Departments of 1Pediatrics and 2Molecular Pharmacology, Northwestern University Medical School, Chicago, Illinois 60611-3008
Submitted 16 September 2002 ; accepted in final form 6 December 2002
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ABSTRACT |
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reactive oxygen species
Increasing evidence suggests that reactive oxygen species (ROS), such as superoxide anions and hydrogen peroxide (H2O2), can stimulate vascular SMC growth (33, 41). These ROS appear to be produced by SMC in response to treatment with growth factors known to cause SMC proliferation (18, 41). Recently, we have shown that ET-1 stimulated ovine fetal pulmonary arterial SMC (FPASMC) proliferation via an induction of ROS (45). In addition, elevation of ROS levels was also mitogenic for these cells (45). In contrast, other experiments have shown that antioxidant treatment (44), or overexpression of catalase (10), reduced viability and induced apoptosis in vascular SMC. Similarly, we have shown that antioxidant treatment or inhibition of the superoxide-producing enzyme NADPH oxidase decreased viability and induced apoptosis in FPASMC (45). Overall, these data suggest that increased levels of ROS stimulate SMC proliferation, whereas decreased ROS levels can prevent proliferation and induce apoptosis. In PPHN, elevated levels of ET-1 may trigger vascular remodeling via ROS-mediated SMC growth. Antioxidant therapy may, therefore, prove to be an effective therapy for diseases arising from excessive vascular muscularization.
Salen-manganese complexes are low-molecular-weight synthetic compounds that possess superoxide dismutase (SOD) and catalase activities, catalytically removing superoxide and H2O2, respectively (4, 17). These compounds are thought to exhibit better stability and bioavailability than proteinaceous antioxidant enzymes. Furthermore, their catalytic mode of action may prove more effective than low-molecular-weight antioxidant compounds. EUK-134 is one such SOD/catalase mimetic (3). This compound has been demonstrated to reduce brain infarct size in a rat model of stroke (3), a condition thought to arise due to increased ROS production. In addition, EUK-134 has been used successfully to reduce the effects of oxidative stress in several other models, including kainate-induced neuropathology (36), amyotrophic lateral sclerosis (22), Alzheimer's disease (2), spongiform encepalopathy (28), and endotoxin-induced multiple organ failure (6). Because superoxide and H2O2 are required to maintain vascular SMC viability, treatment with EUK-134 may stimulate apoptosis in these cells.
In this study we determined the effects of EUK-134 treatment on FPASMC ROS production, proliferation, and viability. We demonstrate that removal of ROS by EUK-134 slows proliferation and, at higher doses, induces programmed FPASMC death. SOD/catalase mimetics may, therefore, represent useful therapeutic agents in the prevention or reversal of the excessive vascular muscularization seen in patients with PPHN.
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MATERIALS AND METHODS |
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Cell culture. Primary cultures of FPASMCs from sheep were isolated
by the explant technique as described previously
(45). Briefly, a segment of
the main pulmonary artery from 136-day-old fetal lambs was excised and placed
in a sterile 10-cm dish containing DMEM supplemented with 1 g/l of glucose.
The segment was stripped of adventitia with a sterile forceps. The main
pulmonary artery segment was then cut longitudinally to open the vessel, and
the endothelial layer was removed by gentle rubbing with a cell scraper. The
vessel was then cut into 2-mm segments, inverted, and placed on a
collagen-coated 35-mm tissue culture dish. A drop of DMEM containing 10% fetal
bovine serum (HyClone), antibiotics (MediaTech), and antimycotics (MediaTech)
was then added, and the cells were grown overnight at 37°C in a humidified
atmosphere with 5% CO2-95% air. The next day, an additional 2 ml of
complete medium were added. The growth medium was subsequently changed every 2
days. When SMC islands could be observed under the microscope, the tissue
segment was removed, and the individual cell islands were subcloned. Identity
was confirmed as FPASMCs by immunostaining (>99% positive) with antibodies
against -smooth muscle actin, calponin, and caldesmon. This was taken
as evidence that cultures were not contaminated with fibroblasts or with
endothelial cells. All cultures for subsequent experiments were maintained in
DMEM supplemented with 10% fetal calf serum (Hyclone), antibiotics
(MediaTech), and antimycotics (MediaTech) at 37°C in a humidified
atmosphere with 5% CO2-95% air. Cells were utilized between
passages 3 and 10.
