Depolarization of Endothelial Cells Enhances Platelet Aggregation Through Oxidative Inactivation of Endothelial NTPDase
Objective— The objective of this study was to investigate whether depolarization of cultured endothelial cells (human umbilical vein endothelial cells, HUVECs) affects their ectonucleotidase activity through superoxide (O2−) production.
Methods and Results— Depolarization by the cation channel gramicidin (100 nmol/L) or tetrabutylammonium chloride (1 mmol/L) induced O2− release from HUVECs (n=4), which was decreased by superoxide dismutase (SOD, 500 U/mL). The activity of endothelial ectonucleotidases was assessed by the production of inorganic phosphate from ADP, which was decreased by chronic depolarization by 25% (n=6, P<0.05) and the amount of AMP derived from ADP in the presence of the 5′-nucleotidase inhibitor α,β-methylene-5′-diphosphate (100 μmol/L). AMP was decreased by chronic depolarization from 0.54±0.16 to 0.39±0.11 μmol/min/mg protein (n=6, P<0.05). This was abolished in the continuous presence of SOD (n=6). NTPDase protein was predominantly expressed in HUVECs (n=4). Protein abundance, viability of cells, and apoptosis rates were not altered by depolarization (n=10). Only in the presence of depolarized HUVECs, but not with control cells or with HUVECs depolarized in the presence of SOD, did 5 μmol/L of ADP cause irreversible platelet aggregation. Increases in transmural pressure induced endothelial depolarization in intact hamster small arterioles.
Conclusions— Depolarization causes the endothelial production of O2−, which inhibits the activity of endothelial ectonucleotidases. Increases in transmural pressure induce endothelial depolarization. In chronically hypertensive diseases, depolarization might favor platelet aggregation.
Platelet adhesion initiates a series of events leading to firm thrombus formation,1,2⇓ for instance, release of platelet granule contents.3 Among the substances released, ADP is a major effector for recruitment of further platelets.4,5⇓ Accordingly, the metabolism of ADP, which co-controls local ADP concentrations, is of substantial importance for primary hemostasis and the center of recent interest in attempts to therapeutically target thrombosis.6–8⇓⇓ In vitro, platelet aggregation induced by various stimuli is effectively inhibited by potato apyrase, an ectonucleotidase (ecto-NTPDase) that catalyzes ADP hydrolysis into free phosphate and AMP.9 Endothelial cells are known to express various families of ecto-NTPDases, among which the E-NTPDases (ecto-nucleoside triphosphate diphosphohydrolases or ecto-apyrases) have gained attention because of their substantial contribution to the inhibition of platelet aggregation.6,7,10⇓⇓ They can be classified according to their preferential substrates.9 CD39 (NTPDase 1) has been recognized as the major NTPDase present in endothelial cells.11 It exhibits the highest ADP:ATP substrate ratio in favor of ADP and is therefore a focus of antiplatelet research.12,13⇓ Indeed, soluble CD39 effectively abolishes platelet aggregation in response to various agonists6 and reduces the cerebral infarct volume that is increased in CD39 −/− mice.8
Endothelial NTPDases have been suggested to be inactivated by exposure to oxidative stress in vitro14 and during ischemia/reperfusion injury in rat glomeruli in vivo.15 Recently, we have reported that depolarization of human umbilical vein endothelial cells (HUVECs) leads to the activation of endothelial NAD(P)H-oxidase, resulting in enhanced O2− release.16 This might be of importance in cardiovascular disease because hypertensive,17 diabetic,18 and injured19 vessels are thought to be chronically depolarized. In isolated vessels, increases in transmural pressure induce depolarization of smooth muscle cells20; however, whether pressure-induced depolarization also occurs in the endothelium is not yet clear. In this study, we investigated whether O2− release induced by endothelial cell membrane depolarization is sufficient to inactivate endothelial NTPDase activity, thereby leading to enhanced ADP-dependent platelet aggregation. In isolated resistance arteries, we further studied whether an increase in transmural pressure leads to endothelial depolarization.
