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Keywords:

  • antioxidants;
  • endothelium;
  • gene regulation;
  • inflammation;
  • reactive oxygen species

Abstract

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References

Plasminogen activator inhibitor type 1 (PAI-1) is induced by many proinflammatory and pro-oxidant factors. Among them, tumor necrosis factor α (TNFα), a pivotal early mediator that regulates and amplifies the development of inflammation, is one of the strongest PAI-1 synthesis activators. Location of the TNFα response element in the PAI-1 promoter is still ambiguous. In this study, we attempted to evaluate the significance of the element located in the 4G/5G site of the PAI-1 promoter in the TNFα stimulation of PAI-1 expression in endothelial cells. PAI-1 expression was monitored at: (a) the level of mRNA using real-time PCR, (b) PAI-1 gene transcription by transfection reporter assays, and (c) protein synthesis using the enzyme immunoassay. NF-κB activity was monitored using the electrophoretic mobility shift assay. Its activity was modified by either antisense oligonucleotides or transfection of endothelial cells with the wild-type or mutated IκBα. We have shown that TNFα-induced expression and gene transcription of PAI-1 involves a regulatory region present in segment −664/−680 of the PAI-1 promoter. This reaction involves the TNFα-induced generation of superoxide leading to activation of NF-κB, and can be abolished by antioxidants and by overexpression of a super-suppressor phosphorylation-resistant IκBα. Stimulation of PAI-1 under these conditions involves the motif of the PAI-1 promoter adjacent to the 4G/5G site, which can directly interact with NF-κB. We show that activation of PAI-1 gene by TNFα and reactive oxygen species is mediated by interaction of NF-κB with the cis-acting element located in the −675 4G/5G insertion/deletion in the PAI-1 promoter.

Abbreviations
DCF

dichlorofluorescin

DCFH-DA

2′7′-dichlorofluorescein diacetate

EMSA

electrophoretic mobility shift assay

H2O2

hydrogen peroxide

HUVEC

human umbilical vein endothelial cells

IKK

IκB kinase

IL

interleukin

NAC

N-acetylcysteine

O2

superoxide anion

PAI-1

plasminogen activator inhibitor-1

PEG

poly(ethylene glycol)

ROS

reactive oxygen species

TGFβ

transforming growth factor β

TNFα

tumor necrosis factor α

Plasminogen activator inhibitor type-1 (PAI-1) is mainly identified as the primary physiological inhibitor of both urokinase-type (uPA) and tissue-type (tPA) plasminogen activators, and plays an important role in regulation of the fibrinolytic system. Under normal conditions, PAI-1 is present in plasma at low concentrations, although high levels are seen in a variety of clinical settings [1]. PAI-1 is an early response gene product known to be activated by numerous factors, including transforming growth factor β (TGFβ) and interleukin (IL)-1β[2], platelet-derived growth factor and β fibroblast growth factor [3], thrombin [4], tumor necrosis factor α (TNFα) [5], insulin [6], angiotensin II [7] and oxidation products [8].

PAI-1 is inherently unstable and readily converts from an active to a latent form [9,10]. Thus, self-inactivation of PAI-1 is a crucial regulatory mechanism, by which this protein functions in circulation. Another mechanism of PAI-1 regulation results from the transcriptional control of its expression. Several regulatory elements have been localized in the human PAI-1 promoter and include two Sp1 elements (−73 and −42 bp) mediating glucose responsiveness [11], a hypoxia-responsive element (−194 bp) [12], a very low-density lipoprotein-responsive site (−672/−657 bp) [13], SMAD 3 and -4 protein-binding sites that mediate TGFβ responsiveness (−730, −580, and −280 bp) [14], oxidative stress and thymosin β4 responsive AP-1 site at −60/52 [15,16], and a 5′ distal TNFα responsive enhancer of the PAI-1 gene located 15 kb upstream of the transcription start site containing a conserved NF-κB-binding site [17].

