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1Institute of Infections and Immunity, Queen's Medical Centre, C-floor West Block, Nottingham NG7 2UH, UK 2Division of Gastroenterology, University Hospital, Nottingham NG7 2UH, UK 3School of Pharmaceutical Sciences, Nottingham University, Nottingham NG7 2RD, UK#dReceived 10 September 2002 Accepted 21 January 2003
Correspondence: Kim Hardie (kim.hardie{at}nottingham.ac.uk)
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Abstract |
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INTRODUCTION |
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TOPAbstractINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Neutrophil-activating protein, NapA (Evans et al., 1995a; Satin et al., 2000), was so designated because of its ability to mediate neutrophil adhesion to endothelial cells (Yoshida et al., 1993) and to bind to both mucin and neutrophil glycosphingolipids (Namavar et al., 1998; Teneberg et al., 1997). However, recently, NapA was shown to bind iron in vitro (Tonello et al., 1999) and to adopt a dodecameric structure consistent with its homology to the Dps family of proteins (Grant et al., 1998). Furthermore, Bijlsma et al. (2000) demonstrated binding of recombinant NapA to DNA in an ELISA in addition to showing that mutation of napA resulted in increased DNA damage in H. pylori.
Many proteins involved in iron acquisition and storage are regulated by the presence of iron in the environment. Dedicated regulatory pathways exist to achieve this, including the ferric uptake regulator (fur) (Braun, 2001; Hantke, 2001). In H. pylori, Fur not only performs its classical role in the regulation of iron uptake (Delany et al., 2001), but also regulates iron storage (Bereswill et al., 2000) and is implicated in iron-independent gene regulation (van Vliet et al., 2001; Bijlsma et al., 2002; Waidner et al., 2002). Since NapA has been shown to be capable of interacting with iron (Tonello et al., 1999) and is implicated in the oxidative stress response in H. pylori (Olczak et al., 2002), and as the presence of free radicals increases in the presence of iron, it is likely that napA is regulated by Fur, in order to sequester iron when abundant and thereby prevent subsequent DNA damage. Lending further weight to this notion, Fur has been shown to be regulated by oxidative stress (Zheng et al., 1999).
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METHODS |
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Stress tests.
H. pylori strains were resuspended to an OD600 of 0.35 in PBS containing 50 mM H2O2, 50 mM H2O2/6.25 µM ethylenediamine-N,N'-diacetic acid (EDDA), 50 mM H2O2/200 µM 2,2'-dipyridyl (DPP) or 50 µM paraquat. Incubation was performed at 37 °C in the VAIN cabinet and duplicate samples were removed for analysis at 0, 0.5, 1, 2 and 4 h. Dilutions between 10-1 and 10-5 were performed in PBS containing 2000 U catalase and aliquots of 8 µl were removed, serially diluted and plated onto blood-agar plates for enumeration. The number of viable cells present before imposition of stress-inducing conditions was taken as 100 % survival and subsequent survival values were calculated relative to this.
Analysis of iron-regulated proteins.
H. pylori strains were subjected to iron stress as described by Bereswill et al. (2000). E. coli strains were grown overnight in LB containing either 6.25 µM ferric sulphate, 0.227 mM EDDA or 0.2 mM DPP and were subjected to fractionation to isolate outer membranes, which were analysed by SDS-PAGE.
Fractionation of H. pylori outer membranes.
Pelleted cells from 1 ml culture were resuspended in 0.1 ml ice-cold 10 mM Tris/HCl (pH 7.4)/20 % (w/v) sucrose containing 0.1 mg lysozyme ml-1. An equal volume of ice-cold 10 mM Tris/HCl (pH 7.4)/1 mM EDTA was then added and the mixture was incubated for 30 min on ice with gentle shaking. Magnesium sulphate was added to a final concentration of 0.2 M and spheroplasts were harvested by centrifugation at 3000 g for 3 min. Following resuspension in 75 µl 20 mM Tris/HCl (pH 7.4)/0.7 % (w/v) sodium lauryl sarcosine and incubation at room temperature for 25 min, outer membranes were pelleted by centrifugation at 3000 g for 30 min, washed in 20 mM Tris/HCl (pH 7.4) and resuspended in the same buffer. Outer-membrane proteins were precipitated with 10 % (v/v) trichloroacetic acid for 60 min on ice prior to SDS-PAGE analysis.
