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1 Division of Environmental Science and Ecological Engineering, Korea University, Anam-Dong 5 Ga, Seoul, South Korea
2 Department of Biotechnology, Yonsei University, Sinchon-Dong, Seoul, South Korea
3 Environmental Biotechnology National Core Research Center, Gyeongsang National University, Jinju, South Korea
4 Department of Microbiology, Cornell University, Ithaca, NY 14853-8101, USA
Correspondence
Woojun Park
wpark{at}korea.ac.kr
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Price et al., 2000; Stohs et al., 2002), and even to naphthalene-metabolizing micro-organisms (Ahn et al., 1998; Garcia et al., 1998; Murphy & Stone, 1955; Park et al., 2004; Pumphrey & Madsen, 2007). Evidence has begun to accumulate showing that inefficient naphthalene biodegradation, either by stationary-phase cells or in the presence of toxic compounds, can generate toxic metabolic intermediates or reactive oxygen species (ROS) (Kang et al., 2006; Park et al., 2004). Many iron–sulfur proteins are involved in naphthalene biodegradation (Karlsson et al., 2003). Under oxidative stress conditions, the iron in haem and iron–sulfur proteins can be oxidized, which may lead to protein inactivation (Imlay, 2003). In our previous work (Kang et al., 2006), the antioxidants ascorbate and ferrous ion were found to lessen the inhibitory effects of ROS generated during naphthalene metabolism.
Among the microbial defence systems against oxidative stress are the antioxidant enzymes superoxide dismutase (SOD) and ferredoxin-NADP+ reductase (Fpr). SOD catalyses the conversion of superoxide to H2O2 and O2 (Giro et al., 2006; Hausladen & Fridovich, 1994). Fpr mediates reversible redox reactions between a single molecule of NADP+/NADPH and two molecules of one-electron carriers, such as ferredoxin or flavodoxin (Bianchi et al., 1995; Lee et al., 2006b). Fpr is involved not only in scavenging ROS but also in the repair of iron–sulfur centres under oxidative stress conditions. Djaman et al. (2004) reported that Escherichia coli mutants lacking SOD or Fpr repaired their iron–sulfur centre-containing enzymes more slowly than the wild-type strain. Thus, the fpr gene confers resistance to oxidative stress in some bacteria (Giro et al., 2006; Lee et al., 2006b). It has been reported that both fpr and sodA are inducible under oxidative stress in Pseudomonas strains (Kim et al., 2000; Lee et al., 2006b). Therefore, our hypothesis is that the overexpression of these antioxidant enzymes may be effective in scavenging the ROS generated during naphthalene degradation. To gain insights into the roles of antioxidant enzymes in the defensive mechanism against oxidative stress, the sod, fpr and ahpC genes were cloned into plasmid vectors that can replicate in Pseudomonas sp. strain As1, which was recently isolated from a pollutant-contaminated site (Kang et al., 2006). In addition, reporter strains prepared by fusing the green fluorescent protein gene (gfp) to the fpr promoter region were used to confirm that oxidative stress occurs during growth on naphthalene and can be lessened by overexpression of Fpr and SOD.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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. Bacteria were grown at 37 °C (E. coli) or 30 °C (Pseudomonas sp. strain As1) in Luria–Bertani (LB) broth or minimal salts basal medium (MSB; Stanier et al., 1966) with vigorous aeration by shaking at 220 r.p.m. MSB medium was supplemented with different carbon sources [naphthalene (0.5 %, w/v), salicylate (2 mM), glucose (2 mM) or pyruvate (2 mM)]. When required, antibiotics were added at the following concentrations: 100 µg ampicillin ml–1; 100 µg kanamycin ml–1; 50 µg tetracycline ml–1. Paraquat (PQ) was added at 300 µM. Growth was monitored by measuring the OD600 of the cultures using a Biophotometer (Eppendorf). Plasmid isolation, gel electrophoresis, transformation and PCR DNA amplification were performed as described previously (Kang et al., 2006).
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), was used for constructing the recombinant plasmids, pBBRfpr, pBBRfpr-R, pBBRsodA and pBBRsodA-R (Fig. 1a). In this expression plasmid, fpr was placed under control of both the lacZ and fpr promoters (pBBRfpr) or only under the control of the native promoter (pBBRfpr-R). Similarly, sodA was placed under control of both the lacZ and sodA promoters (pBBRsodA) or only under the control of the native promoter (pBBRsodA-R). Each plasmid was transformed into E. coli TOP10. Conjugation was performed by triparental filter mating with E. coli HB101(pRK2013) (Figurski & Helinski, 1979) and Pseudomonas sp. strain As1, thus creating strains As1(fpr), As1(fpr-R), As1(sodA) and As1(sodA-R). The transconjugants were selected on LB agar plates, containing 100 µg ampicillin ml–1 and 100 µg kanamycin ml–1, at 30 °C.
