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Journal of Bacteriology, July 2009, p. 4605-4614, Vol. 191, No. 14
0021-9193/09/$08.00+0     doi:10.1128/JB.00144-09
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Redundant Hydrogen Peroxide Scavengers Contribute to Salmonella Virulence and Oxidative Stress Resistance ^,

Magali Hébrard,^1,3 Julie P. M. Viala,^1,3 Stéphane Méresse,^2,3 Frédéric Barras,^1,3 and Laurent Aussel^1,3*

CNRS, Laboratoire de Chimie Bactérienne (UPR 9043), Institut de Microbiologie de la Méditerranée, IFR 88, 31 Chemin Joseph Aiguier, 13402 Marseille Cedex 20, France,¹ Centre d'Immunologie de Marseille-Luminy, CNRS-INSERM-Univ. Méditerranée, Parc Scientifique de Luminy, Case 906, 13288 Marseille Cedex 9, France,² Aix-Marseille University, Marseille, France³

Received 3 February 2009/ Accepted 9 May 2009

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Salmonella enterica serovar Typhimurium is an intracellular pathogen that can survive and replicate within macrophages. One of the host defense mechanisms that Salmonella encounters during infection is the production of reactive oxygen species by the phagocyte NADPH oxidase. Among them, hydrogen peroxide (H₂O₂) can diffuse across bacterial membranes and damage biomolecules. Genome analysis allowed us to identify five genes encoding H₂O₂ degrading enzymes: three catalases (KatE, KatG, and KatN) and two alkyl hydroperoxide reductases (AhpC and TsaA). Inactivation of the five cognate structural genes yielded the HpxF^– mutant, which exhibited a high sensitivity to exogenous H₂O₂ and a severe survival defect within macrophages. When the phagocyte NADPH oxidase was inhibited, its proliferation index increased 3.7-fold. Moreover, the overexpression of katG or tsaA in the HpxF^– background was sufficient to confer a proliferation index similar to that of the wild type in macrophages and a resistance to millimolar H₂O₂ in rich medium. The HpxF^– mutant also showed an attenuated virulence in a mouse model. These data indicate that Salmonella catalases and alkyl hydroperoxide reductases are required to degrade H₂O₂ and contribute to the virulence. This enzymatic redundancy highlights the evolutionary strategies developed by bacterial pathogens to survive within hostile environments.

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Salmonella is a facultative intracellular pathogen that is associated with gastroenteritis, septicemia, and typhoid fever. This gram-negative bacterium survives and replicates in macrophages during the course of infection and can be exposed to a number of stressful environments during its life cycle (16). One of the host defense mechanisms that Salmonella encounters upon infection is the production of superoxide anion O₂^– by the phagocyte NADPH oxidase (1, 25). This radical can pass the outer membrane of the bacteria and represents one of the major weapons used by the macrophage to kill engulfed pathogens (18). Evidence that phagocyte-produced superoxide is a key mechanism for avoiding Salmonella infection is clear: mice and humans who are genetically defective in superoxide production are significantly more susceptible to infection (36, 38). Superoxide dismutases, located in the bacterial periplasm and in the cytoplasm, dismutate superoxide O₂^– to hydrogen peroxide H₂O₂ and molecular oxygen. Unlike superoxide, hydrogen peroxide can diffuse readily across bacterial membranes and form HO hydroxyl radicals in the presence of Fe(II) (18). These reactive oxygen species (ROS) can oxidize and damage proteins, nucleic acids, and cell membranes.

To scavenge and degrade H₂O₂ molecules generated either as a by-product of aerobic metabolism or by the phagocyte NADPH oxidase, Salmonella has evolved numerous defense mechanisms. The KatE and KatG catalases are involved in H₂O₂ degradation, with katE being described as a member of the RpoS regulon (17, 22) and katG being OxyR dependent (26, 39). Both enzymes share the ability to reduce hydrogen peroxide to water and molecular oxygen, and their role was shown to be predominant at millimolar concentrations of H₂O₂ since they do not require any reductant (32). This observation is of particular importance, since these enzymes are not limited by the availability of a reductant, such as NADH, which cannot be generated fast enough to face a burst of H₂O₂. However, the katG and katE simple mutants, as well as the katE katG double mutant, did not show any increased susceptibility in macrophage or virulence attenuation in mice (5, 27). A possible reason could be the presence of a third nonheme and manganese-dependent catalase called KatN (30). This enzyme may contribute to hydrogen peroxide resistance under certain environmental conditions, but its involvement in virulence remains unknown. Moreover, katE, katG, and katN single mutants did not show any susceptibility to exogenous millimolar H₂O₂, essentially due to the compensatory function of the remaining catalases (5, 30).

Another family of enzymes was shown to play an alternative role in H₂O₂ scavenging: the alkyl hydroperoxide reductases. These proteins directly convert organic hydroperoxides to alcohols, e.g., hydrogen peroxide to water. The alkyl hydroperoxide reductase AhpC belongs to the two-cysteine peroxiredoxin family, and the gene encoding this enzyme was identified as a member of the OxyR regulon (26, 39). The redox system consists of two proteins, AhpC and AhpF, with the latter being a thioredoxin reductase-like protein that contains two disulfide centers and transfers electrons from NADH to AhpC (13). AhpC was shown to be a predominant scavenger at low concentrations of H₂O₂, mainly because its catalytic efficiency was better than those of catalases (32). Recently the alkyl hydroperoxide reductase from Helicobacter hepaticus, TsaA (Thiol-Specific Antioxidant), was characterized (24). The tsaA mutant was found to be more sensitive to oxidizing agents like superoxide anion or t-butyl hydroperoxide. Surprisingly, this mutant was more resistant than the wild-type to H₂O₂, essentially because the level of catalase was increased in this background (24). In gastric pathogens, TsaA plays a critical role in the defense against oxygen toxicity that is essential for survival and growth (2). Interestingly, Salmonella contains two genes encoding alkyl hydroperoxide reductases, ahpC and tsaA, whereas a single copy was found in Escherichia coli (ahpC) or in Helicobacter pylori (tsaA).

