1 Research Service and 4 Department of Internal Medicine, Iowa City Veterans Affairs Medical Center, Iowa City 52246; and 2 Department of Internal Medicine and 3 Free Radical and Radiation Biology Program, Department of Radiation Oncology, University of Iowa Roy J. and Lucille A. Carver College of Medicine, Iowa City, Iowa 52242
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ABSTRACT |
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The Pseudomonas aeruginosa secretory product pyocyanin damages lung epithelium, likely due to redox cycling of pyocyanin and resultant superoxide and H2O2 generation. Subcellular site(s) of pyocyanin redox cycling and toxicity have not been well studied. Therefore, pyocyanin's effects on subcellular parameters in the A549 human type II alveolar epithelial cell line were examined. Confocal and electron microscopy studies suggested mitochondrial redox cycling of pyocyanin and extracellular H2O2 release, respectively. Pyocyanin decreased mitochondrial and cytoplasmic aconitase activity, ATP levels, cellular reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide, and mitochondrial membrane potential. These effects were transient at low pyocyanin concentrations and were linked to apparent cell-mediated metabolism of pyocyanin. Overexpression of MnSOD, but not CuZnSOD or catalase, protected cellular aconitase, but not ATP, from pyocyanin-mediated depletion. This suggests that loss of aconitase activity is not responsible for ATP depletion. How pyocyanin leads to ATP depletion, the mechanism of cellular metabolism of pyocyanin, and the impact of mitochondrial pyocyanin redox cycling on other cellular events are important areas for future study.
mitochondria; superoxide; hydrogen peroxide; aconitase
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INTRODUCTION |
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PSEUDOMONAS AERUGINOSA CAUSES acute severe necrotizing pneumonia with high mortality (13, 34) as well as chronic lung infections in patients with cystic fibrosis (CF) or chronic bronchiectasis (12, 13, 34, 44). Chronic lung injury is currently the primary cause of death in CF and has been linked to coexistent P. aeruginosa infection (12, 13). The pathogenic mechanism(s) involved in P. aeruginosa-mediated tissue damage in the lung remain uncertain (12, 13, 34, 44).
Most P. aeruginosa strains secrete pyocyanin
(N-methyl-1-hydroxyphenazine, mw 210) (54).
Pyocyanin is among the P. aeruginosa cytotoxic secretory
factors that are felt to contribute to organism virulence (12,
13, 33, 34, 44, 51, 52). Pyocyanin's cytotoxicity has been
linked to its propensity to undergo cell-mediated redox cycling with
resultant formation of superoxide (O
Pyocyanin can be detected in pulmonary secretions of CF patients and other individuals with chronic bronchiectasis who are infected with P. aeruginosa at concentrations of up to 100 µM (56). These concentrations of pyocyanin cause a variety of deleterious effects on pulmonary cells and tissues in vitro and in vivo (1, 10, 27, 28, 39, 47, 55, 56). Protection by pharmacological agents suggests that at least some of these deleterious effects result from pyocyanin-mediated depletion of cellular cAMP and ATP (11, 28) that occurs via oxidant production (27, 47).
Perhaps linked to cellular depletion of ATP, experiments using either
respiratory tissue or the A549 type II alveolar cell line suggest that
pyocyanin may damage mitochondria (11, 14, 17). In the
case of the cell line, data suggested that pyocyanin-mediated inhibition of cellular aconitase might be important (14,
17). Inhibition of aconitase is not surprising given its known
susceptibility to inactivation by O
The site(s) of O
Therefore, we conducted a series of experiments whose goal was to further define the site of pyocyanin redox cycling within airway epithelial cells with a particular emphasis on the impact of such events on cellular mitochondrial function and energy (ATP) production.
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MATERIALS AND METHODS |
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Cell culture. The human alveolar type II cell line A549 [CL-185; American Type Culture Collection (ATCC), Rockville, MD] was cultured in Dulbecco's modified Eagle's medium (Cellgro) containing 5% heat-inactivated fetal bovine serum (FBS), 2 mM glutamine, and 500 U/ml each of penicillin and streptomycin. Passages from 73 to 100 were used. Experiments were performed when the cell cultures were 95% confluent.