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). The constructed plasmid was introduced into strain As1.
Reporter strains to monitor oxidative stress were prepared by fusing the gfp gene to the fpr promoter. The promoter region of the fpr gene, including the gene encoding FinR, a regulator protein, from Pseudomonas syringae DC3000 was amplified with the Pfnr-gfp1/Pfnr-gfp2 primer pair (CGCGAATTCGGCCATCGACCGCATTTCT/CGCGGTACCTGGACGCTGAGGACACG) (yielding a 1234 bp fragment), and the fragment was cloned into the EcoRI and KpnI cloning site of the pRK415gfp vector (Yin et al., 2003), generating pDCFPRgfp. Conjugation was performed by triparental filter mating with helper strain E. coli HB101(pRK2013) and the above-described recombinant strains. The transconjugants were selected on LB agar plates, containing kanamycin (100 µg ml–1) and tetracycline (50 µg ml–1), at 30 °C. Transformation of the plasmid was confirmed by antibiotic resistance and PCR, using appropriate primer pairs (data not shown).
Northern blot analysis.
Total RNA of cells cultured with or without PQ (300 µM) was extracted at the early-exponential phase (OD600 
0.3) and isolated using an RNeasy kit (Qiagen), following the manufacturer's instructions. RNA concentrations were estimated by absorbance at 260 nm. Samples of total RNA (10 µg) were loaded on denaturing agarose gels containing 0.25 M formaldehyde, separated, and stained with ethidium bromide to visualize 23S and 16S rRNA. The fractionated RNA was transferred to nylon membranes (Schleicher & Schuell) using a Turboblotter (Schleicher & Schuell). The amount of fpr mRNA was determined by hybridizing the membrane with a specific 32P-labelled probe (Invitrogen) prepared by PCR amplification from P. putida strain KT2440 with the primer pair fpr-K01/fpr-K02 (CGCGAATTCCAGGCGTTTGGTTTTCTCAC/CGCGAATTCAGCCCTGCCGCCTTCTCC, 356 bp). The probes for sodA and ahpC mRNA were prepared by PCR amplification with the primer pairs sodA-F/sodA-R [primers for As1(sodA) overexpression] and ahpC-F/ahpC-R, respectively. The band intensity of Northern blot data was measured using a densitometry instrument (800x1600 d.p.i., UMAX, UTA 2100XL, USA) as previously described (Kim et al., 2006b).
HPLC analysis.
A saturated (30 p.p.m.) aqueous solution of naphthalene was used for HPLC analysis. Saturated naphthalene solution was mixed (1 : 1 ratio) with 2x MSB medium before adding 107 cells ml–1 (glucose-grown overnight cultures) to culture media. Residual naphthalene concentration was analysed by HPLC as previously described (Park et al., 2004) except for the following modifications. Samples (0.4 ml) of culture medium were collected at various time points (5 h intervals) and immediately diluted with an equal volume of methanol. Samples were filtered through nylon 66 filters (0.45 µm, 13 mm syringe filter, Whatman). Analytes were separated using a J'sphere ODS-H80 column (150x4.6 mm, YMC) at room temperature with a Younglin model SP930D pump. The mobile phase was 70 % acetonitrile at a flow rate of 1.0 ml min–1. Constituents eluting from the column were detected at 254 nm using a Younglin UV730D detector.
Survival assay.
PQ survival experiments were carried out in LB medium, with shaking at 220 r.p.m. at 30 °C. Stationary-phase cultures (50 µl) were inoculated into 5 ml LB liquid medium at 30 °C with agitation (220 r.p.m.). Then PQ (300 µM final concentration) was added during the exponential phase (OD600 0.3–0.5). Cells were harvested from each culture at 1 h intervals and washed with autoclaved phosphate-buffered saline (PBS; pH 7.5). The harvested cells were diluted and serially plated on LB agar. Agar plates were incubated, lid down, at 30 °C for 16 h before colonies were counted.
Enzyme assays.
