Departments of 1 Internal Medicine and 2 Microbiology, Veterans Affairs Medical Center and The University of Iowa, Iowa City, Iowa, 52242
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ABSTRACT |
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Pseudomonas aeruginosa, an opportunistic human pathogen, causes both acute and chronic lung disease. P. aeruginosa exerts many of its pathophysiological effects by secreting virulence factors, including pyocyanine, a redox-active compound that increases intracellular oxidant stress. Because oxidant stress has been shown to affect cytosolic Ca2+ concentration ([Ca2+]c) in other cell types, we studied the effect of pyocyanine on [Ca2+]c in human airway epithelial cells (A549 and HBE). At lower concentrations, pyocyanine inhibits inositol 1,4,5-trisphosphate formation and [Ca2+]c increases in response to G protein-coupled receptor agonists. Conversely, at higher concentrations, pyocyanine itself increases [Ca2+]c. The pyocyanine-dependent [Ca2+]c increase appears to be oxidant dependent and to result from increased inositol trisphosphate and release of Ca2+ from intracellular stores. Ca2+ plays a central role in epithelial cell function, including regulation of ion transport, mucus secretion, and ciliary beat frequency. By disrupting Ca2+ homeostasis, pyocyanine could interfere with these critical functions and contribute to the pathophysiological effects observed in Pseudomonas-associated lung disease.
oxidants; inositol phosphates; A549 cells; HBE cells; G protein-coupled receptors
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INTRODUCTION |
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THE GRAM-NEGATIVE bacterium Pseudomonas aeruginosa causes acute lung disease with high mortality in patients with hospital-acquired pneumonias (23) and is commonly associated with the chronic lung disease observed in individuals with cystic fibrosis (9). Lung disease is currently the leading cause of morbidity and mortality in cystic fibrosis (26). During infection, P. aeruginosa secretes numerous virulence factors that contribute to its pathophysiological effects. Among these factors is the phenazine derivative pyocyanine (19), a redox-active compound that increases intracellular oxidant stress. Currently, the mechanisms by which these factors exert their effects are poorly understood.
Pyocyanine has been shown to have pathophysiological effects in numerous cell types. Most, if not all, of these effects appear to be due to increased intracellular oxidant formation and can be blocked by exogenous addition of antioxidants. In airway epithelial cells, pyocyanine inhibits ciliary beat frequency (16). This inhibition correlates with decreased cellular levels of ATP and cAMP.
Our studies were designed to identify other potentially pathophysiological effects by pyocyanine in human airway epithelial cells. Previous studies demonstrated that pyocyanine increases oxidant formation in these cells (3, 10). Because oxidant stress has been shown to affect Ca2+ homeostasis in other cell types (8, 20, 24), we examined the effect of pyocyanine on cytosolic Ca2+ concentration ([Ca2+]c) using two human airway epithelial cell lines, A549 and HBE. We found that pyocyanine increases [Ca2+]c under some conditions while inhibiting subsequent [Ca2+]c increases in response to G protein-coupled receptor agonists such as the purinergic receptor agonist ATP. Additional studies were then performed to explore the mechanisms that underlie these effects.
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MATERIALS AND METHODS |
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Materials. Buthionine sulfoximine (BSO), glutathione reductase, NADPH, human placental collagen, N-acetylcysteine (NAC), ATP, 5,5'-dithio-bis(2-nitrobenzoic acid) (DTNB), and carbamylcholine chloride (carbachol) were purchased from Sigma (St. Louis, MO). Pyocyanine was generously provided by Dr. Cox.
Cell culture. The human alveolar type II cell line A549 (American Type Culture Collection CL-185) (18) was cultured in MEM with Earle's salts (GIBCO BRL, Gaithersburg, MD) supplemented with 10% fetal calf serum, 2 mM glutamine, and 500 U/ml each of penicillin and streptomycin. Passages from 85 to 120 were used in these studies. Stock but not experimental cultures of the human bronchial epithelial cell line HBE (14) were cultured in collagen-coated tissue cultureware ( passages 8-25) in the same medium. Experiments were performed on cultures that were 80-90% confluent.