SDS-PAGE and Western blotting.
SDS-PAGE and Western blotting were performed as described in Hardie et al. (1996), except that PBS with 0.5 % (v/v) Tween 20 (PBST) replaced TBST. The primary antibodies anti-NapA (Evans et al., 1995b), kindly provided by Doyle Evans (VA Medical Center, Houston, TX, USA), and our anti-NapA antiserum (see below) were respectively used at dilutions of 1 : 10 000 and 1 : 1000. Western blots were developed using enhanced chemiluminescence (ECL; Amersham) according to the manufacturer's instructions. Whole-cell extracts were prepared by collecting cells by centrifugation for 3 min in a microfuge and resuspending in SDS-PAGE loading buffer. Samples were sonicated for 10 s and boiled for 5 min before being adjusted to equal total protein content according to the OD600 of the harvested bacterial culture, and were then subjected to SDS-PAGE.
DNA manipulation.
DNA was manipulated by standard methods (Sambrook et al., 1989). Restriction enzymes (Promega) were used according to the supplier's instructions. For isolation of plasmid DNA from E. coli, the Qiagen Mini and Midi kits were used. Genomic DNA was extracted from H. pylori according to Atherton (1997). Standard methods were used for preparation of competent cells and for electroporation of plasmids into E. coli (Sambrook et al., 1989). Southern blots were performed using DIG-labelled probes (prepared according to the manufacturer's instructions; Boehringer Mannheim) following PCR amplification from H. pylori strain N6 and pNap2 (see below) using primer pairs nap1 (5'-TGCGATCGTGTTGTTTATG)/nap3 (5'-ATGAGCTTCTAGCATCCAA) and Kan374 (5'-ATAGAA GAAACCCAGGACAATAACC)/KanR778 (5'-ATAGAAGTTCCACAT CATAGGTGG), respectively.
Mutant strain construction.
Following PCR amplification (5 min at 95 °C, 30 cycles of 30 s at 95 °C, 30 s at 50 °C and 2 min at 72 °C, 5 min at 72 °C) of genomic DNA from H. pylori strain 26695 with primers nap1 and nap3, the majority of open reading frame HP0243 (Tomb et al., 1997) was cloned into the pTAg vector using the LigATor kit (Novagen) according to the manufacturer's instructions. The cloned fragment was then recloned into the plasmid vector pUC19 (Amersham Pharmacia Biotech) using the restriction enzymes PstI and XbaI (creating pNap1). A 25 bp deletion and unique BglII site were then engineered into napA by inverse PCR mutagenesis (IPCRM) using primers nap6 (5'-GGC AGATCTAGAATTTCTTTAAAGAT) and nap7 (5'-GGCAGATCTGA ATTTAAAGAGCTCTC) as described previously (Wren et al., 1994). A BamHI fragment containing a 1.4 kb kanamycin-resistance cassette (aph-3) (Trieu-Cuot et al., 1985) was cloned into the unique BglII site generated by IPCRM, creating pNap2.
A mutant in napA was created by natural transformation and allelic exchange mutagenesis as follows. H. pylori strain N6 (Ferrero et al., 1992) was subcultured twice for 24 h on blood-agar plates and the cells were then harvested into 1 ml sterile cold 5 % (w/v) sucrose/15 % (v/v) glycerol. Cells were then washed three times in this buffer and finally resuspended in 30 µl of the same buffer. Plasmid DNA (pNap2) was then added to the cells and electroporation was carried out at 2.4 V, 200 
. Ice-cold ISOSENS broth was then added and cells were incubated on a blood-agar plate for 24 h at 37 °C in the VAIN cabinet. Mutants that had undergone allelic exchange were then selected on plates containing 100 µg kanamycin ml-1 and denoted as H. pylori strain N6 : : napA.
