Biochemical and Biophysical Research Communications 269, 713–717 (2000) doi:10.1006/bbrc.2000.2354, available online at http://www.idealibrary.com on
Endothelin-1 Induces NAD(P)H Oxidase in Human Endothelial Cells Nicole Duerrschmidt, Nico Wippich, Winfried Goettsch, Hans-Juergen Broemme, and Henning Morawietz 1 Institute of Pathophysiology, Martin Luther University Halle-Wittenberg, Magdeburger Strasse 18, D-06097 Halle, Germany
Received February 21, 2000
Superoxide anions (O 2•ⴚ) induce oxidative stress and reduce endothelial NO availability by peroxynitrite formation. In human endothelial cells gp91 phox was identified as the limiting subunit of the forming NAD(P)H oxidase. Because endothelin-1 (ET-1) is considered as a pro-atherosclerotic stimulus, we analyzed the effect of ET-1 on gp91 phox expression and O 2•ⴚ generation in primary cultures of human umbilical vein endothelial cells (HUVECs). The gp91 phox mRNA expression was quantified by standard calibrated competitive reverse transcriptase–polymerase chain reaction. ET-1 induces gp91 phox mRNA expression in HUVEC (max. after 1 h). The induction of gp91 phox expression was dose-dependent, reaching its maximum at 10 nmol/L ET-1. The increased gp91 phox expression is mediated by endothelial receptor type B (ET B). Furthermore, ET-1 augments O 2•ⴚ generation in human endothelial cells as measured by coelenterazine chemiluminescence. These data support a new mechanism: how ET-1 increases oxidative stress in the vessel wall leading to endothelial dysfunction and enhanced susceptibility to atherosclerosis. © 2000 Academic Press
Key Words: endothelin-1; NAD(P)H oxidase; superoxide anion; endothelial cells; atherosclerosis.
Elevated endothelin-1 (ET-1) plasma levels have been shown in a variety of pathophysiological conditions, including atherosclerosis, hypertension, and diabetes (1). The potent vasoconstrictor ET-1 (2) binds to endothelin type A (ET A) and type B (ET B) receptors (3). ET A receptors are present on vascular smooth muscle cells and induce ET-1 mediated vasoconstriction (4). Abbreviations used: CL, chemiluminescence; ET-1, endothelin-1; ET B, endothelin receptor type B; HUVECs, human umbilical vein endothelial cells; RT-PCR, reverse transcriptase–polymerase chain reaction; SOD, superoxide dismutase. 1 To whom correspondence should be addressed. Fax: ⫹49-3455571404. E-mail:
[email protected].
Endothelial cells express only the ET B receptor (5, 6). This endothelial receptor mediates vasodilation and is sensitive to the ET B selective antagonist BQ-788. Superoxide anion (O 2•⫺) generation has been implicated in vascular dysfunction seen in atherosclerosis, hypertension, diabetes and chronic nitrate tolerance (7–10). O 2•⫺ induces oxidative stress and reduces endothelial NO availability by peroxynitrite formation (11). Furthermore, the O 2•⫺ generation is modulated by superoxide dismutases (SODs), the enzyme family that catalyze the formation of hydrogen peroxide and molecular oxygen from O 2•⫺. The major source of endothelial O 2•⫺ generation is the NAD(P)H oxidase (12, 13). The NAD(P)H oxidase consists of 2 membrane-bound subunits (gp91 phox, p22 phox) and 2 cytosolic subunits (p47 phox, p67 phox). In human endothelial cells gp91 phox was identified as the limiting subunit of the O 2•⫺ forming NAD(P)H oxidase (14). The effect of the vasoactive and potential proatherogenic stimulus ET-1 on endothelial superoxide anion generation is currently unknown. Therefore we analyzed the effect of ET-1 on mRNA expression of the limiting NAD(P)H oxidase subunit gp91 phox by competitive RT-PCR and on O 2•⫺ generation by coelenterazineenhanced chemiluminescence (CL) in primary cultures of human umbilical vein endothelial cells (HUVECs). MATERIALS AND METHODS Cell culture. Cell culture reagents and chemicals were purchased from Sigma Chemicals Corp. except when otherwise specified. Primary cultures of human umbilical vein endothelial cells (HUVECs) were isolated with collagenase IV as described previously (15). In order to minimize variations of primary cultures, each day the isolated endothelial cells were pooled and subsequently separated in medium M199 with 1.25 g/L sodium bicarbonate, 100 mg/L L-glutamine (Life Technologies), supplemented with 20% calf serum, 15 mmol/L HEPES, 100.000 U/L penicillin, 100 mg/L streptomycin, 250 mg/L fungizone (Life Technologies), and 16.7 g/L endothelial cell growth supplement (C. C. Pro, Neustadt, Germany). Confluent cell cultures were incubated with medium containing 0.5% calf serum for 24 h and subsequently treated with ET-1 (1 nmol/L to 100
