INTRODUCTION

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VASCULAR SMOOTH MUSCLE CELLS (VSMC) are subjected to hemodynamic forces in vivo that dynamically modulate cellular responses. One such force, cyclic strain, which results from the increase in the circumference of the vessel wall between diastole and systole (6, 28), achieves pathophysiological significance in vascular disease states such as hypertension (7) and atherosclerosis (33), where mechanical forces on the arterial wall may be persistently increased. Mechanical forces have been shown to mediate the synthesis and/or secretion of platelet-derived growth factor (37), fibroblast growth factor (5), and angiotensin II (38), which, in turn, modulate protein synthesis and cell proliferation. In addition, elevated levels of cyclic strain have been shown to modulate the formation of reactive oxygen species (ROS) in human aortic endothelial cells (12) and coronary artery smooth muscle cells (13); however, the influence of cyclic strain on vascular smooth cell superoxide production has not yet been fully elucidated.

Once formed, ROS are scavenged via several intracellular pathways. In the cell cytoplasm, a pathway that utilizes glutathione to maintain the cellular redox state is favored. Glutathione may react directly with ROS as well as serve as an electron donor for glutathione peroxidase, which reduces hydrogen and lipid peroxides (9, 20). As peroxides are metabolized by glutathione peroxidase, reduced glutathione (GSH) is oxidized to glutathione disulfide (GSSG) (9, 19, 20). Glutathione reductase restores intracellular GSH levels by reducing GSSG in a reaction that requires NADPH reducing equivalents. The principle source(s) of NADPH in the cell is the pentose phosphate pathway (9, 17).

Glucose-6-phosphate dehydrogenase (G6PDH) is the first and rate-limiting enzyme in the pentose phosphate pathway, which produces sugar moieties for nucleic acid synthesis (17). G6PDH converts glucose-6-phosphate into 6-phosphogluconate in an NADP⁺-dependent process, thereby generating NADPH. The importance of G6PDH in the defense against oxidant stress has long been recognized in erythrocytes and was recently demonstrated in nucleated eukaryotic cells (35). Saccharomyces cerevisiae G6PDH-deficient mutants are uniquely susceptible to oxidant stress (15), as are murine ES cells containing a G6PDH null mutation (27). In contrast, HeLa cell clones transfected with the G6PDH gene maintain elevated GSH levels in the setting of oxidant stress (34). Furthermore, G6PDH expression is enhanced in human cells treated with agents that increase ROS formation or deplete intracellular GSH pools (30, 34).

The role of mechanical forces on G6PDH expression and activity has not been previously investigated. Here we have demonstrated that pathophysiologically relevant levels of cyclic strain promote ROS formation and GSH depletion in neonatal rat VSMC, which, in turn, induce G6PDH expression and increase G6PDH activity. We further observed that, in G6PDH-deficient states, GSH repletion is impaired and an imbalance in the cellular redox potential is prolonged.

	METHODS

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

Cell culture and application of cyclic strain. Neonatal rat VSMC were a generous gift of Dr. Herbert Kagan. Cells were grown in RPMI supplemented with 10% fetal bovine serum, 100 U/ml penicillin, and 100 µg/ml streptomycin. Cells were passaged twice weekly by harvesting with 0.5% trypsin-EDTA. For cyclic strain experiments, cells were grown on collagen-coated plates and utilized at confluence between passages 5 and 30. In some experiments, cells were treated with dehydroepiandrosterone (DHEA; 100 µM) for 24 h. VSMC were subjected to either 0, 21, or 27% strain, which corresponds to 0, 15, or 20 kPa at a frequency of 1 Hz (60 cycles/min) for 0, 12, 18, 24, or 30 h, as previously described (3).