Pyocyanin purification. Pyocyanin was extracted from the broth culture of P. aeruginosa as previously described (8). Briefly, P. aeruginosa strain PAO1 (15692; ATCC) was grown in glycerol-alanine minimal medium. The bacteria were removed by centrifugation. Pyocyanin in the culture supernatant was extracted into chloroform. The blue pigment in chloroform phase was back extracted into acidified water (10 mM HCl). The water was then adjusted to neutrality (pH 7) by the addition of NaOH. This process was repeated at least five times. The pyocyanin (in acidified form) was stored frozen or at 4°C protected from light. The purity of the pyocyanin solution was confirmed by HPLC using a Beckman System Gold apparatus and a reverse-phase C18 column (Microsorb-MV-C18, 250 × 4.6 mm; Varian, Walnut Creek, CA). The solvent system consisted of 0.05% trifluoracidic acid (TFA) in water and 0.05% TFA in acetonitrile using 25-30-min runs. This pyocyanin purification procedure has been previously shown to result in LPS concentrations in our experimental systems of <0.1 U/ml (9).
Confocal microscopy. A549 cells were cultured to confluency in six-well plates. Cells were incubated with 20 µM 5-(and-6)-chloromethyl-2',7'-dichlorodihydrofluorescein diacetate (CM-H2DCFDA) probe and/or 500 nM MitoTracker red probe (Molecular Probes) for 30 min at 37°C. Pyocyanin was then added at 50 and 100 µM and incubated for 2 h at 37°C. Cells were then studied using confocal microscopy. Controls included cells incubated with pyocyanin only, probe only, and buffer only. The cells were then examined using a Bio-Rad MRC-1024, Kr/Ar laser attached to a Nikon E600 confocal microscope.
Electron microscopy. A549 cells were cultured to confluency on glass coverslips in six-well plates. Cells were treated with 50 and 100 µM pyocyanin for 2 h in Hanks' balanced salt solution (HBSS). A fresh standard buffer was made of 0.1 M Tris-maleate buffer, pH 7.4, with 7% sucrose (TMB/S). Cells were incubated with TMB/S with 1 mM cerium chloride (CeCl3; Sigma), 10 mM aminotriazole, and 0.71 mM NADH for 1 h at 37°C. All solutions were sterilized by filtration through a 0.45-µm Millipore membrane. After this incubation, the cells were fixed in 2% glutaraldehyde in 0.1 mM cacodylate buffer, pH 7.4, at 4°C for 60 min. After being washed with TMB/S, cells were postfixed in 1% OsO4 at 4°C for 60 min, dehydrated in graded ethanol, and embedded in Spurr's resin. Ultrathin sections were prepared, stained with uranyl acetate and lead citrate, and examined by electron microscopy under a Hitachi H-600 electron microscope. Controls included cells treated with pyocyanin alone, CeCl3 alone, and buffer alone.
Aconitase and succinate dehydrogenase activity. For determinations of total aconitase and SDH activity, the cells were collected and sonicated with Tris buffer (50 mM, pH 7.4) containing protease inhibitor cocktail tablet (Boehringer Mannheim, Mannheim, Germany), and the cell lysate was separated from cellular debris by centrifugation (14,000 g, 1 min). For differentiation of cytosolic from mitochondrial aconitase activity, the cells were suspended in Tris (50 mM, pH 7.4) buffer containing 70 mM sucrose, 210 mM mannitol, and protease inhibitor cocktail tablet (Boehringer) and placed on ice. The cell membrane was disrupted by nitrogen cavitation (200 psi for 5 min). The supernatant (cytosolic aconitase fraction) and pellet (mitochondrial aconitase fraction) were separated by centrifugation (10,000 g, 10 min). The separation of cytosol and mitochondria was confirmed by immunoblotting the fractions for the presence of MnSOD (mitochondrial marker) and CuZnSOD (cytosolic marker).