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nmol/L) or with ET-1 (10 nmol/L) and the endothelin receptor type B antagonist BQ-788 (1 mol/L, Alexis). RNA isolation and competitive RT-PCR. Total RNA from HUVECs was isolated by guanidinium thiocyanate/cesium chloride centrifugation (16). The mRNA expression of NAD(P)H oxidase subunit gp91 phox was quantified by standard calibrated competitive reverse transcriptase-polymerase chain reaction (RT-PCR) by use of a linker primer, PCR-generated, internal-deleted, and in vitro-transcribed gp91 phox standard cRNA. In brief, a human gp91 phox specific cDNA fragment (17) of 404 bp was amplified from RNA of human endothelial cells by RT-PCR. The gp91 phox specific cDNA fragment was subsequently cloned into the pCR-Script Amp SK(⫹) Cloning Vector (Stratagene) and its identity confirmed by DNA sequencing (ABI PRISM Dye Terminator Cycle Sequencing Ready Reaction Kit with AmpliTaq DNA Polymerase, FS, Perkin-Elmer-Corp.; ABI 373 DNA Sequencer). DNA sequence was analyzed using Gene Runner software (Hastings Software, Inc.). Database searches of GenBank were performed using BLASTN (18). Subsequently an internal-deleted cRNA standard of 304 bp was constructed by linker primer PCR. The identity of standard was confirmed by cloning and DNA sequencing. The internal-deleted gp91 phox cDNA standard was in vitro transcribed into cRNA (RNA Transcription Kit, Stratagene), and standard cRNA was quantified spectrophotometrically. In competitive RT-PCR experiments equal amounts of total RNA (200 ng) were incubated in separate reactions with defined amounts of gp91 phox standard cRNA for 3 min at 70°C, and subsequently reverse transcribed into cDNA using random hexamer primers and SuperScript II RNase H ⫺ reverse transcriptase (Life Technologies) for 30 min at 42°C. Afterwards, 25% of each reverse transcription reaction were amplified in separate reactions with gp91 phox sense and antisense PCR primers. PCR primers compete for sample-specific and standard molecules in the amplification reaction. The PCR reactions were separated by standard agarose gel electrophoresis, stained with ethidium bromide and documented by photography using Polaroid film type 665. The optical density of standard and sample specific PCR fragment was estimated by a Personal Densitometer (Molecular Dynamics, Sunnyvale, CA). Optical density of standard PCR fragments was normalized with a correction coefficient (gp91 phox: 404 bp/304 bp ⫽ 1.329), and logarithm of the quotient of normalized standard and sample specific PCR fragment density was graphically plotted vs. amount of standard RNA molecules, using the SigmaPlot scientific graphing software (Jandel Corp.). In the graph, equal amounts of RNA molecules in sample and standard were present at equivalence point. Xanthine oxidase activity measurements. The reaction mixture for the determination of the xanthine oxidase/xanthine (XO/X)derived chemiluminescence (CL) consists of 960 L of Tris-HCl buffer (20 mmol/L, pH 7.4), 10 L of XO from buttermilk (5 munits/mL mixture), 10 L of lucigenin (1 mmol/L in Tris-HCl buffer) or 10 L of coelenterazine (Molecular Probes Inc., Eugene, OR, 1 mmol/L in 96% v/v ethanol, stored under nitrogen at ⫺20°C) respectively. The superoxide anion generating reaction was initiated by addition of 20 L of xanthine (10 mmol/L in 0.01 mmol/L NaOH). Lucigenin or coelenterazine CL was recorded