The vascular endothelial monolayer serves as a barrier between the bloodstream and the vascular wall. Apoptotic endothelial cell death may critically disturb the integrity of the endothelial monolayer and thereby contribute to vascular injury and atherosclerosis. Apoptosis has become increasingly recognized as a mechanism of cell death during myocardial ischemia reperfusion injury (IRI), although the relative contribution of necrosis and apoptosis to total cardiac cell loss during IRI remains controversial. Endothelial cell apoptosis has been shown to precede myocyte cell apoptosis in the setting of myocardial IRI (1). The latter study suggests that circulatory proapoptotic inflammatory cytokines (such as tumor necrosis factor [TNF]-α) and reactive oxygen species (ROS), that are increased during myocardial IRI and atherosclerosis, promote myocyte apoptosis subsequent to the induction of endothelial cell apoptosis.
Propofol, 2,6-diisopropylphenol, an IV anesthetic drug frequently used during cardiac surgery and in postoperative sedation (2), enhances red blood cell and tissue antioxidant capacity both in vitro and in vivo (3,4). We (5) recently demonstrated that propofol enhances myocardial antioxidant capacity and results in improved postischemic cardiac function in the isolated rat heart, in a dose-dependent manner. In addition, several studies show that aging enhances the sensitivity of human endothelial cells toward apoptotic stimuli (6). Interestingly, propofol, when applied at a clinically achievable large concentration, enhances the ischemic tolerance of middle-aged rat hearts (7). This finding prompted us to postulate that propofol may produce a cardioprotective effect that is attributable to its ability to enhance endothelial cell resistance toward apoptotic stimuli.
We hypothesized that propofol could inhibit TNF-α-induced human umbilical vein endothelial cells (HUVECs) apoptosis by resuming a proper ratio of the antiapoptotic Bcl-2 protein over the proapoptotic Bax protein expression and that the propofol antiapoptotic effect is related to its antioxidant capacity and its ability to enhance the generation of nitric oxide (NO), an important endothelial cell survival factor.
HUVECs were isolated according to the method of Jaffe et al. (8). Cells were cultured in Dulbecco’s modified Eagle medium (Gibco) supplemented with 20% bovine calf serum (Gibco), maintained at 37°C in 5% CO2, and used at passage 2–3 to avoid “age-dependent” variations in levels of apoptosis.
When the cells were at 70% confluence, the cultured HUVECs were divided into 7 groups: HUVECs in untreated group (control) and propofol treatment control (P25) group were further cultured at 37°C for 24 h, respectively, in the absence (control) or presence of 25 μM propofol (AstraZeneca China, Beijing) in the medium. TNF-α was not applied in the two control groups throughout the experiment. HUVECs in the TNF-α (TNF) group and TNF-α plus propofol treatment groups were initially cultured for 30 min in the presence of zero (TNF group), 12.5 (P12.5 + TNF group), 25 (P25 + TNF group), 50 (P50 + TNF group), and 100 (P100 + TNF group) μM propofol, respectively. The cells were then further cultured for 24 h with the addition of TNF at 40 ng/mL into the medium. The concentration of TNF used to induce apoptosis in the present study was chosen on the basis of previously published literature (9) in addition to preliminary studies. Experiments were repeated a minimum of eight times per group. Our preliminary study results indicated that intralipid, the solvent for propofol, at concentrations needed to dissolve up to 50 or 100 μM propofol did not have a significant effect on HUVEC apoptotic cell death. Therefore, we did not include intralipid groups in the ensuing studies.
Apoptosis was detected using DNA in situ terminal deoxynucleotidyl transferase (TdT)-mediated deoxyuridine triphosphate (dUTP)-biotin nick end-labeling (TUNEL) staining as per the manufacturer’s protocol (Boster Biotech, Wuhan, China). In brief, after equilibration, end-labeling with digoxigenin-11-dUTP by TdT enzyme in buffer was performed for 1 h at 37°C in a humidifying chamber. After treatment with stop/wash buffer, sections were incubated with anti-digoxigenin antibody-peroxidase conjugate, rinsed, and stained with diaminobenzidine tetrahydrochloride. Negative controls were incubated with phosphate buffered saline (PBS) instead of TdT enzyme, and positive controls were treated with DNase1. Sections were counterstained with Mayer’s hematoxylin and mounted. All experiments were repeated on at least six independent occasions with consistent results.