Overproduction of rNapA in E. coli.
For high expression, the napA open reading frame was amplified by PCR from H. pylori strain N6 (5 min at 95 °C, 30 cycles of 30 s at 95 °C, 30 s at 50 °C and 2 min at 72 °C, 5 min at 72 °C) with primers napAFNde (5'-CCCATC CATATGAAAACATTTGAAATTCTAAAACAT) and napAR BamHI (5'-ATGGCAGGATCCTTAAGCCAAATGGGC) and cloned into pET3a (Novagen), creating pCC2. Clones were confirmed by restriction digestion and sequencing. Automated non-radioactive sequencing reactions were carried out using the BigDye terminator cycle-sequencing kit in conjunction with a 373A automated sequencer (Perkin Elmer Applied Biosystems).
Antibody generation.
To supplement the limited supply of anti-NapA (Evans et al., 1995b), a rabbit polyclonal anti-NapA antiserum was generated as follows. H. pylori N6 cells were resuspended in water and incubated for 20 min at room temperature. After centrifugation for 15 min at 17 000 g, the pellet was again incubated in water for 20 min and then centrifuged for 15 min at 25 000 g. The proteins in the resulting supernatant were separated through 15 % SDS-PAGE. NapA was excised from the gel and electroeluted into 50 mM ammonium bicarbonate/0.1 % (w/v) SDS. Following confirmation of the identity of the purified protein by N-terminal sequencing, rabbit polyclonal antibodies were raised according to a protocol based on Harlow & Lane (1988). New Zealand White rabbits were immunized subcutaneously biweekly with between 50 and 400 µg antigen, mixed 50 : 50 with Freund's complete adjuvant on the first immunization and subsequently with Freund's incomplete adjuvant. A test bleed was carried out following the third immunization. A fourth immunization was carried out before finally obtaining complete bleeds from the rabbits. Adsorption of contaminating anti-E. coli antibodies was carried out by incubation at 37 °C for 60 min of 50 µl serum with 1 ml of lysates of E. coli DH5
(pBluescript) and E. coli BL21(DE3)(pLysS,pET3a) prepared in PBSA.
Immunofluorescence.
E. coli strains BL21(DE3)(pLysS,pCC2) and BL21(DE3)(pLysS) were grown in LB containing carbenicillin and chloramphenicol for 3 h. IPTG (1 mM final concentration) was then added and the incubation continued for a further 8 h. One millilitre of culture was added to 10 ml of 80 % (v/v) methanol, mixed very gently and left overnight at 4 °C. Following centrifugation at 2700 r.p.m. for 5 min at 4 °C, cells were resuspended in 1 ml of 80 % (v/v) methanol. An aliquot (10 µl) of this cell suspension was air-dried onto a polylysine-coated slide for 20 min and then covered with 50 µl of 25 mM Tris/HCl (pH 8)/50 mM glucose/10 mM EDTA containing 2 mg lysozyme ml-1 and incubated at room temperature for 5 min. The slides were then covered sequentially in 4 ml of 99 % (v/v) methanol for 1 min and 4 ml acetone for 1 min before air-drying. Fifty microlitres PBST containing 2 % (w/v) BSA was then applied and covered with a cover slip. Following a 15 min incubation at room temperature, 50 µl PBST/BSA containing a 1 : 200 dilution of our anti-NapA antiserum was applied and the slide was incubated for 1 h in a moist chamber. Following three washes in PBST, 50 µl PBST/BSA containing a 1 : 500 dilution of FITC-conjugated protein A (Sigma) was applied and the slide was incubated for 1 h in a moist chamber. Following three washes in 5 ml PBST, 10 µl 10 µg 4,6-diaminidino-2-phenylindole (DAPI) ml-1 was added and the slide was incubated for 1 min before removal and air-drying. Samples were analysed by phase-contrast and fluorescence microscopy.