at 25°C for 5 min using a Lumat LB 9501 chemiluminometer (EG & G Berthold, Wildbad, Germany). Addition of superoxide dismutase (SOD) from bovine erythrocytes (40 g/mL) immediately stops lucigenin CL as well as coelenterazine CL. The XO/X triggered lucigenin and coelenterazine CL was expressed as relative light units per second (rLU 䡠 s ⫺1) (mean ⫾ SEM of 5 parallel experiments). NAD(P)H oxidase activity measurements. HUVECs were seeded in Pyrex tubes (Dunn Corp., Asbach, Germany). Confluent HUVEC cultures were incubated in medium M199 containing 0.5% calf serum for 24 h, stimulated with ET-1 and subsequently, after buffer exchange, used directly for CL measurements in the presence of coelenterazine. Immediately before starting CL measurements HUVECs were washed with PBS (10 mmol/L, pH 7.4, 37°C). Subse-
FIG. 1. Determination of gp91 phox mRNA expression by competitive RT-PCR in human endothelial cells. The method compares the amplification of a gp91 phox-specific cDNA fragment from reversetranscribed total RNA of HUVECs (size: 404 bp) vs different concentrations of an internal-deleted and reverse-transcribed cRNA standard (size: 304 bp) by PCR. The PCR fragments were separated on agarose gels and stained with ethidium bromide.
quently, Pyrex tubes with adherent cells were refilled with 970 L PBS-buffer and placed in a LUMAT LB 9501 chemiluminometer. CL measurement was initiated by addition of 10 L coelenterazine (1 mmol/L in 96% ethanol). Three minutes after starting CL recording, 10 L NADPH (10 mmol/L in PBS) or 10 L SOD (4 mg/mL PBS), respectively, were added to the mixture and CL registration was continued for additional 5 min. In experiments with diphenyliodonium chloride (500 mol/L, ICN Biomedical GmbH, Eschwege, Germany), HUVECs were preincubated with this flavin-containing enzyme inhibitor 90 min before CL measurements. All experiments were carried out at 37°C. The cellular protein concentration was determined with BCA Protein Assay Reagent (Pierce Corp., Rockford, IL). The coelenterazine CL was expressed as relative light units per minute and per g protein (rLU 䡠 min ⫺1 䡠 g protein ⫺1). Statistics. Data are shown as mean ⫾ SEM. Statistical analysis was performed with the ANOVA procedure followed by Bonferroni’s method (multiple comparison) or Student t test (SigmaStat software, Jandel Corp). Differences were taken as statistically significant at P ⬍ 0.05.
RESULTS AND DISCUSSION ET-1 Induces NAD(P)H Oxidase Subunit gp91 phox mRNA in Human Endothelial Cells In human endothelial cells the O2•⫺ forming NAD(P)H oxidase subunit gp91 phox essential for the electron transfer from NAD(P)H to oxygen was identified as the limiting subunit (14). The mRNA expression of the NAD(P)H oxidase subunit gp91 phox was quantified by standard calibrated competitive RT-PCR (Fig. 1). ET-1 induces gp91 phox mRNA expression in HUVEC. Maximal gp91 phox induction was found after 1 h (10 nmol/L ET-1, 163% ⫾ 21 of control, n ⫽ 6, P ⬍ 0.05) (Fig. 2). The increased gp91 phox mRNA expression was transient and returned to baseline after 2 h. The induction of gp91 phox expression was dosedependent, reaching its maximum at 10 nmol/L ET-1. The ET-1-induced gp91 phox mRNA expression is mediated by the endothelial receptor type B (ET B). The ET B-specific inhibitor BQ-788 had no effect on basal gp91 phox mRNA expression (105% ⫾ 13 of control, n ⫽ 3), but prevented ET-1-mediated upregulation of gp91 phox mRNA expression (97% ⫾ 7 of control, n ⫽ 3, P ⬍ 0.05 vs. ET-1) (Fig. 3). These data suggest that the potent vasoconstrictor ET-1 can augment the
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FIG. 2. Time course of gp91 phox mRNA expression in response to ET-1 in HUVECs. HUVECs were incubated in medium containing 0.5% calf serum with 10 nmol/L ET-1 for the indicated times. Total RNA was isolated and gp91 phox mRNA expression determined in equal amounts of RNA by standard calibrated competitive RT-PCR. Values are given as means ⫾ SEM in percentage of control; n ⱖ 3 each.