Catalase activity was measured by monitoring the decrease in A240 resulting from the elimination of H2O2, using a UV–visible spectrophotometer (Optizen 2120, Mecasys, Korea). The absorption coefficient (
) for H2O2 at 240 nm was 43.6 M–1cm–1 (Beers & Sizer, 1952). The standard reaction mixture for the assay contained 50 mM potassium phosphate buffer (pH 7.2), 20 mM H2O2 and 20 µl crude extract in a total volume of 3.0 ml. The reaction was performed at 25 °C. The amount of enzyme activity that decomposed 1 µM H2O2 min–1 was defined as 1 unit (U) of activity (Beers & Sizer 1952). Superoxide dismutase (SOD) activity staining was performed as described previously (Beauchamp & Fridovich, 1971; Kang et al., 2006). Supernatants of cell extracts were loaded onto a 7 % native polyacrylamide gel in a running buffer made of 25 mM Tris and 192 mM glycine. Proteins (15 µg samples) were resolved at 20 mA for 2 h. Subsequently, the gels were processed for SOD activity. The gels were first soaked in 2.5 mM nitro blue tetrazolium (NBT) for 10 min in darkness under gentle shaking. They were then incubated in 50 mM potassium phosphate buffer (pH 7.2) containing 28 mM N,N,N',N'-tetramethylethylenediamine (TEMED) and 28 µM riboflavin for 15 min, in the dark and under constant agitation. The SOD activity appeared as a white band on the blue background gel. Aconitase activity (Giro et al., 2006) was measured at 25 °C in 90 mM Tris/HCl, pH 7.5, 20 mM sodium isocitrate, by following the formation of cis-aconitate at 240 nm (240=3.6 mM–1 cm–1), using a UV–visible spectrophotometer (Optizen 2120, Mecasys). The wild-type and recombinant strains were treated with PQ for 30 min, then transferred to PQ-free LB medium and the aconitase activity measured in a time-dependent manner.
Quantification of GFP fluorescence.
Bacterial cells containing gfp plasmids (Table 1) were grown in LB broth (+ 300 µM PQ) or MSB medium supplemented with glucose (2 mM), pyruvate (2 mM), salicylate (2 mM) or naphthalene (0.5 %). After exposure to PQ for 3 h, a 2 ml cell suspension was collected using a microcentrifuge (13 000 r.p.m.) and washed twice with 1 ml PBS. The optical density of resuspended cultures was then measured and GFP fluorescence intensities were quantified using a microtitre plate reader (VICTOR3, BioRad). The reporter strain expresses a stable GFP variant that absorbs at 488 nm. The microscope was equipped with a fluorescence filter cube for detecting GFP (filter set 38 GFP, Carl Zeiss). The AxioVision software was used to acquire images.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). The expression of these antioxidant enzyme genes in each recombinant strain was confirmed by Northern blot analysis (Fig. 1b). As expected, the plasmids led to elevated expression of both fpr and sodA. fpr expression was clearly highest with the lac promoter, while expression of sodA was high with both promoters (Fig. 1b). The addition of PQ induced both genes (Fig. 1b). The transcript levels of strains As1(fpr) and As1(fpr-R) grown in the presence of PQ increased 1.9- and 1.8-fold compared to cells without PQ. Strain As1(sodA) expressed more sodA (1.6-fold), compared to strain As1(sodA-R) in LB medium. In the presence of PQ, sodA expression in strain As1(sodA-R) increased 1.7-fold, compared to cells grown in LB medium. However, PQ addition only slightly induced sodA expression in strain As1(sodA), probably because high expression of sodA already prevailed in the absence of PQ. The data showed that the fpr and sodA genes were highly inducible by PQ. A much lower mRNA level of indigenous sodA and fpr from wild-type strain As1 was detected, possibly due to the probes, which were designed based on the genes present in P. putida KT2440, whose genome has been completely sequenced (NCBI, NC002947).