In this report, we provide evidence that activation of PAI-1 gene by TNFα and reactive oxygen species (ROS) is mediated by interaction of NF-κB with the cis-acting element located in the −675 4G/5G insertion/deletion in the PAI-1 promoter. Because TNFα is a pivotal early mediator that regulates and amplifies the development of inflammation, this mechanism can be used primarily under diverse inflammatory conditions, including ischemia, traumatic injury, allograft rejection, cytokine stimulation and activation by bacterial components [18,19]. TNFα is known to induce the generation of ROS [20], which constitute primary signals transduced into the cytoplasm and ultimately alter the expression of specific genes [21–23]. Because ligand-stimulated NF-κB activation can be blocked by antioxidants, it appears that the generation of ROS may be involved in the induction of PAI-1 expression by TNFα via the activation of NF-κB.

Results

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References

Upregulation of PAI-1 expression in endothelial cells by ROS

In preliminary experiments, we evaluated the role of ROS during the stimulation of PAI-1 expression in endothelial cells by TNFα. For this purpose, endothelial cells were treated with 10 mmN-acetylcysteine (NAC) for 30 min, followed by incubation with TNFα (50 ng·mL−1) and the released PAI-1 antigen was determined by ELISA. Figure 1A shows that the treatment of endothelial cells with TNFα increased PAI-1 antigen by almost twofold (180 ± 10%, P < 0.05). Cotreatment with NAC inhibited TNFα-induced PAI-1 antigen accumulation by 48 ± 7% (P < 0.05). Similarly, incubation of endothelial cells with 100 or 200 µm H2O2 for 30 min increased PAI-1 by 160 ± 4 and 180 ± 8%, respectively (P < 0.05 for both). Also, in this case, cotreatment with NAC inhibited H2O2-induced PAI-1 antigen release by 38 ± 3 and 26 ± 3% observed at 100 and 200 µm H2O2, respectively (P < 0.05 for both).

image

Figure 1. Effect of TNFα and H2O2 on PAI-1 expression in vascular endothelial cells. Expression of PAI-1 was analyzed at the level of protein synthesis (A) in the presence or absence of NAC (10 mm). The PAI-1 antigen was determined using the ELISA test. (B) PAI-1 mRNA expression analyzed by real-time PCR in endothelial cells induced by TNFα in the absence or presence of different antioxidants. For this purpose, endothelial cells were preincubated for 30 min with NAC (10 mm), catalase (500 U·mL−1), pyrrolidine dithiocarbamate (PDTC) (100 µm), or vitamin C (100 µm) and stimulated with TNFα (50 ng·mL−1) for 4 h. Data are shown as mean ± SD obtained during three separate experiments.

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To determine whether the changes in PAI-1 antigen release were due to modulation of PAI-1 mRNA expression, we performed real-time PCR using endothelial cells treated with TNFα in the presence or absence of NAC. In addition to NAC, other antioxidants, such as PDTC (100 µm), catalase (500 U·mL−1) or vitamin C (100 µm) attenuated TNFα-induced PAI-1 mRNA expression (Fig. 1B).

NF-κB regulates PAI-1 gene transcription

Under basal culture conditions, endothelial cells exhibit little or no dichlorofluorescin (DCF) fluorescence. Treatment for 10 min with TNFα used at 50 ng·mL−1 increased DCF fluorescence, which was attenuated by cotreatment with NAC (10 mm) (Fig. 2A,B). Similarly, incubation of endothelial cells with 100 µm H2O2 for 10 min increased DCF fluorescence, which was then suppressed by cotreatment with NAC. These findings indicate that both TNFα and H2O2 increased intracellular oxidative stress, which was then partially abolished by the antioxidant NAC. Figure 2C shows the effect of poly(ethylene glycol) (PEG) and PEG-catalase on endothelial cells stimulated with TNFα. Results were obtained after 2 h of TNFα stimulation in the presence of PEG alone or PEG-catalase (100–1000 U·mL−1). Experiments were performed three times with similar results.

image

Figure 2. (A, B) The effect of TNFα (50 ng·mL−1) on intracellular oxidation analyzed by DCF fluorescence in vascular endothelial cells with and without of NAC (10 mm). (C) The effect of PEG and PEG-catalase on endothelial cells stimulated with TNFα. Results were obtained after 2 h of TNFα stimulation in the presence of PEG alone or PEG-catalase (100–1000 U·mL−1). Experiments were performed three times with similar results.