Superoxide anion production. Superoxide anion production was measured as previously described (11) with some modifications. Briefly, VSMC were harvested from the plates with type I collagenase (1 mg/ml), soybean trypsin inhibitor (1 mg/ml), and bovine serum albumin (2 mg/ml) and were then centrifuged (500 g, 4°C, 10 min). The cell pellet was resuspended in an assay buffer (in mM: 130 NaCl, 5 KCl, 1 MgCl₂, 1.5 CaCl₂, 35 phosphoric acid, and 20 HEPES, pH 7.4), centrifuged again, resuspended in assay buffer with 10 mM glucose and 1 mg/ml bovine serum albumin, and then stored on ice until use. To measure superoxide anion production, we incubated cells at room temperature for 5 min, added dark-adapted lucigenin (5 µM), and measured photon emission every 15 s for 10 min in a luminometer (model 20E, Turner Designs). A lucigenin blank (<5% of the cell signal) was run with each experiment to ensure that autoxidation did not occur (8) and was subtracted from each time point before transformation of the data. A standard curve for comparison was generated with the use of xanthine/xanthine oxidase (24) to determine the amount of superoxide produced at each time point. Protein content was determined in an aliquot of cells (18), and enzyme activities were standardized to protein to facilitate comparison between treatment groups. To determine the relative contribution of the flavin-containing enzymes NADH/NADPH oxidase, xanthine oxidase, nitric oxide synthase, and cyclooxygenase to superoxide production in this system, we added diphenylene iodonium (10 µM), oxypurinol (100 µM), nitro-L-arginine methyl ester (L-NAME; 10 µM), or indomethacin (10 µM), respectively, to selected experiments and measured superoxide generation.

NADH/NADPH oxidase assay. NADH/NADPH oxidase(s) activity was measured as previously described (11) with some modifications. Cells were washed with ice-cold phosphate-buffered saline and scraped from the plate. Cells from individual wells per cyclic strain plate were combined before centrifugation. Cells were then centrifuged at 1,000 g at 4°C for 10 min. The pellet was resuspended in a lysis buffer containing protease inhibitors (20 mM KH₂PO₄, pH 7.0, 1 mM EGTA, 0.5 µg/ml leupeptin, 0.7 µg/ml pepstatin, and 0.5 mM phenylmethylsulfonyl fluoride) and manually homogenized on ice. NADH or NADPH oxidase activity was measured by a luminescence assay in a 50 mM phosphate buffer, pH 7.0, containing 1 mM EGTA, 150 mM sucrose, 5 µM dark-adapted lucigenin as the electron acceptor, and 100 µM of either NADH or NADPH as the substrate in a final volume of 450 µl. The reaction was started by the addition of 100 µl of homogenate, and luminescence measurements were obtained every 15 s for 10 min. Protein content was determined in an aliquot of the homogenate (18), and the results were standardized to this measurement.

G6PDH activity. G6PDH activity was measured as described previously (31). Cells were washed with phosphate-buffered saline (0.9%), scraped from the plate in 1 ml of assay buffer (50 mM Tris, 1 mM MgCl, pH 8.1), and centrifuged at 2,000 g at 4°C for 10 min. Enzyme activity was determined using a plate-reader spectrophotometer (ThermoMax Microplate Reader; Molecular Devices, Menlo Park, CA) by measuring the rate of increase of absorbance at 340 nm due to the conversion of NADP⁺ to NADPH by either G6PDH or 6-phosphogluconate dehydrogenase (6-PGDH). To determine total dehydrogenase activity, we added 20 µl of supernatant to a well that contained 180 µl of assay buffer and substrates for both enzymes. In a second well, substrates for the enzyme 6-PGDH were added to determine the activity of this enzyme. The rate of change was measured over a 6-min period. G6PDH activity was then determined by subtracting 6-PGDH activity from total dehydrogenase activity. Substrate concentrations were glucose-6-phosphate (200 µM), 6-phosphogluconate (200 µM), and NADP⁺ (100 µM). Protein levels were determined for each sample (18), and activity results were standardized to protein concentration.

Western blot analysis. VSMC from stretched plates were harvested and then centrifuged at 1,000 g for 10 min at 4°C, after which the supernatant was discarded and the samples were frozen at 80°C overnight. The pellet was then homogenized, protein concentration was determined, and 25 µg of protein were added per lane. Protein was size-fractionated electrophoretically with the use of SDS-polyacrylamide gel electrophoresis on a 10% gel and transferred to nitrocellulose membranes blocked with 5% casein solution. The membranes were incubated with a 1:1,000 dilution of a polyclonal rabbit anti-G6PDH antibody (Sigma) and visualized with a 1:2,000 dilution of a goat anti-rabbit secondary antibody conjugated to horseradish peroxidase (Amersham). Signals were detected using enhanced chemiluminescence (ECL system; Amersham).

Intracellular glutathione measurement. Intracellular GSH and total glutathione (GSH + GSSG) levels were measured by utilizing the Bioxytech GSH-400 enzymatic method (OXIS) according to the manufacturer's instructions. Briefly, total cellular protein was precipitated in 5% metaphosphoric acid and centrifuged, and the supernatant was isolated for measurements. The level of GSH or total glutathione was determined from a standard curve of GSH. Total glutathione was calculated by using an apparent molar extinction coefficient at 356 nm (M¹ · cm¹) of 17,400 for glutathione. Protein content (18) was determined before precipitation by metaphosphoric acid.

