Effects of Propofol on Cyclic Strain-induced Endothelin-1 Expression in Human Umbilical Vein Endothelial Cells

This study was approved by the Research Ethics Committee of China Medical University (Taichung, Taiwan). Imubind ET-1 enzyme-linked immunosorbent assay kits were purchased from Amersham-Pharmacia (Amersham, United Kingdom). 2′,7′-dichlorofluorescin diacetate was obtained from Serva Co. (Heidelberg, Germany). Propofol was obtained from B. Braun (Melsungen, Germany), and stock solutions of up to 1 mm propofol in phosphate-buffered saline (PBS) were prepared by diluting 100 mm propofol (dissolved in 100% dimethylsulfoxide). All other chemicals of reagent grade were obtained from Sigma-Aldrich (St. Louis, MO).

Human umbilical vein endothelial cells (HUVECs) were obtained from PromoCell (Heidelberg, Germany) as cryopreserved cells. After thawing, the cells were plated into cultured flasks and cultured to confluence in endothelial cells medium (MCDB 131 medium, PromoCell) containing hydroxyethylpiperazine ethanesulfonic acid (28 mm), fetal calf serum (2%), human recombinant epidermal growth factor (0.1 ng/ml), human recombinant basic fibroblast growth factor (1.0 ng/ml), gentamycin (50 μg/ml), amphotericin B (50 ng/ml), and synthetic hydrocortisone (1.0 μg/ml), and were supplemented with supplement mix (PromoCell) containing endothelial cell growth factor and heparin. Cells were grown at 37°C in a humidified 5% CO²atmosphere for 3 to 4 days. Confluent cultures between passages 2 and 10 were used for all experiments. The cultured HUVECs were randomly assigned to one of the following groups: Group 1 for ET-1 mRNA analysis: HUVECs were either controls or treated with different concentrations of propofol (1–30 μm) in the absence or the presence of strain treatment for 6 h. Group 2 for ET-1 secretion analysis: HUVECs were treated with propofol (1–30 μm) in the absence or the presence of strain treatment for 24 h. Group 3 for ROS detection: HUVECs were treated with propofol (1–30 μm) in the absence or the presence of strain treatment for 1 h, or HUVECs were either controls or treated with cyclic strain or hydrogen peroxide (H²O², 25 μm) in the presence of propofol (30 μm), or antioxidant Trolox (200 μm) for 1 h. Group 4 for ERK phosphorylation analysis: HUVECs were treated with propofol (1–30 μm) in the absence or the presence of strain treatment for 30 min, or HUVECs were either controls or treated with cyclic strain or H²O²(25 μm) in the presence of propofol (30 μm), or Trolox (200 μm) for 30 min. Group 5 for nitric oxide measurement: HUVECs were in control condition; treated with strain alone for 10, 30, 60, or 120 min; or treated with strain in the presence of propofol (30 μm) for 10, 30, 60, or 120 min; or HUVECs were either controls or treated with cyclic strain in the absence or presence of propofol (1–30 μm) for 60 min. Group 6 for eNOS and protein kinase B (Akt) phosphorylation analysis: HUVECs were in control condition; treated with strain alone for 5, 15, 30, or 60 min; or treated with strain in the presence of propofol (30 μm) for 5, 15, 30, or 60 min.

Endothelial cells cultured on the flexible membrane base were subjected to cyclic strain produced by a computer-controlled application of sinusoidal negative pressure, as described previously.23 

Preparation of total RNA and Northern blot analyses of ET-1 and 18S RNA were performed as described previously.23 

ET-1 levels were measured in culture medium using a commercial enzyme-linked immunosorbent assay kit (Amersham-Pharmacia). Results were normalized to cellular protein content in all experiments, and expressed as a percentage relative to the cells incubated with the vehicle.