The redundancy of these antioxidant proteins could explain the extremely high resistance of Salmonella to hydrogen peroxide. It has been shown by Imlay and coworkers that in E. coli, three genes were involved in H₂O₂ scavenging: two catalase genes (katE and katG) and an alkyl hydroperoxide reductase gene (ahpC) (32). Simultaneous inactivation of the katE, katG, and ahpCF genes negated H₂O₂ degradation. As a consequence, this triple mutant, called the Hpx^– mutant, accumulates intracellular H₂O₂ (32). Moreover, H₂O₂ generated by aerobic metabolism was found to be sufficient to create toxic levels of DNA damage in such a background (28). In the present study, we deleted the Salmonella katE, katG, and ahpCF genes and two more genes absent in E. coli, katN and tsaA, to obtain the HpxF^– mutant, which lacks three catalases and two alkyl hydroperoxide reductases. HpxF^– cells exhibited the incapacity to degrade micromolar concentrations of H₂O₂, whereas this phenotype was not observed for the Kat^– (katE katG katN) and Ahp^– (ahpCF tsaA) mutants. Therefore, the HpxF^– mutant exhibited a high sensitivity to this compound. Moreover, this mutant did not show any proliferation within macrophages and presented reduced virulence in mice, suggesting that Salmonella catalases and alkyl hydroperoxide reductases form a redundant antioxidant arsenal essential for survival and replication within host cells.

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Bacterial strains and growth conditions. The bacterial strains used in this study are listed in Table 1. Strains were routinely grown in Luria-Bertani (LB) medium at 37°C. Minimal medium (M9) contained MgSO₄ (1 mM), CaCl₂ (200 µM), and thiamine (1 µg/ml). As a carbon source, glucose was added at a final concentration of 0.2%. Ampicillin (50 µg/ml), kanamycin (25 µg/ml), and chloramphenicol (25 µg/ml) were added when necessary. Deletions of various genes and concomitant insertion of an antibiotic resistance cassette were carried out using lambda-Red-mediated recombination (11). Mutations were moved to the wild-type strain 12023 by P22 transductions. To avoid the outgrowth of suppressed strains, katE, katG, katN, ahpCF and tsaA mutations were selected anaerobically on LB agar supplemented with bovine liver catalase (2,000 U/plate; Sigma-Aldrich). In some cases, antibiotic resistance cassettes were removed by using the temperature-sensitive plasmid pCP20 carrying the FLP recombinase (11).

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TABLE 1. Bacteria and plasmids used in this study

In silico genome analysis. The genome analyzed was that of Salmonella enterica serovar Typhimurium LT2 (23) (www.genome.jp/dbget-bin/www_bfind?S.typhimurium). STM4106, STM1318, and STM0608 correspond to the catalases KatG and KatE and the alkyl hydroperoxide reductase AhpC, respectively. STM1731 was characterized as a third catalase, KatN (30). STM0402 was annotated as a putative thiol-alkyl hydroperoxide reductase, and we renamed it TsaA (thiol-specific antioxidant) for homology with Helicobacter hepaticus TsaA (58.5% identity) (24).

Measurement of hydrogen peroxide accumulation in cell culture. Cells were grown overnight in M9 minimal medium containing 0.2% glucose and supplemented with Casamino Acids (1 mg/ml), diluted to an optical density at 600 nm (OD₆₀₀) of 0.05, and grown for four generations. These cells were then washed in fresh minimal medium containing 0.05% glucose and incubated with shaking at 37°C for 25 min. Aliquots were removed every 5 min, and hydrogen peroxide levels were determined using the Amplex red/horseradish peroxidase method (Invitrogen). The low glucose concentration was used in order to minimize H₂O₂ production by catalyzed glucose autooxidation.

Hydrogen peroxide scavenging by whole cells. Bacterial cultures were grown overnight aerobically in M9 minimal medium supplemented with Casamino Acids, diluted to an OD₆₀₀ of 0.01, and grown to an OD₆₀₀ of 0.3. Cells were diluted in fresh prepared minimal medium at an OD₆₀₀ of 0.1. H₂O₂ was added at a final concentration of 10 µM. Each 90 s, aliquots were removed and assayed immediately for H₂O₂ content by the Amplex red/horseradish peroxidase method (Invitrogen).

Hydrogen peroxide sensitivity assay. Bacterial cultures were grown overnight in LB, diluted to an OD₆₀₀ of 0.05, grown to an OD₆₀₀ of 0.3 in LB, and treated with 1 mM hydrogen peroxide. To determine viability, aliquots were taken immediately before treatment and 1 h and 2 h after treatment, serially diluted, and plated onto LB supplemented with catalase.