The growth rate of cells overexpressing antioxidant enzymes was measured during naphthalene metabolism. Cells were grown at 30 °C in MSB medium supplemented with 0.5 % naphthalene. The growth rates (h–1) of the wild-type (As1) and four recombinant strains [As1(fpr), As1(fpr-R), As1(sodA) and As1(sodA-R)] were 0.17±0.01, 0.29±0.02, 0.23±0.04, 0.26±0.03 and 0.25±0.04, respectively. In separate flasks, the rate of aqueous-phase naphthalene metabolism was monitored using HPLC analysis (data not shown). Naphthalene consumption was most rapid (20 % remaining after 15 h) by strains As1(fpr) and As1(sodA) and slowest for the wild-type (40 % remaining after 15 h). Thus, the growth-rate and HPLC analyses indicated that naphthalene degradation efficiency was improved in all recombinant cells, compared to the wild-type. In addition, the expression of the nahAc gene (encoding naphthalene dioxygenase) was increased in all recombinant cells (Fig. 1c). The nahAc mRNA levels of strains As1(fpr), As1(fpr-R), As1(sodA) and As1(sodA-R) increased 3.0-, 3.2-, 2.9- and 3.1-fold compared to wild-type, respectively. This was consistent with previous results showing that the nahAc expression levels in Pseudomonas sp. strain O1 were highest after the addition of antioxidants (Kang et al., 2006).
Survival of bacterial cells exposed to PQ
The resistance of recombinant cells to PQ, a superoxide generator, was investigated (Fig. 2a). As expected, the parental strain (As1), was inhibited by PQ treatment after 5 h of growth. Strains As1(fpr), As1(fpr-R) (not shown) and As1(sodA-R) (Fig. 2a) were resistant to 300 µM PQ, as they maintained high growth rates and achieved high cell densities. Surprisingly, the growth of strain As1(sodA) was also severely inhibited by the addition of PQ (Fig. 2a).
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). Strain As1 harbouring an empty vector (pBBR1MCS2) was used as a control. The wild-type cells, and strains As1(pBBR1MCS2) and As1(sodA) were sensitive to PQ, as no growth was observed on the plate, while strains As1(fpr) and As1(fpr-R) were resistant to PQ. Survival of wild-type and recombinant cells exposed to PQ was assessed in LB medium (Fig. 2c): growth of strain As1(sodA-R) was not affected by PQ (Fig. 2c), whereas death of the wild-type and As1(sodA) strains occurred in the presence of PQ, possibly due to accumulation of the SOD by-product, hydrogen peroxide.
We used strain As1(ahpCsodA) (Fig. 1a) to test the hypothesis that PQ-induced death of strain As1(sodA) was due to excess intracellular hydrogen peroxide. We reasoned that overexpression of alkyl hydroperoxide reductase (AhpC) would consume peroxides, thereby compensating for SOD overactivity. We constructed strain As1(ahpCsodA) overexpressing AhpC and confirmed this strain using Northern blot analysis. As expected, the inserted ahpC gene was overexpressed in both the absence and presence of PQ (Fig. 2d). Indeed, the data from strain As1(ahpCsodA; Fig. 2c) show that overexpression of AhpC rendered cells more resistant to PQ than strain As1(sodA).
Analysis of SOD, catalase and aconitase activities
In order to confirm the above-shown relationships between SOD and alkyl hydroperoxide reductase, these enzymes were measured directly. Elevated SOD activity was measured in strains As1(sodA-R) and As1(sodA), which have additional copies of the SOD gene. After growth in LB medium without exposure to PQ, the wild-type and strains As1(fpr), and As1(fpr-R) showed low levels of SOD activity (Fig. 3a). However, as expected, the SOD activity in strains As1(sodA-R) and As1(sodA) was substantially increased (Fig. 3a): for strain As1(sodA), the increase was 4.3-, 3.0- and 3.7-fold [compared to wild-type, As1(fpr), and As1(fpr-R), respectively] and for strain As1(sodA-R), the increase was 2.7-, 1.9- and 2.3-fold [compared to wild-type, As1(fpr) and As1(fpr-R), respectively]. The additional band exhibiting SOD activity occurred because of the higher mass of the protein produced by the introduced sod gene. In the presence of PQ, strains As1(sodA) and As1(sodA-R) showed even higher SOD expression (Fig. 3a), likely alleviating the oxidative stress generated by PQ. The SOD levels of strains As1(fpr), As1(fpr-R), As1(sodA) and As1(sodA-R) increased 4.6-, 3.9-, 10.5- and 9.8-fold, respectively, compared to wild-type.
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). Interestingly, both the SOD and catalase activities were highly induced in all recombinant strains during naphthalene metabolism (Fig. 3b). The SOD expression of strains As1(fpr), As1(fpr-R), As1(sodA) and As1(sodA-R) was induced 1.4-, 1.4-, 2.4- and 2.1-fold compared to wild-type. These results indicate that elevated levels of these enzymes likely routinely counteract the oxidative stress generated during naphthalene metabolism.