Spin trapping and electron paramagnetic resonance. Electron paramagnetic resonance (EPR) experiments were performed on airway epithelial cells cultured in 24-well tissue culture plates. Reaction mixture (0.5 ml/well) consisting of Hanks' balanced salt solution containing 100 mM 5,5-dimethyl-1-pyrroline N-oxide (DMPO; OMRF Spin Trap Source, Oklahoma City, OK) and 100 µM diethylenetriamine pentaacetic acid (DTPA; Aldrich Chemical, Milwaukee, WI) with or without the indicated additions was placed on the cell monolayer. Samples were incubated for 15 min at room temperature, and then the reaction mixture was transferred to a quartz flat cell. All spectra are the result of seven signal-averaged scans and were obtained at room temperature using a Bruker model ESP300 EPR spectrometer (Karlsruhe, Germany). Instrument settings were as follows: microwave power, 20 mW; modulation frequency, 100 kHz; receiver gain, 4 × 105; modulation amplitude, 0.501 G; time constant, 327.68 ms; sweep rate, 335.5 s.
Measurement of
[Ca2+]c.
Ca2+ measurements were performed
at the Cell Fluorescence Core Facility (Veterans Affairs Medical
Center, Iowa City, IA). Cells were cultured on collagen-coated 25-mm
round glass coverslips. Cells were loaded in complete medium with fura
2 by direct addition of the cell-permeant form, fura 2-AM (Molecular
Probes, Eugene, OR), to the culture dish containing the coverslip and
incubation for 30 min at 37°C. Cells were washed with
HEPES-buffered saline [in mM: 135 NaCl, 5 KOH, 10 HEPES, 1.2 CaCl2, 1.2 MgCl2, and 10 glucose
(HBS-G)], and measurements of the apparent
[Ca2+]c
were done in HBS-G using the Photoscan II spectrofluorometer (Photon
Technologies International, New Brunswick, NJ) with a Nikon microscope
(Nikon, Niles, IL). Final Ca2+
concentration ([Ca2+])
values were determined using a PTI software package from the ratio of
emission intensities [emission wavelength
(em) 510 nm] for
excitation wavelengths (
ex)
of 340 and 380 nm. Briefly, background fluorescence
intensities for each
ex were
obtained using unloaded cells and were subtracted from the raw data.
The ratios of the corrected fluorescence intensities were then
converted to [Ca2+]
values using the formula
[Ca2+] = Kd · (R
Rmin)/(Rmax
R), where the maximum and minimum ratios
(Rmax and
Rmin, respectively) as well as the
apparent dissociation constant
(Kd) were
empirically derived from
[Ca2+] curves
generated using the instrument. In earlier work (6), we found that this
method gives similar values for basal
[Ca2+] as well as for
Ca2+ increases in response to
agonists as does the more laborious method that involves determining
Rmax and
Rmin using ionomycin followed by
EGTA (7).
Fluorescence measurements of intracellular oxidant formation. To measure intracellular oxidant formation, we used an oxidant-sensitive dichlorodihydrofluorescein derivative (Iowa probe) generously provided by Dr. Stephen Hempel (Dept. of Internal Medicine, Veterans Affairs Medical Center and the Univ. of Iowa, Iowa City, IA). In cell-free studies (15), the Iowa probe behaved identically to commercially available dichlorodihydrofluorescein compounds (Molecular Probes). However, in cell culture studies, the Iowa probe was significantly more sensitive in detecting oxidant formation in response to pyocyanine as well as to other redox-active compounds (S. Hempel, unpublished observations). Although the exact mechanism by which these compounds detect oxidant formation is not fully understood, reaction of these probes with H2O2 appears to require peroxidase activity or iron (25).