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To determine whether TNFα-induced PAI-1 expression involves the activation of NF-κB, we performed electrophoretic mobility shift assay (EMSA) studies using the consensus oligonucleotide for NF-κB (Fig. 3A), and PCR product containing wild-type sequence from −664 to −680 of PAI-1 promoter (Fig. 3B), 17-bp fragment (−664 to −680) of the PAI-1 promoter containing the putative κB-binding site (Fig. 3C) [24]. The treatment of endothelial cells with TNFα or H2O2 resulted in NF-κB activation which was inhibited in competition experiment using either the κB consensus oligonucleotide or fragment −664 to −680 of PAI-1 promoter containing the κB-binding site but not by mutated fragment −664 to −680 of PAI-1 promoter. NAC abolished NF-κB activation, indicating that TNFα- and H2O2-induced activation of NF-κB involves the generation of ROS. The identity of NF-κB present in the nuclear complex formed with the NF-κB consensus oligonucleotide was proved by supershift produced with anti-p65 sera (Fig. 3C). The effect of oxidative stress-induced NF-κB activation in TNFα-activated cells was confirmed using an antioxidant NAC. The cells were transfected with pNF-κB-SEAP plasmid and treated with TNFα in the presence or absence of NAC. NF-κB promoter activation by TNFα was inhibited by antioxidant, NAC (Fig. 3D). To confirm the role of fragment (−664 to −680) of PAI-1 promoter in this reaction, endothelial cells were analyzed after transfection with either wild-type PAI-1 promoter or its mutated version. Mutation of the NF-κB putative binding site within the PAI-1 promoter abolished its sensitivity to induction by TNFα(Fig. 3E).

image

Figure 3. Increased expression of NF-κB in endothelial cells with upregulated PAI-1 upon treatment with TNFα. EMSAs using oligonucleotides derived from consensus sequence of NF-κB (A) and containing the putative of κB binding site of PAI-1 promoter (B). Endothelial cells were stimulated with TNFα (50 ng·mL−1) or H2O2 (100 and 200 µm) in the presence of NAC (10 mm). (C) EMSAs using oligonucleotides (ACGTGGGGGAGTCAGCC) containing putative of κB binding site of PAI-1 promoter. Endothelial cells were stimulated with TNFα (50 ng·mL−1) or H2O2 (200 µm) in the presence or absence of NAC (10 mm). In the competition experiment, an excess amount unlabeled oligonucleotide containing the putative NF-κB binding site of PAI-1 promoter or unlabeled mutanted oligonucleotide or unlabeled consensus of NF-κB oligonucleotide was added to binding system 10 min prior to adding labeled oligonucleotide. The nuclear complex produced with the NF-κB probe is supershifted by the p65 antibody. Endothelial cells were transfected with pNF-κB-SEAP in the presence of the pSEAP as a control vector. The cells were preincubated with antioxidant NAC (10 mm) for 30 min and then treated with TNFα (50 ng·mL−1). Secreted alkaline phosphatase (SEAP) activities in the cells were determined by Chemiluminescence Detection Kit (D).  (E) A mutation of the NF-κB putative binding site (−664 to −680) within PAI-1 promoter abolishes its sensitivity to TNFα-induced transcriptional activity. In this experiment endothelial cells were transfected either with p800Luc or its mutated version, in which the −664 to −680 fragment (ACGTGGGGGAGTCAGCC) was substituted by the mutated sequence (ACATGGGCCAGTCAGCC). Mutation of the wild-type sequence was done on a PAI-1 promoter cloned into pLuc vector. Transfected cells were treated with TNFα (50 ng·mL−1) for 24 h and the cells harvested 12 h later. Luciferase activity was determined by the Dual-Luciferase assay kit. Data are shown as the mean of three separate experiments ± SD.