For a detailed methods section, please see the online version of this article, which can be accessed at http://atvb.ahajournals.org.
Endothelial Cell Culture and Platelet Preparation
Human umbilical vein endothelial cells (HUVECs) were cultured as previously described.21 Platelet-rich plasma (PRP) was obtained from citrate-anticoagulated blood as previously described.22 Platelet counts were obtained with a resistance particle counter (Coulter Z2).
Measurements of Superoxide 0(O2−) and of HUVEC Membrane Potential
O2− was measured using the fluorescent dye 2′,7′-dichlorodihydrofluorescein diacetate. Cells were preincubated with the NO-synthase inhibitor N-nitro-l-arginine (l-NA; 50 μmol/L), washed three times, and incubated with 10 μmol/L 2′,7′-dichlorodihydrofluorescein diacetate for 15 minutes. In separate experiments, membrane potential was measured using the fluorescent dye bis-[1,3-dibutylbarbituric acid] trimethineoxonol (DiBac43) as described previously.16 Fluorescence intensities were recorded on a confocal microscope (Zeiss LSM 410).
Platelet aggregation was measured turbidimetrically in PRP adjusted to 200,000 platelets/μL as previously described.22 In additional experiments, platelets (2×108/mL) were stimulated with ADP or control substances in the presence of endothelial cells (105/mL) and aggregation measured turbidimetrically as described by Marcus et al.10
Preparation of Supernatants for Ecto-NTPDase Activity Measurements
HUVECs were washed 3 times in phosphate-free modified tyrode buffer and incubated with 100 μmol/L ADP for 15 minutes at 37°C. Experiments were performed in the presence of APCP (100 μmol/L) to avoid additional cleavage of inorganic phosphate (Pi) from the AMP formed by degradation of ADP. The supernatants were then collected and stored at −20°C.
Measurement of Free Phosphate and of Nucleotides
Pi in supernatants was measured by the malachite green assay described by Baykov et al.23 AMP and ADP were measured by high-pressure liquid chromatography as described22 with an EC 250/4 nucleosil carbohydrate column. Retention time for AMP was 4.5 minutes and for ADP 10.5 minutes.
Assessment of Cell Viability and Apoptosis
A trypan blue exclusion assay was performed to assess cell viability. Cells able to exclude the dye were assumed viable. In additional experiments LDH release was assessed using a CytoTox 96® assay (Promega). Apoptosis was determined flow cytometrically using an annexin-V apoptosis detection kit (BD Pharmingen).
HUVECs were washed once in PBS and lysed in %1 buffer Triton-X 100. Protein content was assessed, and equal amounts were separated by SDS-PAGE by using standard techniques of blotting and chemiluminescent visualization as previously described.22
Measurement of Endothelial Membrane Potential Changes in Small- Resistance Arteries
Animal care and the conduct of the experiments were in strict accordance with the German animal protection laws. Preparation of vessels from the gracilis muscle of Golden Syrian hamsters was previously described.24 Cannulated small-resistance arteries were analyzed on a confocal microscope. The endothelium was loaded from the luminal side with the voltage-sensitive dye di-8-ANEPPS as described elsewhere.25 After subtraction of the background signal, the ratio of the two signals (>570 nm/525 to 565 nm) was calculated.24
DiBac43 and di-8-ANEPPS were from Molecular Probes (Netherlands). Polyclonal rabbit anti-NTPDase antibody (anti-CD39) was a kind gift of Prof. Adrien Beaudoin (Canada). All other substances were obtained from Sigma Chemicals Co., Germany.
All data are expressed as means±SEM. Data were analyzed using one-way ANOVA or Student’s t test for paired or unpaired data. Differences were considered significant when the error probability level was P<0.05.