Cells were washed in Dulbecco’s modified Eagle medium without bovine serum albumin, and cytospins were performed (650 rpm for 6 min) on saline-coated slides at 1 × 106 cells/mL. Slides were fixed in 2% paraformaldehyde for 15 min at room temperature and washed 5 times in PBS. Cells were permeabilized for 10 min at room temperature in blocking buffer (3% bovine serum albumin in PBS) plus 0.1% Triton X-100 followed by blocking of nonspecific binding in blocking buffer for 1 h at room temperature. They were then incubated with the primary antibodies (anti-Bcl-2 1:50 dilution, and, anti-Bax 1:50 dilution) (Boster Biotech). After overnight incubation at 4°C, cells were washed with PBS and incubated for 30 min at room temperature with the biotinylated secondary antibody. After washing, the cells were incubated for 30 min in Vectastain Elite ABC reagent (Boster Biotech). After another wash, cells were incubated in peroxidase substrate solution until the desired stain intensity was developed. The cells were then washed, dehydrated in increasing concentrations of ethanol, and cover-slipped using permount. Control sections were either incubated with the secondary antibody alone or immunoabsorbed with an excess of blocking peptide. Random fields (20–30 per slide) were examined at a high magnification (×400) to calculate the prevalence of DNA fragmentation and Bcl-2/Bax expression. The percentage of TUNEL-positive cells (termed apoptotic index, AI) was determined by dividing the number of positive-staining nuclei by the total number of nuclei of the cell and multiplying that value by 100. The percentage of Bcl-2/Bax expression was also determined. The density of Bcl-2 and Bax protein expression was determined using an automatic computer-assisted image analyzing system (IBAS-2000; Kontron, Germany), which automatically measures the density of 100 HUVECs from 4 to 6 random fields. The density of Bcl-2 and Bax protein expression was expressed in arbitrary units.
Electron microscopy was performed to confirm that the ultrastructural features of apoptosis were present in cells exposed to TNF. Endothelial cells exposed to the conditions outlined above were fixed in 2.5% glutaraldehyde (pH 7.3) buffered with 0.1 mol/L sodium cacodylate overnight at 4°C and then washed with 0.1 mol/L sodium cacodylate buffer for 15 min before postfixation with 1% osmium tetroxide buffered with 0.1 mol/L sodium cacodylate for 1 h on ice. After another wash with 0.1 mol/L sodium cacodylate buffer for 15 min, cells were dehydrated with increasing concentrations of alcohol. Next, cells were infiltrated with propylene oxide for 15 min, followed by 1:1 propylene oxide/epoxy resin for 1 h, 1:2 propylene oxide/epoxy resin for 2 h, and finally 100% epoxy resin for 2 h. Cells were embedded with fresh epoxy resin into molds and placed in a 60°C oven for 2 h. Ultrathin sections were stained with uranyl acetate and lead citrate and were examined with the use of a Hitachi H-600 electron microscope (Hitachi, Japan).
Media of the cultured endothelial cells were collected 24 h after their respective treatments. The concentration of nitrites (NO2 −) and nitrates (NO3 −), stable end products of NO, was determined by the Griess reaction, as follows. After deproteination by a solution of zinc sulfate, samples were incubated with cadmium granules to reduce nitrate to nitrite. The total nitrite was measured at 540-nm absorbance by diazotization with Griess reagent (Boshide Biotech Ltd., Wuhan). Nitrite concentrations were calculated by comparison with a standard calibration curve with sodium nitrite.
Results were expressed as mean ± sd. Data were tested for normal distribution by the Kolmogorov-Smirnov test. Significance was evaluated using analysis of variance followed by Tukey’s post hoc test. The correlation relationships were evaluated by the Pearson test. P < 0.05 was considered significant.