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To further evidence whether NF-κB activation is required for TNFα- and H2O2-induced PAI-1 expression, in subsequent experiments NF-κB expression was downregulated with antisense oligodeoxynucleotide (5′-GGGGAACAGTTCGTCCATGGC-3′) that is specific to the NF-κB subunit, RelA p65. In parallel, endothelial cells incubated with sense oligonucleotide were used as a control. In contrast to the sense RelA oligonucleotide, the addition of the antisense RelA oligonucleotide to endothelial cells resulted in strong inhibition of both TNFα- and H2O2-induced PAI-1 mRNA expression (Fig. 4A) and promoter activity (Fig. 4B), proving that NF-κB is required for both stimulated pathways. The efficacy of antisense oligonucleotides to reduce the expression of NF-κB is shown in Fig. 4C.

image

Figure 4. Effect of oligonucleotides antisense to RelA p65 on PAI-1 mRNA expression and promoter activity in endothelial cells. Endothelial cells were preincubated with the sense or antisense oligonucleotides and then treated with TNFα (50 ng·mL−1) or H2O2 (100 µm). (A) Expression of PAI-1 mRNA determined by real-time PCR and (B) activity of PAI-1 promoter. (C) Treatment of endothelial cells with the oligonucleotide antisense to RelA p65 abolished formation of the nuclear complex with the labeled probe containing NF-κB consensus sequence, as determined by EMSA.

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Effects of NF-κB inhibition on PAI-1 expression

The NF-κB complex is retained in the cytoplasm by IκB proteins. Activation of NF-κB involves its phosphorylation by IκB kinase (IKK) and subsequent degradation of IκB by 26S proteasome. To inhibit NF-κB, we treated endothelial cells with a relatively specific IκBα 26S proteasome inhibitor, MG132 used in the concentration range 0–1000 nm to prevent degradation of IκBα. In the presence of MG132, TNFα-induced expression of PAI-1 detected at the level of mRNA (Fig. 5C) or protein synthesis (Fig. 5A) was blocked in a concentration-dependent manner. Similarly, the stimulating effect of TNFα was abolished by salicylate, which is an IKK inhibitor (Fig. 5B,D). The data are consistent with observations presenting the effect of TNFα on PAI-1 promoter activity (Fig. 5E) and mRNA expression (Fig. 5F) tested in endothelial cells transfected either with empty vector (pCMV4), or containing wild-type IκBα (WT) or mutated IκBα (MT). Transfection with pCMV4 or WT-IκBα had little or no effect on TNFα-induced PAI-1 mRNA expression or promoter activity. However, overexpression of MT-IκBα, which cannot be phosphorylated at Ser32 and Ser36, resulted in a substantial decrease in TNFα-induced PAI-1 expression and promoter activity. These findings indicate that signaling pathways proximal to IκBα phosphorylation (i.e. IKK) are potential targets for activation by ROS.

image

Figure 5. Dependence of PAI-1 expression upon NF-κB activation. (A, B) Inhibition of PAI-1 antigen expression produced by incubation of TNFα-stimulated endothelial cells with increasing concentrations of 26S proteasome inhibitor, MG132 and sodium salicylate, respectively. (C, D) Similar reduction in PAI-1 mRNA expression in the same cells as analyzed by real-time PCR. All experiments were performed at least 3 times with reproducible results and data are shown as mean ± SD. (E, F) Changes in PAI-1 expression as measured at the level of PAI-1 promoter activity and PAI-1 mRNA evaluated by real-time PCR, respectively. In these experiments, endothelial cells were transfected with the empty vector (pCMV4), and the vector expressing either a wild-type IκBα (WT) or its nonphosphorylable mutant IκBα (MT). Results were standardized to cotransfection RSV-β-Gal expression plasmid (internal control). All data represent the mean of three separate experiments ± SD.

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Discussion

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References

In this study, we have shown that TNFα-induced expression and gene transcription of PAI-1 involve a regulatory region present in segment −664/−680 of the PAI-1 promoter, corresponding to the previously reported IL-1α-inducible site located between −675 and −669 [24]. This is supported by the following observations: (a) the oligonucleotide containing the putative κB-binding site of PAI-1 promoter (−664/−680) can bind transcriptional factor subunits p50 and p65. This binding is specific and can be abolished by triple mutation of the oligonucleotide, as seen both in the direct binding and during competitive inhibition experiments. (b) Mutation of the regulatory region abolished its responsiveness in the PAI-1 promoter to both TNFα and ROS, as demonstrated after the transfection of cells with p800Luc or its mutated version, respectively. Furthermore, responsiveness of this element to activation of endothelial cells by TNFα or ROS was also demonstrated by EMSA experiments. Interestingly, this regulatory element is adjacent to the polymorphic locus, −675 4G/5G insertion/deletion, which has stimulated increased interest in PAI-1 as a risk factor for venous thrombosis [25]. In particular, the 4G allele in this polymorphism has been associated with higher levels of plasma PAI-1 antigen and PAI-1 activity [26]. Furthermore, in the presence of the 4G allele not only was the PAI-1 response more pronounced, but so too was the response of other acute-phase reactants, which implies that the increases in these reactants are secondary to the increase in PAI-1 [27].