Cellular glutathione peroxidase and glutathione reductase activities. Cellular glutathione peroxidase and glutathione reductase activities were determined by using the Bioxytech GPx-340 assay (OXIS) and the GR-340 assay (OXIS), respectively, in accordance with the manufacturer's instructions. Briefly, to determine glutathione peroxidase activity, cells were scraped from the plate and homogenized in cold assay buffer (50 mM Tris · HCl, pH 7.5, 5 mM EDTA, and 1 mM dithiothreitol). Homogenates were centrifuged, and the supernatant was stored on ice. Glutathione peroxidase activity was determined indirectly by measuring the oxidation of NADPH to NADP⁺ at 340 nm (Beckman DU 640B spectrophotometer; Beckman Coulter, Fullerton, CA) over time that occurs with recycling oxidized glutathione. To determine glutathione reductase activity, we scraped cells from the plate and homogenized them in cold assay buffer (50 mM potassium phosphate, pH 7.5, and 1 mM EDTA). The supernatant was isolated and stored on ice. Glutathione reductase activity was measured by the oxidation of NADPH to NADP⁺ at 340 nm over time. Activity results were standardized to cell protein levels (18).

Lactate dehydrogenase release. Cells were treated according to experimental protocol, and lactate dehydrogenase (LDH) release was measured in the medium of cells after treatment with an LDH detection kit (Sigma) according to the manufacturer's protocol. Results were compared with measurements in media from untreated cells (negative control) and cell lysates (positive control).

Antisense phosphorothioate oligodeoxynucleotide transfection. Cells were grown to 80% confluence and transfected with an antisense oligodeoxynucleotide to G6PDH mRNA (5'-AGGUCACCCGAUGCACCCAUGAUGA-3') (Sequitor, Natick, MA) for 14 h, by using Oligofectin G (1.25 µl/ml medium; Sequitor) as the vehicle, in serum-free, antibiotic-free Opti-MEM medium (GIBCO-BRL), or Oligofectin G alone. Cells were then washed twice in serum-free medium and transferred to full-growth medium for an additional 24 h before the application of cyclic strain. Cells were assayed according to the specific treatment protocol.

Statistical analysis. Data are expressed as means ± SE. Comparison between groups was performed by Student's paired two-tailed t-test.

	RESULTS

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

Cyclic strain and superoxide production. VSMC were subjected to 0, 15 (physiological), or 20 (pathophysiological) kPa of cyclic strain at 1 Hz for 24 h. While there was no difference in superoxide production in cells exposed to 0 or 15 kPa of strain [11.3 ± 0.2 vs. 10.6 ± 0.4 nmol O₂ · 10 min¹ · mg protein¹, P = not significant (NS), n = 3], superoxide production was significantly increased in smooth muscle cells subjected to 20 kPa of cyclic strain (11.3 ± 0.2 vs. 14.4 ± 0.2 nmol O₂ · 10 min¹ · mg protein¹, P < 0.0001, n = 3) (Fig. 1A.). On the basis of these observations, cells were subjected to 0 (control) or 20 kPa of cyclic strain in all subsequent experiments. To examine the time course of superoxide production, we subjected VSMC to cyclic strain for 0, 12, 18, or 24 h. Compared with levels in unstretched cells, superoxide levels were significantly elevated after 18 h of strain (10.5 ± 0.2 vs. 13.2 ± 0.1 nmol O₂ · 10 min¹ · mg protein¹, P < 0.0001, n = 3) and increased further by 24 h (10.9 ± 0.2 vs. 14.3 ± 0.1 nmol O₂ · 10 min¹ · mg protein¹, P < 0.0001, n = 3), after which time a plateau was reached (Fig. 1B). Accordingly, a 24-h time point was chosen for subsequent experiments. Superoxide production was significantly attenuated by the addition of superoxide dismutase (100 U/ml) in VSMC subjected to 0 (10.2 ± 0.5 vs. 5.1 ± 0.4 nmol O₂ · 10 min¹ · mg protein¹, P < 0.002, n = 3) and 20 kPa (16.2 ± 1.2 vs. 4.1 ± 1.2 nmol O₂ · 10 min¹ · mg protein¹, P < 0.007, n = 3) of cyclic strain.

