• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of jbacterPermissionsJournals.ASM.orgJournalJB ArticleJournal InfoAuthorsReviewers
J Bacteriol. May 2008; 190(10): 3658–3669.
Published online Feb 29, 2008. doi:  10.1128/JB.00022-08
PMCID: PMC2395008

The Peptidoglycan-Associated Lipoprotein OprL Helps Protect a Pseudomonas aeruginosa Mutant Devoid of the Transactivator OxyR from Hydrogen Peroxide-Mediated Killing during Planktonic and Biofilm Culture [down-pointing small open triangle]

Abstract

OxyR controls H2O2-dependent gene expression in Pseudomonas aeruginosa. Without OxyR, diluted (<107/ml) organisms are easily killed by micromolar H2O2. The goal of this study was to define proteins that contribute to oxyR mutant survival in the presence of H2O2. We identified proteins in an oxyR mutant that were oxidized by using 2,4-dinitrophenylhydrazine for protein carbonyl detection, followed by identification using a two-dimensional gel/matrix-assisted laser desorption ionization-time of flight approach. Among these was the peptidoglycan-associated lipoprotein, OprL. A double oxyR oprL mutant was constructed and was found to be more sensitive to H2O2 than the oxyR mutant. Provision of the OxyR-regulated alkyl hydroperoxide reductase, AhpCF, but not AhpB or the catalase, KatB, helped protect this strain against H2O2. Given the sensitivity of oxyR oprL bacteria to planktonic H2O2, we next tested the hypothesis that the biofilm mode of growth might protect such organisms from H2O2-mediated killing. Surprisingly, biofilm-grown oxyR oprL mutants, which (in contrast to planktonic cells) possessed no differences in catalase activity compared to the oxyR mutant, were sensitive to killing by as little as 0.5 mM H2O2. Transmission electron microscopy studies revealed that the integrity of both cytoplasmic and outer membranes of oxyR and oxyR oprL mutants were compromised. These studies suggest that sensitivity to the important physiological oxidant H2O2 in the exquisitely sensitive oxyR mutant bacteria is based not only upon the presence and location of OxyR-controlled antioxidant enzymes such as AhpCF but also on structural reinforcement by the peptidoglycan-associated lipoprotein OprL, especially during growth in biofilms.

Pseudomonas aeruginosa is an important opportunistic pathogen and a leading cause of global nosocomial infections (46). In human disease, the organism is most frequently encountered in patients whose immune system has been compromised, including individuals suffering from burns (30), cancer chemotherapy (55), organ transplantation (16), and various pneumonias (34). However, the organism receives its greatest notoriety because it is the predominant pathogen during airway infection of patients afflicted with the inherited disease, cystic fibrosis (CF) (3, 23). Within the thick CF airway mucus, P. aeruginosa forms highly refractory communities known as biofilms. In this remarkably dynamic niche, the bacteria are tightly packed against one another and encased in oblong spheroid-shaped biofilms composed of both human and bacterial products (63). Arguably, the biofilm mode of growth is one that inherently resists killing by conventional antibiotics regimens and professional phagocytes that include alveolar macrophages. However, during CF airway disease, it is the neutrophil whose titers rise nearly 1,500-fold in the chronically infected airways (5).

During infection, P. aeruginosa faces potential death when the organism encounters stimulated neutrophils that have undergone what is commonly referred to as the oxidative (or respiratory) burst. One toxic product of the respiratory burst is hydrogen peroxide (H2O2) (27), which is generated through protonation of O2 in the acidic milieu of the phagolysosomal vacuole. Within this vacuole, H2O2 concentrations have been estimated to be as high as 100 mM (32), a level that easily causes the death of both planktonic and biofilm P. aeruginosa (7, 26, 28, 38). In fact, stimulated neutrophils release even significant H2O2 (~12 μM) in the extracellular milieu (59). Despite these oxidative perils, P. aeruginosa is remarkably equipped with a powerful battery of either constitutive or inducible enzymatic defenses to help collaboratively detoxify H2O2 and/or organic peroxides. These include three catalases (KatA, KatB, and KatC) (7, 38, 42), and at least four alkyl hydroperoxide reductases, AhpA, AhpB, and AhpCF (45), and Ohr (organic hydroperoxide reductase, (43). Thus, given these advantageous defenses, the organism is inherently able to respond quickly to sudden changes in H2O2 levels, especially in the context of extracellular versus intraphagosomal concentrations, to avoid death. However, the most robust genetic response to H2O2 in P. aeruginosa is governed by the global transactivator OxyR (25, 45).

Activation of OxyR in Escherichia coli is initiated upon exposure to low levels of endogenous or exogenous H2O2. The transcriptionally dormant OxyR becomes an active transactivator when two sulfhydryl groups on cysteines 199 and 208 are oxidized in an H2O2-dependent fashion. This event allows OxyR and RNA polymerase (61) to transcribe genes whose products are specifically designed to cope with H2O2-mediated stress (see the recent review by Hassett and Imlay [27]). The three major OxyR-regulated antioxidant enzymes include the tetrameric catalase KatB, the alkylhydroperoxide reductase AhpCF, and periplasmic AhpB (25, 45). However, bacteria lacking OxyR exhibit truly remarkable aerobic phenotypes. First, oxyR mutant bacteria are unable to form isolated colonies on rich media (e.g., Luria-Bertani agar [L-agar] plates) (25). Second, when 5 μl of overnight aerobic cultures was serially diluted and spotted onto L-agar plates, survivors grow from only undiluted cultures (25). An even more striking defect is highlighted when a katA mutation is introduced into an oxyR mutant background. This strain was unable to grow even from undiluted cultures on aerobic L agar (25). The mechanistic basis for this exquisite sensitivity is that L broth is known to contain autoxidizable components that are capable of generating ~1.2 μM H2O2 per min (25), an amount that is sufficient to kill ≤107 oxyR mutant bacteria per ml. Finally, we also showed that oxyR mutant bacteria have impaired virulence properties in mouse and fruit fly (Drosophila melanogaster) infection models and show increased susceptibility to killing by human neutrophils (33).

In the present study, we examined the mechanistic basis underlying the limited resistance of a P. aeruginosa oxyR mutant to H2O2 under aerobic conditions. Using both proteomic and genetic approaches, we discovered that the peptidoglycan-associated lipoprotein OprL is involved in what modest protection is afforded an oxyR mutant when exposed to exogenous H2O2. Transmission electron microscopy (TEM) studies suggest that it is likely that the altered membrane integrity of the oxyR oprL double mutant compromises H2O2-sensitive respiratory membrane components and, surprisingly, the cytoplasmic AhpCF provides some protection when provided in trans.

MATERIALS AND METHODS

Bacterial strains, plasmids and planktonic growth conditions.

All bacteria and plasmids used in the present study are listed in Table Table1.1. Organisms were grown in either Luria-Bertani broth (L broth), L broth containing 100 mM KNO3, or 1% Trypticase soy broth (TSB). Cultures were grown at 37°C with shaking at 300 rpm. Culture volumes were 1/10 of the total Erlenmeyer flask volume to ensure maximum aeration. Media were solidified with 1.5% Bacto agar. Frozen bacterial stocks were stored at −80°C in a 1:1 mixture of 30% glycerol and stationary-phase bacterial suspension.

TABLE 1.
Strains and plasmids in this study

Manipulation of recombinant DNA and genetic techniques.