Membrane Potential in Cultured HUVECs
In a previous work we measured the resting membrane potential of HUVEC by DiBAC43, and it amounted to −47±2 mV.16 Depolarization using gramicidin (100 nmol/L) increased basal DiBac43 fluorescence by 39±21% (n=4, P<0.05, Figure 1B), corresponding to membrane potential values of about −34±2 mV. The potassium channel blocker tetrabutylammonium chloride (TBA) (1 mmol/L) increased fluorescence by 28±5% (n=4, P<0.01).
Chronic depolarization of HUVECs for 24 hours (gramicidin, 100 nmol/L) increased O2− production (in the presence of l-NA) by 54±17% over control cells (P>0.01). This increase was prevented by superoxide dismutase (SOD, U/mL) whereas boiled SOD had no effect (Figure 1A and online Figure I, which can be accessed at http://atvb.ahajournals.org, n=10). SOD (500 U/mL) plus catalase (1000 U/mL) had no additional effect. O2− production was completely abolished by the diphenyleneiodoniumchloride (30 μmol/L; P<0.01), an unspecific inhibitor of NAD(P)H-oxidase.
The Pi concentration [Pi] in supernatants of HUVECs incubated with 100 μmol/L ADP in the presence of 5′-nucleotidase inhibitor α,β-methylene-5′-diphosphate (APCP, 100 μmol/L) for 15 minutes was 77.8±22.8 μmol/L (n=6). In the supernatants of HUVEC that had been depolarized for long periods by gramicidin (100 nmol/L), [Pi] was decreased by 25.1±6.8% (P<0.05), similar as after depolarization using TBA (1 mmol/L), which decreased it by 29.7±11.4% (P<0.05, n=5). [Pi] in supernatants immediately after depolarization was not altered. [Pi] in supernatants of gramicidin-treated cells in the presence of SOD was increased by 33.6±23.2% compared with gramicidin alone (P<0.05 versus gramicidin) and by 38.8±20.4% with gramicidin in the presence of SOD and catalase (P<0.01 versus gramicidin, n=6, all data Figure 2A), which was not significantly different from depolarization in the presence of SOD only. When depolarization was induced in the presence of catalase only, [Pi] was not different from gramicidin alone (not shown).
Endothelial ectonucleotidase activity was further assessed by the transformation of ADP to AMP in the presence of the 5′-nucleotidase inhibitor APCP (100 μmol/L). The AMP concentration in supernatants was reduced by chronic depolarization for 24 hours from 0.54±0.16 to 0.39±0.11 μmol AMP/min/mg protein. This effect was abolished when SOD alone (Figure 3A, n=6; P<0.05) or SOD with catalase were present (not shown). NTPDase activity calculated on the basis of ADP metabolized per minute and per amount of protein behaved similarly: ADP degradation was inhibited for gramicidin-treated cells but not in the presence of SOD or SOD with catalase (Figure 3B, n=6).
When 4 μL of supernatants of depolarized cells (100 μL/100 μmol/L ADP) were used as a stimulus, aggregation reached 68.4±6% of the aggregation achieved by 4 μmol/L of ADP (set as maximal aggregation). This was significantly more than the response caused by supernatants of nondepolarized cells (52.9±4%, P<0.05, n=6). Depolarization-induced loss of ADP degradation was prevented when SOD was present. These supernatants caused 53.5±5% of maximum aggregation (P<0.05, n=6), similar as control supernatants (Figure III, which can be accessed at http://atvb.ahajournals.org).
Aggregation experiments were also performed in mixed suspensions containing HUVECs and washed platelets. In the presence of HUVECs, ADP in concentrations that otherwise induced irreversible aggregation (5 μmol/L) caused transient aggregation only. This effect was lost when HUVECs had been chronically depolarized but was again observed when depolarization was performed with SOD (n=3). Such differences were not found when other platelet stimuli, such as collagen, thrombin receptor-activating peptide (TRAP), or epinephrine, were used as stimuli (n=2, Figure 4).