TUNEL staining was rare in control (3.1% ± 0.5%) and propofol (P25)-treated HUVECs (3.0% ± 0.6%) (Fig. 1, A and B). Stimulation of HUVECs with TNF resulted in a dramatic increase in the AI to 45.5% ± 1.2% (Fig. 1, C and E). Propofol dose-dependently reduced TNF-induced apoptosis. More profound reduction in AI was seen in P25 (35.0% ± 0.7%; Fig. 1, D and E) and P50 (25.2% ± 0.8%). P100 did not significantly further decrease AI (22.6% ± 0.5%) compared with that of P50.
As shown in Figures 2 and 3, stimulation of HUVECs with TNF leads to a significant reduction in Bcl-2 protein expression and a significant increase in Bax protein expression as compared with untreated controls. Propofol (25 μM) did not affect either Bcl-2 or Bax protein expression in the absence of TNF stimulation. However, propofol, at ≥12.5 μM, significantly and dose-dependently attenuated TNF-induced reduction in Bcl-2 protein expression (Fig. 2E). The maximal effect was seen at P100. The Bcl-2 density in the P100 + TNF group was significantly higher than that in the P50 + TNF group. Propofol also significantly attenuated the TNF-induced increase in Bax protein expression in a dose-dependent manner at the range 12.5–50 μM. There was no difference in effect between P50 and P100. TNF reduced the ratio of Bcl-2 expression over Bax expression (Bcl-2/Bax) as compared with control (Fig. 4A). Propofol dose-dependently attenuated the TNF-induced reduction in Bcl-2/Bax ratio (Fig. 4A).
As shown in Figure 5, stimulation of HUVECs with TNF led to significantly increased production of NO as compared with untreated control. Interestingly, addition of propofol (at 25 μM) to the culture medium also resulted in a significantly increased production of NO in the absence of a noxious stimulus (such as TNF). P12.5 and P25, in a dose-dependent manner, further increased TNF-induced release of NO. However, P50 and P100 did not result in a more profound increase in HUVECs’ NO release in the presence of TNF, as compared with P25.
By qualitative electron microscopic analysis, typical features of apoptosis could hardly be seen in HUVECs from control and P25 groups (Fig. 6), but they were apparent in HUVECs from the TNF group (Fig. 6C). Apoptotic morphologic changes were also seen in HUVECs from TNF plus propofol treatment groups (Fig. 6D shows a typical endothelial cell from P25 + T group), but to a much lesser degree in terms of severity.
A tight inverse correlation existed between the ratio of Bcl-2/Bax protein expression and AI in untreated control HUVECs, and TNF-treated HUVECs with or without concomitant administration of propofol (r = −0.9520, P = 0.0009, Fig. 4B). There was no relation between NO production and AI in HUVECs from control and the propofol (25 μM) treated groups. This is because propofol increased NO production without affecting the AI in the absence of TNF stimulation. A trend of inverse relation between NO production and AI was seen in HUVECs stimulated with TNF alone or TNF with varying concentrations of propofol, but this was not statistically significant (r = −0.85, P = 0.07). However, there was a weak but significant positive correlation between NO production and the rate of Bcl-2/Bax protein expression (r = 0.93, P = 0.02) in HUVECs stimulated with TNF and without propofol treatments.
HUVECs are an abundant and easily accessible endothelial cell type. Study indicates that HUVECs and human coronary microvascular endothelial cells have similar sensitivities to the harmful effects of inflammatory cytokines (10), including TNF and oxidative damage. Hence, HUVECs are used as a tool in exploring the mechanisms involved in the pathogenesis of cardiovascular diseases.