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RESULTS |
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TOPAbstractINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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To determine the temporal production of NapA, H. pylori strain N6 was grown until late stationary phase and whole-cell samples were removed at selected time-points for Western blot analysis following adjustment for total cell number (Fig. 1). NapA production occurred throughout the growth of H. pylori, but was enriched in stationary phase, consistent with its proposed functions in iron storage and DNA protection.
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Mutation of napA decreases the survival of H. pylori in the presence of oxidative stress
In order to assess the function of NapA in vivo, a mutant of napA was created in H. pylori strain N6 (Ferrero et al., 1992) as described in Methods and confirmed by PCR, Southern blot and Western blot analysis (data not shown, see Fig. 4b). Mutation of napA did not result in a severe growth impediment, as evidenced by the similar OD600 reached after growth for 48 h in Brucella broth supplemented with FCS and either 500 µM NaCl or 100 or 500 µM iron chloride (OD600 = 1.04, 1.46, 1.36, respectively), compared with the parent strain H. pylori N6 (OD600 = 1.08, 1.1, 1.2, respectively; see Fig. 4 for description of methods employed).
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Although mutation of napA was well tolerated when H. pylori is grown in nutrient-rich conditions, the survival of the mutant was reduced in adverse conditions. The wild-type H. pylori strain N6 and the napA mutant N6 : : napA were exposed to hydrogen peroxide in broth culture and viable counts were taken over the subsequent 4 h (see Methods). Fig. 2 shows that the wild-type strain survived significantly better under these conditions. Similar results were observed when the wild-type and mutant strains were challenged in broth cultures with 50 µM paraquat, which generates superoxide ions rather than hydroxyl free radicals (Fig. 2). One of the major effectors of DNA damage (leading ultimately to cell death) in oxidative conditions is free intracellular ferrous iron. The homology of NapA to the iron-storage protein bacterioferritin led us to hypothesize that, in the absence of iron (generated by iron chelators), the lack of NapA would not be as detrimental. However, addition of the iron chelators DPP and EDDA to the growth medium simultaneously with hydrogen peroxide did not improve the viability of the H. pylori napA mutant reproducibly (data not shown).
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Identification of putative fur boxes within the napA promoter
Despite its small size, the H. pylori genome encodes a number of systems involved in iron uptake and storage, including the regulatory protein Fur (Tomb et al., 1997). We observed that there are several A+T-rich regions of dyad symmetry immediately upstream of the start codon of napA that could function as a fur box. We used the consensus H. pylori fur box sequence proposed recently by Delany et al. (2001) to identify putative fur boxes centred between 20 and 180 bp upstream of genes in H. pylori strain J99 (Alm et al., 1999) involved in iron storage and detoxification using the analysis tools available at http://rsat.ulb.ac.be/rsat/. Searches of the 200 bp upstream of napA with this consensus (NtaTNaN5ttTT aatNAaAATNataAaANatT) identified two regions (Fig. 3a), each with 8/10 of the absolutely conserved bases (indicated in upper case) plus 11 of the pyrimidine bases (indicated in lower case). The two missing conserved bases were replaced by a pyrimidine. One of the putative fur boxes was centred at position -167 and the other at -111 from the ATG. The spacing from the mapped transcription start site (Olczak et al., 2002) is also indicated in Fig. 3(a), and parallel searches revealed the fur boxes located experimentally by Delany et al. (2001) upstream of frpB1 (Fig. 3b).