Fluorescence analysis. FPASMCs were seeded onto 96-well plates (Costar) and allowed to adhere for at least 18 h. Cells were then washed in PBS and incubated in DMEM containing 050 µM EUK-134, 50 U/ml of polyethylene glycol (PEG)SOD (Sigma), or 25 U/ml of PEG-catalase (Sigma) for 30 min. Dihydroethidium (DHE; 20 µM, Molecular Probes) or 2',7'-dichlorodihydrofluorescein diacetate (H2DCF-DA; 20 µM, Molecular Probes) was added to the media 15 min before the end of the experiment. Cells were washed with PBS and imaged using a Nikon Eclipse TE-300 fluorescent microscope. DHE-stained cells were observed after excitation at 518 nm and emission at 605 nm. H2DCF-DA-stained cells were observed using excitation at 485 nm and emission at 530 nm. Fluorescent images were captured using a CoolSnap digital camera, and the average fluorescent intensities (to correct for differences in cell number) were quantified using Metamorph imaging software (Fryer). Statistical analyses between treatments were carried out as detailed in Statistical analysis.
Cell proliferation assays. FPASMC at 2,500 cells/well were
seeded onto 96-well plates (Costar;
25% confluence) and allowed to adhere
for at least 18 h. The initial number of viable cells was then determined to
correct for differences in starting cell number between experiments and to
monitor changes in cell number over time. This was determined using the Cell
Titer 96 AQueous One Solution kit (Promega), the basis of which has
been shown to be a reliable alternative to [3H]thymidine
incorporation (14). The
tetrazolium reagent is bioreduced to a colored product, the quantity of which
is proportional to the number of metabolically active cells. Twenty
microliters of reagent was added directly to cells in 100 µl of medium, and
following a 2-h incubation period at 37°C, the absorbance at 492 nm was
read using a Labsystems Multiskan EX plate reader (Fisher). For the
proliferation assay, cells were washed with PBS and incubated with media
containing 10% fetal calf serum and 050 µM EUK-134 or a combination
of PEG-SOD and PEG-catalase with activities equivalent to 50 µM EUK-134.
The Cell Titer 96 AQueous One assay was repeated as described above
to determine the number of viable cells at 18, 30, and 48 h after
treatment.
Loss of mitochondrial membrane potential. Mitochondrial membrane potential has been analyzed previously using the lipophilic cation 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide (15). This dye fluoresces red in its multimeric form in healthy mitochondria and is the active reagent in the DePsipher Mitochondrial Potential Assay kit (Trevigen). FPASMCs were seeded onto 96-well plates and incubated with 0 or 50 µM EUK-134 as described above. DePsipher reagent (25 µg/ml) was added at 024 h after treatment with EUK-134 and incubated for a further 20 min. The aggregate red form was observed by fluorescence microscopy after excitation at 518 nm and emission at 605 nm as described above.