Our data show that activation of endothelial cells with TNFα to produce PAI-1 is mediated by a ROS-stimulated increase in NF-κB activity. Treatment with H2O2 increased PAI-1 expression and the effect was reduced by cotreatment with the antioxidant, NAC. Indeed, direct inhibition of NF-κB, either by antisense oligonucleotide or by overexpression of IκBα, attenuated both TNFα- and H2O2-induced PAI-1 expression. TNFα was reported to induce both superoxide anion and H2O2 production [28,29]. Because TNFα and ROS have similar effects on PAI-1 expression, these findings suggest a common mechanism by which these signaling pathways lead to the induction of PAI-1 in endothelial cells. Interestingly, PAI-1 upregulation by TGF-β has also been shown to involve ROS production in mesangial cells [30]. Furthermore, high glucose [31] and cyclic strain [32] can upregulate PAI-1 expression in endothelial cells through ROS generation. Indeed, PAI-1 is regulated by a redox-sensitive mechanism after the exposure to ionizing radiation in renal tubular epithelial cells [33]. In addition, IL-1-mediated upregulation of PAI-1 expression in cardiac microvascular endothelial cells also appears to be ROS dependent [34].

In this study, we also characterized the inhibitory mechanism produced by antioxidants on TNFα-induced PAI-1 expression in endothelial cells. Our findings indicate that this inhibition occurs via the suppression of NF-κB. Although TNFα could activate NF-κB by a ROS-independent mechanism, the ability of ROS to stimulate NF-κB activity may provide a synergistic effect in this activation [35]. TNFα is known to mediate the PAI-1 acute phase response in vivo and a κB-like sequence has been found in a number of promoters of acute phase-regulated genes [36]. Involvement of the NF-κB signaling pathway in PAI-1 upregulation has been reported in human endothelial cells stimulated by lipopolysaccharide [37] and in proximal tubular cells exposed to uremic toxins [38]. NF-κB also contributes to Chlamydia pneumoniae-induced overexpression of PAI-1 in vascular smooth muscle cells and endothelial cells [39]. Furthermore, NF-κB-like binding site in the PAI-1 promoter is responsible for IL-1-induced PAI-1 expression and this element is involved in the control of plasma levels of PAI-1 [24]. Thus, multiple cytokines and infectious agents can upregulate PAI-1 expression, leading to altered vascular hemostasis and proliferation.

Experimental procedures

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References

Reagents

All standard tissue culture reagents including M199 and fetal bovine serum were obtained from Gibco-BRL (Bethesda, MD, USA). TNFα were obtained from Promega (Madison, WI, USA). All other reagents were obtained from Sigma (St. Louis, MO, USA), unless indicated otherwise. The inhibitor Z-Leu-Leu-Leu-CHO (MG132) was from BioMol (Plymouth Meeting, PA, USA). The chemiluminescence detection reagent for western blotting was from Pierce (Rockford, IL, USA). The protein determination assay reagent, acrylamide, TEMED, and ammonium persulfate were from Bio-Rad (Hercules, CA, USA). LipofectAMINE Plus reagent was obtained from Gibco-BRL. Plasmid p800LUC with PAI-1 promoter was obtained as a gift from D. J. Loskutoff (Scripps Institute, La Jolla, CA, USA). Plasmids with wild-type IκBα (WT) and mutated IκBα (MT) were obtained as a gift from J. K. Liao (Harvard Medical School, Boston, MA, USA). [32P]dATP[γP] (6000 Ci·mmol−1) and T7 Sequenase (v.2.0) were purchased from Amersham (Piscataway, NJ). Protein assay reagents and polyacrylamide gel chemicals were from Bio-Rad. P-NF-κB-SEAP vector and SEAP Chemiluminescence Detection Kit was from BD Bioscience Clontech (Mountain View, CA, USA).