The protein content of each sample was determined using the BCA protein assay (Pierce, Rockford, IL). For aconitase determinations, 50 µg of protein from the supernatant were mixed with reaction buffer [50 mM Tris (pH 7.4), 1 U/ml isocitrate dehydrogenase, 0.6 mM MnCl2, 20 µM fluorocitrate, 400 µM cis-aconitate, 1 µM phenazine methosulfate (PMS), 200 µM cytochrome c, and 200 µM NADP], and the mixture was incubated at 25°C for 30 min. The optical density (OD) was measured at 550 nm. In some assays for mitochondrial aconitase activity, 1 mM NaCN was added to the reaction mixture to prevent reoxidation of cytochrome c by mitochondrial cytochrome oxidase. For SDH determinations, 50 µg of protein from the supernatant were mixed with a different reaction buffer [50 mM Tris (pH 7.4), 20 mM sodium succinate, 1 mM KCN, 1 µM PMS, and 5 mM cytochrome c], and the OD was measured at 550 nm after 5 min at 25°C.Immunoblotting. Cells were rinsed twice with PBS and lysed by addition of Tris buffer (50 mM, pH 7.4) containing 1% Nonidet P-40 (Amresco, Solon, OH). Cell lysates were collected into Eppendorf tubes and sonicated, and cellular debris was removed by centrifugation (14,000 g, 1 min). Samples (20-30 µg protein) were mixed 1:1 with sample buffer (1.25 M Tris, pH 6.8, 20% glycerol, 4% SDS, 10% 2-mercaptoethanol, and 0.05% bromphenol blue), and proteins were separated by 10% SDS-PAGE. The protein was transferred to a nitrocellulose membrane overnight at 30 V. The membrane was blocked with 5% skim milk in Tris-buffered saline with 0.1% Tween (TBST) for 1 h and incubated with the primary antibody (1:1,000 dilution) for 1-2 h. The blot was washed with TBST and incubated with a 1:10,000 dilution of the secondary antibody (horseradish peroxidase-conjugated anti-IgG; Amersham Pharmacia Biotech, Piscataway, NJ). The immunoreactive protein was detected with an enhanced chemiluminescence detection kit (Amersham Pharmacia Biotech).
ATP determination. The procedure was adapted from that of Takahashi et al. (50) with some modification. We obtained the cell extract by adding 100 µl of ice-cold perchloric acid (6%) to cells. The perchloric acid solution was collected in a test tube and neutralized to pH 7 with 22.5 µl of KOH (4 M) and 10 µl of Tris buffer (2 M). ATP was measured by the luciferin-luciferase method with a luminometer (Analytical Luminescence Laboratories, Cockeysville, MD) according to the manufacturer's instructions. Results were normalized to cellular protein content. Results were similar when cellular DNA was used as the denominator.
DNA quantification. The cells were lysed with Tris buffer (50 mM, pH 7) containing 10 mM NaCl and 0.1% SDS. The cell solution (1 ml) was mixed with 10 µl of 100 µg/ml 4',6-diamidino-2-phenylindole and incubated at RT for 10 min. The fluorescent intensity was measured at an excitation wavelength of 360 nm and emission wavelength of 460 nm (30).
3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide reduction. 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) was dissolved in MEM without phenol red. A549 cells were incubated with MTT (0.5 mg/ml) at 37°C for 2 h. They were then lysed with 100% propanol. We then determined the presence of blue formazan resulting from cell-mediated reduction of MTT by measuring absorbance at 550 nm.
SOD and catalase overexpression.
Recombinant adenoviral vectors expressing MnSOD (Ad CMV MnSOD), CuZnSOD
(Ad CMV CuZnSOD), catalase (Ad CMV catalase), or -galactosidase (Ad
CMV LacZ) were constructed by and purchased from the Vector Core
Facility of The University of Iowa. Each adenoviral stock (4-6 × 1010 DNA particles/ml) was stored in 3%
sucrose at
80°C. Multiplicities of infection (MOI) ranging from 10 to 100 were routinely employed. Cells were exposed to adenovirus in
media containing 5% FBS at 37°C for 24h. We confirmed successful
transfection by measuring expression of each antioxidant enzyme at both
the protein and activity level by immunoblot and by a native gel
activity assay staining, respectively (3, 4, 35).
Pyocyanin metabolism. Increasing concentrations of pyocyanin were added to A549 monolayers or cell-free media, and samples were incubated for 4, 8, 12, and 24 h. Cells and medium were harvested from the wells, pyocyanin was extracted with chloroform, and the amount of pyocyanin recovered at each time point was then quantified by HPLC as described in Pyocyanin purification.