Caspase activation analysis. FPASMCs were seeded onto 96-well plates and incubated with 0 or 50 µM EUK-134 as described above. Caspase activation was visualized by cotreating cells with 1 µM CaspACE FITC-VAD-FMK In Situ Marker (Promega). This is a fluorescent analog of the pancaspase inhibitor Z-VAD-FMK, which readily enters cells and binds irreversibly to activated caspases (8). Fluorescent cells were observed using excitation at 485 nm and emission at 530 nm. Activation of specific caspase isoforms was determined by immunocytochemistry. After 24 h of EUK-134 treatment, cells were washed in PBS and fixed in 4% (vol/vol) paraformaldehyde at 4°C for 20 min. Cells were washed three times in PBS for 5 min and then treated with 0.1% IGEPAL CA-630 (Sigma) for 1 min at room temperature. After three PBS washes, cells were blocked in 5% (wt/vol) nonfat dry milk (Challenge Dairy Products) and 0.05% Tween (Fisher Biotech) in PBS for 60 min at room temperature. Cells were washed three times in PBS and incubated with primary antibodies against cleaved caspase-3 (1:250; Trevigen), cleaved caspase-8 (1:100; Cell Signaling), and cleaved caspase-9 (1:100; Cell Signaling) in 5% BSA (Sigma) and 0.05% Tween in PBS at 4°C for 16 h. Cells were washed three times in PBS and incubated with goat anti-rabbit (caspase-3 and caspase-9) or goat anti-mouse (caspase-8) IgG conjugated to rhodamine red (both from Molecular Probes) in 5% nonfat dry milk and 0.05% Tween in PBS for 60 min in the dark at room temperature. Cells were washed three times in PBS and visualized using fluorescence microscopy with excitation at 518 nm and emission at 605 nm as described above.
TdT-mediated dUTP nick end labeling analysis. TdT-mediated dUTP nick end labeling (TUNEL) analysis (13) was performed on EUK-134-treated FPASMC using the DeadEnd Colorimetric Apoptosis Detection System (Promega). FPASMCs were seeded onto 96-well plates and incubated with0or50 µM EUK-134 as described above. After 30 h, cells were washed in sterile PBS and fixed in 4% (vol/vol) paraformaldehyde for 20 min at 4°C. Cells were washed twice in PBS and then incubated with TdT and reaction mix, including fluorescein-12-dUTP, for 1 h at 37°C. Cells were washed for 30 min in 2x saline-sodium citrate buffer and then incubated with PBS plus 4',6-diamidino-2-phenylindole dihydrochloride (DAPI; 5 µM) for 15 min at room temperature. DAPI is a blue fluorescent nuclear stain, and this step ensured that approximately equal cells were imaged in each slide. The cells were visualized by indirect immunofluorescence with excitation at 485 nm and emission at 530 nm as described above.
Statistical analysis. The relative fluorescent intensity was calculated for DHE, dichlorofluorescein, FITC-VAD-FMK, and secondary antibodies to cleaved caspases and then expressed as means ± SD. The relative change in cell number was calculated for the treatment groups and expressed as means ± SD. Comparisons between treatment groups were made by ANOVA using the GB-STAT software program. P < 0.05 was considered statistically significant.
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RESULTS |
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We then determined the effects of EUK-134 on FPASMC proliferation utilizing an assay sensitive to the number of viable cells. An increase in viable cell number in the untreated samples (10% serum only) relative to the initial number of cells at time 0 is indicative of serum-induced proliferation; after 48 h, there was an increase of 110% (Fig. 2). Fewer viable cells in the antioxidant-treated samples relative to untreated samples indicates a decrease in proliferation. EUK-134 (5 µM) inhibited proliferation with an increase of 61% at 48 h, whereas a combination of SOD and catalase with predicted activities equivalent to 50 µM EUK-134 resulted in an increase of 5% (Fig. 2). Fewer viable cells in the antioxidant-treated samples relative to the initial number of cells at time 0 indicates cell death. Incubation with 50 µM EUK-134 resulted in 45% of the viable cell number relative to time 0 at 48 h, although SOD and catalase in combination gave no significant decrease (Fig. 2).
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To identify the mechanism of EUK-134-induced FPASMC death, we performed assays indicative of apoptosis. First, we examined whether EUK-134 treatment induced a loss of mitochondrial membrane potential. This was determined using DePsipher, a dye that readily enters cells and fluoresces bright red in its multimeric form within healthy mitochondria. In apoptotic cells, the electrochemical gradient across the mitochondrial membrane collapses, and the reagent cannot accumulate within the mitochondria. Figure 3 illustrates the loss of mitochondrial membrane potential in FPASMC following treatment with 50 µM EUK-134, with a significant decrease after 18 h.