Cell culture

Human umbilical vein endothelial cells (HUVEC) were cultured in growth medium containing M199 and 20% (v/v) fetal bovine serum in a 90–95% humidified atmosphere of 5% (v/v) CO2 at 37 °C. To evaluate the effect of different compounds on PAI-1 expression at the level of protein synthesis, cells were grown on 48-well microplates. To determine PAI-1 mRNA levels, endothelial cells were cultured in 10 cm dishes. In all experiments, cellular viability was assessed by Trypan Blue dye-excluding cells. Relatively pure (> 95%) vascular endothelial cells were confirmed by their morphological features using phase-contact microscopy and immunofluorescent staining with PECAM-1 antibody (data not shown). At the concentrations used, there were no observable adverse effects of TNFα, superoxide dismutase, catalase or other antioxidants on cellular viability for all treatment conditions.

Treatment conditions

Prior to treatment with TNFα, endothelial cells were starved for ≈ 12 h in M199 supplemented with 0.1% (v/v) fetal bovine serum. Endothelial cells were then stimulated with TNFα (50 ng·mL−1) or H2O2 (100 and 200 µm) in the presence and absence of NAC (10 mm), pyrrolidinedithiocarbamate (100 µm), vitamin C (100 µm) and catalase (500 U·mL−1). In some experiments, the 26S proteasome inhibitor, MG132 (10 nm to µm), was added 30 min prior to TNFα stimulation. Sodium salicylate (2 and 4 mm) was added 1 h before the TNFα stimulation.

Real-time quantitative RT-PCR

Total RNA (1 µg) was extracted from endothelial cells using Trizol reagent (Life Technologies Inc, Rockville, MD, USA) and processed directly to cDNA synthesis using the TaqMan Reverse Transcription Reagents kit (Applied Biosystems, Foster City, CA, USA), according to the manufacturer's protocol. The PAI-1 and β-actin expression was quantified by real-time RT-PCR using ABI Prism 7000 Sequence Detection System (Applied Biosystems), according to the manufacturer's protocol. Briefly, 2.5, 2.0; 1.5, 1.0; 0.5 and 0.25 µL of synthesized cDNA were amplified in triplicate for both β-actin and each of the target genes to create a standard curve. Likewise, 2 µL of cDNA was amplified in triplicate in all isolated samples for each primer/probe combination and β-actin. Each sample was supplemented with both respective 0.3 µm forward and reverse primers, fluorescent probe, and made up to 50 µL using qPCR™ Mastermix for SYBR Green I (Eurogentec, Seraing, Belgium). All the following PCR primers were designed using software primerexpress (Applied Biosystems) forward 5′-TGCTGGTGAATGCCCTCTACT-3′, reverse 5′-CGGTCATTCCCAGGTTCTCTA-3′ forward 5′-CGTACCACTGGCATCGTGAT-3′, reverse 5′-GTGTTGGCGTACAGGTCTTTG-3′ specific for mRNAs of PAI-1 and β-actin, respectively. β-Actin was used as an active and endogenous reference to correct for differences in the amount of total RNA added to the reaction and to compensate for different levels of inhibition during reverse transcription of RNA and during PCR. Each target probe was amplified in a separate 96-well plate. All samples were incubated at 50 °C for 2 min and at 95 °C for 10 min, and then cycled at 95 °C for 30 s, 56 °C for 1 min and 72 °C for 1 min for 40 cycles. SYBR Green I fluorescence emission data were captured and mRNA levels were quantified using the critical threshold (Ct) value. Analyses were performed with abi prism 7000 (SDS Software). Controls without reverse transcription and with no template cDNA were performed with each assay. To compensate for variations in input RNA amounts, and efficiency of reverse transcription, β-actin mRNA was quantified and results were normalized to these values. Relative gene expression levels were obtained using the ΔCt method. Amplification specific transcripts were further confirmed by obtaining melting curve profiles.