Mitochondrial potential. Mitochondrial membrane potential was measured fluorometrically by previously described methods (7, 53). Briefly, the cells were grown in 96-well plates and incubated with or without pyocyanin for 24 h. The medium was removed, and the wells were rinsed twice with HBSS. The fluorescent probe JC-1 dissolved in HBSS (2 µg/ml; Molecular Probes, Eugene, OR) was incubated with the cells for 4 h at 37°C. Fluorescence intensity was measured at excitation and emission wavelengths of 544/590 nm (red) and 485/530 nm (green). Mitochondrial membrane potential is proportional to the red/green fluorescence intensity ratio.
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RESULTS |
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Subcellular localization of pyocyanin redox cycling.
Previous work indicates that pyocyanin redox cycles on exposure to
either eukaryotic or prokaryotic cells, resulting in the production of
O
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Pyocyanin depletes cellular ATP and alters mitochondrial electron
transport.
Because mitochondrial activity is critical to optimal production of
cellular ATP, we sought to confirm previous reports (11, 28) that pyocyanin exposure depletes cells of ATP. Consistent with these previous reports, we found a time- and
concentration-dependent depletion of the ATP content of cells exposed
to pyocyanin (Fig. 4).
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Pyocyanin inhibits mitochondrial and cytosolic aconitase activity. Given that mitochondrial electron transport is critical to cellular ATP production and that pyocyanin redox cycling appeared to occur at/near mitochondria, and that pyocyanin decreased cellular MTT reduction, we sought to explore further whether mitochondria were targets of pyocyanin's effects and, if so, what mitochondrial components were affected.
Previous work by Gardner and colleagues (14, 17) demonstrates that aconitase activity is inhibited in A549 cells by pyocyanin. This is not surprising given that the Fe-S center of aconitase is extremely sensitive to the effects of O
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Pyocyanin depolarizes mitochondrial membranes.
To further assess the effect of pyocyanin on mitochondria, we
measured mitochondrial membrane potential. Alteration in mitochondrial membrane potential has been shown to be a sensitive indicator of
oxidant damage (53). Using a fluorescent probe-based
assay, we observed that exposure of cells to pyocyanin resulted in a decrease in mitochondrial membrane potential (Fig.
8).
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Pyocyanin is metabolized by A549 cells.
A somewhat surprising observation made during the course of the above
experiments was that when the cells were exposed to lower
concentrations of pyocyanin, aconitase activity was only transiently
decreased, rebounding to normal levels even though the pyocyanin was
not washed off the cells following the initial addition (Fig.
9A). This was not due to
increases in cellular SOD content (data not shown). These data suggest
the possibility that pyocyanin might be undergoing metabolism to a less
bioactive compound. Accordingly, pyocyanin was added to A549 monolayers and tissue culture wells lacking cells. At increasing times, the well
contents were removed and analyzed for the presence of pyocyanin. In
the presence of A549 cells, the pyocyanin concentration decreased, with
a half-life of ~12 h (Fig. 9B). This appeared to be a
saturable process, as at concentrations >20 µM pyocyanin, the effect
was less pronounced (Fig. 9C).
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Pyocyanin-induced mitochondrial superoxide activity is linked to
aconitase inhibition.
Although previous work suggests that pyocyanin-induced
O
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DISCUSSION |
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Pyocyanin is one of several virulence factors secreted by P. aeruginosa (13, 34, 44, 51). Although the ability of pyocyanin to redox cycle and generate ROS has been linked to its cytotoxicity for both eukaryotic and prokaryotic cells, the subcellular targets impacted by pyocyanin remain ill defined. Previous studies suggested mitochondria as one of the targets through which pyocyanin disrupts lung epithelial cell functions and contributes to pathogenesis of lung injury resulting from both acute and chronic P. aeruginosa lung infection (14, 17).