Aconitase is a dehydratase enzyme containing a [4Fe–4S] cluster. Aconitase was chosen to investigate whether overexpressed Fpr was involved in the recovery of oxidative damage. Recovery of aconitase activity in all recombinant strains was accelerated compared to the wild-type (Fig. 3c). The aconitase activities of strains As1(fpr) and As1(sodA-R) returned to their initial levels very rapidly – within 10 min. The aconitase activities of strains As1(fpr-R) and As1(sodA) were also regenerated to about 80 % of their original levels within 20 min of incubation. The wild-type strain showed a very slow aconitase activity regeneration rate ( 50 % of the original value in 30 min) after being challenged by oxidative stress.
Responses of oxidative-stress reporter strains (fpr–gfp fusions) under different carbon source and oxidative stresses
The effect of different carbon sources on the degree of oxidative stress in Pseudomonas sp. strain As1 was examined using several GFP-fused reporter strains. This reporter assay is based on the inducibility of the fpr promoter in P. syringae DC3000. Previously, our data showed that the fpr promoter of P. syringae DC3000 is highly inducible in an oxidant-concentration-dependent manner (Lee et al., 2006b). The normalized GFP intensity (GFP intensity/OD600; Lee et al., 2006a) of the wild-type Pseudomonas sp. strain As1 cells grown in naphthalene-amended medium increased markedly (14-fold) compared to pyruvate- or glucose-grown cells (Fig. 4a). The GFP level in salicylate-amended media also increased 21-fold. GFP fluorescence was also measured in wild-type and recombinant strains during naphthalene degradation. The GFP intensity of all recombinant cells was below that of wild-type (Fig. 4b). This result shows that cells of strain As1 exhibit oxidative stress during naphthalene or salicylate degradation. GFP expression levels were measured in cells overexpressing antioxidant enzymes treated with PQ (Fig. 4c). The GFP level in the wild-type showed a significant increase compared to all the recombinant strains in the absence of PQ (Fig. 4c, white bars), indicating that overexpressed antioxidant enzymes play a role in detoxifying the ROS generated under normal growth conditions. The GFP levels of the wild-type and strain As1(sodA) were dramatically increased in the presence of PQ compared to the other recombinant strains, as shown by GFP intensity patterns in fluorescence photomicrographs (Fig. 4c).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Kim et al., 2006a; Lee, 1999) and simultaneously lead to the expression of native antioxidant enzymes such as those examined here. In our previous study using Pseudomonas sp. strain O1 (Kang et al., 2006) we demonstrated that SOD and catalase activities were higher in naphthalene- vs glucose-grown cells. These studies indicate that cells have defence systems that partially guard against oxidative stress.
Furthermore, addition of two antioxidants (ascorbate and ferrous ion) to cells growing on naphthalene had three effects: (i) accelerated cell growth; (ii) increased rate of naphthalene consumption; and (iii) higher expressed levels of naphthalene dioxygenase (nahAc) transcripts (Kang et al., 2006). The present study confirmed and extended the work of Kang et al. (2006) by demonstrating elevated SOD and catalase levels in naphthalene-grown cells (Fig. 3) and GFP intensity in oxidative-stress reporter strains of Pseudomonas sp. strain As1 grown on naphthalene and salicylate (Fig. 4). Strain As1 was chosen in this study because we have shown that it grows well in the presence of both naphthalene and salicylate (Kang et al., 2006). Furthermore, strain As1 has higher tolerance to salicylate than many other isolates. This property may be an important feature in microbial biodegradation.
Salicylate is known to cause uncoupling of oxidative phosphorylation (Saxena et al., 1995), and is also toxic as a naphthalene metabolite (Kang et al., 2006; Price et al., 2000). In eukaryotes, salicylic acid causes a significant increase in oxygen uptake, which then generates hydrogen peroxide or other ROS (Battaglia et al., 2005). We hypothesized that the overexpression of antioxidant enzymes (Fpr or SodA) may be effective in relieving oxidative stress generated during naphthalene degradation. Antioxidant enzymes (Fpr or SodA) were overproduced in the naphthalene-degrading bacterium Pseudomonas sp. strain As1 (Fig. 1). As we expected, growth rates, naphthalene-degradation efficiency and naphthalene-catabolic gene expression were improved in all recombinant cells, compared to the wild-type (Fig. 1). These data provide physiological evidence that oxidative stress during naphthalene metabolism can be alleviated by the overproduction of antioxidant enzymes.