Depolarization did not result in any detectable changes in cell viability assessed by a trypan blue exclusion assay. After 24 hours, 99.1±0.8% of control cells were viable, similar as after treatment with gramicidin (99.4±0.4%) or TBA (99.1±0.5%, n=9, NS). Likewise, there were no differences in LDH release between depolarized and control cells (see online Figure IIA, which can be accessed at http://atvb.ahajournals.org). Furthermore, there was no difference in the relative amount of apoptotic cells because 9.8±1.1% of nonnecrotic cells chronically treated with gramicidin showed apoptosis as compared with 9.3±1.1% of control cells (n=16, NS, Figure IIB).
NTPDase Protein Expression
Western blots performed using a polyclonal NTPDase antibody (raised against porcine pancreatic NTPDase and provided by Prof. A. Beaudoin) did not reveal an effect of depolarization on the expression of this protein by HUVECs (Figure 5A). Densitometric analysis of 3 experiments showed equal expression of the protein under the conditions tested, that is control, gramicidin (100 nmol/L), gramicidin with SOD (500 U/mL), and gramicidin with SOD and catalase (1000 U/mL, n=3 each, see online Figure IV, which can be accessed at http://atvb.ahajournals.org).
Comparison of protein expression levels in HUVECs, PRP, and platelet-poor plasma (PPP) showed that HUVECs contained the highest levels of NTPDase protein, followed by platelets. There were no signals when PPP was used as substrate (Figure 5B).
Increases in Transmural Pressure Depolarize the Endothelium of Isolated Arteries
Perfusion of isolated arterioles with di-8-ANEPPS selectively loaded the endothelium as demonstrated by confocal microscopy (Figure 6B). High potassium buffer (125 mmol/L, corrected for osmolarity by reducing sodium) induced a depolarization that shifted emission leftwards and increased the ratio of the dual-wavelength recording (Figure 6A, upper trace). Washout of the high potassium buffer brought the ratio back to baseline within a few minutes. Increasing transmural pressure from 45 mm Hg to 100 mm Hg also resulted in a rise of the ratio (Figure 6A, middle trace), which returned to baseline after renewed decrease of transmural pressure. This was not caused by a change in focal plane because vessels that were already depolarized by high potassium buffer (absence of calcium) did not show an increase in fluorescence ratio after an additional increase in transmural pressure (Figure 6C, lower trace, n=4 each).
ADP is the major mediator of platelet recruitment after platelet activation. We and others have shown that after collagen stimulation of platelets, ADP is released in large amounts.22 Because ecto-NTPDases rapidly hydrolyze ADP, they are considered to be of major significance in platelet recruitment.8,26⇓ We have recently observed that the activity of platelet ecto-NTPDase(s) might be oxidatively influenced.22 However, in quantitative terms, the ecto-NTPDase activity of endothelial cells seems to be the dominant contributor to the rate of ADP degradation in the vasculature.26 In comparison with platelets, endothelial cells express higher amounts of one specific ecto-NTPDase, CD39,27 which is thought to be the ecto-NTPDase predominantly responsible for vascular nucleotide metabolism and, therefore, inhibition of platelet activation.9–11⇓⇓ It was recently demonstrated in mouse microvessels that the endothelium mainly contains NTPDase 1 (CD39),12 whereas NTPDase 2 (CD39L1), a preferential NTPDase that should rather increase ADP concentrations and, consequently, platelet aggregation, is expressed in the adventitia. The activity of CD39 has been observed to be lost after endothelial cell activation by tumor necrosis factor-α, an effect that was assigned to oxidative inactivation of the enzyme rather than to a change in protein content.14 Bakker and colleagues28 early observed a loss of rat glomerular NTPDase activity on infusion of the redox cycling compound doxorubicin, which was not caused by cytotoxicity and a reduction of inflammation-induced loss of rat kidney NTPDase activity on addition of SOD and catalase.29 Because thiol oxidation and disulfide bond formation are established biological effects of oxidants,30 the formation of CD39 oligomers mediated by disulfide bonds represents a possible molecular target for oxidative modulation.31,32⇓ Although others have suggested that exposure to oxidants decreases CD39 protein in CD39-transfected COS cells,11 we could not observe a change in NTPDase protein expression caused by depolarization-induced endothelial reactive oxygen species (ROS) production or by depolarization itself in HUVECs.