All plasmid and chromosomal nucleic acid manipulations were by standard techniques (50). Plasmid DNA was transformed into E. coli strain DH5α-MCR (Protein Express, Cincinnati, OH). To detect the presence of insert DNA, X-Gal (5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside; 40 μg/ml) was added to agar media. Restriction endonucleases, the Klenow fragment of DNA polymerase I, T4 DNA polymerase, and T4 DNA ligase were used as specified by the vendor (Invitrogen/Gibco-BRL Corp., Gaithersburg, MD). Plasmid DNA was isolated by using plasmid miniprep isolation kits (Qiagen), and restriction fragments were recovered from agarose gels by using SeaPlaque low-melting-point agarose (FMC BioProducts, Rockland, ME). PCRs were performed by using Pfu DNA polymerase (BRL) and appropriate primers in an MJ Research thermal cycler, with 30 cycles of denaturation (2 min, 94°C), annealing (1 min, 54°C), and extension (1 min 30 s, 72°C). Amplified DNA fragments were gel purified, cloned into pCR2.1 (Invitrogen), and sequenced.

Construction of P. aeruginosa mutants.

The strategy for insertional inactivation of some of the genes listed in Table Table11 was facilitated by gene disruption with either an 850-bp gentamicin resistance (Gmr) cassette from pUCGM (52), and the gene replacement vector pEX100T (29), the latter of which allowed for selection of double-crossover events within putative recombinants cultured on agar containing 5% sucrose. All mutants were confirmed by PCR.

Semiquantitative expression of oprL by RT-PCR.

Total RNA from stationary- and exponential-phase PAO1 and oxyR mutant bacteria was purified by using a RiboPure-Bacteria kit as specified by the manufacturer (Ambion). After treatment with DNase, 250 ng of total RNA was used to amplify the oprL and omlA, the latter of which is a constitutively expressed gene and internal control (44) by using one-step reverse transcription-PCR (RT-PCR) kit (Qiagen). Both RT-PCR analyses of oprL and omlA were performed using 25 PCR cycles. PCR products were separated by electrophoresis on 1.2% agarose gels. Band intensities were determined by using AlphaEase FC StandAlone software (Alpha Innotech).

H2O2 sensitivity measurements.

Bacteria were grown aerobically in L broth for 17 h at 37°C. Cultures were diluted in 5 ml of 0.8% low-melting-point agarose (SeaPlaque) in an L-broth base to a final optical density at 600 nm of 0.01. Suspensions were distributed evenly on L agar plates, and the top agarose was allowed to solidify. Filter paper disks (7 mm) containing 10 μl of either 4.4 M H2O2 or 1 M t-butyl hydroperoxide (t-BOOH) were placed on the top agar surface, and the plates were incubated at 37°C for 24 h. The zones of growth inhibition were measured from at least three experiments in triplicate. Using yet another measure of quantitative H2O2 susceptibility, the same titers of 17-h-old suspensions were treated with 5 mM H2O2 for 0, 20, 30, and 40 min, respectively. Serial dilutions were spotted on L agar plates containing filter-sterilized bovine liver catalase (Boehringer Mannheim), which allows the oxyR mutant to survive the ~1.2 μM H2O2 produced in aerobic L agar plates by autoxidation (25). Surviving bacteria were enumerated after a 24-h incubation at 37°C.

Catalase assays.

Cell extracts prepared from sonicated stationary-phase bacteria were prepared in 50 mM potassium phosphate buffer (pH 7.0; KPi). Catalase activity was measured by the decomposition of 19.5 mM H2O2 in KPi at 240 nm (7, 38). One unit of activity is defined as that which decomposes 1 μmol of H2O2 min−1 mg of protein−1. Protein concentrations were estimated by the method of Bradford (4) using bovine serum albumin fraction V (Sigma) as a standard. Where applicable, statistics were performed by using a Student t test, with all assays being performed in triplicate.

Detection of protein carbonyl formation using DNPH derivatization and two-dimensional (2-D) gel electrophoresis after H2O2 exposure.

The oxidation of specific amino acids is considered an appropriate indicator of the level of oxidative stress mediated by H2O2 and other oxidants in aerobic cells. Upon exposure to such oxidants, carbonyl groups in specific proteins are formed which can react with 2,4-dinitrophenylhydrazine (DNPH). An antibody specific for DNP can then bind to the adduct, and protein carbonyl formation can be quantified. First, the P. aeruginosa oxyR mutant was grown aerobically overnight in L broth and then diluted 1:100 in fresh medium. Bacteria were then grown to mid-logarithmic phase and treated with 10 mM H2O2 for 30 min. The organisms were harvested by centrifugation at 13,000 × g for 10 min at 4°C and subsequently washed twice in 10 mM Tris-HCl (pH 7.8). Cells were then disrupted with lysis buffer (9.5 M urea, 2% CHAPS {3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate}, 0.8% ampholytes [Amersham], 1 mM EDTA, 50 mM dithiothreitol [DTT]) at room temperature for 30 min. Immobiline Drystrips (Amersham) were used for isoelectric focusing of 80 μg of cell extract in the first dimension using an IPGphor isoelectric focusing system (Pharmacia Biotech). The strips were then immediately subjected to “in-strip” DNPH derivatization as described previously (10, 13, 47). Briefly, the strips were incubated for 20 min in 2 N HCl-10 mM DNPH at room temperature, followed by equilibration in 50 mM Tris-HCl containing 6 M urea, 2% (wt/vol) sodium dodecyl sulfate (SDS), 30% (vol/vol) glycerol, and 1% DTT (pH 6.8) for 15 min. The strips were then reequilibrated in the same buffer containing 2.5% iodoacetamide, thereby replacing the DTT. Strip proteins were then separated in the second dimension by SDS-12% polyacrylamide gel electrophoresis using a vertical gel electrophoresis unit (SE 400; Hoefer). After electrophoresis, proteins were electroblotted onto polyvinylidene difluoride membranes (Hybond-C; Amersham) and the carbonylated proteins were detected by using an OxyBlot kit (Chemicon International, Germany). Briefly, polyvinylidene difluoride membranes were incubated with (i) primary antibody (rabbit immunoglobulin G, 1:150 dilution) specific to the DNP moiety on the proteins, (ii) secondary antibody, and finally (iii) goat anti-rabbit immunoglobulin G (horseradish peroxidase conjugated, 1:300 dilution). Detection of oxidized proteins was assessed by chemiluminescence (RPN2109; Amersham). The membranes were exposed to Hybond-ECL film (Amersham) for either 3 s, 10 min, or 24 h, and the film was developed. Quantification of protein spots was performed by using ImageJ for the Macintosh (http://www.versiontracker.com/dyn/moreinfo/macosx/19224).

Protein identification by MALDI-TOF mass spectrometric protein analyses.

Protein spots were excised from AgNO3-stained polyacrylamide gels (54). Prior to the “in-gel” trypsin digestion step, the AgNO3 was removed from protein spots by using H2O2 and ammonium bicarbonate as previously described (57). Next, selected proteins were digested with 125 ng of trypsin (Promega)/μl for 16 to 20 h, and the peptides obtained were analyzed by matrix-assisted laser desorption ionization-time of flight (MALDI-TOF) mass spectrometry. The proteins were identified by searching a P. aeruginosa mass spectrometric database (Swiss-Prot). Briefly, the generated mass lists composed of monoisotopic [M+H]+ masses were first filtered for common contaminants (e.g., keratin) and subsequently used for database searches by using the ProFound search algorithm (64).

Flow-cell biofilm experiments.

Biofilms were grown in confocal “friendly” flow chambers as previously described (36). Briefly, bacteria were grown aerobically in L broth at 37°C until the stationary growth phase, diluted 1:50 into 1% TSB, and a 0.2-ml suspension was used to inoculate flow cells (Stovall Life Sciences, Inc., Greensboro, NC). Flow was initiated at a rate of 0.17 ml min−1 after the bacteria were allowed to attach for 1 h. After a 3-day incubation at room temperature (22 to 23°C), biofilms were treated with various concentrations of H2O2 for 30 min. The biofilms were then stained with a live/dead viability stain composed of SYTO 9 and propidium iodine (Molecular Probes, Inc., Eugene, OR). Biofilm images were obtained by using an LSM 510 confocal microscope (Carl Zeiss, Inc., Germany). The excitation and emission wavelengths for green fluorescence were 488 and 500 nm, respectively, while those for red fluorescence were 490 and 635 nm, respectively. All biofilm experiments were repeated at least three times. The live/dead ratios of the biofilms were calculated by using 3D for LSM (v.1.4.2) software (Carl Zeiss).