Bacterial infection of macrophages and survival assays. RAW 264.7 macrophages were seeded at a density of 4 x 10⁵ cells per well in six-well tissue culture plates containing Dulbecco's modified Eagle medium (DMEM) with 10% fetal calf serum (FCS) (HyClone) and supplemented with gamma interferon (IFN-) (10 U/ml; ImmunoTools) 24 h before use. Bone marrow cells were isolated from femurs of 8- to 10-week-old C57BL/6 female mice and differentiated into macrophages as described previously (12). Bacteria were cultured overnight at 37°C with shaking and opsonized in DMEM containing FCS and normal mouse serum (10%; Perbio) for 30 min. The macrophages were activated with 0.2 µM phorbol 12-myristate 13-acetate (PMA) (Sigma Aldrich) before infection. Where indicated, diphenyleneiodonium (DPI) (Sigma Aldrich) was added at various concentrations to inhibit NADPH oxidase. Bacteria were added to the monolayers at a MOI of 70:1, centrifuged at 400 x g for 5 min at 4°C, and incubated for 30 min at 37°C in 5% CO₂. The macrophages were washed three times and incubated with DMEM containing FCS and 100 µg/ml gentamicin for 90 min, after which the gentamicin concentration was decreased to 10 µg/ml for the remainder of the experiment. For enumeration of intracellular bacteria, macrophages were washed two times with phosphate-buffered saline and lysed with 0.1% Triton X-100, and a dilution series was plated on LB agar supplemented with catalase.

Competition assays. Eight- to 10-week-old C57/B6 mice were inoculated intraperitoneally with equal amounts of two bacterial strains for a total of 10⁵ bacteria per mouse. The spleens were harvested 48 h after inoculation and homogenized. Bacteria were recovered and enumerated after plating a dilution series onto LB agar and LB agar with the appropriate antibiotics, both supplemented with catalase. Competitive indexes (CIs) were determined for each mouse (3). The CI is defined as the ratio between the mutant and wild-type strains within the output (bacteria recovered from the mouse after infection) divided by their ratios within the input (initial inoculum). A one-sample t test was used to determine whether the CI was significantly different from 1. All statistical analyses were performed by using the Prism software program (GraphPad, San Diego, CA). The two-tailed P value was calculated.

Competition assays in LB medium were performed as described previously (10). Briefly, bacterial mixtures were incubated with equal amounts at 37°C for 24 h under shaking. Cells were diluted and plated on LB agar supplemented with catalase, and the resulting colonies were plated on the selective medium to determine the relative percentage of each strain recovered. The CI was calculated as described above.

RNA extraction and quantitative PCR. RNA extraction from intracellular bacteria was adapted from a previously described method (14). In brief, activated RAW 264.7 macrophages were seeded in 10 tissue-culture-treated plates (128 by 86 mm). The infection was carried out as described above with few modifications: bacteria were added to the monolayers at a MOI of 50:1, and supplemented DMEM containing 66 µg/ml gentamicin was added for 45 min following the three washes. Six hours postinfection, cells were washed twice with phosphate-buffered saline and lysed, and RNA was stabilized with a solution of 0.3% sodium dodecyl sulfate, 1% acidic phenol, and 19% ethanol for 30 min on ice. RNA extraction was performed with the SV Total RNA isolation system (Promega). cDNAs were synthesized from 1 µg RNA with the Superscript II reverse transcriptase and random primers (Invitrogen). Real-time PCR was performed on a Mastercycler ep realplex instrument (Eppendorf) by using the SYBR Premix Ex Taq (Perfect Real Time) PCR kit (Takara Bio Group, Japan). Specific primers are described in Table S1 in the supplemental material. Melting curves were analyzed to control for the specificities of the PCRs. Copy numbers were calculated from a standard curve plotting four different dilutions of genomic DNA against the PCR cycle number at which the measured fluorescence intensity reached the threshold, specified so that it was significantly above the noise band of the baseline (10 times the standard deviation).

Plasmid construction. The cloning vector used was pACYC184. The insert carrying the 200-bp-upstream katG start codon and the katG gene was PCR amplified from S. Typhimurium 12023 by using the forward primer 5'-CCCAAGCTTCCGGGAGCTTTATTACAACTC-3' and the reverse primer 5'-GGGGGATCCCTGGTTGTGCATAACATAGGC-3'. PCR products were digested using BamHI and HindIII and cloned into the pACYC184 vector to generate pkatG, and the insert was verified by sequencing. The insert carrying the 200-bp-upstream tsaA start codon, and the tsaA gene was PCR amplified by using the forward primer 5'-CCCTCTAGAAGCGAGGGCGTGCGTCAGATC-3' and the reverse primer 5'-GGGGGATCCGAAGTGGCGCTGGAAGCGTTG-3'. PCR products were digested using BamHI and XbaI and cloned into the pACYC184 vector to generate ptsaA, and the insert was verified by sequencing.

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Three catalases and two alkyl hydroperoxide reductases allow Salmonella to tolerate an anaerobic/aerobic shift. To assay the importance of Salmonella catalases and alkyl hydroperoxide reductases during aerobic metabolism, two mutants were built: a katE katG ahpCF mutant (HpxT^–, for Hpx_three^–) and a katE katG katN ahpCF tsaA mutant (HpxF^–, for Hpx_five^–). These strains were precultured in LB anaerobically and then diluted in aerobic medium, and their phenotypes were compared to that of the E. coli katE katG ahpCF mutant (Hpx^–). Salmonella HpxT^– cells did not present any growth defect, whereas the HpxF^– mutant initially grew and then stalled for 2 to 3 h, finally catching up at the end of the stationary phase (Fig. 1A). At OD₆₀₀s of 0.1, 0.5, and 1, aliquots of each culture were removed and observed under a microscope. Whereas Salmonella wild-type and HpxT^– cells appeared rod shaped, HpxF^– cells formed long filaments (Fig. 1C). This trend was emphasized at an OD₆₀₀ of 0.5 and correlated with the lag observed during growth but decreased at an OD₆₀₀ of 1 when the cells escaped from the lag and resumed growing. Similar observations were made with the E. coli Hpx^– mutant, which presented a growth defect and filamentation when the cells were submitted to an anaerobic/aerobic shift (Fig. 1B and C). Presumably, upon the switch to aerobiosis, cells experience a sudden burst in ROS that catalases and alkyl hydroperoxide reductases are able to scavenge. These results are consistent with the characterization of the E. coli Hpx^– strain already reported (28). Taken together, these data indicated that inactivation of three catalases and two alkyl hydroperoxide reductases led to a growth lag and filamentation of Salmonella under an aerobic shift.