For these studies, cells in 12-well tissue culture plates were washed twice with warm HBS-G and preincubated at 37°C for 30 min in HBS-G containing 5 µM Iowa probe. At the end of the preincubation period, the indicated concentration of pyocyanine was added, and the cells were incubated for 1 h at 37°C. To measure cell-associated fluorescence, the cells were washed twice with ice-cold PBS and incubated on ice with PBS containing 0.2% Triton X-100. The cell extract was removed from the cells, and the relative fluorescence intensity of the extract (Altering cellular antioxidant capacity. To reduce glutathione levels, cells were incubated with 100 µM BSO in complete medium for 48 h. To measure glutathione, cells were washed twice with ice-cold PBS, scraped into 0.01 N HCl, frozen overnight, and thawed. Samples were centrifuged at 12,000 rpm for 5 min to remove cellular debris, and total glutathione in the cell extract was determined by assaying the rate of DTNB reduction in the presence of NADPH and glutathione reductase (11). Oxidized glutathione was measured in cell extracts treated for 1 h with 2-vinylpyridine (Aldrich Chemical). Reduced glutathione was determined by subtracting oxidized glutathione from total. Values were generated by comparison with a reduced glutathione standard curve and were normalized to total cell protein, measured using the micro bicinchoninic acid assay (Pierce, Rockford, IL). For studies with NAC, cells were pretreated with the indicated concentration of NAC for 2-4 h before experiments were performed. NAC at these concentrations markedly acidifies the medium. Thus all NAC-containing solutions were adjusted to pH 7.3-7.5 before use.
Turnover of inositol phosphates. Cells were seeded into six-well tissue culture plates, allowed to attach overnight, and then cultured for 48 h in complete medium containing 1 µCi/ml of myo-[3H]inositol (Amersham, Arlington Heights, IL). At the end of the labeling period, turnover of inositol phosphates (IPs) was measured as previously described (7). Briefly, cells were washed with HBS-G and incubated in HBS-G for 20 min at 37°C. Cultures were then incubated for 20 min with HBS-G containing 10 mM LiCl. Finally, cells were stimulated with the indicated agonist for 20 min or with pyocyanine for 10 min and then agonist for 10 min. IPs were extracted overnight at 4°C with 0.5 M perchloric acid. The acid extract was neutralized with 2.5 M KOH and 0.5 M HEPES (pH 7.4) and centrifuged to remove the precipitate. The IP species were then collected using anion-exchange column chromatography (Dowex AG, 1-8X, 100-200 mesh, formate form; Bio-Rad, Hercules, CA) as previously described (22).
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RESULTS AND DISCUSSION |
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Pyocyanine increases oxidant formation in A549 and
HBE cells.
Previous work in our laboratories (3) and by Gardner (10) using two
different assay techniques demonstrates that pyocyanine stimulates
oxidant formation in A549 cells. To verify these results in our A549
cultures and to determine whether pyocyanine increases superoxide anion
(· O2) formation in HBE
cells, we used the spin-trapping agent DMPO and EPR spectroscopy.
Figure 1 shows pyocyanine-dependent
formation of the DMPO-· OH adduct (aN = aH = 14.9 G, where
aN and aH are the
splitting constants for nitrogen and hydrogen, respectively) in HBE
(Fig. 1A) and A549 (Fig.
1B) cells. Formation of
this adduct could result from a reaction between DMPO and either
· O
2 or the hydroxyl radical
(· OH). Addition of superoxide dismutase markedly attenuates
or abolishes the signal (Fig. 1, C and
D), demonstrating that
· O
2 is required for adduct formation. In addition, catalase has little or no effect (Fig. 1,
E and
F) on the signal. This result argues
against · OH formation, since its formation requires
H2O2
and hence is inhibited by catalase. Consistent with this conclusion is
our observation that the · OH scavenger DMSO, which reacts
with · OH to form the methyl radical (· CH3) and
subsequently DMPO-· CH3,
did not alter the response to pyocyanine (data not shown). Together,
these results indicate that the spectra shown in Fig. 1 reflect
pyocyanine-induced · O
2
formation.