Assay of intracellular oxidative stress

HUVEC of fewer than three passages were cultured in 35 mm dishes (Corning, NY, USA) coated with 0.1% (w/v) gelatin. Phenol-free M199 medium + 15 mm Hepes (pH 7.4) were used. Before seeding the endothelial cells, a sterile cover-slip was placed on the bottom of each dish. To subtract background fluorescent activity from intracellular fluorescence, these cover-slips were eliminated before measurement in order to have a cell-free control field. The preincubation period with antioxidants was 30 min. Intracellular generation of ROS was quantified using 2′7′-dichlorofluorescein diacetate (DCFH-DA) (Molecular Probes, Eugene, OR, USA). This esterified form is cell membrane permeable and undergoes deacetylation by intracellular esterases. Upon oxidation, DCFH is converted to dichlorofluorescin (DCF), a fluorescent compound. Confluent endothelial cell monolayers were incubated 30 min with 30 µm DCFH-DA before stimulation with TNFα. Briefly, PEG and PEG-catalase were dissolved in phenol-free culture medium and applied to endothelial cells 1 h before stimulation with TNFα. Results were obtained after 2 h of TNFα stimulation in the presence of PEG alone or PEG-catalase (100–1000 U·mL−1). Fluorescence was monitored using an inverted microscope (Zeiss Axiovert 405M, Oberkochen, Germany) with a specimen stage for 35 mm dishes. This custom-design stage was equipped with a temperature and gas control, thus allowing incubator conditions [i.e. 5% (v/v) CO2, 37 °C] under the microscope. Culture medium pH was 7.4 under microscopy conditions. A mercury lamp with a 490 nm filter was used as a light source for excitation. Excitation time (3 s) was constant for all conditions. The emission wavelength was set to 525 nm. Images were acquired using a CCD camera (Photometrics CH 250, Tucson, CA, USA) with a 512 × 512 pixel format. Digitalized images were transferred to a Sun SPARC workstation IPX (Sun Microsystems, Mountain View, CA, USA). Analysis was performed with isee software v. 3.6 (Inovision, Durham, NC, USA). For each condition, data were acquired from six different and representative fields (three separate regions of interest within two separate microscopic fields). Fluorescence intensity (excitation wavelength 490 nm; emission wave length 520 nm; excitation time 3 s) was recorded on a gray scale from 0 to 16 384. Background fluorescence (cell-free area) was subtracted from total fluorescent intensity.

EMSA

Confluent HUVECs were stimulated with TNFα (50 ng·mL−1) for 2 h. The double-stranded oligonucleotides: (a) transcription factor consensus oligonucleotide for NF-κB (AGTTGAGGGCACTTTCCCAGG) obtained from Integrated DNA Technologies, Inc (Coralville, IA, USA), (b) PCR products containing wild-type sequence from −664 to −680 of PAI-1 promoter (ACGTGGGGGAGTCAGCC), and (c) an oligonucleotide from the sequence −664 to −680 of PAI-1 promoter (ACGTGGGGGAGTCAGCC) were used as labeled probe. The introduction the mutations to wild-type sequence from −664 to −680 of PAI-1 promoter cloned into pLUC vector was carried out using a Quick Change site-directed mutagenesis kit (Stratagene, La Jolla, CA, USA) with mutagenic primers: 5′-ACATGGGGGAGTCAGCC-3, 5′-GGCTGACTCCCCCATGT-3′; 5′-ACATGGGGCAGTCAGCC-3, 5′-GGCTGACTGCCCCATGT-3′; 5′-ACATGGGCCAGTCAGCC-3, 5′-GGCTGACTGGCCCATGT-3′. The PCR product and the oligonucleotides were labeled to high specific activity with T4 polynucleotide kinase (Promega) using [32P]dATP[γP] (Amersham Biotech) 37 °C for 10 min and subsequently purified by electrophoresis in 7% polyacrylamide gels using 0.5× Tris/borate/EDTA, pH 7.8. We used 6–7 × 104 c.p.m. for EMSA. Binding reactions were performed in the binding buffer 20 mm Hepes/KOH, pH 7.5, 32 mm KCL, 5 mm MgCl2, 1 mm dithiothreitol, 10% (v/v) glycerol) in the presence of nonspecific competitor poly(dI:dC) (Amersham Biotech). Crude nuclear extracts (15 µg) from control cells and stimulated with TNFα (50 ng·mL−1) cells were incubated with 0.01 pmol of γ32P-labeled oligonucleotides and PCR products for 20 min at room temperature in a total volume of 20 µL. For competition experiments, 200-fold molar excess of unlabeled double-stranded competitor oligonucleotides (consensus of NF-κB, the sequence from −664 to −680 of PAI-1 promoter, mutated fragment −664 to −680 of PAI-1 promoter) were added to the reaction mixtures. To identify a transcription factor, 4 µg of polyclonal antibodies against NF-κB subunits (p50, p65) were introduced to the binding reactions for 30 min prior to the addition of the radioactive probe. DNA–protein complexes were resolved from unbound oligonucleotides and PCR products by elecrophoresis on 5% polyacrylamide gel using 0.5× Tris/borate/EDTA buffer (150 V for 2 h). Gels were vacuum-dried and autoradiographed with intensifying screens for 2–24 h at −20 °C.