View larger version (14K): [in this window] [in a new window]   Fig. 1.   Superoxide (O2) production by vascular smooth muscle cells (VSMC) subjected to cyclic strain is threshold and time dependent. A: VSMC were subjected to 0, 15, or 20 kPa of cyclic strain at 1 Hz for 24 h (n = 3). B: superoxide production was measured in VSMC subjected to 0 () or 20 kPa () cyclic strain at 1 Hz for 0, 12, 18, or 24 h (n = 3). Superoxide is expressed as nmol O2 · 10 min1 · mg protein1, and data are given as means ± SE. *P < 0.00001 vs. 0 kPa of cyclic strain.

Superoxide production results from enhanced NADH/NADPH oxidase(s) activity. To determine the source of superoxide production, we treated cells with diphenylene iodonium (100 µM), oxypurinol (100 µM), L-NAME (10 µM), or indomethacin (10 µM) to investigate the role of the flavin-containing enzymes NADH/NADPH oxidase, xanthine oxidase, nitric oxide synthase, and cyclooxygenase, respectively. Only the addition of diphenylene iodonium significantly inhibited superoxide production in both cells subjected to 0 (11.3 ± 0.2 vs. 8.6 ± 0.3 nmol O₂ · 10 min¹ · mg protein¹, P < 0.00001, n = 3) and 20 kPa (14.4 ± 0.2 vs. 8.7 ± 0.2 nmol O₂ · 10 min¹ · mg protein¹, P < 0.00001, n = 3) of cyclic strain (Table 1). This result suggests that NADH/NADPH oxidase(s) contributed significantly to superoxide formation in VSMC exposed to cyclic strain. To confirm these observations, we measured NADH oxidase and NADPH oxidase activities directly (Fig. 2). Compared with cells exposed to no strain, VSMC subjected to 20 kPa of cyclic strain demonstrated increased NADH oxidase activity (5.3 ± 1.0 vs. 23.5 ± 3.3 µmol O₂ · 10 min¹ · mg protein¹, P < 0.05, n = 4) and NADPH oxidase activity (3.5 ± 0.5 vs. 6.5 ± 0.5 µmol O₂ · 10 min¹ · mg protein¹, P < 0.006, n = 4).

View larger version (13K): [in this window] [in a new window]   Fig. 2.   Cyclic strain increases NADH and NADPH oxidase activity. VSMC were exposed to 0 (open bars) or 20 kPa (filled bars) of cyclic strain for 24 h, and oxidase activity was measured in cell homogenates (n = 4). NADH oxidase and NADPH oxidase activity are measured as µmol O2 · 10 min1 · mg protein1 and data are reported as means ± SE. *P < 0.05 vs. 0 kPa of cyclic strain. #P < 0.006 vs. 0 kPa of cyclic strain.

Cyclic strain and intracellular GSH levels. To examine the influence of increased superoxide production on cellular redox state, we measured total glutathione (GSH + GSSG) and GSH levels. Concomitant with an increase in superoxide production, 20 kPa of cyclic strain produced a significant increase in total glutathione levels by 18 h compared with that at baseline (0 h) (123.3 ± 1.5 vs. 101.3 ± 1.0% control, P < 0.0001, n = 4), but a significant reduction in GSH levels (56.5 ± 3.2 vs. 93.0 ± 7.3% control, P < 0.004, n = 4), indicating an increase in GSSG formation. Total glutathione levels remained elevated in cells exposed to 20 kPa of cyclic strain for 24 h compared with levels at baseline (188.0 ± 9.8 vs. 101.3 ± 1.0% control, P < 0.0001, n = 4), and GSH levels were increased (196.0 ± 8.8 vs. 93 ± 7.34% control, P < 0.0001, n = 4) (Fig. 3), suggesting enhanced glutathione recycling.

View larger version (21K): [in this window] [in a new window]   Fig. 3.   Cyclic strain influences reduced glutathione (GSH) and total glutathione [GSH + oxidized glutathione disulfide (GSSG)] levels in a time-dependent manner. VSMC were exposed to 0 or 20 kPa cyclic strain for 0, 12, 18, or 24 h, and GSH (open bars) and total glutathione (GSH + GSSG) levels (filled bars) were determined (n = 4). GSH and total glutathione levels are measured as µmol · l1 · mg protein1, and results are expressed as %control, with 0 kPa of strain serving as the control for 20 kPa of strain at each time point. Data are presented as means ± SE. *P < 0.004 vs. 0 h. #P < 0.0001 vs. 0 h.