NAD(P)H oxidase expression in human endothelial cells. To further validate these findings on the functional level, we measured the effect of ET-1 on NAD(P)H oxidase activity in human endothelial cells. Augmented NAD(P)H Oxidase Activity by ET-1 in Human Endothelial Cells Lucigenin can undergo a cycle of univalent reduction followed by autooxidation and thus has the capacity to
FIG. 3. Dose-dependent expression of gp91 phox mRNA in response to ET-1 in HUVECs. The upregulation of gp91 phox mRNA can be inhibited by the ET B-specific inhibitor BQ-788. HUVECs were incubated in medium containing 0.5% calf serum with ET-1 (1–100 nmol/L), BQ-788 (1 mol/L), or ET-1 (100 nmol/L) and BQ-788 for 1 h. Total RNA was isolated and gp91 phox mRNA expression determined in equal amounts of RNA by standard calibrated competitive RT-PCR. Values are given as means ⫾ SEM in percentage of control; n ⱖ 3 each; *P ⬍ 0.05 vs con, #P ⬍ 0.05 vs ET-1 (100 nmol/L).
FIG. 4. (A) Comparison of sensitivity of equimolar concentration (10 mol/L) of lucigenin and coelenterazine for detection of superoxide anion formation using the XO/X reaction as a model system. The XO/X-generated CL can be inhibited by SOD. (B) The XO/X-mediated superoxide anion formation measured by coelenterazine CL can be inhibited by the flavin-containing enzyme inhibitor diphenyliodonium chloride.
mediate the production of superoxide anions. Consequently, lucigenin can not be used as a reliable indicator for the presence of O 2•⫺ (19, 20). Recently, coelenterazine was introduced as a useful and sensitive chemiluminescent probe for the measurement of generation of O 2•⫺ in neutrophils (21). Furthermore, it was shown that coelenterazine in contrast to lucigenin does not enhance superoxide formation (22). The sensitivity of both chemiluminescent probes for detection of superoxide anion formation was compared using the XO/X reaction as a model system (Fig. 4). As seen from data presented in Fig. 4A, the sensitivity of coelenterazine for O 2•⫺ detection is considerably higher compared with an equimolar concentration of lucigenin. The immediate inhibition of CL by SOD indicates that the source of the detected CL were actually superoxide anions. Furthermore, the coelenterazine-mediated CL can be completely inhibited by the flavincontaining enzyme inhibitor diphenyliodonium chloride in this model system (Fig. 4B). Human endothelial cells are able to produce O 2•⫺ (12, 13, 23–26). We adapted the coelenterazine CL method
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FIG. 5. ET-1 increases superoxide anion formation in HUVECs. HUVECs were stimulated in medium containing 0.5% serum with ET-1 (10 nmol/L; 90 min). Superoxide anion formation was determined by coelenterazine-enhanced CL (relative light units 䡠 min ⫺1 䡠 g protein ⫺1, n ⫽ 9, P ⬍ 0.05).