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Pyocyanine increases [Ca2+]c. Previous studies (8, 24) show that oxidants increase [Ca2+]c in a variety of cell types. In early studies, we found that pyocyanine causes an acute, transient increase in [Ca2+]c. A representative example of this response in A549 cells is shown in Fig. 2A. In these studies, the apparent basal [Ca2+]c values were ~100-150 nM for both A549 and HBE cells and were similar from experiment to experiment. In contrast, the magnitude of the maximal [Ca2+]c increase in response to pyocyanine, although similar within an experiment, varied considerably from experiment to experiment, ranging from 50 to over 300 nM. Moreover, concentration-dependence studies indicate that a threshold concentration of pyocyanine is required to elicit the response. This threshold concentration also varied considerably from experiment to experiment (ranging from 80 to 250 µM). Responses to other Ca2+ agonists in these same studies did not exhibit the same degree of variability, suggesting that this is a specific characteristic of the pyocyanine response. The basis for this variability remains unclear but did not appear to depend on the preparation of pyocyanine used or on the confluency state of the cells. We speculate that the variability may reflect differences in uptake of pyocyanine, via a mechanism currently unknown, or differences in the level of reducing sources (NADPH, NADH). Further experiments will be necessary to test these and other possibilities.
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Pyocyanine stimulates IP turnover. Release of Ca2+ from intracellular Ca2+ stores can result from formation of the second messenger inositol 1,4,5-trisphosphate (IP3). To determine whether pyocyanine increases [Ca2+]c by increasing IP3, we used an assay that measures turnover of IPs. This assay has been shown to be the most sensitive measure of increased IP3. Moreover, in previous work (17), we found that results using this method accurately reflected increased IP3 formation.
Using this approach, we found that pyocyanine increases IP turnover in a concentration-dependent manner in both A549 and HBE cells. In three independent experiments, the maximum increase observed in response to pyocyanine ranged from 116 ± 6 to 169 ± 11% of control value (means ± SD for triplicate samples). As a positive control in these studies, we used the purinergic receptor agonist ATP. We observed that 1 mM ATP stimulated IP turnover in the range of 141 ± 10 to 206 ± 18% of control value in these experiments. Moreover, as with our [Ca2+]c measurements, we observed that the minimum pyocyanine concentration required to stimulate IP turnover varied markedly. As stated above, the basis for this variability remains unknown.Cellular antioxidant capacity alters the response
to pyocyanine.
To determine whether oxidants contribute to the pyocyanine-dependent
increase in
[Ca2+]c,
we decreased the antioxidant capacity of the cells by decreasing intracellular levels of the thiol antioxidant glutathione. To do this,
cells were treated for 48 h with 100 µM BSO, an inhibitor of
-glutamylcysteine synthetase (21). Total glutathione
levels in control cells were 79 ± 7 and 42 ± 6 nmol/mg protein
(means ± SE; combined data from 6 independent experiments with
triplicate samples for each cell type) for A549 and HBE cells,
respectively. In each case, the majority (84-90%) of the
glutathione was in the reduced form. A 48-h treatment with BSO
decreased total glutathione levels by 80-98% (A549, 2.2 ± 0.44 ng/mg protein; HBE, 9.5 ± 3.0 ng/mg protein; means ± SE; combined data from two independent experiments with
triplicate samples for each cell type).
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Pyocyanine inhibits the response to Ca2+ agonists. Because pyocyanine itself increases [Ca2+]c, we wondered whether pyocyanine would affect the subsequent response to Ca2+ agonists. The purinergic receptor agonist ATP was used for these experiments, since it increases [Ca2+]c in both A549 (Fig. 5A) and HBE (Fig. 5B) cells.