Cyclic strain and glutathione peroxidase and glutathione reductase activities. To evaluate whether the observed flux in GSH levels was the result of increased cycling to GSSG or decreased recycling of GSSG, we measured the activities of glutathione peroxidase and glutathione reductase. Compared with activities in unstretched cells, there was a significant increase in both glutathione peroxidase activity (384.3 ± 42.0 vs. 730.0 ± 96.0 mU · ml¹ · mg protein¹, P < 0.03, n = 3) and glutathione reductase activity (29.4 ± 2.1 vs. 72.1 ± 4.5 mU · ml¹ · mg protein¹, P < 0.02, n = 3) at 18 h in cells exposed to 20 kPa of cyclic strain, and these activities remained elevated after 24 h of cyclic strain (Table 2).

Cyclic strain and G6PDH activity and expression. Because G6PDH is an important source of NADPH, which is required as a cofactor for the conversion of GSSG to GSH by glutathione reductase to restore the intracellular thiol redox state, we investigated the influence of cyclic strain on G6PDH activity and expression in cells subjected to cyclic strain. G6PDH activity was enhanced after 18 h of cyclic strain (622.6 ± 27.0 vs. 807.9 ± 32.2 U · 6 min¹ · mg protein¹, P < 0.001, n = 6) and achieved maximal activity by 24 h (602.1 ± 52.7 vs. 1,167.5 ± 88.0 U · 6 min¹ · mg protein¹, P < 0.001, n = 6) (Fig. 4). Western blot analysis demonstrated an increase in G6PDH protein that correlated with the increase in enzyme activity (Fig. 5).

View larger version (16K): [in this window] [in a new window]   Fig. 4.   Cyclic strain increases glucose-6-phosphate dehydrogenase (G6PDH) activity. VSMC were exposed to 0 or 20 kPa of cyclic strain for 24 h, and G6PDH activity was determined. G6PDH activity was measured as U · 6 min1 · mg protein1 (n = 6), and data are presented as means ± SE. *P < 0.001 vs. 0 kPa.

View larger version (20K): [in this window] [in a new window]   Fig. 5.   Cyclic strain increases G6PDH expression. Top: Western blot analysis was performed to examine G6PDH expression (59 kDa) in cells subjected to 0 or 20 kPa of cyclic strain for 24 h. Optical density (OD units) is reported below each band. B: corresponding G6PDH activity was measured as U · 6 min1 · mg protein1 (n = 6) in VSMC subjected to 0 or 20 kPa of cyclic strain for 24 h. Data are presented as means ± SE. *P < 0.001 vs. 0 kPa.

1

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Inhibition of G6PDH activity and intracellular GSH levels. To confirm the importance of G6PDH as a source of NADPH to restore intracellular GSH, we inhibited G6PDH enzyme activity by treatment with DHEA (100 µM), a recognized noncompetitive inhibitor of the enzyme. Superoxide production in VSMC treated with DHEA for 24 h was not significantly different in cells exposed to 0 kPa of strain (11.0 ± 0.4 vs. 10.5 ± 0.3 nmol O₂ · 10 min¹ · mg protein¹, P = NS, n = 3); however, compared with untreated cells, VSMC treated with DHEA and subjected to 20 kPa of strain demonstrated a modest reduction in superoxide generation (15.3 ± 1.0 vs. 12.3 ± 0.3 nmol O₂ · 10 min¹ · mg protein¹, P < 0.05, n = 3). Treatment with DHEA for 24 h significantly inhibited G6PDH activity in VSMC exposed to 0 (602.1 ± 52.7 vs. 10.2 ± 1.0 U · 6 min¹ · mg protein¹, P < 0.00001, n = 6) and 20 kPa (1,167.5 ± 88.0 vs. 15.7 ± 3.2 U · 6 min¹ · mg protein¹, P < 0.00001, n = 6) of cyclic strain (Fig. 6A). This inhibition of G6PDH activity was not due to a decrease in G6PDH protein expression, as shown by Western blotting (Fig. 6B), or an increase in cell death, as demonstrated by LDH release from cells treated with DHEA subjected to 0 or 20 kPa of cyclic strain compared with that from a cell lysate, respectively (61.2 ± 0.1 vs. 98.9 ± 7.2 vs. 1,786.5 ± 2.0 units LDH/ml, P < 1 × 10¹² vs. cell lysate, n = 4). This reduction in G6PDH activity was accompanied by a significant decrease in total glutathione levels in DHEA-treated compared with untreated cells (70.4 ± 3.7 vs. 158.3 ± 9.0% control, P < 0.0008, n = 3) after 24 h of cyclic strain as well as a marked reduction in GSH levels in DHEA-treated compared with untreated cells subjected to cyclic strain (55.9 ± 7.5 vs. 183.9 ± 6.9% control, P < 0.0003, n = 3) (Fig. 6C). Glutathione peroxidase activity was significantly elevated in DHEA-treated compared with untreated cells exposed to 0 (717.3 ± 60.7 vs. 283.7 ± 34.2 mU · ml¹ · mg protein¹, P < 0.004, n = 3) and 20 kPa (1,164.6 ± 200.8 vs. 522.2 ± 46.3 mU · ml¹ · mg protein¹, P < 0.006, n = 3) of cyclic strain. In contrast, glutathione reductase activity was not increased in DHEA-treated cells subjected to 0 or 20 kPa of cyclic strain (40.9 ± 5.5 vs. 41.2 ± 4.5 mU · ml¹ · mg protein¹, P = NS, n = 3).