Measurement of intracellular ROS formation in HUVECs was recorded by monitoring changes in diclorofluorescein fluorescence as described previously.22 

Western blot analysis was performed as described previously.23Membranes were blocked with 5% nonfat dry milk (NFDM) in PBS-0.1% Tween for 90 min and washed 3 times with PBS-0.1% Tween-1% NFDM, and then incubated with a 1:500 dilution of antiphospho-ERK antibodies (New England BioLabs; Beverly, MA), anti-ERK antibodies (Santa Cruz Biotechnology; Santa Cruz, CA), anti-Ser1177 phospho-eNOS antibodies, anti-Ser473 phospho-Akt antibodies, anti-Akt antibodies (Cell Signaling Technology; Beverly, MA); or anti-eNOS antibodies (BD Bioscience; San Jose, CA) in PBS-1% NFDM for 1 h and washed 3 times with PBS-0.1% Tween-1% NFDM; and then incubated with a 1:4,000 dilution of the horseradish peroxidase-coupled appropriate secondary antibody (Pierce; Rockford, IL) in PBS-0.1% Tween-1% NFDM and washed 3 times with PBS-0.3% Tween-1% NFDM.

The culture medium was stored at −70°C until use. After the medium had been thawed, the sample was deproteined with 2 volumes of 4°C 99% ethanol and centrifuged (3 000 g for 10 min). These medium samples (100 μl) were injected into a collection chamber containing 5% vanadium trichloride. This strong reducing environment converts both nitrate and nitrite to nitric oxide. A constant stream of helium gas carried nitric oxide into a nitric oxide analyzer (Seivers 270B NOA; Seivers Instruments Inc.; Boulder, CO), where the nitric oxide was reacted with ozone, resulting in the emission of light. Light emission is proportional to the nitric oxide formed; standard amounts of nitrate were used for calibration.

To measure nitric oxide synthase (NOS) activity, L-arginine to L-citrulline conversion was assayed in cells with a NOS detection assay kit (Calbiochem; San Diego, CA) according to the manufacturer’s instructions. Briefly, cells were lysed with a buffer containing 1% NP-40, 0.5% sodium deoxycholate, and 0.1% sodium dodecyl sulfate. After incubation on ice for 30 min, cell extracts were centrifuged to remove cell debris. Protein extracts were then incubated for 60 min at 37°C in a solution of 10 μm L-[³H]arginine, 1 mm nicotinamide adenine dinucleotide phosphate, 1 μm flavin adenine dinucleotide, 1 μm flavin mononucleotide, 100 nm calmodulin, 600 μm CaCl², and 3 μm tetrahydrobiopterin in a final volume of 40 μl. The reaction was stopped by the addition of 400 μl of stop buffer (10 mm EDTA, 50 mm HEPES buffer, pH 5.5) to the reaction mixture. Then 100 μl of equilibrated resin was added to each mixture. Reaction samples were transferred to spin cups and centrifuged at 10,000 g for 30 s. The radioactivity of the flow-through was measured by liquid scintillation counting. Enzyme activity was expressed as citrulline production in pmol · min · mg^-1· protein⁻¹.

Data are presented as mean ± SEM. Statistical analysis was performed as appropriate using Student’s t  test or ANOVA, followed by a Dunnett multiple comparison test using Prism version 3.00 for Windows (GraphPad Software; San Diego, CA). P  values < 0.05 were considered to be statistically significant.

HUVECs cultured on flexible membrane bases were subjected to deformation to produce an average strain of 20%. We examined the effect of propofol on strain-increased ET-1 mRNA levels. HUVECs under cyclic strain for 6 h showed a significant increase in ET-1 mRNA levels (203.8 ± 31.2% over controls); however, treatment with propofol (3–30 μm) significantly reduced the strain-induced ET-1 expression (41.6 ± 13.5, 52.5 ± 12.5, and 56.6 ± 10.7% reduction with 3, 10, and 30 μm propofol, respectively) (fig. 1A). We then examined the effect of propofol on strain-increased ET-1 secretion. ET-1 released into the culture media was measured. As shown in figure 1B, HUVECs under cyclic strain for 24 h increased ET-1 secretion (112.3 ± 11.2% over controls), and also, propofol (3–30 μm) significantly inhibited strain-increased ET-1 secretion (34.8 ± 12.4, 45.8 ± 7.5, and 52.9 ± 8.7% reduction with 3, 10, and 30 μm propofol, respectively). These data indicate that propofol inhibits strain-increased ET-1 secretion in endothelial cells.