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Pyocyanine inhibits IP turnover in response to ATP. To determine whether pyocyanine inhibits the [Ca2+]c increase in response to ATP by inhibiting agonist-dependent IP3 formation, we measured the effect of pyocyanine on ATP-stimulated turnover of IPs. In these experiments, cells were labeled with myo-[3H]inositol for 48 h, washed, and incubated for 20 min in buffer with 10 mM LiCl. The cells were then incubated for 10 min with or without pyocyanine and finally for 10 min with or without 1 mM ATP. We found that pyocyanine inhibits IP turnover in response to ATP. Moreover, inhibition of the response to ATP can occur at pyocyanine concentrations that, within a given experiment, do not by themselves stimulate an increase in IP turnover. As an example of these results, we obtained values of 100 ± 11, 87 ± 2, 160 ± 27, and 86 ± 4% (means ± SD for triplicate samples from a representative experiment) for control, pyocyanine alone (200 µM), ATP alone, and pyocyanine followed by ATP, respectively. Similar results were seen in two other independent experiments. In studies in which we observed a pyocyanine-dependent increase in IP turnover, no additional increase was observed with subsequent exposure to ATP. These data further suggest that the effect of pyocyanine on IP3 formation in response to ATP is independent of its effect on IP3 formation itself.
Activation of protein kinase C (PKC) has been shown to inhibit signaling by G protein-coupled receptors linked to PIP2-PLC (4). In addition, oxidant stress has been shown to activate PKC (27). Thus, to determine whether pyocyanine inhibits IP3 formation in response to ATP by activating PKC, we tested the effect of the PKC inhibitors staurosporine and bisindolylmaleimide. Results from these experiments illustrate several points (Fig. 6). First, ATP stimulates IP turnover, and inhibitor treatment has little or no effect on this response. Second, activation of PKC by the phorbol ester phorbol 12-myristate 13-acetate (PMA) inhibits the ATP response, and this inhibition is prevented by pretreating the cell with either inhibitor. Finally, pyocyanine inhibits IP turnover in response to ATP, but, in contrast to PMA, PKC inhibitors do not block this effect. These data suggest that pyocyanine does not exert its inhibitory effect by activating PMA-sensitive isoforms of PKC. We cannot rule out the possibility, however, that the ATP response is inhibited by pyocyanine through activation of PMA-insensitive isoforms of PKC that are not inhibited by these concentrations of PKC inhibitors.
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Conclusions. These studies are the first to examine the effect of Pseudomonas pyocyanine on [Ca2+]c in human airway epithelial cells. We found that pyocyanine has two such effects. Pyocyanine alone at higher concentrations increases [Ca2+]c, presumably by increasing IP3, which stimulates Ca2+ release from intracellular Ca2+ stores and a subsequent influx of extracellular Ca2+. Conversely, pyocyanine at lower concentrations inhibits IP3 formation and the subsequent increase in [Ca2+]c in response to Ca2+ agonists. Because of differences in the concentration dependence of these two effects and because they reflect opposite effects on IP metabolism, it follows that the two effects are independent and therefore must involve different molecular mechanisms. However, both effects appear to share the common feature of being mediated, at least in part, by pyocyanine-generated oxidants.
These effects occur at concentrations ranging from 80 to 250 µM. Pyocyanine has been detected at concentrations as high as 75-100 µM in sputum (28) and bronchoalveolar lavage fluid from patients with Pseudomonas infections. With consideration that dilution occurs as a result of lavage, it is reasonable to assume that higher concentrations of pyocyanine are present at the site of infection. Thus the concentrations used in our studies are likely to be physiologically relevant. The molecular mechanisms by which pyocyanine increases IP3 and [Ca2+]c remain to be identified. Previous studies suggest several possibilities. One of these involves oxidant-dependent activation of protein tyrosine kinases (PTKs) with subsequent tyrosine phosphorylation and activation of PLC- ![]() |
ACKNOWLEDGEMENTS |
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This work was supported in part by Veterans Affairs Merit Review Grants (awarded to G. M. Denning and B. E. Britigan by the Office of Research and Development, Medical Research Service, Department of Veterans Affairs) as well as by National Institutes of Health Grant AI-34954 and the Cystic Fibrosis Foundation.
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FOOTNOTES |
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This work was performed during the tenure of B. E. Britigan as an Established Investigator of the American Heart Association.
Address for reprint requests: G. M. Denning, Bldg. 3, Rm. 139, Veterans Affairs Medical Center, Iowa City, IA 52246.
Received 2 June 1997; accepted in final form 2 February 1998.
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