View larger version (21K): [in this window] [in a new window]   Fig. 6.   Dehydroepiandrosterone (DHEA) inhibits G6PDH activity and GSH repletion in cells subjected to cyclic strain without altering G6PDH expression. A: cells were treated with DHEA (100 µM) for 24 h and subjected to 0 (open bars) or 20 (shaded bars) kPa of cyclic strain. G6PDH activity (U · 6 min1 · mg protein1) was determined (n = 6), and data are presented as means ± SE. *P < 0.001 vs. 0 kPa. **P < 0.00001 vs. 0 kPa (DHEA). #P < 0.00001 vs. 20 kPa (DHEA). B: Western blotting reveals that the decline is G6PDH activity seen with DHEA treatment is not the result of decreased protein expression. Densitometry (OD units) is reported below each band. C: total glutathione and GSH levels (n = 4) were measured as µmol · l1 · mg protein1, and results are expressed as %control with 0 kPa of strain serving as the control for 20 kPa of cyclic strain. Data are reported as means ± SE. *P < 0.000001.

Inhibition of G6PDH expression and intracellular GSH levels. To establish further the role of G6PDH in maintaining cellular redox state, G6PDH expression was diminished by transfection with an antisense phosphorothioate oligodeoxynucleotide to G6PDH mRNA. After transfection for 14 h, there was a 69% reduction of G6PDH protein expression as demonstrated by Western blot analysis (Fig. 7A, top), and G6PDH expression was only minimally enhanced after 20 kPa of cyclic strain for 24 h (Fig. 7A, bottom). This degree of inhibition of G6PDH expression was not associated with a significant reduction in superoxide production in cells subjected to 20 kPa of cyclic strain for 24 h (13.9 ± 0.8 vs. 12.6 ± 1.0 nmol O₂ · 10 min¹ · mg protein¹, P = NS, n = 3). G6PDH activity was decreased by 26% in transfected cells subjected to 0 kPa of cyclic strain (775.0 ± 31.2 vs. 572.0 ± 5.3 U · 6 min¹ · mg protein¹, P < 0.0002, n = 4) and by 45% in cells that experienced 24 h of 20 kPa of cyclic strain (1,124.0 ± 70.7 vs. 621.5 ± 23.7 U · 6 min¹ · mg protein¹, P < 0.001, n = 4) (Fig. 7B). This decrease in G6PDH activity was not the result of cell death, as demonstrated by LDH release from transfected cells subjected to 0 or 20 kPa of cyclic strain for 24 h compared with that from a cell lysate preparation (60.7 ± 11.2 vs. 38.6 ± 10.8 vs. 1,585.3 ± 2.6 units LDH/ml, P < 1 × 10¹¹ vs. cell lysate, n = 4). The reduction in G6PDH activity was accompanied by a significant reduction in total glutathione levels in transfected compared with nontransfected cells subjected to cyclic strain for 24 h (103.3 ± 2.0 vs. 188.0 ± 10% control, P < 0.0002, n = 4). GSH levels were also significantly reduced in transfected compared with nontransfected cells subjected to cyclic strain for 24 h (48.2 ± 7.6 vs. 200.8 ± 12.4% control, P < 0.00005, n = 4) (Fig. 7C). The decrease in G6PDH expression was additionally associated with an increase in glutathione peroxidase activity in cells subjected to 0 kPa of cyclic strain, which was augmented further by 20 kPa of cyclic strain (421.6 ± 73.5 vs. 1,117.12 ± 207.4 mU · ml¹ · mg protein¹, P < 0.03, n = 3). In contrast, glutathione reductase activity was not increased in transfected cells subjected to 0 or 20 kPa of cyclic strain for 24 h (30.2 ± 5.5 vs. 32.5 ± 3.6 mU · ml¹ · mg protein¹, P = NS, n = 3) and, in fact, was decreased compared with nontransfected cells subjected to 20 kPa of cyclic strain for 24 h (32.5 ± 3.6 vs. 70.9 ± 9.0 mU · ml¹ · mg protein¹, P < 0.02, n = 3).