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NapA is regulated by fur
Since putative fur boxes are located upstream of napA (see above), we investigated whether the production of NapA (like that of the H. pylori bacterioferritin, Pfr; Bereswill et al., 1998) was regulated by iron via Fur. The H. pylori fur mutant 11638 : : fur and its parent, NCTC 11638 (Bereswill et al., 2000; Bijlsma et al., 2002), were grown overnight in Brucella broth containing 10 % FCS and supplemented with either 100 µM FeCl3, 500 µM FeCl3, 40 µM of the iron chelator desferal or 500 µM NaCl to exclude any effect of osmotic stress. To assess the effect of NapA upon survival under these conditions, H. pylori strains N6 and N6 : : napA were analysed similarly. Whole-cell extracts containing matched total protein contents were prepared and SDS-PAGE revealed modulation of the production of a 19 kDa protein, presumed to be Pfr by analogy with the data reported by Bereswill et al. (2000), in response to iron availability (Fig. 4a). As reported previously, when the growth medium was supplemented with desferal, the level of Pfr was higher in a fur mutant than in the parent strain. Also in accordance with this previous study, the OD of cultures grown in the presence of desferal was considerably reduced (OD600 = 0.3, 0.3, 0.2 and 0.1 for H. pylori strains N6, N6 : : napA, NCTC 11638 and 11638 : : fur, respectively, compared with 1.1–1.4 in the absence of desferal). Although the levels of many proteins appeared to stay the same or to fall in the presence of desferal, one in particular, at 66 kDa, displayed a marked increase under these conditions in both strains. Western blots demonstrated that NapA production in H. pylori strains N6 and NCTC 11638 is regulated by iron in a manner similar to that demonstrated previously for Pfr. In H. pylori strain 11638 : : fur, NapA levels were at least twofold higher under all growth conditions and increased most significantly (approximately fivefold) in the presence of desferal (Bereswill et al., 2000; Fig. 4b). NapA is absent from N6 : : napA (Fig. 4b).
Since NapA is predicted by homology to bacterioferritin to bind iron, although it does not possess an active ferroxidase centre, we hypothesized that its production might influence the expression of other iron-regulated proteins. However, no effect was observed on the expression of iron-regulated proteins such as the visible 19 kDa (presumed Pfr) protein when the H. pylori napA mutant was compared with its parent strain (data not shown). Similarly, high-level production of soluble recombinant NapA in E. coli did not induce the expression of iron-regulated proteins, despite the possibility that this would lower intracellular iron concentrations (data not shown).
NapA co-localizes with cellular DNA in vivo
The use of fluorescent microscopy has allowed us to visualize the co-localization of rNapA with DNA directly within the living E. coli cell and also demonstrates that, in so doing, the DNA is concentrated to one part of the cell. E. coli BL21(DE3)(pLysS) harbouring either pCC2 (encoding napA) or the empty control plasmid vector (pET3a) was grown for 8 h and the production of rNapA by the former was verified by SDS-PAGE (Fig. 5) and Western blot analysis (data not shown). When cells were lysed in a French pressure cell at 12 000 p.s.i. and insoluble material was pelleted by centrifugation at 10 000 g for 10 min, more than 90 % of the rNapA detected was present in soluble cell fractions and, thus, the high-level production was not leading to the formation of inclusion bodies (data not shown). DNA was stained with DAPI and the cells were probed with anti-NapA antisera and FITC-conjugated protein A. Microscopy was then performed to visualize the cells by phase-contrast, plus the distribution of DAPI and FITC. Fig. 5 shows that, in BL21(DE3)(pLysS,pCC2), the DAPI (Fig. 5b) and FITC (Fig. 5c) co-localized to a region of visually increased density seen in phase-contrast images (Fig. 5a). In contrast, with strain BL21(DE3)(pLysS,pET3a), no areas of increased density were seen by phase-contrast (Fig. 5d), the DNA appeared to be generally spread throughout the cell, except for occasional polar concentration in some cells, and the FITC staining was at background levels. Similar results were obtained on three separate occasions, suggesting that the overproduced rNapA protein co-localizes with DNA and may indeed induce compartmentalization of the DNA within the cell. The observed compartmentalization of the DNA was not due to rNapA simultaneously forming insoluble inclusion bodies. The two strains grew comparably, as judged by OD600 measurements taken throughout growth (data not shown). Unfortunately, technical problems prevented us from observing this effect in H. pylori.