TNF stimulation resulted in a reduced Bcl-2/Bax ratio in HUVECs. Propofol dose-dependently enhanced the ratio of the antiapoptotic Bcl-2 protein over the proapoptotic Bax protein expression in this system. This was associated with graded suppression of TNF-induced apoptosis as assessed by TUNEL assay and confirmed by characteristic apoptotic morphologic changes. At a dose range from 12.5 to 50 μM, propofol enhancement of Bcl-2/Bax ratio was achieved through an increase in expression of Bcl-2 and a decrease in the expression of Bax. However, this effect seems dose-limited because the largest concentration of propofol (P100) did not significantly reduce Bax expression more than P50. However, P100 did increase Bcl-2 expression more than P50. This could be explained on the basis that P100 completely abolished TNF-induced increases in Bax expression while maintaining baseline levels (Fig 3E). This is likely an important mechanism of protection. Similar to other important molecules, notably NO, Bax has a dual role. Under pathological conditions, Bax over-expression may induce mitochondrial depolarization and cytochrome c release (11), resulting in the downstream activation of executioner caspases (12) to augment apoptosis. The formation Bax-Bax homodimer serves to induce apoptosis, whereas the Bax-Bcl-2 heterodimer formed under physiological conditions is an important inhibitor of apoptosis (13).
It is noteworthy that propofol treatment primarily restores the expression of Bcl-2 and the Bcl-2/Bax ratio toward normal values and does not result in an over-expression of the antiapoptotic Bcl-2 protein. This effect is not what we expected, but may represent a promising therapeutic approach. Bcl-2 is localized to intracellular sites of ROS generation including mitochondria and may function in an antioxidant pathway to prevent apoptosis (14). After an apoptotic signal, cells sustain progressive lipid peroxidation. Over-expression of Bcl-2 functions to suppress lipid peroxidation (14). However, in lymphocytes, the level of Bcl-2 expression may determine the balance between apoptosis and necrosis, but does not prevent cell death induced by oxidized low density lipoproteins (oxLDLs) (15). In cells expressing relatively high levels of Bcl-2, oxLDLs induced mainly necrosis. In cells expressing relatively low levels of Bcl-2, the rate of oxLDL-induced apoptosis was more rapid than that of primary necrosis (15). One study demonstrated that over-expression of Bcl-2 paradoxically exerted a proapoptotic effect in the reperfused liver (16). This study, together with ours, suggest that stabilizing Bcl-2 and Bax expression, rather than induction of the antiapoptotic Bcl-2 and/or suppression of the proapoptotic Bax protein, represents a more meaningful approach.
Propofol (P25) treatment, in the absence of TNF, enhances the production and release of NO from HUVECs. This is similar in nature to a previous report that application of propofol stimulates the production of NO from cultured porcine aortic endothelial cells (17). Interestingly, TNF enhances the production of NO to a similar degree as propofol (Fig. 5), but, in contrast, this is accompanied by an increase in apoptosis in HUVECs. It seems plausible that the TNF-induced NO overproduction relative to control levels (in the absence of antioxidant intervention) would result in increased production of peroxynitrite (ONOO−) which may promote apoptosis by increasing Bax expression (18). Coculture of HUVECs with TNF and propofol led to profound overproduction of NO compared with TNF or propofol alone. This was associated with reduced Bax expression and enhanced Bcl-2 production compared with the TNF group. This suggests that overproduction of NO by endothelial cells in response to TNF stimulation is initially intended to protect the cells, rather than produce cell injury. This is manifested by the significant positive correlation between NO production and the rate of Bcl-2/Bax protein expression in HUVECs stimulated with TNF, with and without propofol in increasing concentrations. Indeed, one study showed that NO confers resistance to apoptosis in HUVECs (19), likely secondary to the attenuation of caspase activity by nitrosylating caspase 3 and stabilizing Bcl-2. The question arises: How does propofol confer its protective effect on endothelial cells?