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Next, we looked for evidence of caspase activation, another event characteristic of apoptosis. An FITC-conjugated pan-caspase inhibitor (FITC-VAD-FMK) was used as an in situ marker of programmed cell death. This cell-permeable molecule binds irreversibly to activated caspases and can be visualized by fluorescence microscopy. By visualizing cells at different time points, we detected an increase in FITC-VAD-FMK fluorescence in FPASMC at 24 h after the addition of 50 µM EUK-134, but not at 18 h (Fig. 4A). To identify specific members of the caspase family that become activated in FPASMC following exposure to EUK-134, we used antibodies against the cleaved isoforms of caspase-8, caspase-9, and caspase-3. Immunohistochemistry and fluorescence microscopy revealed an increase in active caspase-9 and caspase-3, but not in active caspase-8, 24 h after treatment with 50 µM EUK-134 (Fig. 4B).
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We also looked for DNA fragmentation as an additional late marker of programmed cell death. TUNEL analysis identified apoptotic nuclei in FPASMC at 30 h after treatment with 50 µM EUK-134, but not at 24 h (Fig. 5). SOD and catalase activity equivalent to 50 µM EUK-134 inhibited FPASMC proliferation but did not decrease the number of viable cells relative to time 0 (Fig. 2). Accordingly, we failed to detect TUNEL-positive nuclei in cells treated with PEG-SOD and PEG-catalase combined (data not shown).
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DISCUSSION |
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In this study, we demonstrate a decrease in serum-induced FPASMC proliferation in response to treatment with 5 µM EUK-134 or with a combination of SOD and catalase. The mechanisms that regulate vascular SMC growth are complex, and potential redoxsensitive molecules that mediate FPASMC growth are currently unknown. Recent studies have identified a role for the cyclin-dependent kinase inhibitor p27Kip1 in regulating rat pulmonary arterial SMC proliferation (16). Furthermore, prostacyclin analogs inhibited the serum-induced proliferation of human pulmonary arterial SMCs (12), whereas the 3-hydroxy-3-methylglutaryl-coenzyme A reductase inhibitor simvastatin attenuated monocrotaline-induced vascular remodeling in a rat model of pulmonary hypertension (30). However, these studies were conducted in adult models and cells and may have limited relevance to the fetal pulmonary vasculature. In the developing rat lung, maximal SMC proliferation occurred during the embryonic period, followed by a decline in the fetal period and a steady decrease after birth (5). Furthermore, the decrease in proliferation was associated with an increase in the expression of perlecan, which was predominant in nonreplicating cells. Perlecan, a basement membrane heparan sulfate proteoglycan, was shown to decrease vascular SMC proliferation by downregulating growth-promoting transcription factors (47). Nitric oxide (NO), produced in endothelial cells by the enzyme endothelial nitric oxide synthase (eNOS), may also play a role in the proliferation of adjacent vascular SMC. NO donors had a biphasic effect on rat FPASMC growth: DNA synthesis was stimulated by micromolar concentrations but inhibited at higher levels (43). Recently, we have demonstrated differences in NO and eNOS levels between fetal and adult pulmonary arterial endothelial cells (46), which may influence FPASMC growth in the developing lung. Additional studies are, therefore, warranted to identify the redoxsensitive mechanisms involved in FPASMC proliferation.
EUK-134 (50 µM) induced FPASMC apoptosis, characterized by a loss of mitochondrial membrane potential, activation of caspase-3 and caspase-9, and DNA fragmentation. Our findings that equivalent activities of SOD and catalase inhibited FPASMC proliferation but did not induce apoptosis suggest that a prolonged antioxidant capability is required. Indeed, one advantage of salen-manganese compounds is their increased stability and bioavailability relative to proteinaceous antioxidant enzymes (4, 17). Accordingly, DHE and H2DCF-DA fluorescence levels in FPASMCs treated with SOD and catalase were identical to controls after 30 h (data not shown). Conversely, fluorescence levels in FPASMC incubated with 50 µM EUK-134 were higher than in untreated cells after 30 h (data not shown). However, this apparent paradox can be explained by the release of ROS by mitochondria in cells undergoing apoptosis (26), an event we have demonstrated previously in apoptotic FPASMC (45).