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FIG. 1. E. coli Hpx– and Salmonella HpxF– strains share the same growth defect linked to filamentation. S. Typhimurium 12023 (wild type; filled squares), E. coli MG1655 (wild type; filled circles), S. Typhimurium HpxF– (katE katG katN aphCF tsaA; open triangles), S. Typhimurium HpxT– (katE katG aphCF; open squares), and E. coli Hpx– (katE katG aphCF; open circles) cells were grown in LB medium anaerobically until early log phase. The cells were then diluted in aerobic LB medium to an OD600 of 0.05, and aerobic growth was monitored at 600 nm (A and B). At OD600s of 0.1, 0.5, and 1, cells from the five strains were removed for microscopic observations (C) (magnification, x100).

Viability of the HpxF^– mutant is not affected by endogenous hydrogen peroxide. To establish the impact of endogenous H₂O₂ on Salmonella fitness, we built two supplementary mutants, which we referred to as the Kat^– mutant, which lacks three catalases (katE, katG, and katN), and the Ahp^– mutant, which lacks two alkyl hydroperoxide reductases (ahpCF and tsaA). Then, we evaluated their ability to eliminate endogenously produced H₂O₂. In order to monitor H₂O₂ accumulation generated during aerobic growth, the strains were grown in minimal medium supplemented with Casamino Acids for at least four generations and immediately resuspended in fresh medium. H₂O₂ was then quantified every 5 min over a period of 25 min. In wild-type and Kat^– strains, the H₂O₂ levels remained constant, accumulating at a rate of 1.2 nM/min (Fig. 2A). The Ahp^– mutant also showed only a minor accumulation of H₂O₂ (2 nM/min), reaching a final concentration of 0.14 µM. Within HpxF^– cells, the H₂O₂ concentration increased 2.6-fold in 25 min to reach 0.36 µM, with a rate of 8.8 nM/min (Fig. 2A). Thus, a HpxF^– mutant accumulated much more endogenous H₂O₂ than Kat^– or Ahp^– mutants, underlying the synergistic action of both type of enzymes on H₂O₂ degradation. Then, to estimate the impact of endogenous H₂O₂ accumulation on their fitness, the four bacterial strains were grown aerobically in LB and viability was assayed at various times. Growth was monitored at 600 nm, and the growth curves of the four strains were found to be identical (data not shown). Then, aliquots from each culture were removed at OD₆₀₀s of 0.5 and 2 and plated on LB agar medium supplemented with catalase. The results showed that the wild type and the three mutants shared identical plating recovery, indicating that none of the mutants presented any viability defect in comparison to its wild-type parent (Fig. 2B). Thus, the HpxF^– mutant clearly accumulated H₂O₂ during growth, but the concentrations reached in LB medium were not high enough to compromise viability. Finally, the wild-type strain and the three mutants (the Ahp^–, Kat^–, and HpxF^– mutants) were grown in minimal medium under aerated conditions. The growth of all the mutants was slowed down, with HpxF^– being slightly more affected (Fig. 2C). Thereby, the nutrient limitation led to a growth rate reduction for the Ahp^–, Kat^–, and HpxF^– mutants. Surprisingly, addition of Casamino Acids did not significantly change the growth rate of the HpxF^– mutant (data not shown). This last observation ruled out the possibility that this mutation led to amino acid auxotrophy as described for the E. coli sodA sodB mutant (6).

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FIG. 2. Endogenous H2O2 does not affect HpxF– mutant viability. (A) Cultures of Salmonella Typhimurium 12023 (wild type; filled squares), the Ahp– mutant (aphCF tsaA; filled circles), Kat– (katE katG katN, open squares), and the HpxF– mutant (katE katG katN aphCF tsaA; open circles) were grown overnight aerobically in minimal medium supplemented with Casamino Acids, diluted to an OD of 0.05, and grown for four generations. The cells were then washed in fresh medium and incubated, and aliquots were removed every 5 min to measure H2O2 concentrations as described in Materials and Methods. (B) Wild-type, Kat–, Ahp–, and HpxF– strains were grown in LB medium. At OD600s of 0.5 (white bars) and 2 (gray bars), aliquots were removed from each culture and plated on LB agar supplemented with catalase to determine cell viability. Results are the means ± standard deviations for three independent experiments, each in triplicate. (C) Wild-type, Kat–, Ahp–, and HpxF– cells were grown in minimal medium overnight. The cells were then diluted to an OD600 of 0.1, and aerobic growth was monitored at 600 nm.