TEM.

All bacteria were grown in L-broth until the stationary growth phase, clarified by centrifugation at 13,000 × g for 10 min, and 0.5 ml suspensions treated with 50 mM H2O2 in fresh L-broth for 30 min. After a second centrifugation step, cell pellets were fixed with 2.5% glutaraldehyde and 4% formaldehyde (vol/vol) in 0.1 M cacodylate buffer, postfixed with OsO4 in the present of 2% (wt/vol) potassium ferrocyanide, and embedded in Eponate 12. Thin sections were poststained with (2%) uranyl acetate and lead citrate and examined with a Zeiss transmission electron microscope at an accelerating voltage of 80 kV. The TEM micrographs were taken at two different magnifications, ×40,000 and ×100,000, to indicate the overall cellular size and possible membrane integrity alterations.

RESULTS

Remarkable sensitivity of oxyR mutant bacteria to merely aerobic growth on rich media and protection by anaerobic growth.

We have previously shown that oxyR mutant bacteria are exquisitely susceptible to micromolar levels of H2O2 based upon the fascinating observation that L broth generates ~1.2 μM H2O2/min through autoxidation of medium components (25). For example, note that when serial dilutions of stationary-phase oxyR mutant organisms are spotted onto L agar plates under aerobic conditions (+O2), only the undiluted (UN) sample survives (Fig. (Fig.1A,1A, lane 2), while wild-type organisms show the characteristic serial dilution pattern of cells that harbor the full OxyR-controlled antioxidant enzymatic repertoire, including KatB-AnkB (PA4613-PA4612), AhpB (PA0848), and AhpCF (PA0139-PA0140) (Fig. (Fig.1A,1A, lane 1). Complementation in cis in the unique attB site on the P. aeruginosa chromosome with the wild-type oxyR gene restored the serial dilution capacity and growth on aerobic L agar of wild-type bacteria (Fig. (Fig.1A,1A, lane 3). Similarly, serial dilution of the oxyR mutant in the filter-sterilized supernatant from stationary-phase wild-type bacteria, which contains the major, protease-resistant catalase KatA from lysed cells (25), also showed a protective pattern (Fig. (Fig.1A,1A, lane 4). By far the most H2O2-sensitive strain was an oxyR katA double mutant, where organisms could not even survive spotting of undiluted cells onto aerobic L agar (Fig. (Fig.1A,1A, lane 5). Interestingly, when the same serial dilutions were plated under anaerobic conditions where there is no possibility of medium H2O2 generation, the bacteria grew as wild-type organisms with negligible differences (Fig. (Fig.1B,1B, −O2).

FIG. 1.
Sensitivity of oxyR mutant bacteria to serial dilution on aerobic but not anaerobic media. Bacteria were grown aerobically in L-broth until stationary phase. Organisms were serially diluted in either L broth or sterile-filtered spent culture supernatant, ...

Because of the remarkable aerobic sensitivity of the P. aeruginosa oxyR mutant to H2O2, our next goal was to identify proteins that protect the aerobic oxyR mutant from H2O2. We were particularly interested in proteins that are not involved in what might be predicted to be a classical antioxidant enzymatic response to oxidative stress (e.g., OxyR regulon) that has been demonstrated in the most heavily researched bacteria that include E. coli (65), Salmonella enterica serovar Typhimurium (19), and Xanthomonas campestris (9).

Determination of protein carbonylation by DNP derivatization of H2O2-treated oxyR mutant bacteria.

Our next goal was to evaluate the role of proteins in the oxyR mutant that serve to provide limited protection to this extremely sensitive organism when faced with H2O2. Realizing that the oxidation of specific amino acids is considered an appropriate indicator of the level of oxidative stress mediated by H2O2 and other oxidants in aerobic cells, we embarked on the determination of carbonyl groups in proteins that can be formed upon oxidation by H2O2 that can react with DNPH. By using an antibody specific for DNP, protein carbonyl formation can be quantified. For example, using this technique, glyceraldehyde-3-phosphate dehydrogenases (both Tdh2p and Tdh3p) were found to play a role in defense against H2O2 in Saccharomyces cerevisiae (14).

Because wild-type bacteria have their full antioxidant enzymatic gamut (e.g., isozymes of catalase, peroxidase, Ahp, and superoxide dismutase [SOD]), we first discovered that the difference in carbonylated proteins between H2O2--treated and control organisms were only a few spots. Of these, we found that three hypothetical proteins (spots 1, 2, and 6) and also oprD and aguA were more susceptible to protein carbonylation in wild-type bacteria treated with H2O2 than in untreated bacteria (data not shown). In contrast, the oxyR mutant lacks (i) the H2O2-inducible catalase KatB, (ii) the cytoplasmic AhpCF, and (iii) the periplasmic AhpB. Thus, it is not surprising that this organism is severely impaired in its ability to cope with even micromolar levels of H2O2 (25, 45), Therefore, the oxyR mutant strain was used in the following experiments. First, the organism was exposed to H2O2 at a concentration that would minimally affect wild-type bacteria (10 mM H2O2, 30 min) to determine carbonylated proteins in H2O2-treated versus untreated bacteria using a coupled 2-D gel/MALDI-TOF mass spectrometric approach. Figure Figure2A2A depicts a silver-stained 2-D gel of untreated oxyR mutant bacteria, while Fig. Fig.2C2C shows a similar gel of protein lysates derived from organisms treated with 10 mM H2O2. Figure 2B and D are representative Western blots of Fig. 2A and C obtained by using the Oxyblot technique (see Materials and Methods). Note the differences in the overall amount and intensity of spots 8 and 9, respectively (also refer to Fig. 2E and F). Upon observation of distinct differences in oxidized protein profiles between control and H2O2-treated oxyR bacteria (Fig. 2B and D), we next used MALDI-TOF mass spectrometry to identify proteins with increased oxidation patterns. Table Table22 lists the 13 proteins that were identified in the present study, including the PA number, function (if known), the molecular mass and pI of each protein, and several mass spectrometric parameters. Several of these were the outer membrane porins OprF and OprD and the peptidoglycan-associated lipoprotein OprL. Others included four hypothetical proteins, a probable peptidyl-prolyl cis-trans isomerase (FkbP-type), 30S ribosomal protein Si, agmatine deiminase, succinate dehydrogenase (SDH; β-subunit), a thiol peroxidase, and DnaK. Unfortunately, the identification of many of the proteins that were clearly different in DNPH reactivity between oxyR control and H2O2-treated organisms were not represented in sufficient quantity or quality for identification, even after overloading the gels.

FIG. 2.
Protein oxidation profile of P. aeruginosa oxyR mutant bacteria exposed to H2O2. (A to D) AgNO3-stained 2-D gels of control (A) and H2O2-treated (10 mM for 30 min) (C) bacteria; Western blot and DNP staining of control (B) and H2O2-treated (10 mM for ...
TABLE 2.
Proteins in H2O2-treated P. aeruginosa oxyR mutant that were reproducibly oxidized after DNP derivatization and identified by MALDI-TOF mass spectrometric analysisa

The peptidoglycan-associated lipoprotein OprL is essential for the limited H2O2 resistance of an oxyR mutant.