Although we cannot ascribe our findings to endothelial CD39, an oxidative inactivation of which is discussed controversially, they support the concept of NTPDases as targets of redox regulation, a regulatory mechanism, which can also be found in NTPDases of other species.33,34⇓ Therefore, we cannot exclude that ectonucleotidases other than CD39, for example, N-PPases (nucleotidepyrophosphatases) were also affected. However, because the reaction catalyzed by these enzymes seems to predominantly result in the hydrolysis of 2 moles of Pi,9,35⇓ their involvement seems improbable because of the complementary behavior of AMP and Pi (parallel increases) in our study. A significant influence of alkaline phosphatase is unlikely as a result of the physiological pH conditions prevailing in our experiments, and 5′-nucleotidase was blocked throughout. Hence, the enzymatic activity investigated in this study is most likely that of CD39.
Inactivation of endothelial NTPDase activity was assessed by two independent methods: AMP formation and Pi production from ADP. Because endothelial 5′-nucleotidase normally further degrades AMP to adenosine, thereby releasing another Pi,9,36⇓ it was necessary to perform the experiments in the presence of APCP, which, at the concentration used (100 μmol/L), completely inhibits endothelial 5′-nucleotidase activity.36 Under more physiological conditions, when 5′-nucleotidase is active, the generation of adenosine would even supply another antiaggregatory agent.36 This should yet increase the proaggregatory effects of an inhibition of NTPDase. Pertinently, there was no significant hydrolysis of ADP in the absence of endothelial cells, which themselves did not release relevant amounts of ADP.
The depolarization-induced effects were reversed completely by addition of SOD, and this was not enhanced by catalase, which alone did also not influence the rate of formation of Pi. This indicates that O2− is the predominant ROS involved in inactivation of endothelial ecto-NTPDases. Cellular viability or apoptosis rates, which are both known to increase cellular O2− release,30,37⇓ were not altered by depolarization. Furthermore, platelet reactivity to other stimuli, such as collagen, epinephrine, or TRAP, was not affected by depolarization. In two different bioassays, we demonstrated that endothelial NTPDase activity is lost on chronic depolarization and indeed results in enhanced platelet aggregation. In addition, comparison of expression of NTPDase in HUVECs, PPP, or PRP showed the highest protein expression in HUVEC, little in PRP, and none in PPP. This confirmed previous observations27 and made confounding contributions of NTPDases from other sources unlikely.
By directly measuring membrane potential changes in intact resistance vessels selectively loaded with a membrane-potential sensitive, ratiometric dye from their luminal side, we, for the first time, demonstrate that increases in transmural pressure depolarize the endothelium. Because chronic depolarization of vascular cells or attenuated endothelial hyperpolarization have been observed in spontaneously hypertensive rats,17 in endothelial injury after balloon angioplasty,19 and in diabetic patients,18 chronic or repeated depolarization on elevated pressure is likely a mechanism of endothelial cell activation in vivo. The mechanisms underlying such endothelial depolarization remain unknown. Although there is a possibility of direct ion channel activation,38 a mechanism more likely to occur in vivo is electrical coupling through myoendothelial gap junctions that conduct electrical changes from vascular smooth muscle to endothelial cells.39,40⇓ Evidence for a role of ROS in depolarization-induced deterioration of endothelial function comes from the observation that delivery of SOD decreases blood pressure in spontaneously hypertensive rats (SHR),41 and by increased O2− production in internal mammary arteries and saphenous veins from diabetic patients.42 We have previously observed that depolarization of HUVECs was associated with enhanced NAD(P)H-oxidase-dependent O2− production.16 As shown here, this production is sustained for 24 hours and is sufficient to inactivate endothelial ecto-NTPDases. Two substances causing depolarization by independent mechanisms induced endothelial O2− production and inactivated NTPDases. Thus, it is unlikely that the effects of gramicidin, which was used in most experiments, are caused by effects on cellular ion concentration and pH that have been reported.43
We conclude that oxidative inactivation of endothelial ecto-NTPDases as a result of chronic depolarization can result in altered ADP-dependent platelet recruitment. Such depolarization might occur in hypertensive vessels because increases in transmural pressure induce endothelial depolarization in isolated hamster arterioles. As platelet aggregation initiated by contact with ruptured endothelium depends on the release of autoactivating substances like ADP, the membrane potential and consequently the NTPDase-activity of the surrounding endothelial cells might pivotally influence further growth of the local thrombus. Moreover, in light of the observation that certain fatty acids might increase endothelial NTPDase activity,44 a potential novel role for an endothelium-derived hyperpolarizing factor, which is widely assumed to be an epoxyeicosatrienoic acid,45,46⇓ could be postulated. Chronic vessel depolarization and the release of substances, such as endothelium-derived hyperpolarizing factor influencing endothelial membrane potential, might therefore be involved in the regulation of thrombus formation in vivo.