for detection of O 2•⫺ generated by human umbilical vein endothelial cells (HUVECs). Coelenterazine CL was significantly augmented above control levels after ET-1 stimulation of HUVECs (max. after 90 min: 235 ⫾ 44 rLU 䡠 min ⫺1 䡠 g protein ⫺1 vs. con: 170 ⫾ 27 rLU 䡠 min ⫺1 䡠 g protein ⫺1, n ⫽ 9, P ⬍ 0.05) (Fig. 5). The NAD(P)H oxidase responsible for O 2•⫺ formation has been suggested to be mainly localized in the plasma membrane, but also microsomal localization of NAD(P)H oxidase has been discussed (27). It is generally accepted that nucleotides does not cross the cell membrane (28, 29). Addition of NADPH caused only a minor stimulation of coelenterazine CL in HUVECs, which did not reach the level of statistical significance (not shown). The absence of a stimulation of coelenterazine CL by addition of NADPH could be interpreted as an indication for integrity of plasma membrane of cultured human endothelial cells. Since coelenterazine as a lipophilic compound is able to penetrate the cell membrane, our data support the view that the detected coelenterazine CL in both controls and ET-1 stimulated HUVECs is mainly mediated by intracellular sources of NADPH and O 2•⫺. Furthermore, SOD and diphenyliodonium chloride had only a minor inhibitory effect on coelenterazine CL mediated by intact HUVECs (not shown). These data further support an intracellular source of O 2•⫺ formation, because SOD can not penetrate the cell membrane due to its size. We assume that diphenyliodonium chloride due to its polar properties cannot enter the cell in concentrations mediating complete inhibition of O 2•⫺ formation during the incubation period. These data suggest a NAD(P)H oxidase as main source of ET-1-stimulated endothelial O 2•⫺ formation, but additional O 2•⫺ generating enzymes might be involved as well. Endothelin has been implicated in the development and progression of cardiovascular diseases such as ath-
erosclerosis, hypertension, coronary heart disease, or heart failure (1, 30). One pathophysiological mechanism explaining the role of ET-1 in these diseases could be the ET-1-mediated augmented endothelial superoxide anion formation described in this study. Augmented superoxide anion formation could lead to increased peroxynitrite formation reducing the endothelial NO availability, or increased oxidative modification of low-density lipoproteins (LDL). In addition, the increased vascular superoxide anion production in nitrate tolerance can be partially normalized by the nonselective ET-1 receptor blocker bosentan (31). In summary our data are the first evidence that ET-1 induces oxidative stress in human endothelial cells. These findings suggest a new pathophysiological mechanism how elevated ET-1 levels mediate development and progression of endothelial dysfunction and cardiovascular diseases. ACKNOWLEDGMENTS We are grateful to J. Holtz for continuous support and R. Gall and R. Busath for excellent technical assistance. This study was supported by the Novartis Foundation for Therapeutic Research.