First, isogenic mutants were constructed in each of the 13 candidate genes encoding the oxidized proteins detected in the H2O2-treated oxyR mutant (Fig. 2B and D) to assess their possible contribution to H2O2 resistance or susceptibility. There were no differences in the H2O2 sensitivity of each of these mutants compared to wild-type bacteria (data not shown). Therefore, we next constructed 13 isogenic double mutants in an unmarked oxyR mutant background. Surprisingly, we found that only the oxyR oprL double mutant was more sensitive to H2O2 than the oxyR mutant (Fig. (Fig.3A,3A, lane 5 versus lane 2). Representative L agar plates are included on the top of Fig. 3A and B to reveal a visual representation of these data. In contrast, the oxyR oprL mutant with AhpB (lane 7 versus plasmid control [lane 6]) or KatB (lane 9 versus plasmid control [lane 6]) provided in trans could not contribute to protection of the oxyR oprL mutant in this assay system. Only provision of AhpCF (lane 8) afforded some protection of the oxyR oprL mutant against H2O2-mediated killing. In contrast, the oxyR oprL mutant was no more sensitive to t-butyl hydroperoxide (Fig. (Fig.3B,3B, lane 5 versus lane 2) or cumene hydroperoxide (data not shown) than the single oxyR mutant. This indicates that OprL seems to confer a level of resistance that is specific for only H2O2 and not for organic peroxides.

FIG. 3.
Sensitivity of P. aeruginosa compared to mutants strains to H2O2 or t-BOOH. All bacteria were growth aerobically in L broth until achieving stationary growth phase, and identical CFU were added to 0.8% molten top-agarose and poured onto L agar ...

To clarify the H2O2 sensitivity differences in oxyR versus oxyR oprL mutant bacteria, we also examined survival profiles upon exposure of these bacteria to 5 mM H2O2 for 40 min in planktonic culture, except that we used stationary-phase planktonic cultures that have ~109 CFU/ml. At these high cell titers, the oxyR mutant shows an H2O2 sensitivity profile identical to that of wild-type bacteria (Fig. (Fig.3C).3C). Interestingly, the single oprL and the double oxyR oprL mutants showed increased sensitivity to H2O2. In fact, the oxyR oprL double mutant was nearly 2 logs more sensitive to H2O2 than the oxyR mutant. This reinforces the premise that the oxyR mutant alone at high cell densities on planktonic culture has enhanced resistance to H2O2.

Increased sensitivity of oxyR oprL versus oxyR mutants to H2O2 in biofilms.

We have previously shown that P. aeruginosa biofilms are inherently resistant to concentrations of H2O2 that easily kill planktonic bacteria (26, 28). In particular, estimates range from 10 to 75 mM, depending on the growth and treatment parameters (7, 28, 45). Despite the sensitivity of the oxyR oprL mutant to H2O2 in free-living, planktonic culture, we tested the hypothesis that the biofilm mode of growth might still afford this severely compromised organism resistance to H2O2-mediated killing. This could be due to the formation of a mature biofilm and the deposition of matrix-associated factors (e.g., the protease-resistant catalase KatA [25]) that could detoxify it, thereby protecting susceptible organisms in the immediate surroundings. Mature biofilms were grown for a period of 3 days, after which they were exposed to different concentrations of H2O2. First, we exposed the wild type and the oprL mutant to H2O2 when cultured in biofilms. Mature, 3-day-old biofilms were then stained with a viability stain, where green cells are alive and red cells are dead (see Fig. S1 in the supplemental material, top view [A] and sagittal view [B]). In slight contrast to the results with planktonic cells, oprL mutant biofilms were no different in overall architecture or sensitivity to H2O2 compared to wild-type bacteria (see Fig. S1 in the supplemental material). Note that the biofilm mode of growth protected wild-type and oprL mutant bacteria from H2O2 concentrations of ~70 mM, a finding consistent with previous results (28).

We next performed similar experiments with wild-type and oxyR mutant bacteria using our flow biofilm system. We first observed that oxyR mutant biofilms were not nearly as thick and robust as those formed by wild-type organisms (Fig. (Fig.4,4, lower panel). Recall that the oxyR mutant cannot grow as single colonies on aerobic L agar plates because of ~1.2 mM H2O2 being generated in the media through autoxidation of components in it (25). The medium used for our biofilm studies, TSB, also does not support the aerobic growth of oxyR mutant bacteria as isolated colonies (data not shown), which is similar to the results obtained on L agar (Fig. (Fig.1A,1A, lane 2). Therefore, we believe that the oxyR mutant biofilm was not nearly as robust and differentiated as wild-type bacteria because of the low, yet significant peroxigenic stress in our flow biofilm system. To test this hypothesis experimentally, we performed flow biofilm analyses of the wild type and oxyR mutant in the presence or absence of added bovine liver catalase. Consistent with the dramatic H2O2 sensitivity of oxyR mutant bacteria, the addition of bovine liver catalase to the growth medium allowed for complete restoration of mature biofilm formation by the oxyR mutant (see Fig. S2 in the supplemental material).

FIG. 4.
Confocal laser scanning microscopic images of H2O2 treated wild-type P. aeruginosa and oxyR mutant biofilms. Biofilms were grown on glass substrata for 3 days, and the wild-type and oxyR mutant were treated with H2O2 at 0, 30, 50, 70, and 100 mM. The ...

Unlike wild-type and oprL mutant bacteria, the oxyR mutant was easily killed by 30 mM H2O2 (Fig. (Fig.4,4, lower panel) relative to wild-type organisms (Fig. (Fig.4,4, top panel). In addition, H2O2 caused a mechanical disruption of the biofilms because of the vigorous bubbling due to inherently high catalase activity in the biofilms (18). This is the reason behind the “web”-like biofilm structures observed in both wild-type and oxyR mutant biofilms treated with >30 mM H2O2.

We next assessed the H2O2 sensitivity of oxyR relative to oxyR oprL mutants in biofilms using very low concentrations of H2O2 to help reveal potential subtle differences in biofilm susceptibility patterns between these two strains. Although the oxyR mutant was resistant to 5 mM H2O2 (Fig. (Fig.5,5, top panel), as little as 0.5 mM H2O2 killed the vast majority of oxyR oprL bacteria in biofilms (Fig. (Fig.5,5, bottom panel). Moreover, the dead/live ratios/mm3 of the oxyR oprL mutant biofilm (light gray bars, Fig. Fig.6)6) were also higher than the oxyR mutant (black bars, Fig. Fig.6)6) at the same concentration of H2O2, while the oxyR dead/live ratio was greater than that of the wild-type strain PAO1 (dark gray bars, Fig. Fig.66).

FIG. 5.
Confocal laser scanning microscopic images of H2O2-treated oxyR and oxyR oprL mutant biofilms. Biofilms were grown on glass substrata for 3 days as in Fig. Fig.4.4. oxyR and oxyR oprL mutant biofilms were treated with 0.05, 0.5, and 5 mM H2O2 ...
FIG. 6.
Calculation of dead/live ratios of H2O2-treated P. aeruginosa biofilms. The ratio of live versus dead wild-type (dark gray bars), oxyR (black bars), and oxyR oprL (light gray bars) biofilm bacteria are calculated from optical sections of confocal images ...

The sensitivity of the oxyR oprL versus oxyR mutant to H2O2 in both planktonic cells and biofilm is not based upon significant differences in catalase activity.

Even though the oxyR oprL mutant was more sensitive to H2O2 than the oxyR mutant, we suspected that this could be due to overall differences in total catalase activity. Recall that the oxyR mutant possesses only the protective activity of the major catalase, KatA (45). However, in planktonic cells, the oxyR oprL mutant was found to have has slightly increased catalase activity compared to the oxyR mutant (Fig. (Fig.7A,7A, lane 5 versus lane 2), yet the double mutant was paradoxically more sensitive to H2O2 (recall Fig. 3A and C). However, during biofilm culture, we found no differences in catalase activity between the two mutant strains (Fig. (Fig.7C).7C). Moreover, the catalase activity of both strains during anaerobic culture was also similar (Fig. (Fig.7B,7B, lanes 5 versus lane 2). However, the catalase activity of all bacteria was inexplicably higher (~5-fold) during anaerobic growth (Fig. 7B and A), and yet the oxyR and oxyR oprL strains possessed nearly 2,000 U less than did the wild-type bacteria (Fig. (Fig.7B,7B, lanes 2 and 5 versus lane 1). We offer an explanation for this apparent paradox in the Discussion.