The appearance of TUNEL-positive nuclei after 30 h of incubation with 50 µM EUK-134 is in agreement with our previous findings when looking at FPASMC apoptosis in response to treatment with ascorbic acid and with inhibitors of NADPH oxidase (45). However, this late onset of programmed cell death is unusual, since many proapoptotic stimuli generate DNA fragmentation much earlier. For example, Tsai et al. (44) found evidence of apoptosis in rat aortic SMCs treated with the antioxidants N-acetylcysteine and pyrrolidinedithiocarbamate after 6 h. This suggests that FPASMC may be more resistant to antioxidants in the short term or that different apoptotic pathways are activated in these cells. Certain apoptotic stimuli trigger a loss of mitochondrial membrane potential, resulting in the release of cytochrome c (42). Cytochrome c is then free to bind apoptotic protease-activating factor 1 (49), resulting in the activation of caspase-9 (35), which in turn activates caspase-3 (23). In the final stages of apoptosis, caspase-3 stimulates the cleavage of cytoskeletal, nuclear scaffold, DNA repair, and cell cycle proteins and initiates DNA fragmentation (34). Because EUK-134-induced FPASMC apoptosis involved the loss of mitochondrial membrane potential followed by caspase-9 and caspase-3 activation, it is likely that the pathway is regulated by mitochondrial-associated proteins (42). However, the influence of ROS on the expression of these proteins is poorly understood. A recent study demonstrated that the proapoptotic Bcl-2 proteins Bax and Bak were required for programmed cell death in rat 1a cells exposed to hyperoxia, a condition that increases levels of ROS (11). Surprisingly, ROS were not required to stimulate hyperoxia-induced apoptosis because antioxidant treatment did not prevent cell death. Furthermore, overexpression of Bcl-xL inhibited hyperoxia-induced apoptosis, suggesting that it is not regulated by ROS. Overall, these data illustrate the complexity of the mechanisms involved. Additional studies are, therefore, required to identify molecules that induce FPASMC apoptosis after prolonged ROS removal.
Many studies have looked at the potential of antioxidant treatment to prevent ROS-induced programmed cell death. Indeed, EUK-134 has been used successfully to inhibit ROS-induced apoptosis (25, 32). This suggests a unique phenotype of vascular SMC in which ROS are required for cell viability. The effects of antioxidants on vascular endothelial cell proliferation and survival are also poorly understood. One study found that antioxidants decreased the viability of aortic smooth muscle cells at a concentration that had no effect on the viability of endothelial cells from the same source vessels (44). Since PPHN is characterized by impaired endothelial activity (7), it is important that any treatment strategy does not further disrupt this function. Studies to determine fetal pulmonary arterial endothelial cell viability in response to antioxidant treatment are, therefore, required. Our findings that equivalent units of SOD and catalase to that in 50 µM EUK-134 prevented FPASMC proliferation, but did not induce apoptosis, highlights the advantages of using synthetic salen-manganese compounds over proteinaceous enzymes. Higher doses of SOD and catalase, or repeated treatments, may be required to induce apoptosis using these enzymes. The effects of EUK-134 on vascular remodeling and pulmonary hypertension in the ductal ligation model of PPHN may reveal the potential of this antioxidant as a therapeutic drug.
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ACKNOWLEDGMENTS |
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This research was supported in part by National Institutes of Health Grants HL-60190, HL-67841, and HD-398110 and March of Dimes Grant FY00-98 (all to S. M. Black).
S. M. Black is a member of the Feinberg Cardiovascular Research Institute.
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FOOTNOTES |
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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.
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REFERENCES |
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