This work was supported by a grant from the Friedrich-Baur-Foundation of the Ludwigs-Maximilians-University. The authors thank D. Goessel and D. Kiesl for excellent technical assistance.
↵*These authors contributed equally to this work.
Received September 4, 2002; revision accepted October 1, 2002.
- ↵Siess W. Molecular mechanisms of platelet activation. Physiol Rev. 1989; 69: 58–178.
- ↵Valles J, Santos MT, Aznar J, Martinez M, Moscardo A, Pinon M, Broekman MJ, Marcus AJ. Platelet-erythrocyte interactions enhance alpha(IIb)beta(3) integrin receptor activation and P-selectin expression during platelet recruitment: down-regulation by aspirin ex vivo. Blood. 2002; 99: 3978–3984.
- ↵Santos MT, Valles J, Marcus AJ, Safier LB, Broekman MJ, Islam N, Ullman HL, Eiroa AM, Aznar J. Enhancement of platelet reactivity and modulation of eicosanoid production by intact erythrocytes: a new approach to platelet activation and recruitment. J Clin Invest. 1991; 87: 571–580.
- ↵Enjyoji K, Sevigny J, Lin Y, Frenette PS, Christie PD, Esch JS, Imai M, Edelberg JM, Rayburn H, Lech M, Beeler DL, Csizmadia E, Wagner DD, Robson SC, Rosenberg RD. Targeted disruption of cd39/ATP diphosphohydrolase results in disordered hemostasis and thromboregulation. Nat Med. 1999; 5: 1010–1017.
- ↵Pinsky DJ, Broekman MJ, Peschon JJ, Stocking KL, Fujita T, Ramasamy R, Connolly ES Jr, Huang J, Kiss S, Zhang Y, Choudhri TF, McTaggart RA, Liao H, Drosopoulos JH, Price VL, Marcus AJ, Maliszewski CR. Elucidation of the thromboregulatory role of CD39/ectoapyrase in the ischemic brain. J Clin Invest. 2002; 109: 1031–1040.
- ↵Kaczmarek E, Koziak K, Sevigny J, Siegel JB, Anrather J, Beaudoin AR, Bach FH, Robson SC. Identification and characterization of CD39/vascular ATP diphosphohydrolase. J Biol Chem. 1996; 271: 33116–33122.
- ↵Sevigny J, Sundberg C, Braun N, Guckelberger O, Csizmadia E, Qawi I, Imai M, Zimmermann H, Robson SC. Differential catalytic properties and vascular topography of murine nucleoside triphosphate diphosphohydrolase 1 (NTPDase1) and NTPDase2 have implications for thromboregulation. Blood. 2002; 99: 2801–2809.