FIG. 7.
Catalase activity of planktonic versus biofilm-grown bacteria. Planktonic bacteria were grown in aerobic L broth (A) or anaerobic L broth with 100 mM KNO3 (B) at 37°C until the stationary phase. Cell extracts were then assayed for catalase activity ...

TEM evidence H2O2-mediated structural damage in oxyR and oxyR oprL mutant P. aeruginosa.

The oprL gene was previously shown to be (i) regulated by RegA and iron levels (17), (ii) maximally expressed in the stationary growth phase (17), and (iii) overexpressed during long-term anaerobic pyruvate fermentation (51). It is also essential for normal cell shape in the related organism P. putida (49). We next tested the hypothesis that OxyR may play a role in the expression of the oprL gene. We based this hypothesis on the fact that the promoter region of the oprL gene possessed 10 of 13 of the canonical OxyR promoter recognition sequence (ATAG-N6-CTAT-N6-ATAG-N6-CTAT) (65). To test this postulate experimentally, oprL gene expression was measured by RT-PCR. We found no difference in oprL gene expression in the wild type versus the oxyR mutant in both exponential- and stationary-phase organisms (data not shown), thereby refuting our hypothesis.

Because OprL has been shown to play a role in overall cell envelope structure in the related strain P. putida (49), we tested the hypothesis that the sensitivity of the oxyR oprL mutant to H2O2 compared to the oxyR mutant might be because of structural alterations in the membrane(s) between these two strains. To test this hypothesis, we elected to perform an ultrastructural analysis of wild-type, oxyR mutant, and oxyR oprL mutant bacteria using TEM to determine whether the oxyR or oprL mutations, either singly or in tandem, affected cell morphology in the presence versus the absence of H2O2. We used high levels of H2O2 (50 mM) in these studies because the morphology of wild-type organisms appeared to be unaltered when organisms were treated with 10 mM H2O2 (data not shown). Untreated bacteria were characterized by intact, smooth, and uninterrupted membranes with a clearly visible periplasmic space (Fig. (Fig.8A,8A, arrow). Treatment with H2O2 caused a slight swelling of the bacteria, and yet the membranes remained intact (Fig. (Fig.8B).8B). Similarly, the untreated oxyR mutant also had intact, yet not as clearly defined membranes (Fig. (Fig.8C).8C). However, the H2O2-treated oxyR mutant possessed an invaginated membrane organization with no clearly defined periplasmic space (Fig. (Fig.8D).8D). In contrast, the oxyR oprL double mutant also possessed wavy, invaginated membranes, more ghost cells, and probably “peeling” of membranes (Fig. (Fig.8E).8E). In contrast, H2O2 treatment caused intermittent membrane fusion and the periplasmic space to appear beaded, with numerous invaginations and evidence of membrane-membrane fusion (Fig. (Fig.8F8F).

FIG. 8.
Ultrastructure of P. aeruginosa oxyR and oxyR oprL control and H2O2-treated bacteria. All strains were grown aerobically in L broth until the stationary growth phase. The bacteria were then treated with 50 mM H2O2 for 30 min at room temperature prior ...

We also collected pictures of wild-type, oxyR mutant, and oxyR oprL mutant strains in the absence (Fig. 8G, I, and K) versus the presence (Fig. 8H, J, and L) of H2O2. Note that the majority of H2O2-treated organisms are slightly swollen relative to control bacteria. Photographs of these bacteria were taken at ×40,000 and ×100,000 magnification for statistical purposes that are meant to define (i) differences in cell size and (ii) changes in membrane integrity, respectively. About 70 bacteria photographed at ×40,000 magnification were used to compare the structural nature of the membranes under control and H2O2-treated conditions. An evaluation of untreated PAO1 showed that ca. 77% of cells have an intact, smooth, and uninterrupted membrane with a clearly visible periplasmid space (Fig. 8A and G). Similarly, nearly 86% of the untreated oxyR mutant also had intact membrane (Fig. 8C and I). In contract, only 42% of the oxyR oprL mutant membranes had intact membranes, whereas the remainder possessed wavy, invaginated membranes and clear evidence of peeling (Fig. 8E and K). In contrast, treatment with H2O2 caused membrane alterations even in wild-type strain PAO1, for which ~67% of the treated cells have a wavy membrane (Fig. 8B and H). However, we found that 72% of H2O2-treated oxyR mutants possessed invaginated membranes with no clearly defined periplasmic space (Fig. 8D and J), whereas 82% of the oxyR oprL mutant H2O2 revealed intermittent membrane fusion while the periplasmic space appeared beaded, with numerous invaginations and evidence of membrane-membrane fusion events (Fig. 8F and L).

DISCUSSION

Approximately 7 years ago, we demonstrated that an absence of the global H2O2-responsive regulator OxyR in P. aeruginosa renders such organisms exquisitely susceptible to micromolar H2O2 (25, 45). Given that H2O2 is a significant component of the antimicrobial armament of human phagocytic cells (e.g., neutrophils and macrophages), an in-depth analysis of oxyR mutant phenotypes became a major research focus. Toward this end, one of the most peculiar and intriguing phenotypes of an oxyR mutant is an inability to form isolated colonies on aerobic (Fig. (Fig.1A)1A) but not anaerobic L agar (25) (Fig. (Fig.1B).1B). This was found to be due to the fact that diluted organisms cannot cope with the paltry 1.2 μM H2O2 produced per min in aerobic L broth by autoxidation (25), levels that are consistent with those produced by human erythrocytes (22, 60). Proteins in the oxyR mutant would predictably be most susceptible to H2O2-mediated oxidation at the level of outer/cytoplasmic membranes and periplasm, especially when diluted to ≤107 cells per ml, but also affecting the overall metabolic properties of the organisms.

Effects of H2O2 on P. aeruginosa metabolism.

In the face of potentially toxic levels of H2O2, bacteria such as P. aeruginosa encounter significant metabolic problems. For example, Tamarit et al. (58) showed that in E. coli H2O2 caused oxidation of the β-subunit of the F1Fo-ATPase. In doing so, the ability of such organism to generate the vast majority of its cellular ATP is compromised. Furthermore, Farr et al. (20) experimentally reinforced this premise in demonstrating that an H2O2-mediated loss in ΔP membrane potential significantly impacts ATP-dependent anabolic activities that include the synthesis of essential cellular components (e.g., DNA, RNA, and protein). However, two of the oxidized proteins of the H2O2-treated oxyR mutant discovered in the present study were the β-subunit of the tricarboxylic acid cycle enzyme, SDH (SdhB) and DnaK, an Hsp70 molecular chaperone (1). Because the cytoplasmic membrane-bound SDH is involved in the production of reducing power in the form of FADH2, its electrons could drive the formation of the vastly more reactive oxygen reduction product, HO. HO. reacts with virtually all know biomolecules at rates approaching the diffusion limit (62). SDH also has Fe-S clusters that have been shown to be involved in the production of superoxide (O2) in E. coli (39). However, formation of O2 by SDH is likely a result of the formation of a flavosemiquinone intermediate that can react with molecular oxygen (35), forming O2. Thus, in SOD-proficient organisms, of which virtually all P. aeruginosa express at least Fe-SOD (6), H2O2 would still be formed in aerobic bacteria. SdhB is also known to be Fur-activated in E. coli (24) and also inducible by nitric oxide (NO) (48). In contrast, DnaK is known to be involved in the H2O2-mediated stress response of Salmonella enterica serovar Typhimurium by a mechanism involving chaperone-like activity (41). However, oxyR sdhB and oxyR dnaK mutants were no more susceptible to H2O2 than the oxyR mutant.