Our previous studies showed that cyclic strain increases ROS formation in endothelial cells, which is involved in ET-1 induction.22,23We next examined whether propofol prevents the strain-increased ROS formation. HUVECs were treated with propofol (1–30 μm) in the absence or presence of strain treatment. The addition of propofol (3–30 μm) to cultured HUVECs significantly inhibited strain-induced ROS formation, as measured after strain treatment for 1 h (fig. 2A). The treatment of propofol (10 μm) or antioxidant Trolox (200 μm) to cultured HUVECs also significantly inhibited strain- or H²O²-induced ROS formation (fig. 2B). These findings clearly demonstrate that propofol inhibits strain-increased intracellular ROS levels in endothelial cells.

To gain insight into the mechanism of action of propofol, we thus examined whether propofol affects intracellular protein kinase signaling pathways. Given that the ERK signaling pathway is involved in strain-induced ET-1 expression,22,23we further investigated whether propofol inhibits the ERK pathway in strain-treated endothelial cells. We examined the phosphorylation of ERK in HUVECs exposed to propofol (1–30 μm) in the absence or presence of strain treatment. As shown in figure 3, HUVEC exposure to strain treatment for 30 min rapidly activated phosphorylation of ERK. However, HUVECs treated with propofol (10 and 30 μm) showed significantly decreased strain-induced ERK phosphorylation (fig. 3A). Moreover, HUVECs treated with propofol (10 μm) or Trolox (200 μm) also showed significantly decreased strain- or H²O²-induced ERK phosphorylation (fig. 3B). These findings imply that propofol inhibits strain-activated ERK signaling pathways via  attenuation of ROS formation in endothelial cells.

To determine propofol’s mechanisms of action, NOS, NOS activity, phosphor-eNOS, and phospho-Akt (for serine 473 and threonine 308 isoforms) were detected. Exposure of HUVECs to propofol time- and dose-dependently enhanced the strain increase of nitric oxide generation (fig. 4, A and B). All three NOS isoforms characterized to date depend on calmodulin activation, but are distinguished from each other by their calcium sensitivity and enzymatic activity. A constitutive NOS activity was detected and quantified in whole extracts of cells, and this activity was dependent on the presence of calcium (fig. 4C). Exposure of HUVECs to propofol (10 and 30 μm) also enhanced the strain-increased NOS activity at 60 min after stimulation (fig. 4C). In addition, as shown in figure 5, HUVEC exposure to strain treatment activated phosphorylation of eNOS and Akt. Propofol treatment in HUVECs under cyclic strain significantly enhanced strain-increased phospho-eNOS (fig. 5A) and phospho-Akt (Ser 473 and Thr 308) (fig. 5, B and C). Exposure of HUVECs to propofol (10 and 30 μm) alone induced increase of nitric oxide generation, NOS activity, and eNOS phosphorylation. Overall, the magnitude of the effect of propofol on HUVECS not subjected to strain on the NO production was weaker, as compared with propofol increase in nitric oxide production in HUVECs under strain treatment (data not shown).

To examine whether propofol antagonizes strain-induced ET-1 release through nitric oxide production and protein kinase (PKG) pathways, we used PTIO(100 μm), a nitric oxide scavenger, and KT5823 (1 μm), a specific inhibitor of PKG, to determine the involvement of nitric oxide/PKG pathway in propofol modulation of strain-induced ERK phosphorylation and ET-1 release. In the presence of PTIO(100 μm), or KT5823 (1 μm), the inhibitory effect of propofol on strain-increased ERK phosphorylation and ET-1 release was reversed (fig. 6, A and B). In contrast, HUVECs exposed to cyclic strain in the presence of the nitric oxide donor S-Nitroso-N-acetylpenicillamine (100 μm) or cyclic guanosine monophosphate (cGMP) analogs, 8-bromoguanosine-3′,5′-cyclic monophosphate (100 μm), which activates PKG, showed significant inhibition of strain-stimulated ERK phosphorylation and ET-1 release. These results indicate the involvement of nitric oxide/PKG pathway in propofol modulation of strain-stimulated ET-1 release.