TNF stimulates up-regulation of NO synthase activity and NO production in HUVECs which can be accompanied by a burst in production (three- to fourfold increase) of intracellular ROS including superoxide anion and H2O2 (20). The reaction of NO with superoxide anion can increase the generation of ONOO−. Exposure of endothelial NO synthase to oxidants (including ONOO−) causes increased enzymatic uncoupling and the generation of superoxide anion rather than NO (21), resulting in increased oxidant stress and a net decrease in NO production. Propofol can directly scavenge ROS including ONOO− (22), therefore blocking the vicious cycle. Our recent finding that the addition of H2O2 exaggerates TNF-induced HUVEC apoptosis (Luo and Xia, unpublished data) supports the interplay between ROS and TNF.
The release of cytochrome c from mitochondria into the cytosol is considered an important event leading to irreversible cell death and the opening of mitochondrial transition pore (MPTP) which may be an important mechanism for mitochondrial cytochrome c release (23). Inhibition of MPTP reduced TNF-induced release of mitochondrial cytochrome c in HUVECs (24). Furthermore, it has been suggested that propofol may offer myocardial protection by inhibiting MPTP, at concentrations as small as 11–22 μM (2–4 μg/mL), likely acting indirectly through its antioxidant properties because ROS is a major mediator of MPTP opening (25). Propofol, however, may directly inhibit myocyte MPTP at concentrations ≥50 μM (26). Interestingly, the most profound inhibition of TNF-induced HUVEC apoptosis was manifested at propofol concentrations ≥50 μM (P50 and P100) in this study, suggesting inhibition of MPTP in HUVECs as one of the mechanisms of propofol’s protection.
Our study provides evidence for the first time that propofol, a frequently used anesthetic for cardiac surgery and for postoperative sedation, attenuates TNF-induced HUVEC apoptosis in a dose-dependent manner. Clinically relevant blood concentrations of propofol include 0.8–1.0 μg/mL on awakening from propofol anesthesia, 1–2 μg/mL on long-term sedation in the intensive care unit, at least 2.5 μg/mL for satisfactory hypnosis, and 3–11 μg/mL (approximately 17–62 μM) for maintenance of satisfactory anesthesia (27). Because the release of inflammatory cytokines (including TNF) is increased during and after cardiac surgery and because serum from patients with acute coronary syndromes displays a proapoptotic effect on endothelial cells (28), application of propofol, in large concentrations (approximately 50–60 μM) (3), may prove to be a promising approach in reducing peri- and postoperative IRI and related cardiovascular symptoms.
1. Scarabelli T, Stephanou A, Rayment N, et al. Apoptosis of endothelial cells precedes myocyte cell apoptosis in ischemia/reperfusion injury. Circulation 2001;104:253–6.
2. Bryson HM, Fulton BR, Faulds D. Propofol: an update of its use in anaesthesia and conscious sedation. Drugs 1995;50:513–59.
3. Ansley DM, Sun J, Visser WA, et al. High dose propofol enhances red cell antioxidant capacity during CPB in humans. Can J Anaesth 1999;46:641–8.
4. Runzer TD, Ansley DM, Godin DV, et al. Tissue antioxidant capacity during anesthesia: propofol enhances in vivo red cell and tissue antioxidant capacity in a rat model. Anesth Analg 2002;94:89–93.
5. Xia Z, Godin DV, Chang TK, et al. Dose-dependent protection of cardiac function by propofol during ischemia and early reperfusion in rats: effects on 15-F2t-isoprostane formation. Can J Physiol Pharmacol 2003;81:14–21.
6. Hoffmann J, Haendeler J, Aicher A, et al. Aging enhances the sensitivity of endothelial cells toward apoptotic stimuli: important role of nitric oxide. Circ Res 2001;89:709–15.
7. Xia Z, Godin DV, Ansley DM. Propofol enhances ischemic tolerance of middle-aged rat hearts: effects on 15-F(2t)-isoprostane formation and tissue antioxidant capacity. Cardiovasc Res 2003;59:113–21.
8. Jaffe EA, Nachman RL, Becker CG, et al. Synthesis of antihemophilic factor antigen by cultured human endothelial cells. J Clin Invest 1973;52:2745–56.
9. Polunovsky VA, Wendt CH, Ingbar DH, et al. Induction of endothelial cell apoptosis by TNFα: modulation by inhibitors of protein synthesis. Exp Cell Res 1994;214:584–94.