View larger version (20K): [in this window] [in a new window]   Fig. 7.   Transfection with an antisense phosphorothioate oligodeoxynucleotide to G6PDH mRNA inhibits G6PDH expression, activity, and GSH repletion in VSMC subjected to cyclic strain. A: cells were transfected with an antisense oligodeoxynucleotide to G6PDH mRNA (+AS) for 14 h or treated with the Oligofectin G vehicle (AS), and G6PDH expression was determined by Western blot analysis (top). Cells were then subjected to 0 or 20 kPa of cyclic strain for 24 h (bottom). Band density (OD units) is reported below each band. B: G6PDH activity (U · 6 min1 · mg protein1) was measured (n = 6) in transfected cells (+AS) and in cells treated with Oligofectin G vehicle (AS), and data are presented as means ± SE. *P < 0.0002 vs. AS. **P < 0.001 vs. 0 kPa (AS). #P < 0.001 vs. 20 kPa (AS). C: total glutathione and GSH levels (n = 4) were measured as µmol · l1 · mg protein1, and results are expressed as %control with 0 kPa of strain serving as the control for 20 kPa of cyclic strain. Data are presented as means ± SE. *P < 0.003 vs. AS. **P < 0.0001 vs. 20 kPa.

	DISCUSSION

TOP ABSTRACT INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

In these studies, we found that pathophysiologically relevant levels of cyclic strain applied to VSMC in vitro stimulated superoxide production by increased activity of NADH/NADPH oxidase(s) and, concomitant with this response, enhanced G6PDH expression and activity. Increased formation of superoxide imposed an oxidant stress on the cell as demonstrated by the decrease in cellular GSH levels and increase in GSSG formation. This change in cellular thiol redox status was accompanied by enhanced activity of glutathione peroxidase and glutathione reductase, indicating that there is an increase in GSH/GSSG cycling. Furthermore, cyclic strain induced G6PDH expression and enhanced enzyme activity. The role of G6PDH in modulating the cellular response to oxidant stress and maintaining intracellular GSH levels was confirmed by inhibiting enzyme activity with the noncompetitive inhibitor DHEA. Similarly, when G6PDH expression was reduced by an antisense phosphorothioate oligodeoxynucleotide to G6PDH mRNA, enzyme activity was diminished and GSH levels failed to be restored to baseline values. Inhibition of G6PDH expression and activity was associated with an increase in glutathione peroxidase activity, suggesting that G6PDH modulates basal antioxidant defense. The accompanying failure to augment glutathione reductase activity in the setting of enhanced GSSG formation demonstrates further the role of G6PDH as a source of NADPH to maintain cellular thiol redox homeostasis.

It is not entirely surprising that cyclic strain stimulates superoxide production via NADH/NADPH oxidase(s) in VSMC. Cyclic strain-mediated superoxide generation has been demonstrated previously in human aortic endothelial cells (12) and coronary artery smooth muscle cells (13), and this response, in part, was attributed to an increase in NADPH oxidase activity. Although one group of investigators failed to demonstrate an increase in cyclic strain-mediated superoxide production in VSMC, review of their data shows that they exposed cells to a subthreshold level of strain (14).

The NADH/NADPH oxidase(s) system appears to play a predominant role in superoxide production in vascular cells (22, 23, 25). For example, VSMC treated with angiotensin II produce abundant superoxide by increasing NADH/NADPH oxidase activity, and infusion of angiotensin II to produce hypertension in the rat enhances superoxide generation via a similar mechanism. (11, 29). Similarly, the importance of the NAD(P)H oxidase enzyme has been described in rabbit aortic fibroblasts (25, 26).