The major new finding of this work is that propofol inhibits strain-induced ET-1 secretion and enhances nitric oxide production in endothelial cells. It is supported by the observations that propofol inhibits strain-induced ET-1 expression in part via  the attenuation of ROS formation and the enhancement of nitric oxide production in endothelial cells under strain. Previous studies, including ours, have indicated that hemodynamic forces, including shear flow and pressure-induced strain,22–25can stimulate the ET-1 gene expression. Recent studies provide evidence that ROS may act as second messengers in cells exposed to various stimuli.22–24Previous studies from our collaborating laboratory and others have shown that cyclic-strain treatment of endothelial cells can induce intracellular ROS generation.21–23This ROS generation is sustained at an elevated level as long as the mechanical forces are maintained, and returns to basal levels with the removal of the forces.21Elevated ROS levels are involved in the release of ET-1,22,23and this gene induction can be attenuated by antioxidant pretreatment of cells.22,23,26,27 

Some of the plant-derived agents possess antioxidative effects,22,26–32and may have potential therapeutic benefit in clinical use. The plant-derived propofol is a potent intravenous hypnotic agent commonly administered for induction and maintenance of anesthesia, and for sedation in the intensive care unit.^1-3It is also a potent antioxidant.3,7,8The results of our present study further demonstrated that propofol reduced the strain-induced ROS generation in the endothelial cells. In particular, it has been demonstrated that activation of ERK is redox-sensitive,21–23and that suppression of ROS inhibits strain-induced ET-1 gene expression.22,23One possible explanation for the inhibitory effect of propofol on strain-induced ET-1 expression may thus be its ability to attenuate ROS formation. Alternatively, propofol may inhibit strain-induced ET-1 gene expression by virtue of its increasing nitric oxide production.5,7We found that propofol treatment, in the absence of cyclic strain, enhances the production and release of nitric oxide from HUVECs (data not shown). The magnitude of the effect of propofol on HUVECS not subjected to strain on the nitric oxide production was weaker, as compared with propofol increase in nitric oxide production in HUVECs under strain treatment. This is similar in nature to previous reports that application of propofol alone stimulates the production of nitric oxide from cultured vascular endothelial cells.33,34Our findings also indicate that propofol via  nitric oxide production suppresses the strain-stimulated ERK activation and ET-1 expression. Nitric oxide acting through soluble guanylyl cyclase and cGMP formation is a negative regulator of ET-1 gene induction.10,35–37Our findings further support that nitric oxide acts as a negative regulator in strain-induced ET-1 expression under propofol treatment. We also demonstrated that HUVEC exposure to strain treatment activated phosphorylation of eNOS and Akt. Nitric oxide synthesized by eNOS is regarded as an endothelial cell survival factor.38The serine-threonine kinase Akt activation has been reported to activate eNOS, which leads to nitric oxide production, promoting cell survival. The Akt/eNOS/nitric oxide pathway has been identified as an important survival pathway in endothelial cells.39,40Previously, Wang et al.  reported that propofol treatment, in the presence of H²O², significantly increased eNOS expression but not Akt phosphorylation.41In contrast, in the present study, we found that propofol treatment in HUVECs under cyclic strain significantly enhanced strain-increased phospho-eNOS and phospho-Akt. In addition, HUVECs exposed to cyclic strain in the presence of the nitric oxide donor S-Nitroso-N-acetylpenicillamine or cGMP analogs, 8-bromoguanosine-3, 5′-cyclic monophosphate, which activates PKG, showed significant inhibition of strain-stimulated ERK phosphorylation and ET-1 release. These results further suggest the involvement of nitric oxide/PKG pathways in propofol modulation of strain-stimulated ET-1 release.

In conclusion, the data obtained in the present study suggest that the propofol-induced nitric oxide production and suppression of cyclic strain-induced ET-1 expression can be considered as one of the mechanisms responsible for the protective effect of propofol in vascular vessels. These findings have highlighted the therapeutic potentials of using plant-derived propofol for the treatment of arteriosclerosis and hypertension. In light of our findings, propofol can be considered as a potentially active compound for use in conditions where ROS are implicated, although clinical studies are needed to establish the usefulness of this natural product in the treatment or prevention of human diseases.

The authors thank David Chiu-Yin Kwan, Ph.D. (Professor, Department of Medicine, Faculty of Health Sciences, McMaster University, Hamilton, Ontario, Canada), for his helpful comments during the preparation of this manuscript.