REFERENCES 1. Miyauchi, T., and Masaki, T. (1999) Annu. Rev. Physiol. 61, 391– 415. 2. Yanagisawa, M., Kurihara, H., Kimura, S., Tomobe, Y., Kobayashi, M., Mitsui, Y., Yazaki, Y., Goto, K., and Masaki, T. (1988) Nature 332, 411– 415. 3. Huggins, J. P., Pelton, J. T., and Miller, R. C. (1993) Pharmacol. Ther. 59, 55–123. 4. Hosoda, K., Nakao, K., Hiroshi, A., Suga, S., Ogawa, Y., Mukoyama, M., Shirakami, G., Saito, Y., Nakanishi, S., and Imura, H. (1991) FEBS Lett. 287, 23–26. 5. Ogawa, Y., Nakao, K., Arai, H., Nakagawa, O., Hosoda, K., Suga, S., Nakanishi, S., and Imura, H. (1991) Biochem. Biophys. Res. Commun. 178, 248 –255. 6. Heinroth-Hoffmann, I., Vogelsang, M., Schiewe, P., Morawietz, H., Holtz, J., Ponicke, K., and Brodde, O. E. (1998) Br. J. Pharmacol. 125, 1202–1211. 7. White, C. R., Brock, T. A., Chang, L. Y., Crapo, J., Briscoe, P., Ku, D., Bradley, W. A., Gianturco, S. H., Gore, J., Freeman, B. A., and Tarpey, M. M. (1994) Proc. Natl. Acad. Sci. USA 91, 1044 –1048. 8. Nakazono, K., Watanabe, N., Matsuno, K., Sasaki, J., Sato, T., and Inoue, M. (1991) Proc. Natl. Acad. Sci. USA 88, 10045–10048. 9. Mullarkey, C. J., Edelstein, D., and Brownlee, M. (1990) Biochem. Biophys. Res. Commun. 173, 932–939. 10. Munzel, T., Sayegh, H., Freeman, B. A., Tarpey, M. M., and Harrison, D. G. (1995) J. Clin. Invest. 95, 187–194. 11. Rubanyi, G. M., and Vanhoutte, P. M. (1986) Am. J. Physiol. 250, H822– 827. 12. Mohazzab, K. M., Kaminski, P. M., and Wolin, M. S. (1994) Am. J. Physiol. 266, H2568 –2572. 13. Jones, S. A., O’Donnell, V. B., Wood, J. D., Broughton, J. P., Hughes, E. J., and Jones, O. T. (1996) Am. J. Physiol. 271, H1626 –H1634. 14. Rueckschloss, U., Wippich, N., Broemme, H.-J., Holtz, J., and Morawietz, H. (1998) Circulation 98, I734 –735.
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15. Morawietz, H., Rueckschloss, U., Niemann, B., Duerrschmidt, N., Galle, J., Hakim, K., Zerkowski, H. R., Sawamura, T., and Holtz, J. (1999) Circulation 100, 899 –902. 16. Chirgwin, J. M., Przybyla, A. E., MacDonald, R. J., and Rutter, W. J. (1979) Biochemistry 18, 5294 –5299. 17. Royer-Pokora, B., Kunkel, L. M., Monaco, A. P., Goff, S. C., Newburger, P. E., Baehner, R. L., Cole, F. S., Curnutte, J. T., and Orkin, S. H. (1986) Nature 322, 32–38. 18. Altschul, S. F., Gish, W., Miller, W., Myers, E. W., and Lipman, D. J. (1990) J. Mol. Biol. 215, 403– 410. 19. Liochev, S. I., and Fridovich, I. (1997) Arch. Biochem. Biophys. 337, 115–120. 20. Vasquez-Vivar, J., Hogg, N., Pritchard, K. A., Jr., Martasek, P., and Kalyanaraman, B. (1997) FEBS Lett. 403, 127–130. 21. Lucas, M., and Solano, F. (1992) Anal. Biochem. 206, 273–277. 22. Tarpey, M. M., White, C. R., Suarez, E., Richardson, G., Radi, R., and Freeman, B. A. (1999) Circ. Res. 84, 1203–1211.
23. Matsubara, T., and Ziff, M. (1986) J. Cell Physiol. 127, 207–210. 24. Matsubara, T., and Ziff, M. (1986) J. Immunol. 137, 3295–3298. 25. Zweier, J. L., Kuppusamy, P., Thompson-Gorman, S., Klunk, D., and Lutty, G. A. (1994) Am. J. Physiol. 266, C700 –708. 26. Zulueta, J. J., Yu, F. S., Hertig, I. A., Thannickal, V. J., and Hassoun, P. M. (1995) Am. J. Respir. Cell Mol. Biol. 12, 41– 49. 27. Mohazzab, K. M., and Wolin, M. S. (1994) Am. J. Physiol. 267, L823– 831. 28. Bishop, C., Rankine, D. M., and Talbott, J. H. (1959) J. Biol. Chem. 234, 1233–1237. 29. Liersch, M., Groteluschen, H., and Decker, K. (1971) Hoppe Seylers Z. Physiol. Chem. 352, 267–274. 30. Schiffrin, E. L., and Touyz, R. M. (1998) J. Cardiovasc. Pharmacol. 32, S2–13. 31. Kurz, S., Hink, U., Nickenig, G., Borthayre, A. B., Harrison, D. G., and Munzel, T. (1999) Circulation 99, 3181–3187.
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