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DISCUSSION |
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The presence of multiple fur boxes within the napA promoter is not unusual (Fig. 3) and has also been noted experimentally (Delany et al., 2001). It is possible that one of the fur boxes upstream of napA serves as a high-affinity, iron-regulated gene regulator, whilst the other binds Fur with a low affinity and is not influenced by iron availability, in a manner similar to that described for frpB1 by Delany et al. (2001). Indeed, Fur has been shown to respond to environmental signals other than iron concentration. For instance, Fur has been implicated in regulation of H. pylori genes in response to a range of conditions unrelated to iron availability, such as the modulation of H. pylori urease in response to nickel (van Vliet et al., 2001), differential regulation of pfr in the presence of a range of different metals (Bereswill et al., 2000) and H. pylori acid resistance (Bijlsma et al., 2002). The sequence variation within the proposed fur boxes is not unusual, since Fur appears to have a flexible mode of DNA interaction, providing it with the ability to behave both as a very specific repressor and as a more general regulator (de Lorenzo et al., 1988; Escolar et al., 1999). Experimental confirmation is required to identify these sequences as true fur boxes and to evaluate their relative contribution to recognition of the napA promoter by Fur.
Although it would be predicted that the Pfr ferritin of H. pylori is regulated by iron availability via fur, there is disagreement in the literature. Bereswill et al. (2000) demonstrated, using the chelator desferal, that Pfr was repressed by the removal of iron, whereas Dundon et al. (2001) reported that they could not show this repression. Since Dundon et al. (2001) were also unable to demonstrate an effect of desferal on NapA, they concluded that fur was not involved in napA regulation. The inconsistency in these results may reflect different growth conditions, since Bereswill et al. (2000) grew their H. pylori cultures in broth supplemented with FCS, whilst Dundon et al. (2001) appeared to supplement H. pylori broth cultures with cyclodextrin. Cyclodextrin affects the iron-chelating properties of desferal, negating its effect upon cells and leading to an absence of expected iron-regulation (A. H. M. van Vliet, personal communication). We were able to repeat the results of Bereswill et al. (2000), indicating iron-dependent regulation of Pfr, identify putative fur boxes upstream of the pfr and napA genes (Figs 3b and 4) and, in parallel, demonstrate repression of napA by desferal. This would suggest that fur does regulate napA, and this was confirmed by our demonstration that the effect of desferal upon NapA production is less evident in a fur mutant of H. pylori. There appears to be approximately fivefold more NapA in the presence of desferal when an H. pylori strain deficient in fur is analysed. This difference is not a reflection of growth-phase-dependent NapA production noted earlier, since the optical densities of the cultures analysed were comparable. Moreover, since the level of NapA is elevated under all the growth conditions analysed in the H. pylori fur mutant compared with its parent, it can be concluded that Fur influences the production of NapA. However, since the H. pylori fur mutant retained the ability to modulate the level of NapA in response to the availability of iron, additional regulatory influences must be at work, e.g. aconitases (Hantke, 2001).