The HpxF^– mutant is highly sensitive to exogenous hydrogen peroxide. To evaluate the role of catalases and alkyl hydroperoxide reductases when Salmonella is submitted to exogenous stress, the wild-type strain and the three mutants were exposed to various exogenous H₂O₂ levels. The bacterial strains were first assayed for their ability to scavenge micromolar concentrations of H₂O₂. Bacterial cultures were grown to an OD₆₀₀ of 0.3 and diluted in minimal medium, and H₂O₂ was added at the final concentration of 10 µM. The wild-type, Ahp^–, and Kat^– strains scavenged H₂O₂ in less than 7 min, whereas the HpxF^– mutant failed to do so (Fig. 3A). A wild-type strain heat killed at 80°C displayed a degradation profile similar to that of the HpxF^– mutant (Fig. 3A). These data demonstrated that the combination of mutations in catalase- and alkyl hydroperoxide reductase-encoding genes prevented scavenging of exogenous H₂O₂ at micromolar concentrations. They also highlighted the catalytic efficiency of catalases and alkyl hydroperoxide reductases, since the Ahp^– and Kat^– mutants could completely degrade 10 µM H₂O₂ in a few minutes.

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FIG. 3. The HpxF– mutant is highly sensitive to exogenous hydrogen peroxide. (A) Wild-type (filled squares), Ahp– (filled circles), Kat– (open squares), HpxF– (open circles), and heat-killed wild-type (filled triangles) cells were grown in minimal medium to an OD600 of 0.3 and then diluted at an OD600 of 0.1. H2O2 was added at a final concentration of 10 µM. Aliquots were removed every 90 s to assay H2O2 levels as described in Materials and Methods. (B) Bacterial cultures (wild type, Ahp–, Kat–, and HpxF–) grown to an OD600 of 0.3 in LB medium were treated with 1 mM hydrogen peroxide (arrow). The growth was then monitored at 600 nm. (C) Viability was assayed, respectively, before (white bars) and 1 h (gray bars) and 2 h (black bars) after H2O2 treatment by plating the bacterial cells on LB agar supplemented with catalase. Results are the means ± standard deviations for three independent experiments, each in triplicate.

Next, to evaluate the impact of millimolar concentrations of H₂O₂ on Salmonella fitness, strains were grown to an OD₆₀₀ of 0.3 in LB medium and treated with 1 mM H₂O₂, and growth was monitored at 600 nm over time. The Kat^– and HpxF^– mutant growth was abolished following the addition of exogenous H₂O₂ (Fig. 3B). In contrast, growth of the wild type and the Ahp^– strain was not affected, reflecting the poor efficiency of AhpC and TsaA in scavenging millimolar concentrations of H₂O₂. Finally, we examined cell viability after exposure to 1 mM H₂O₂. Bacterial cells were collected just before and 1 and 2 h after H₂O₂ treatment. We observed that 2 h after stress, plating recovery of the HpxF^– mutant exhibited a 25-fold drop, from 10⁸ to 4 x 10⁶ CFU/ml, whereas plating recovery of the wild-type strain showed a 10-fold increase (Fig. 3C). Viability of the Kat^– strain was also affected, since its plating recovery showed a threefold decrease 2 h after stress, whereas the Ahp^– mutant survived as well as its wild-type parent. Differences in plating recovery between the Kat^– and HpxF^– mutants also assigned a scavenging effect to the Ahp enzymes, observed only when the catalases were inactivated (Fig. 3C). All together, these experiments highlighted the sensitivity of the HpxF^– mutant when subjected to exogenous stress and the importance of the catalases in degrading millimolar concentrations of exogenous hydrogen peroxide.

The HpxF^– mutant is defective for proliferation within macrophages. To investigate the involvement of catalases and alkyl hydroperoxide reductases in intracellular proliferation, we infected two kind of cells: macrophages freshly prepared from murine bone marrow and considered highly oxidant and RAW 264.7 mouse macrophages. Both kind of cells were activated with IFN- and PMA and infected with the wild-type, Ahp^–, Kat^–, and HpxF^– Salmonella strains. Bacterial proliferation was assayed by calculating the proliferation index as a ratio of the intracellular bacteria between 16 and 2 h postinfection. Within murine bone marrow, the proliferation indexes of the wild-type and Kat^– cells were found to be nearly similar (2.38 ± 0.21 and 2.76 ± 0.10, respectively), whereas the Ahp^– and HpxF^– mutants were defective for proliferation (0.92 ± 0.07 and 0.64 ± 0.01, respectively) (Fig. 4A). Within RAW cells, the proliferation index was also very near for the wild-type and Kat^– cells (11.9 ± 0.6 and 11.5 ± 1.9, respectively) whereas the Ahp^– and HpxF^– mutants exhibited proliferation indexes of 5.5 ± 2.1 and 0.5 ± 0.1, respectively (Fig. 4B). These results indicated that the HpxF^– mutant was unable to proliferate within bone marrow or RAW 264.7 macrophages. Moreover, they highlighted the contribution of Ahp enzymes to intracellular replication within macrophages. Next, to test if the lack of proliferation of the HpxF^– mutant was due to exogenous oxidative stress generated by the phagocyte NADPH oxidase, we infected macrophages in the presence of various concentrations of DPI, an inhibitor of this enzymatic complex. The proliferation index of the HpxF^– strain increased 3.7-fold in the presence of 1 µM DPI (Fig. 4C). Conversely, the proliferation index of the wild-type strain decreased from 10.4 ± 2.4 to 3.9 ± 2.1 when DPI was added, indicating that this molecule had an undesirable effect on the bacterial proliferation (Fig. 4C). Thus, we calculated the comparative wild type/HpxF^– proliferation index ratio, which dropped from 28.8 to 2.9 in the presence of the NADPH oxidase inhibitor (Fig. 4C). These results showed that despite the nonspecific effect of DPI, the oxidative stress generated by the NADPH oxidase was partially responsible for the proliferation defect of the HpxF^– mutant.