10. Willam C, Koehne P, Jurgensen JS, et al. Tie2 receptor expression is stimulated by hypoxia and proinflammatory cytokines in human endothelial cells. Circ Res 2000;87:370–7.
11. Jurgensmeier JM, Xie Z, Deveraux Q, et al. Bax directly induces release of cytochrome c from isolated mitochondria. Proc Natl Acad Sci USA 1998;95:4997–5002.
12. Granville DJ, Shaw JR, Leong S, et al. Release of cytochrome c, Bax migration, Bid cleavage, and activation of caspases 2, 3, 6, 7, 8, and 9 during endothelial cell apoptosis. Am J Pathol 1999;155:1021–5.
13. Krajewski S, Krajewska M, Shabaik A, et al. Immunohistochemical determination of in vivo distribution of Bax, a dominant inhibitor of Bcl-2. Am J Pathol 1994;145:1323–36.
14. Hockenbery DM, Oltvai ZN, Yin XM, et al. Bcl-2 functions in an antioxidant pathway to prevent apoptosis. Cell 1993;75:241–51.
15. Meilhac O, Escargueil-Blanc I, Thiers JC, et al. Bcl-2 alters the balance between apoptosis and necrosis, but does not prevent cell death induced by oxidized low density lipoproteins. FASEB J 1999;13:485–94.
16. Oshiro T, Shiraishi M, Muto Y. Adenovirus mediated gene transfer of antiapoptotic protein in hepatic ischemia-reperfusion injury: the paradoxical effect of Bcl-2 expression in the reperfused liver. J Surg Res 2002;103:30–6.
17. Petros AJ, Bogle RG, Pearson JD. Propofol stimulates nitric oxide release from cultured porcine aortic endothelial cells. Br J Pharmacol 1993;109:6–7.
18. Arstall MA, Sawyer DB, Fukazawa R, et al. Cytokine-mediated apoptosis in cardiac myocytes: the role of inducible nitric oxide synthase induction and peroxynitrite generation. Circ Res 1999;85:829–40.
19. Ho FM, Liu SH, Liau CS, et al. Nitric oxide prevents apoptosis of human endothelial cells from high glucose exposure during early stage. Cell Biochem 1999;75:258–63.
20. Deshpande SS, Angkeow P, Huang J, et al. Rac1 inhibits TNF-alpha-induced endothelial cell apoptosis: dual regulation by reactive oxygen species. FASEB J 2000;14:1705–14.
21. Zou MH, Shi C, Cohen RA. Oxidation of the zinc-thiolate complex and uncoupling of endothelial nitric oxide synthase by peroxynitrite. J Clin Invest 2002;109:817–26.
22. Kahraman S, Demiryurek AT. Propofol is a peroxynitrite scavenger. Anesth Analg 1997;84:1127–9.
23. Green DR, Reed JC. Mitochondria and apoptosis. Science 1998;281:1309–12.
24. Walter DH, Haendeler J, Galle J, et al. Cyclosporin A inhibits apoptosis of human endothelial cells by preventing release of cytochrome C from mitochondria. Circulation 1998;98:1153–7.
25. Sztark F, Ichas F, Ouhabi R, et al. Effects of the anaesthetic propofol on the calcium-induced permeability transition of rat heart mitochondria: direct pore inhibition and shift of the gating potential. FEBS Lett 1995;368:101–4.
26. Kowaltowski AJ, Castilho RF, Vercesi AE. Mitochondrial permeability transition and oxidative stress. FEBS Lett 2001;495:12–5.
27. Short TG, Aun CS, Tan P, et al. A prospective evaluation of pharmacokinetic model controlled infusion of propofol in paediatric patients. Br J Anaesth 1994;72:302–6.
28. Valgimigli M, Agnoletti L, Curello S, et al. Serum from patients with acute coronary syndromes displays a proapoptotic effect on human endothelial cells: a possible link to pan-coronary syndromes. Circulation 2003;107:264–70.