Under conditions of excess iron, the fur mutant displayed a smear of proteins at the expected mobility for Pfr (Fig. 4). This is likely to be due to (i) the ability of ferritin to assume a number of different conformational forms depending on the extent of iron loading due to oxidation (Welch et al., 2001), particularly since such isoforms have been described for H. pylori Pfr by Waidner et al. (2002), and (ii) the susceptibility of H. pylori Pfr to post-translational modification (Waidner et al., 2002). In agreement with the former suggestion, the dispersed nature of the 19 kDa protein was less evident when cultures were supplemented with NaCl to control for osmotic effects on protein production and thus contained lower iron concentrations. It is not entirely surprising that overproduction (recombinant NapA in E. coli) or absence (H. pylori napA mutant) of a single iron-binding protein did not result in alteration of the levels of proteins shown to be regulated by iron, since the total iron pool would neither increase nor be depleted thanks to the many systems present to deal with iron, ensuring the provision of iron in an accessible form and avoiding its potentially toxic effects. The presence of multiple iron-recycling pathways and their careful regulation result in compensatory modulation of these interconnected systems, allowing cells to balance iron storage and removal to maintain optimal growth (Abdul-Tehrani et al., 1999).
For the first time, co-localization of NapA with cellular DNA was demonstrated in vivo, as has been shown for the other members of the Dps family of proteins (Azam et al., 2000). Previous attempts to show this in vitro by DNA mobility assays and ELISA have met with varying success (Tonello et al., 1999; Bijlsma et al., 2000). Because in vitro analysis of DNA interactions requires the maintenance of protein function during overproduction and purification, failure to observe binding is not conclusive proof of its absence. Therefore, we chose to investigate whether NapA interacts with DNA in vivo. Indirect evidence exists that suggests that NapA interacts with DNA, resulting in protection against DNA damage (Bijlsma et al., 2000); we were able to support this observation by co-localizing NapA with DNA through fluorescence microscopy using specific antisera. The association between NapA and DNA appeared to cause a condensation of the DNA within one part of the cell, which was not the result of NapA forming insoluble inclusion bodies. This condensation may be due to the high level of rNapA in the cells (approx. 80 % of total protein), however, as Dps was shown previously to be evenly distributed within E. coli cells (Azam et al., 2000). This observation is consistent with DNA wrapping around a hexameric array of NapA in a pattern similar to that proposed for Dps (Grant et al., 1998).
In summary, we have shown that NapA production is affected by Fur and, consistent with its homology to Dps, it is produced maximally in stationary phase and co-localizes with DNA, causing it to accumulate in one area of the bacterial cell. This is likely to protect the DNA from damage by free radicals, a suggestion that is borne out by the loss of viability of an H. pylori napA mutant when exposed to oxidative stress. It will be interesting to investigate whether the protective role of NapA facilitates the pathogenicity of H. pylori.
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Acknowledgments |
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 G. Wang, R. C. Conover, S. Benoit, A. A. Olczak, J. W. Olson, M. K. Johnson, and R. J. Maier Role of a Bacterial Organic Hydroperoxide Detoxification System in Preventing Catalase Inactivation J. Biol. Chem., December 10, 2004; 279(50): 51908 - 51914. [Abstract] [Full Text] [PDF]
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J. Seshu, J. A. Boylan, F. C. Gherardini, and J. T. Skare Dissolved Oxygen Levels Alter Gene Expression and Antigen Profiles in Borrelia burgdorferi Infect. Immun., March 1, 2004; 72(3): 1580 - 1586. [Abstract] [Full Text] [PDF]
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D. S. Merrell, L. J. Thompson, C. C. Kim, H. Mitchell, L. S. Tompkins, A. Lee, and S. Falkow Growth Phase-Dependent Response of Helicobacter pylori to Iron Starvation Infect. Immun., November 1, 2003; 71(11): 6510 - 6525. [Abstract] [Full Text] [PDF]
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, napA mutant) were subjected to oxidative stress induced by the presence of 50 mM hydrogen peroxide (a) or 50 µM paraquat (b). At the times indicated, samples were removed and survival was assessed by calculating the number of c.f.u. present relative to those present initially (see Methods). The graph shows the log10 percentage survival plotted against the time of exposure to hydrogen peroxide or paraquat. Data shown are representative of four independent experiments and the standard errors of the mean c.f.u. are indicated by the error bars.