One of the greatest surprises of this work was the dramatic increase in catalase activity in organisms grown anaerobically via respiratory NO3 reduction (Fig. (Fig.7B).7B). We had initially shown increased catalase activity in anaerobically grown P. aeruginosa (21). In the present study, we show that the anaerobic induction of optimal anaerobic catalase activity actually requires OxyR (Fig. (Fig.7B,7B, lane 2 versus lane 1). Being that multiple prokaryotic and eukaryotic catalases are multifunctional (e.g., catalases/peroxidase) during aerobic growth (11, 12), there is no reason to believe that they may not also possess an anaerobic detoxification function of molecules that have a similar molecular structure, redox properties, and electronic state. Catalase has actually been shown to slowly detoxify NO (8). It is possible that the major P. aeruginosa KatA also possesses this function since we detected significant spectral changes upon the exposure of purified KatA to NO gas (data not shown). However, further experimentation is required to precisely define the mechanism of NO binding to the core heme of KatA.

Role of OxyR-controlled gene products in H2O2 resistance in oxyR oprL mutant bacteria.

One of the unique features of the oxyR mutant was that it paradoxically possessed near-wild-type catalase levels (Fig. (Fig.3)3) (25) and yet was exquisitely sensitive to H2O2 in diluted planktonic and biofilm culture. Thus, as discussed above, it would behoove P. aeruginosa to possess the capacity to deploy antioxidant enzymes such as the periplasmic alkyl hydroperoxide reductase/catalase AhpB (45) for protection from H2O2. The same paradigm holds true in gram-negative bacteria in the strategic deployment of β-lactamase to the periplasm when confronted with the β-lactam antibiotics. Surprisingly, it was another P. aeruginosa OxyR-regulated gene product, AhpCF, that offered some level of protection when provided in trans to the oxyR oprL mutant (Fig. (Fig.3A,3A, lane 8 versus lane 5), and yet neither AhpCF, AhpB, nor KatB could exclusively restore wild-type H2O2 resistance levels. However, the overall protection by AhpCF alone was greater than that of AhpB and KatB against H2O2, and this is consistent with AhpCF protecting an oxyR mutant in the aerobic plate dilution assay (25). Although one of the oxidized proteins in the oxyR mutant was a thiol peroxidase known as Tpx, the oxyR tpx mutant was a “red herring” in the sense that it, too, was no more sensitive to H2O2 than was the oxyR mutant. This suggests that the P. aeruginosa Ahp enzyme class has a range of substrate specificities that include both H2O2 and organic peroxides.

Impact of OprL on biofilm sensitivity to H2O2 and associated structural abnormalities.

However, three of the oxidized proteins in the oxyR mutant were the outer membrane proteins OprF, OprD, and the peptidoglycan-associated lipoprotein OprL. Because two of the aforementioned proteins are surface exposed (OprF and OprD), they are obviously the most readily accessible to H2O2, the diffusion rate of which is only limited by enzymes capable of detoxifying it (e.g., catalases, peroxidases, and AHPs) and certain porins of the eukaryotic aquaporin family (2). However, unlike OprF and OprD, OprL is not on the surface of P. aeruginosa but is reported to be linked to peptidoglycan, which resides between the cytoplasmic and outer membranes. P. putida OprL in P. putida has multiple functions, including the uptake of a number of carbon sources (37), as well as resistance to SDS, EDTA, and deoxycholate (49). However, far less is known of its function in P. aeruginosa. Interestingly, we found that OprL contributes significantly to the overall biofilm sensitivity of the oxyR mutant organism when we discovered a dramatically enhanced sensitivity of the oxyR oprL mutant in biofilms. In addition, we also observed greater structural alterations in oxyR oprL mutants treated with H2O2 relative to the oxyR mutant alone (Fig. (Fig.8).8). Finally, consistent with the theme of structural proteins contributing to H2O2 resistance in bacteria, E. coli porins were found to contribute to resistance to the lactoperoxidase-H2O2-SCN antimicrobial system (15), while optimal H2O2 resistance in serovar Typhimurium requires a 59-kDa outer membrane protein (56).

The other unique feature of cells exposed to H2O2 is its capacity to induce swelling due to the oxygen produced by catalase-mediated decomposition, despite the fact that oxygen diffusion through membranes is through free diffusion. Another classic example of cell swelling in the face of H2O2-generating agents is in the case of paraquat-treated E. coli (40). When exposed to redox-active paraquat under aerobic conditions, the optical density of the suspension actually increased, an event that would possibly have led to the postulate that paraquat did not interrupt cell growth or kill E. coli. However, it was ultimately found that the bacteria were actually perishing, and the increase in optical density of the suspension was attributable to cell swelling (40).

Summary.

In conclusion, we have shown that the limited ability of an oxyR mutant of P. aeruginosa to survive exogenous exposure to H2O2 under aerobic conditions requires not only the support of the putatively bifunctional catalase/Ahp AhpCF but also the structural support of the peptidoglycan-associated lipoprotein OprL. This dynamic is unique in the sense that a classical oxidative stress response (e.g., OxyR) has now been linked to a structural protein, OprL, especially during the biofilm mode of growth. This suggests that translational efforts to combat certain sensitive features of highly recalcitrant pathogens can be multifaceted, attacking transcription factors (e.g., OxyR), organic hydroperoxide reductases (e.g., AhpCF), and membrane or peptidoglycan-associated proteins (e.g., OprL).

Supplementary Material

[Supplemental material]

Acknowledgments

This study was supported by National Institutes of Health grant GM-69845 to D.J.H.

Footnotes

[down-pointing small open triangle]Published ahead of print on 29 February 2008.