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FIG. 4. HpxF– mutations led to a drastic attenuation of intracellular proliferation within bone marrow-derived and RAW 264.7 macrophages. Opsonized bacteria (Wild type or Kat–, Ahp–, or HpxF– mutant) were phagocytosed by freshly prepared bone marrow-derived macrophages (A) or RAW 264.7 cells (B), both activated with IFN- and PMA. Two and sixteen hours postinfection, mouse macrophages were lysed for enumeration of intracellular bacteria (gentamicin protected), determined by CFU counts. The values shown represent the proliferation index, calculated as a ratio of the intracellular bacteria between 16 and 2 h postinfection. (C) The HpxF– mutant and the wild-type strain were tested for infection of activated RAW 267.4 cells without or with 1 µM DPI, which inhibits the phagocyte NADPH oxidase. The proliferation index was calculated as described above. The inset represents the proliferation index ratio (wild type/HpxF–) without or with 1 µM DPI. Results presented in the three panels are the means ± standard deviations for at least three independent experiments, each in triplicate.

The HpxF^– mutant exhibited attenuated virulence in mice. Finally, we tested whether mutation of catalase- and alkyl hydroperoxide reductase-encoding genes contributed to virulence in the animal. To address this issue, we used mixed infections of wild-type and mutant strains in mice to determine the CI, a very sensitive technique to compare the virulence levels of different strains (3). The lack of three catalases and two alkyl hydroperoxide reductases significantly decreased Salmonella virulence, since the CI of the HpxF^– mutant was found to be 0.07 (Table 2). In contrast, the CIs of the Kat^– and Ahp^– mutants did not show any significant alteration in virulence (Table 2). These results indicate that deletion of all catalase- and Ahp-encoding genes strongly affects Salmonella virulence in the mouse model. As a control, a competition assay was performed in LB, in which the mutant and wild-type strains were mixed one to one and grown under aerated conditions. The CIs of the Kat^–, Ahp^–, and HpxF^– mutants were found to be 1.10, 0.86, and 0.69, respectively (Table 3). Thus, the three mutants tested could compete with the wild-type strain in rich culture medium. All in all, these data provided evidence for a contribution of catalases and alkyl hydroperoxide reductases in allowing resistance against oxidative stress generated during the infection process.

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TABLE 2. Competition assays with micea

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TABLE 3. Competition assays with LB aerated culturesa

Gene expression pattern in LB and macrophages. To monitor the expression of Salmonella catalase- and Ahp-encoding genes, we performed quantitative reverse transcription-PCR using a wild-type strain grown either within RAW 264.7 macrophages for 6 h or in LB to an OD₆₀₀ of 2. Within macrophages, ahpC and tsaA were strongly transcribed (3 x 10¹⁰ and 4.7 x 10¹⁰ copies/µg RNA, respectively) (Fig. 5). In comparison, the three catalase-encoding genes presented much lower expression levels (4.3 x 10⁹, 4.7 x 10⁸, and 4.5 x 10⁸ copies/µg RNA for katG, katE, and katN, respectively) (Fig. 5). These data highlight the efficient transcription of ahpC and tsaA, both being strongly expressed in LB and within macrophages.

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FIG. 5. Expression pattern of catalase- and alkyl hydroperoxide reductase-encoding genes in LB and RAW 264.7 macrophages. RNA was extracted from the Salmonella wild-type strain grown either in LB medium to an OD600 of 2 or within activated RAW 264.7 cells during 6 h. cDNA were synthesized from 1 µg RNA, and real-time PCR was performed to amplify the katE, katG, katN, ahpC, tsaA genes. A noncoding (n.c.) DNA domain located between STM0413 and STM0414 was used as a control to monitor the basal level of expression due to residual genomic DNA. The results are the means ± standard deviations for three independent experiments.

The katG and tsaA genes can complement the HpxF^– mutant. To evaluate the contributions of individual genes in H₂O₂ scavenging, we cloned katG and tsaA into the low-copy-number plasmid pACYC184. Wild-type and HpxF^– strains were transformed with the pkatG and ptsaA plasmids, and these strains were grown in LB. When submitted to 1 mM H₂O₂, the HpxF^– strain carrying pkatG grew as well as its wild-type parent (Fig. 6A). In contrast, the HpxF^– strain carrying ptsaA showed a growth defect similar to that of the HpxF^– strain carrying pACYC184 (Fig. 6A). Thus, the presence of katG carried on a low-copy-number plasmid (10 to 15 copies/cell) was sufficient to allow wild-type resistance to exogenously added H₂O₂. On the other hand, the tsaA-encoded enzyme did not support a catalytic efficiency strong enough to allow the HpxF^– mutant to be resistant to 1 mM H₂O₂.

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FIG. 6. pkatG and ptsaA complement the HpxF– mutant with different efficiencies. Bacterial cultures (wild type, filled symbols; HpxF–, open symbols) were transformed with empty vector (squares), the pkatG plasmid (circles), or the ptsaA plasmid (triangles). The cells grown to an OD600 of 0.3 and were treated with 1 mM H2O2 (arrow). Growth was monitored at an OD of 600 nm (A), and the viability was assayed before (white bars) and 1 h (gray bars) and 2 h (black bars) after H2O2 treatment (B). Wild-type and HpxF– strains were transformed with empty vector, pkatG, and ptsaA and were tested for infection within activated RAW 264.7 cells. Bacterial cells were plated and counted on LB agar supplemented with chloramphenicol and catalase. The values shown represent the proliferation index, calculated as a ratio of the intracellular bacteria between 16 and 2 h postinfection (C). Results presented in panels B and C are the means ± standard deviations for at least three independent experiments, each in triplicate.