G6PDH, the first enzyme in the pentose phosphate pathway, is an important source of NADPH and, hence, participates in the defense against oxidant stress (27, 34). G6PDH activity is increased when GSH stores are depleted (30) and, in turn, promotes the formation of GSH from its oxidized form, GSSG, to restore the cellular redox state. This response has been demonstrated previously in cultured cells exposed to hydrogen peroxide (15, 34) or GSH-depleting drugs (30). Here we have shown for the first time that a mechanical force, cyclic strain, enhances G6PDH expression and upregulates G6PDH activity in the setting of GSH depletion, and we have shown that this response occurs concomitantly with an increase in superoxide production. The exact mechanism underlying the translation of the cyclic strain-mediated mechanical signal to a biochemical response for G6PDH remains undetermined and requires further investigation.

It is important to note that G6PDH-derived NADPH is utilized as a cofactor and substrate for several cellular enzyme systems, including glutathione reductase, nitric oxide synthase, dihydrofolate reductase (in the synthesis of tetrahydrobiopterin), and NADPH oxidase (36). While it may seem counterintuitive that the cell can provide substrate for both oxidant (NADPH oxidase) and antioxidant (glutathione reductase) functions, it is likely that the relative flux, or location, of these enzymes vis-à-vis that of NADPH may ultimately determine the net cellular redox effect. In addition, we and others (11) have shown that the predominant superoxide-generating enzyme in VSMC is NADH oxidase; the relative contribution of NADPH oxidase, albeit significant, is far less by comparison.

G6PDH is expressed in all cell types, although to varying degrees (4). The level of expression, and hence activity, may determine the relative contribution of this enzyme to cellular antioxidant defenses. In cells in which G6PDH activity is singularly important for oxidant defense, such as erythrocytes, G6PDH deficiency promotes adverse sequelae in the setting of ROS. We evaluated the relative contribution of G6PDH activity to antioxidant defense in VSMC by using two methods: inhibiting enzyme activity with DHEA and reducing G6PDH expression by transfection with an antisense phosphorothioate oligodeoxynucleotide to G6PDH mRNA.

DHEA inhibits G6PDH activity in mammalian cells by binding to the enzyme-coenzyme substrate ternary complex(es) (10) and may, as a result, promote oxidant stress. Interestingly, in our DHEA-treated cells subjected to cyclic strain, superoxide anion production was modestly diminished. This likely resulted from decreased substrate availability with significant G6PDH inhibition, because it was previously shown that DHEA does not directly influence the activity of NADPH oxidase (21). Despite the decrease in superoxide production, DHEA-treated cells experienced significant oxidant stress as revealed by the depleted intracellular GSH stores after 24 h of 20 kPa of cyclic strain. The decrease in glutathione reductase activity, an NADPH-requiring enzyme, contributes further to the decrease in GSH levels by preventing glutathione recycling. Moreover, the increase in glutathione peroxidase activity following DHEA treatment suggests that G6PDH contributes to VSMC antioxidant defense under basal conditions.

In contrast to our observations, it has been suggested that DHEA acts as an antioxidant agent (32). Many of these studies have been performed in models that utilize hyperglycemia as the oxidant stress (1, 2). Therefore, inhibition of G6PDH would result in an inhibition of glycolysis, and hence a protective effect, in this model. It has also been reported that DHEA increases catalase activity (16), although these observations have not been confirmed in other models. Finally, as a steroid, DHEA may induce synthesis of proteins that could influence cellular redox state.

Because enzyme activity and expression are linked, we chose to inhibit G6PDH expression only partially, because complete inhibition of this enzyme was surmised to be lethal (27). We utilized antisense technology to inhibit G6PDH protein expression to overcome the limitations associated with using a steroid compound to inhibit G6PDH activity. Partial inhibition of G6PDH enzyme expression and activity did not suppress the increase in superoxide production and prevented GSH repletion in cells subjected to 20 kPa of cyclic strain for 24 h. This phenomenon has been described previously in murine embryonic stem cells in which targeted homologous recombination was used to disrupt the G6PDH gene. With the use of this technique, a population of G6PDH null cells was created and was found to be exquisitely sensitive to oxidant stress (27). As we demonstrated with DHEA, transfected VSMC did not increase glutathione reductase activity, presumably because of a lack of NADPH, to recycle GSSG back to GSH.

It was demonstrated previously that G6PDH expression and activity are increased by oxidant stress to restore the intracellular redox state. We have now shown, for the first time, that G6PDH expression and activity are dynamically regulated in response to oxidant stress in vascular cells, specifically, VSMC. Additionally, we have included a mechanical force, cyclic strain, to the list of oxidant stressors that increase G6PDH expression and activity and further suggest that G6PDH may play a significant role in modulating vascular redox state.