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

REFERENCES

1. Allan, B., M. Linseman, L. A. MacDonald, J. S. Lam, and A. M. Kropinski. 1988. Heat shock response of Pseudomonas aeruginosa. J. Bacteriol. 1703668-3674. [PMC free article] [PubMed]
2. Bienert, G. P., A. L. Moller, K. A. Kristiansen, A. Schulz, I. M. Moller, J. K. Schjoerring, and T. P. Jahn. 2007. Specific aquaporins facilitate the diffusion of hydrogen peroxide across membranes. J. Biol. Chem. 2821183-1192. [PubMed]
3. Boucher, R. C. 2007. Cystic fibrosis: a disease of vulnerability to airway surface dehydration. Trends Mol. Med. 13231-240. [PubMed]
4. Bradford, M. M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72248-254. [PubMed]
5. Britigan, B. E., M. B. Hayek, B. N. Doebbeling, and R. B. Fick, Jr. 1993. Transferrin and lactoferrin undergo proteolytic cleavage in the Pseudomonas aeruginosa-infected lungs of patients with cystic fibrosis. Infect. Immun. 615049-5055. [PMC free article] [PubMed]
6. Britigan, B. E., R. A. Miller, D. J. Hassett, M. A. Pfaller, M. L. McCormick, and G. T. Rasmussen. 2001. Antioxidant enzyme expression in clinical isolates of Pseudomonas aeruginosa: identification of an atypical form of manganese superoxide dismutase. Infect. Immun. 697396-7401. [PMC free article] [PubMed]
7. Brown, S. M., M. L. Howell, M. L. Vasil, A. J. Anderson, and D. J. Hassett. 1995. Cloning and characterization of the katB gene of Pseudomonas aeruginosa encoding a hydrogen peroxide-inducible catalase: purification of KatB, cellular localization, and demonstration that it is essential for optimal resistance to hydrogen peroxide. J. Bacteriol. 1776536-6544. [PMC free article] [PubMed]
8. Brunelli, L., V. Yermilov, and J. S. Beckman. 2001. Modulation of catalase peroxidatic and catalatic activity by nitric oxide. Free Radic. Biol. Med. 30709-714. [PubMed]
9. Charoenlap, N., W. Eiamphungporn, N. Chauvatcharin, S. Utamapongchai, P. Vattanaviboon, and S. Mongkolsuk. 2005. OxyR mediated compensatory expression between ahpC and katA and the significance of ahpC in protection from hydrogen peroxide in Xanthomonas campestris. FEMS Microbiol. Lett. 24973-78. [PubMed]
10. Choi, J., A. I. Levey, S. T. Weintraub, H. D. Rees, M. Gearing, L. S. Chin, and L. Li. 2004. Oxidative modifications and down-regulation of ubiquitin carboxyl-terminal hydrolase L1 associated with idiopathic Parkinson's and Alzheimer's diseases. J. Biol. Chem. 27913256-13264. [PubMed]
11. Claiborne, A., and I. Fridovich. 1979. Purification of the o-dianisidine peroxidase from Escherichia coli B: physicochemical characterization and analysis of its dual catalatic and peroxidatic activities. J. Biol. Chem. 2544245-4252. [PubMed]
12. Claiborne, A., D. P. Malinowski, and I. Fridovich. 1979. Purification and characterization of hydroperoxidase II of Escherichia coli B. J. Biol. Chem. 25411664-11668. [PubMed]
13. Conrad, C. C., J. Choi, C. A. Malakowsky, J. M. Talent, R. Dai, P. Marshall, and R. W. Gracy. 2001. Identification of protein carbonyls after two-dimensional electrophoresis. Proteomics 1829-834. [PubMed]
14. Costa, V. M., M. A. Amorin, A. Quintanilha, and P. Moradas-Ferreira. 2002. Hydrogen peroxide-induced carbonylation of key metabolic enzymes in Saccharomyces cerevisiae: the involvement of the oxidative stress response regulators Yap1 and Skn7. Free Radic. Biol. Med. 331507-1515. [PubMed]
15. De Spiegeleer, P., J. Sermon, K. Vanoirbeek, A. Aertsen, and C. W. Michiels. 2005. Role of porins in sensitivity of Escherichia coli to antibacterial activity of the lactoperoxidase enzyme system. Appl. Environ. Microbiol. 713512-3518. [PMC free article] [PubMed]
16. Dua, S., W. Chalermskulrat, M. B. Miller, M. Landers, and R. M. Aris. 2006. Bilateral hematogenous Pseudomonas aeruginosa endophthalmitis after lung transplantation. Am. J. Transplant. 6219-224. [PubMed]
17. Duan, K., E. R. Lafontaine, S. Majumdar, and P. A. Sokol. 2000. RegA, iron, and growth phase regulate expression of the Pseudomonas aeruginosa tol-oprL gene cluster. J. Bacteriol. 1822077-2087. [PMC free article] [PubMed]
18. Elkins, J. G., D. J. Hassett, P. S. Stewart, H. P. Schweizer, and T. R. McDermott. 1999. Protective role of catalase in Pseudomonas aeruginosa biofilm resistance to hydrogen peroxide. Appl. Environ. Microbiol. 654594-4600. [PMC free article] [PubMed]
19. Farr, S. B., and T. Kogoma. 1991. Oxidative stress responses in Escherichia coli and Salmonella typhimurium. Microbiol. Rev. 55561-585. [PMC free article] [PubMed]
20. Farr, S. B., D. Touati, and T. Kogoma. 1988. Effects of oxygen stress on membrane functions in Escherichia coli role of HPI catalase. J. Bacteriol. 1701837-1842. [PMC free article] [PubMed]
21. Frederick, J. R., J. G. Elkins, N. Bollinger, D. J. Hassett, and T. R. McDermott. 2001. Factors affecting catalase expression in Pseudomonas aeruginosa biofilms and planktonic cells. Appl. Environ. Microbiol. 671375-1379. [PMC free article] [PubMed]
22. Giulivi, C., P. Hochstein, and K. J. Davies. 1994. Hydrogen peroxide production by red blood cells. Free Radic. Biol. Med. 16123-129. [PubMed]
23. Govan, J. R. W., and G. S. Harris. 1986. Pseudomonas aeruginosa and cystic fibrosis: unusual bacterial adaptation and pathogenesis. Microbiol. Sci. 3302-308. [PubMed]
24. Hantke, K. 1987. Selection procedure for deregulated iron transport mutants (fur) in Escherichia coli K-12: fur not only affects iron metabolism. Mol. Gen. Genet. 210135-139. [PubMed]
25. Hassett, D. J., E. Alsabbagh, K. Parvatiyar, M. L. Howell, R. W. Wilmott, and U. A. Ochsner. 2000. A protease-resistant catalase, KatA, that is released upon cell lysis during stationary phase, is essential for aerobic survival of a Pseudomonas aeruginosa oxyR mutant at low cell densities. J. Bacteriol. 1824557-4563. [PMC free article] [PubMed]
26. Hassett, D. J., J. G. Elkins, J.-F. Ma, and T. R. McDermott. 1999. Pseudomonas aeruginosa biofilm sensitivity to biocides: use of hydrogen peroxide as model antimicrobial agent for examining resistance mechanisms. Methods Enzymol. 310599-608. [PubMed]
27. Hassett, D. J., and J. A. Imlay. 2006. Oxidative stress systems in bacteria: four model systems, p. 544-574. In C. Nickerson and M. J. Schurr (ed.), Molecular paradigims of infectious disease: a bacterial perspective. Kluwer Academic-Plenum Publishers, New York, NY.
28. Hassett, D. J., J.-F. Ma, J. G. Elkins, T. R. McDermott, U. A. Ochsner, S. E. H. West, C.-T. Huang, J. Fredericks, S. Burnett, P. S. Stewart, G. McPheters, L. Passador, and B. H. Iglewski. 1999. Quorum sensing in Pseudomonas aeruginosa controls expression of catalase and superoxide dismutase genes and mediates biofilm susceptibility to hydrogen peroxide. Mol. Microbiol. 341082-1093. [PubMed]
29. Hoang, T. T., A. J. Kutchma, A. Becher, and H. P. Schweizer. 2000. Integration-proficient plasmids for Pseudomonas aeruginosa: site-specific integration and use for engineering of reporter and expression strains. Plasmid 4359-72. [PubMed]
30. Holder, I. A. 1993. P. aeruginosa burn infections: pathogenesis and treatment, p. 275-295. In M. Campa, M. Bendinelli, and H. Friedman (ed.), Pseudomonas aeruginosa as an opportunistic pathogen. Plenum Press, Inc., New York, NY.
31. Holloway, B. 1955. Genetic recombination in Pseudomonas aeruginosa. J. Gen. Microbiol. 13572-581. [PubMed]
32. Iyer, G. Y. N., M. F. Islam, and J. H. Quastel. 1961. Biochemical aspects of phagocytosis. Nature 192535-541.