We also investigated cell viability after exposure to 1 mM H₂O₂. HpxF^– cells carrying pkatG exhibited a wild-type plating recovery (10⁸ to 2 x 10⁹ CFU/ml 2 h after stress), compared to a 10-fold decrease in viability of the HpxF^–, pACYC184-carrying control (Fig. 6B). Moreover, in a wild-type background, the presence of pkatG yielded an increased plating recovery of 2 x 10⁹ CFU/ml 2 h after stress, compared to 7 x 10⁸ CFU/ml obtained with pACYC184. The contribution of tsaA was less pronounced but increased 4.5-fold the bacterial cell survival level: 54% of the HpxF^– cells carrying ptsaA survived for 2 h after the stress, compared to 12% for the HpxF^– cells carrying pACYC184 (Fig. 6B). Thus, katG was found to restore the viability of the HpxF^– cells, whereas a slight contribution was attributed to tsaA.

The contribution of katG and tsaA was also assayed under infection conditions. All proliferation indexes were found to decrease in the presence of the pACYC184 plasmid and its derivatives, as described previously (19). Transformation of the wild-type strain with pkatG or ptsaA did not significantly change the intracellular proliferation compared to results with the empty vector (1.6-fold and 0.73-fold, respectively) (Fig. 6C). In contrast, the presence of katG or tsaA noticeably increased the proliferation index of the HpxF^– mutant (9.6-fold or 6.7-fold, respectively). Moreover, the proliferation indexes of the HpxF^– strain carrying pkatG and wild-type strain carrying pACYC184 were found to be quite similar (2.21 ± 0.95 versus 2.47 ± 0.31) (Fig. 6C). This observation suggested that an overexpressed single gene suppressed a phenotype generated by the mutation of five different genes. Thus, these experiments assigned a direct role for both the tsaA and katG genes in intracellular bacterial growth, showing a link between resistance to oxidative stress and proliferation within macrophages.

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Resistance to oxidative stress is one of the key processes that allow pathogens to survive within macrophages. Hence, pathogenic bacteria have developed a series of antioxidative defenses which either eliminate ROS or repair their damages (18, 31). In recent years, several attempts have been made to assess the contribution of these defenses to the overall virulence of bacteria. Surprisingly, the outcome was not as straightforward as one would have anticipated. Thus, in several cases, inactivation of genes encoding ROS-eliminating enzymes failed to cause a significant defect in bacterial virulence. For instance, inactivation of catalase-encoding genes had little if any effect on pathogenicity-associated traits of Salmonella Typhimurium (5), Yersinia pestis (15), Francisella tularensis (21), Staphylococcus aureus (9), and Neisseria gonorrhoeae (34). In the present study, we addressed the question of the involvement of cytoplasmic H₂O₂-degrading enzymes in Salmonella virulence. Our study revealed that efficient proliferation of this bacterium in macrophages relies on the activities of three catalases and two alkyl hydroperoxide reductases. It illustrates how Salmonella has evolved numerous and redundant defense mechanisms to survive within such hostile and oxidative environments.

Analysis of the S. Typhimurium genome allowed us to identify five genes encoding H₂O₂ degrading enzymes: two alkyl hydroperoxide reductases, AhpC and TsaA, and three catalases, KatE, KatG, and KatN. Alkyl hydroperoxide reductase activity was found to contribute to macrophage infection, since the proliferation index of a strain lacking both ahpC and tsaA was decreased by twofold. This observation could be linked to the oxidative microenvironment present within a Salmonella-containing vacuole (SCV), where ROS and reactive nitrogen species are massively synthesized after macrophage phagocytosis (18, 36). Stimulation of NADPH oxidase and NO synthase, both located on the SCV membrane of the macrophages, leads to the production of O₂^– and NO, respectively. Both molecules (i) are used directly as a reactive species against Salmonella, (ii) are enzymatically transformed into another reactive species (H₂O₂ or NO₂), or (iii) react with each other to generate a peroxynitrite anion, ONOO^–. In contrast, a Kat^– mutant strain, lacking all three catalases, exhibited a wild-type-like proliferation index within macrophages and mice. This is consistent with previous results in which a Salmonella katE katG mutant retained full murine virulence (5). Alternatively, the wild-type phenotype of the Kat^– mutant could be related to the low expression levels of katE, katG, and katN under infection conditions. However, this possibility can be discarded on the basis that a Kat^– mutant failed to resist to millimolar exogenous H₂O₂. This implies that the level of expression recorded was sufficient for participation in H₂O₂ scavenging. Moreover, one can anticipate that the presence of catalases was the very thing accounting for the resistance of the Ahp^– mutant to millimolar concentrations of H₂O₂. These observations fit with the E. coli model, wherein AhpC was proposed to be of primary importance in scavenging micromolar concentrations of H₂O₂ whereas the role of catalases was dominant at millimolar concentrations of H₂O₂ (32).

