuri and Maier, 2006; Singh and Singh, 2012).

To investigate the role of MsrAB in B. cereus, we constructed a non-polar ΔmsrAB mutant and a ΔmsrAB-complemented strain using a multicopy pHT304-based plasmid (Arantes and Lereclus, 1991). Expression of msrAB in the complemented strain was under the control of its own promoter. We did not detect msrAB mRNA by RT-PCR in the mutant, proving that the genomic disruption of the gene generated an msrAB-null mutant. Figure 1 shows that msrAB was overexpressed in the strain ΔmsrAB/pHT304msrAB at the EE and LE growth phases. Therefore, msrAB expression level was not restored by complementation.

The growth characteristics of the three strains, ΔmsrAB, ΔmsrAB/pHT304msrAB, and the parental wild-type strain (WT), were determined under pH-regulated aerobic respiratory conditions in synthetic MOD medium. Figure 2A shows that the lag phase was 2.5-fold lower in the ΔmsrAB strain (0.7 ± 0.1 h⁻¹) than in the ΔmsrAB/pHT304msrAB (1.8 ± 0.9 h⁻¹) and WT (1.9 ± 0.2 h⁻¹) strains. Exponential growth kinetics were similar in the three strains for the first 6 h. After this initial growth time, WT and ΔmsrAB cultures entered stationary phase. In contrast, ΔmsrAB/pHT304msrAB continued to grow and reached the stationary growth phase at a higher final biomass (2.6 ± 0.1 g.L⁻¹) than ΔmsrAB (1.9 ± 0.1 g.L⁻¹) and WT (1.8 ± 0.2 g.L⁻¹). The viabilities of ΔmsrAB and ΔmsrAB/pHT304msrAB cells, harvested at S growth phase, were similar to the viability of WT after 2 days but declined by more than 100-fold after 5 days of storage at 4°C (Figure 2B). This suggests that msrAB expression impacts the metabolic activity of B. cereus cells at the end of growth (Chubukov and Sauer, 2014). Figure 2C shows that the ΔmsrAB and ΔmsrAB/pHT304msrAB strains consumed higher amounts of glucose than WT at the beginning of exponential growth. The ΔmsrAB/pHT304msrAB culture could be distinguished from the ΔmsrAB culture by continued glucose consumption between the LE and S growth phases (Figure 2C). At the end of growth, ΔmsrAB/pHT304msrAB consumed a higher level of glucose than ΔmsrAB and WT. During aerobic respiratory growth, glucose is catabolized into CO₂ through the TCA cycle, and acetate is excreted as a by-product of overflow metabolism (Madeira et al., 2015; Duport et al., 2016). Figure 2D shows that ΔmsrAB cells, and to a lesser extent ΔmsrAB/pHT304msrAB cells, excreted higher amounts of acetate than WT cells during exponential growth. Acetate accumulation stopped at the LE growth phase in the ΔmsrAB and ΔmsrAB/pHT304msrAB cultures while it continued to accumulate between the LE and S growth phases in the WT culture. Taken together, these results suggest that msrAB expression impacts on the metabolic activity of B. cereus under aerobiosis.

To determine whether the alteration of glucose catabolism was associated with changes in extracellular protein production, extracellular proteins were extracted from culture supernatants of the three B. cereus strains, harvested during the EE, LE, and S growth phases (Madeira et al., 2015). Figure 3A shows that the ΔmsrAB culture supernatant accumulated a higher amount of exoproteins than that of WT at the LE phase. However, ΔmsrAB supernatant had 50 and 90% fewer exoproteins in the EE and S growth phases, respectively, compared with WT. This decreased exoprotein concentration could have resulted from a higher protease activity in the ΔmsrAB culture supernatant. To test this hypothesis, we quantified the protease activity of the ΔmsrAB, ΔmsrAB/pHT304msrAB and WT culture supernatants against casein. Figure 3B shows that the ΔmsrAB culture supernatant sustained a higher protease activity than WT, markedly in the EE and S growth phases. These changes in protease activity were only partially rescued in ΔmsrAB/pHT304msrAB. However, unlike ΔmsrAB/pHT304msrAB, there was no correlation between the protease activity and the amount of exoproteins in ΔmsrAB at LE phase (Figure 3A). This indicates that changes in msrAB expression could be selective for certain extracellular proteases.

To determine if altered metabolism in ΔmsrAB and ΔmsrAB/pHT304msrAB was associated with cellular and exoproteome profile changes, we quantified protein abundance level differences between ΔmsrAB, ΔmsrAB/pHT304msrAB, and WT cells in the EE, LE and S growth phases. Exoproteome and cellular proteome samples were prepared from supernatant cultures and whole-cell lysates, respectively. A total of 200,746 and 71,676 MS/MS spectra were recorded from cellular proteome and exoproteome samples, respectively. A total of 922 proteins were identified in the cellular proteome (Table S1) and 371 proteins were identified in the exoproteome (Table S2), based on the confident detection of at least two different peptides. A two-sample t-test was then conducted separately between WT and ΔmsrAB, and between ΔmsrAB and ΔmsrAB/pHT304msrAB. All proteins with a p ≤ 0.01 and at least a 2-fold change (log₂ fold-change ≥ 1) were considered to be differentially modulated in terms of abundance. A total of 64 and 78 proteins were found to vary in abundance in ΔmsrAB compared with WT in the cellular proteome and exoproteome fractions, respectively. The majority (80%) of these proteins were not rescued in ΔmsrAB/pHT304msrAB (data not shown). The Venn diagrams presented in Figure 4 show the growth phase distribution of the identified proteins. Less than 2% of proteins showed abundance level changes in all three growth stages, indicating that msrAB modulates B. cereus cellular and exoproteome mainly in a growth phase-dependent manner. The impact of msrAB disruption appeared to be more important at the LE and S than the EE growth phase in the cellular proteome (Figure 4A), according to its expression (Figure 1).

We identified peptides with oxidized Met in ΔmsrAB, ΔmsrAB/pHT304msrAB, and WT, in both the cellular proteome and the exoproteome at EE, LE, and S growth phases, as previously described (Madeira et al., 2015). The Met(O) content of both the cellular proteome and the exoproteome was estimated by comparing the number of Met(O) to the total number of Met residues identified in each of the three biological samples obtained for each growth phase in each of the three strains (Tables S4, S5). Figure 5 shows that the Met(O) content of WT and ΔmsrAB/pHT304msrAB decreased similarly in the cellular proteome (Figure 5A) and exoproteome (Figure 5B) during growth. The Met(O) content of the ΔmsrAB intracellular proteome also decreased during exponential growth and was lower than the Met(O) content of WT at LE phase and higher at S growth phase. More importantly, the Met(O) content of the ΔmsrAB exoproteome remained constant during growth and accounted for 38 ± 3% of total Met residues. Taken together, these results indicate that MsrAB regulates the dynamic of the Met(O) content of the proteome, especially at the exoproteome level.

To identify peptides exhibiting significant differences in Met(O) content in ΔmsrAB compared with WT, we conducted a t-statistical analysis. For a robust analysis, we considered a peptide as containing an oxidized Met residue when it was identified in at least two biological replicates. The lists of intra- and extracellular peptides showing significant Met(O) level changes (log₂ fold-change > 1 and p ≤ 0.01) according to growth phase are presented in Tables 3, 4.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.