Antisense oligonucleotide transfection

To transfect antisense oligonucleotides to RelA, cells were cultured in six-well plates and grown in M199 containing 20% (v/v) fetal bovine serum. After 24 h, wells were washed with serum- and antibiotic-free medium. Phosphorothioate oligonucleotides (200 nm) and the oligofectamine reagent (Invitrogen, Life Technologies) were added to the wells. Cells were incubated for 20 h with sense (5′-GCCATGGACGAACTGTTCCCC-3′) or antisense (5′-GGGGAACAGTTCGTCCATGGC-3′) oligonucleotides and then treated for additional 4 h with TNFα or H2O2 and total cellular RNA was isolated by the TRIzol Reagent.

Plasmids construcions

Introduction of mutations to the wild-type sequence from −664 to −680 of PAI-1 promoter cloned into pLUC vector was carried out using a Quick Change site-directed mutagenesis kit (Stratagene) and mutagenic primers: 5′-ACATGGGGGAGTCAGCC-3′, 5′-GGCTGACTCCCCCATGT-3′; 5′-ACATGGGGCAGTCAGCC-3′, 5′-GGCTGACTGCCCCATGT-3′; 5′-ACATGGGCCAGTCAGCC-3′, 5′-GGCTGACTGGCCCATGT-3′. The nickled vector DNA incorporating the desired mutations was than transformed into XL1-Blue cells.

Transfection reporter assays

Semiconfluent cell cultures in six-well tissue culture plates were transfected with DNA constructs (plasmid p800LUC with the PAI-1 promoter, p800LUCmut containing mutated sequence from −664 to −680 of the PAI-1 promoter and pCMV.IκBα expression vector) or with pNF-κB-SEAP vector. The cells were grown in six wells with a density of ≈ 1 × 105 cells·mL−1 and transfected using 1 µg of the plasmid DNA and lipoficatmine (Gibco BRL) according to manufacturer's instructions. As an internal control for transfection efficiency, pRSV.βGAL plasmid (0.5 µg) was cotransfected in all experiments. In parallel experiments, endothelial cells were transfected with luciferase reporter vector pGL3 (Promega) and used as control cells to test whether the effect of inhibitors was specific for the PAI-1 promoter.

Overexpression of IκBα

The following expression plasmids were used: (a) WT IκBα, encoding the full-length protein, was N-terminus FLAG-tagged in pCMV4. (b) MT IκBα, lacking the serine phosphorylation sites (32 and 36) and thus resistant to degradation by the 26S proteasome, was also N-terminus FLAG-tagged in pCMV4. Endothelial cells were transfected with IκBα expression plasmids by lipofectamine method. After 48 h cells were treated with TNFα (50 ng·mL−1) and total cellular RNA isolated 4 h later by using the TRIzol Reagent.

Statistical analysis

All values are expressed as mean ± SE compared with controls and among separate experiments. Paired and unpaired Student's t-tests were employed to determine the significance of changes. A P-value < 0.05 was considered statistically significant.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References

This work was supported by the Polish Committee for Scientific Research (KBN) Grant no. 3 PO4A 01425.

References

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References
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