An HpxF^– mutant lacking the three catalases and the two alkyl hydroperoxide reductases was unable to proliferate or even survive within macrophages. These mutations also contributed to the attenuation of Salmonella virulence, since the HpxF^– cells exhibited a reduced proliferation in mice. Exogenous ROS were found to be partially responsible for the proliferation defect of the HpxF^– mutant, since its intracellular replication increased when NADPH oxidase was inhibited. But the DPI did not allow this mutant to reach a wild-type replication efficiency, and its effects appeared quite difficult to sort out since it altered proliferation of the wild-type strain as well. Still, the fact that DPI increased the replication of the HpxF^– mutant is fully consistent with the "exogenous ROS" hypothesis. Other causes could account for the reduced proliferation of the HpxF^– cells. For instance, this mutant was found to accumulate H₂O₂ during aerobic growth much faster than its wild-type parent. Thus, endogenous ROS generated during aerobic metabolism in the bacterial cytoplasm might contribute to cumulative oxidative damages. Moreover, the HpxF^– mutant was found to grow poorly in minimal medium, and nutrient limitation within the SCV could amplify this trend. Finally, the two detrimental effects could act synergistically and explain the drastic attenuation of proliferation of the HpxF^– mutant within macrophages and mice.

The HpxF^– mutant exhibited much stronger deficiencies than the Ahp^– or Kat^– mutant. The additive effect of the ahpC tsaA and katE katG katN mutations opened the possibility that both types of scavenging systems contribute in a specific way to the replication process. For instance, H₂O₂ concentrations were found to vary during infection, increasing massively just after phagocytosis and decreasing during the next 6 h (36). Then, both type of activity would be exploited at different time points during the whole process. An alternative possibility is that the substrate repertoire of alkyl hydroperoxide reductases is much more expanded than that of catalases: ethyl, t-butyl, cumene, and linoleic acid hydroperoxides, among others, were efficiently reduced by Salmonella AhpC (29). Also, this enzyme was shown to protect Salmonella against reactive nitrogen intermediates and to detoxify peroxynitrite (ONOO^–) to nitrite fast enough to forestall DNA oxidation (4, 8). Hence, catalases could act as an H₂O₂ scavenger while alkyl hydroperoxide reductases could intervene in eliminating micromolar concentrations of H₂O₂ and other hydroperoxides.

The redundancy of Salmonella enzymes is striking even in a comparison with the closely related E. coli, which requires only three genes, ahpC, katE, and katG, to scavenge H₂O₂. It is a classic hypothesis that occurrence of redundancy can be rationalized by genetic regulation. In the present work, the expression levels of Salmonella tsaA and ahpC, quantified 6 h after macrophage infection, were found to be high. Conversely, the three catalase-encoding genes were poorly transcribed. The importance of fine-tuning gene expression was also illustrated by the fact that a moderate increase in gene dosage (15 copies per cell) allowed either katG or tsaA to confer a proliferation index similar to that of the wild type. This demonstrated that increased synthesis of only one type of H₂O₂-degrading activity is sufficient to restore a wild-type replication level within macrophages. However, a certain degree of redundancy also arises at the regulatory level, with OxyR being the transcriptional activator of katG and ahpC whereas katE and katN transcription is RpoS dependent. Our ongoing work aims at assessing a link between redundancy and genetic regulation by carrying out a thorough gene regulation study with Salmonella-infected macrophages.

Previous studies had revealed that an important aspect of Salmonella survival within macrophages was the production of two periplasmic Cu/Zn superoxide dismutases, SodCI and SodCII. These enzymes combat phagocytic superoxide O₂^– by dismutation to H₂O₂ and O₂. SodCI but not SodCII was found to play a role during infection of mice by Salmonella (10, 20, 35). In this context, SodCI and SodCII were considered the front line for combating NADPH-oxidase-generated superoxide. A remaining question was the management of H₂O₂ produced by dismutation of superoxide. The H₂O₂ flux into the bacterial cytoplasm was shown to be rapid, with a high permeability coefficient of the membrane (33). Here we showed that cytoplasmic catalases and alkyl hydroperoxide reductases constitute a second line for combating H₂O₂ produced by SodCI and SodCII in the periplasm. Hence, together with these previous studies, the present work contributes to a better understanding of the strategies used by Salmonella to resist host-produced ROS. It will now be extremely important to bring into the picture the proposal by Fang and coworkers that Salmonella pathogenicity island 2 allows bacteria to avoid NADPH oxidase-dependent killing by macrophages (37). Their data suggested that the NADPH phagocyte oxidase could be excluded from SCV in a Salmonella pathogenicity island 2-dependent manner. Thus, an emerging comprehensive picture would be that Salmonella relies on two alternative ways of coping with oxidative stress: excluding the NADPH phagocyte oxidase and metabolizing ROS produced by the host. Investigation of the interplay between the two strategies represents a major challenge for the future.

 
Thanks are due to the members of the F.B. group, A. Latifi and P. Moreau (LCB, Marseille, France), for fruitful discussions and to R. Stevens (GAFL, Avignon, France) for critical reading of the manuscript.

This work was funded by the ANR (Programme Blanc "Stress Oxydant" and Programme M.I.E. "Salmo-sensor"), the CNRS, and the Université de la Méditerranée. M.H. was supported by the Ministère de la Recherche and J.P.M.V. by the Fondation pour la Recherche Médicale (FRM).

 
* Corresponding author. Mailing address: Laboratoire de Chimie Bactérienne, 31 Chemin Joseph Aiguier, 13402 Marseille Cedex 20, France. Phone: 33 4 91 16 46 59. Fax: 33 4 91 71 89 14. E-mail: aussel{at}ifr88.cnrs-mrs.fr

Published ahead of print on 15 May 2009.

Supplemental material for this article may be found at http://jb.asm.org/.

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Journal of Bacteriology, July 2009, p. 4605-4614, Vol. 191, No. 14
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