- ↵Robson SC, Kaczmarek E, Siegel JB, Candinas D, Koziak K, Millan M, Hancock WW, Bach FH. Loss of ATP diphosphohydrolase activity with endothelial cell activation. J Exp Med. 1997; 185: 153–163.
- ↵Sohn HY, Keller M, Gloe T, Morawietz H, Rueckschloss U, Pohl U. The small G-protein Rac mediates depolarization-induced superoxide formation in human endothelial cells. J Biol Chem. 2000; 275: 18745–18750.
- ↵Wellman GC, Cartin L, Eckman DM, Stevenson AS, Saundry CM, Lederer WJ, Nelson MT. Membrane depolarization, elevated Ca2+ entry, and gene expression in cerebral arteries of hypertensive rats. Am J Physiol Heart Circ Physiol. 2001; 281: H2559–H2567.
- ↵Kohler R, Brakemeier S, Kuhn M, Behrens C, Real R, Degenhardt C, Orzechowski HD, Pries AR, Paul M, Hoyer J. Impaired hyperpolarization in regenerated endothelium after balloon catheter injury. Circ Res. 2001; 89: 174–179.
- ↵Harder DR. Pressure-induced myogenic activation of cat cerebral arteries is dependent on intact endothelium. Circ Res. 1987; 60: 102–107.
- ↵Gloe T, Riedmayr S, Sohn HY, Pohl U. The 67-kDa laminin-binding protein is involved in shear stress-dependent endothelial nitric-oxide synthase expression. J Biol Chem. 1999; 274: 15996–16002.
- ↵Krotz F, Sohn HY, Gloe T, Zahler S, Riexinger T, Schiele TM, Becker BF, Theisen K, Klauss V, Pohl U. NAD(P)H oxidase-dependent platelet superoxide anion release increases platelet recruitment. Blood. 2002; 100: 917–924.
- ↵Wolin MS. Interactions of oxidants with vascular signaling systems. Arterioscler Thromb Vasc Biol. 2000; 20: 1430–1442.
- ↵Asai T, Miura S, Sibley LD, Okabayashi H, Takeuchi T. Biochemical and molecular characterization of nucleoside triphosphate hydrolase isozymes from the parasitic protozoan Toxoplasma gondii. J Biol Chem. 1995; 270: 11391–11397.
- ↵Stommel EW, Cho E, Steide JA, Seguin R, Barchowsky A, Schwartzman JD, Kasper LH. Identification and role of thiols in Toxoplasma gondii egress. Exp Biol Med (Maywood). 2001; 226: 229–236.
- ↵Kawashima Y, Nagasawa T, Ninomiya H. Contribution of ecto-5′-nucleotidase to the inhibition of platelet aggregation by human endothelial cells. Blood. 2000; 96: 2157–2162.
- ↵Sobey CG. Potassium channel function in vascular disease. Arterioscler Thromb Vasc Biol. 2001; 21: 28–38.
- ↵Schnackenberg CG, Welch WJ, Wilcox CS. Normalization of blood pressure and renal vascular resistance in SHR with a membrane-permeable superoxide dismutase mimetic: role of nitric oxide. Hypertension. 1998; 32: 59–64.
- ↵Guzik TJ, Mussa S, Gastaldi D, Sadowski J, Ratnatunga C, Pillai R, Channon KM. Mechanisms of increased vascular superoxide production in human diabetes mellitus: role of NAD(P)H oxidase and endothelial nitric oxide synthase. Circulation. 2002; 105: 1656–1662.
- ↵Campbell WB, Gebremedhin D, Pratt PF, Harder DR. Identification of epoxyeicosatrienoic acids as endothelium-derived hyperpolarizing factors. Circ Res. 1996; 78: 415–423.
- ↵Bolz SS, Fisslthaler B, Pieperhoff S, De Wit C, Fleming I, Busse R, Pohl U. Antisense oligonucleotides against cytochrome P450 2C8 attenuate EDHF-mediated Ca(2+) changes and dilation in isolated resistance arteries. FASEB J. 2000; 14: 255–260.