Treatment of our fpr and sodA recombinant strains with the superoxide-generating reagent PQ led to strain-specific responses. Overexpression of Fpr [strains As1(fpr) and As1(fpr-R)] resulted in resistance to PQ (Fig. 2b). Consistent with these results, Fpr overproduction also boosted cell viability in PQ-exposed E. coli cells (Bianchi et al., 1995; Giro et al., 2006). By contrast, overexpression of SOD [strain As1(sodA)] produced sensitivity to PQ (Fig. 2). SOD overexpression in E. coli has produced variable results: beneficial effects against PQ-mediated oxidative damage (Bhattacharya et al., 2004; Goulielmos et al., 2003); but also increased sensitivity to PQ (Scott et al., 1987). A negative effect of SOD overexpression has also been observed in several mammalian systems (Amstad et al., 1991; Costa et al., 1993). When our data showed a negative side effect of SOD (Fig. 2b), we reasoned that the combination of overproduction of SOD protein in strain As1(sodA) and exogenous PQ generated high toxic concentrations of hydrogen peroxide, resulting in cell death.
To eliminate the negative effects of SOD overexpression in the presence of PQ, AhpC and SOD were simultaneously overexpressed in strain As1 (Fig. 2c). Rodriguez et al. (2000) demonstrated a similar result, where the negative effects caused by SOD overexpression in mammalian cells were neutralized by catalase overproduction. Our results indicated that AhpC likely plays a crucial role in the removal of the hydrogen peroxide generated by SOD overexpression in the presence of PQ. It is likely that superoxide anions produced as a result of PQ treatment created a significant increase in lipid hydroperoxide and/or the production of hydrogen peroxide by cellular SOD. The superoxide anion could easily cause lipid peroxidation, together with the formation of a perhydroxyl radical (Halliwell & Gutteridge, 1999). Protection from these peroxides inside the cell may be facilitated by AhpC, which is involved in the scavenging of intracellular hydrogen peroxide (Seaver & Imlay, 2001) and is also responsible for conversion of lipid hydroperoxides to the corresponding nontoxic alcohols (Halliwell & Gutteridge, 1984; Wang et al., 2006). The experiments here have not addressed the specific mechanism(s) by which AhpC overexpression enhances the survival of strain As1(sodA); however, the impact of AhpC in reducing the toxic effects of SOD was clear.
E. coli aconitase is known to be very sensitive to superoxide (Hausladen & Fridovich, 1994). Superoxide oxidizes the [4Fe–4S]2+ in this dehydratase to the unstable [4Fe–4S]3+, which then rapidly degrades to [3Fe–4S]1+, thereby inactivating the enzyme activity or damaging the metabolic pathway (Flint et al., 1993). When oxidants were removed, the activity of the dehydratase could be restored (Flint et al., 1993). Our results indicate that overexpressed Fpr or SOD in recombinant cells can maintain the aconitase activity under oxidative conditions and allow the enzyme to recover its activity (Fig. 3c). A similar result was obtained when an E. coli mutant lacking Fpr showed slow recovery of the aconitase activity compared to the wild-type (Giro et al., 2006; Li & Demple, 1996); in contrast, E. coli overexpressing Fpr showed dramatically accelerated recovery of aconitase activity to normal levels within a few minutes.
In this study, the overexpression of SOD and Fpr in Pseudomons sp. strain As1 has been shown to enhance both naphthalene degradation and cellular resistance to oxidative stress. We expect that the results of this study will contribute to the development of new strategies for understanding and manipulating the physiology of micro-organisms important for biotechnology and bioremediation.
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ACKNOWLEDGEMENTS |
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Edited by: D. Arp
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Received 6 April 2007;
revised 13 June 2007;
accepted 22 June 2007.
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, wild-type (As1); 
, As1(sodA); 
, As1(sodA-R). The results are means±SD (n=3). (b) Tolerance of cells to PQ. Six strains were grown for 16 h on LB agar in the absence (left) or presence (right) of 300 µM PQ. (c) Survival of wild-type and three recombinant strains in LB medium containing 300 µM PQ. Cells were grown to exponential phase in LB medium and exposed to 300 µM PQ. 
, As1(ahpCsodA). The results are means±SD (n=3). (d) Northern blot analyses of ahpC transcription levels in cells placed under oxidative stress by PQ.
, As1(sodA); 