Results from confocal microscopy provide the most direct evidence to
date that extracellularly administered pyocyanin reaches mitochondria,
where it may enhance ROS generation. It is important to note that
oxidation of DCFH2 can potentially occur through mechanisms
other than ROS production (26, 40, 45). In fact, we have
preliminary evidence that suggests that pyocyanin can directly oxidize
DCFH2 independently of ROS formation. Thus we cannot
definitively attribute the detection of mitochondrial-associated DCF
fluorescence to mitochondrial production of O
Previous electron microscopy work in which pyocyanin, at a
concentration slightly higher than that employed in our studies, was
applied to the mucosal surface of human nasal turbinate cultures for
3 h revealed extensive damage to airway cell mitochondria (11). Marked surface blebbing and other changes were also
noted (11). In contrast, at 2 h we saw only modest
alterations in mitochondrial matrix structure in A549 cells exposed to
the highest concentration of pyocyanin that we employed (100 µM). The
difference in results from the two studies likely reflects the
differences in the model systems, organ culture vs. cell line,
antioxidant defenses of the cells involved, as well as perhaps the
slightly lower pyocyanin concentrations that we employed. In our
experience, A549 cells tend to be more resistant to oxidative injury
than other cell types. Nevertheless, our observation that pyocyanin decreases mitochondrial membrane potential supports an important effect
of the compound on mitochondria. P. aeruginosa exotoxin A
has also been shown to alter the mitochondrial morphology of the 16HBE
human bronchial epithelial cell line and induce mitochondrial membrane
depolarization via a mechanism that involves O
Whether pyocyanin diffuses passively across the cell membrane to reach
mitochondria or is actively and selectively directed to this
subcellular site remains to be fully defined. Regardless of its
mechanism for cellular entry, our studies confirm previous observations
that exposure to pyocyanin results in a time- and concentration-dependent depletion of cellular ATP and decreased activity of cellular aconitase, an enzyme known to be highly sensitive to inactivation by O
We found that overexpression of MnSOD (mitochondrial) but not CuZnSOD
(cytosolic) protected cellular aconitase activity of the cell. This
would suggest that mitochondrial generation of O
Mitochondrial electron transport via the Krebs cycle plays a critical role in cellular ATP production. Because pyocyanin depletes cellular ATP and inhibits a key component of the Krebs cycle (aconitase), it would be logical to link the two events. However, our transfection studies with MnSOD reveal that, despite our ability to protect both mitochondrial and cytosolic aconitase activity from pyocyanin-mediated inhibition, cellular loss of ATP as a consequence of pyocyanin exposure still occurred. However, these data are somewhat difficult to interpret due to the fact that, for unknown reasons, overexpression of MnSOD, as well as CuZnSOD and catalase, by themselves produced a decrease in the ATP levels of the cells.
Nevertheless, these results suggest that other cellular components, besides aconitase, are negatively affected by pyocyanin and are responsible for the loss of cellular ATP resulting from cellular exposure to this agent. Whether this involves pyocyanin-mediated effects on cellular levels of NADH or NADPH, inhibition of glycolysis, or other mitochondrial processes requires further investigation.
An additional and previously unappreciated aspect of the interaction of pyocyanin with airway epithelial cells is that these cells have the capacity to catabolize pyocyanin. The products of this metabolism are currently under study. Cellular recovery of aconitase activity correlates with the extent to which pyocyanin is metabolized, suggesting that the product(s) of pyocyanin metabolism is less cytotoxic than the parent compound.
We have shown that, when administered to airway epithelial cells in vitro, pyocyanin can move to subcellular sites in which mitochondria reside, where it is able to participate in redox chemistry. Pyocyanin exposure leads to a decrease in cellular ATP and inhibition of both mitochondrial and cytosolic aconitase activity, the extent and persistence of which are influenced by cellular metabolism of pyocyanin. Although augmenting MnSOD levels protects cellular aconitase, this does not prevent pyocyanin-mediated depletion of cellular ATP, suggesting the events are not linked. The way in which pyocyanin exposure leads to depletion of ATP, the mechanism of cellular metabolism of pyocyanin, and the impact of mitochondrial pyocyanin redox cycling on other cellular events are important areas for future study.
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ACKNOWLEDGEMENTS |
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We thank Dr. Shankar Iyer for assistance with the HPLC determinations.
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FOOTNOTES |
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This work was supported in part by grants from the Research Service of the Department of Veterans Affairs (to B. E. Britigan, M. L. McCormick, and G. M. Denning), the Public Health Service (RO1 AI-43954, P30 DK5-4759), and the Heartland Affiliate of the American Heart Association (to G. M. Denning, K. J. Reszka).
Address for reprint requests and other correspondence: B. E. Britigan; Univ. of Iowa Hospitals and Clinics; Dept. of Internal Medicine, SW54, GH; Iowa City, IA 52242 (E-mail: bradley-britigan{at}uiowa.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
First published November 1, 2002;10.1152/ajplung.00316.2002
Received 17 September 2002; accepted in final form 30 October 2002.
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