33. Lau, G. W., B. E. Britigan, and D. J. Hassett. 2005. Pseudomonas aeruginosa OxyR is required for full virulence in rodent and insect models of infection and for resistance to human neutrophils. Infect. Immun. 732550-2553. [PMC free article] [PubMed]
34. Lau, G. W., D. J. Hassett, and B. E. Britigan. 2005. Modulation of lung epithelial functions by Pseudomonas aeruginosa. Trends Microbiol. 13389-397. [PubMed]
35. Leger, C., K. Heffron, H. R. Pershad, E. Maklashina, C. Luna-Chavez, G. Cecchini, B. A. Ackrell, and F. A. Armstrong. 2001. Enzyme electrokinetics: energetics of succinate oxidation by fumarate reductase and succinate dehydrogenase. Biochemistry 4011234-11245. [PubMed]
36. Lequette, Y., and E. P. Greenberg. 2005. Timing and localization of rhamnolipid synthesis gene expression in Pseudomonas aeruginosa biofilms. J. Bacteriol. 18737-44. [PMC free article] [PubMed]
37. Llamas, M. A., J. J. Rodriguez-Herva, R. E. Hancock, W. Bitter, J. Tommassen, and J. L. Ramos. 2003. Role of Pseudomonas putida tol-oprL gene products in uptake of solutes through the cytoplasmic membrane. J. Bacteriol. 1854707-4716. [PMC free article] [PubMed]
38. Ma, J.-F., U. A. Ochsner, M. G. Klotz, V. K. Nanayakkara, M. L. Howell, Z. Johnson, J. Posey, M. L. Vasil, J. J. Monaco, and D. J. Hassett. 1999. Bacterioferritin A modulates catalase A (KatA) activity and resistance to hydrogen peroxide in Pseudomonas aeruginosa. J. Bacteriol. 1813730-3742. [PMC free article] [PubMed]
39. Messner, K. R., and J. A. Imlay. 2002. Mechanism of superoxide and hydrogen peroxide formation by fumarate reductase, succinate dehydrogenase, and aspartate oxidase. J. Biol. Chem. 27742563-42571. [PubMed]
40. Minakami, H., and I. Fridovich. 1990. Effects of paraquat on cultures of Escherichia coli: turbidity versus enumeration. Free Radic. Biol. Med. 8387-391. [PubMed]
41. Morgan, R. W., M. F. Christman, F. S. Jacobson, G. Storz, and B. N. Ames. 1986. Hydrogen peroxide-inducible proteins in Salmonella typhimurium overlap with heat shock and other stress proteins. Proc. Natl. Acad. Sci. USA 838059-8063. [PMC free article] [PubMed]
42. Mossialos, D., G. R. Tavankar, J. E. Zlosnik, and H. D. Williams. 2006. Defects in a quinol oxidase lead to loss of KatC catalase activity in Pseudomonas aeruginosa: KatC activity is temperature dependent and it requires an intact disulphide bond formation system. Biochem. Biophys. Res. Commun. 341697-702. [PubMed]
43. Ochsner, U. A., D. J. Hassett, and M. L. Vasil. 2001. Genetic and physiological characterization of ohr, encoding a protein involved in organic hydroperoxide resistance in Pseudomonas aeruginosa. J. Bacteriol. 183773-778. [PMC free article] [PubMed]
44. Ochsner, U. A., A. I. Vasil, Z. Johnson, and M. L. Vasil. 1999. Pseudomonas aeruginosa fur overlaps with a gene encoding a novel outer membrane lipoprotein, OmlA. J. Bacteriol. 1811099-1109. [PMC free article] [PubMed]
45. Ochsner, U. A., M. L. Vasil, E. Alsabbagh, K. Parvatiyar, and D. J. Hassett. 2000. Role of the Pseudomonas aeruginosa oxyR-recG operon in oxidative stress defense and DNA repair: OxyR-dependent regulation of katB, ahpB, and ahpCF. J. Bacteriol. 1824533-4544. [PMC free article] [PubMed]
46. Odeh, R., and J. P. Quinn. 2000. Problem pulmonary pathogens: Pseudomonas aeruginosa. Semin. Respir. Crit. Care Med. 21331-339. [PubMed]
47. Reinheckel, T., S. Korn, S. Mohring, W. Augustin, W. Halangk, and L. Schild. 2000. Adaptation of protein carbonyl detection to the requirements of proteome analysis demonstrated for hypoxia/reoxygenation in isolated rat liver mitochondria. Arch. Biochem. Biophys. 37659-65. [PubMed]
48. Richardson, A. R., P. M. Dunman, and F. C. Fang. 2006. The nitrosative stress response of Staphylococcus aureus is required for resistance to innate immunity. Mol. Microbiol. 61927-939. [PubMed]
49. Rodriguez-Herva, J. J., M. I. Ramos-Gonzalez, and J. L. Ramos. 1996. The Pseudomonas putida peptidoglycan-associated outer membrane lipoprotein is involved in maintenance of the integrity of the cell envelope. J. Bacteriol. 1781699-1706. [PMC free article] [PubMed]
50. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
51. Schreiber, K., N. Boes, M. Eschbach, L. Jaensch, J. Wehland, T. Bjarnsholt, M. Givskov, M. Hentzer, and M. Schobert. 2006. Anaerobic survival of Pseudomonas aeruginosa by pyruvate fermentation requires an Usp-type stress protein. J. Bacteriol. 188659-668. [PMC free article] [PubMed]
52. Schweizer, H. P. 1993. Small broad-host-range gentamicin resistance gene cassettes for site-specific insertion and deletion mutagenesis. BioTechniques 15831-833. [PubMed]
53. Schweizer, H. P., and T. T. Hoang. 1995. An improved system for gene replacement and xylE fusion analysis in Pseudomonas aeruginosa. Gene 15815-22. [PubMed]
54. Shevchenko, A., O. N. Jensen, A. V. Podtelejnikov, F. Sagliocco, M. Wilm, O. Vorm, P. Mortensen, H. Boucherie, and M. Mann. 1996. Linking genome and proteome by mass spectrometry: large-scale identification of yeast proteins from two-dimensional gels. Proc. Natl. Acad. Sci. USA 9314440-14445. [PMC free article] [PubMed]
55. Simon, A., U. Bode, and K. Beutel. 2006. Diagnosis and treatment of catheter-related infections in paediatric oncology: an update. Clin. Microbiol. Infect. 12606-620. [PubMed]
56. Stinavage, P. S., L. E. Martin, and J. K. Spitznagel. 1990. A 59 kilodalton outer membrane protein of Salmonella typhimurium protects against oxidative intraleukocytic killing due to human neutrophils. Mol. Microbiol. 4283-293. [PubMed]
57. Sumner, L. W., B. Wolf-Sumner, L. K. Pannell, and H. M. Fales. 2002. Silver stain removal using H2O2 for enhanced peptide mass mapping by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry. Rap. Commun. Mass Spectrom. 16160-168. [PubMed]
58. Tamarit, J., E. Cabiscol, and J. Ros. 1998. Identification of the major oxidatively damaged proteins in Escherichia coli cells exposed to oxidative stress. J. Biol. Chem. 2733027-3032. [PubMed]
59. Test, S. T., and S. J. Weiss. 1984. Quantitative and temporal characterization of the extracellular H2O2 pool generated by human neutrophils. J. Biol. Chem. 259399-405. [PubMed]
60. Varma, S. D., and P. S. Devamanoharan. 1991. Hydrogen peroxide in human blood. Free Radic. Res. Commun. 14125-131. [PubMed]
61. Wang, X., P. Mukhopadhyay, M. J. Wood, F. W. Outten, J. A. Opdyke, and G. Storz. 2006. Mutational analysis to define an activating region on the redox-sensitive transcriptional regulator OxyR. J. Bacteriol. 1888335-8342. [PMC free article] [PubMed]
62. Wolcott, R. G., B. S. Franks, D. M. Hannum, and J. K. Hurst. 1994. Bactericidal potency of hydroxyl radical in physiological environments. J. Biol. Chem. 2699721-9728. [PubMed]
63. Worlitzsch, D., R. Tarran, M. Ulrich, U. Schwab, A. Cekici, K. C. Meyer, P. Birrer, G. Bellon, J. Berger, T. Wei, K. Botzenhart, J. R. Yankaskas, S. Randell, R. C. Boucher, and G. Doring. 2002. Reduced oxygen concentrations in airway mucus contribute to the early and late pathogenesis of Pseudomonas aeruginosa cystic fibrosis airway infection. J. Clin. Investig. 109317-325. [PMC free article] [PubMed]
64. Zhang, S., and B. T. Chait. 2000. ProFound: an expert system for protein identification using mass spectrometric peptide mapping information. Anal. Chem. 722482-2489. [PubMed]
65. Zheng, M., X. Wang, B. Doan, K. A. Lewis, T. D. Schneider, and G. Storz. 2001. Computation-directed identification of OxyR DNA binding sites in Escherichia coli. J. Bacteriol. 1834571-4579. [PMC free article] [PubMed]

Articles from Journal of Bacteriology are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...