Human endothelial cells cultured under high glucose (HG) conditions were shown before to upregulate several basement membrane proteins, including fibronectin (FN), thus mimicking effects of diabetes. Using human macrovascular (HUVEC) and microvascular (HMEC) endothelial cell lines, we evaluated in the present study some of the key molecular signaling events involved in HG-induced FN overexpression. This expression was shown to be dependent on endogenous endothelin (ET) receptor-mediated signaling. We also examined the roles played by protein kinase C (PKC) and the transcription factors nuclear factor κB (NF-κB) and activating protein (AP)-1 with respect to such changes. HG, PKC activators, and ETs (ET-1 and ET-3) that increased FN expression also caused activation of NF-κB and AP-1. Inhibitors of both NF-κB and AP-1 prevented HG- and ET-induced FN production. ET receptor blockade also prevented these HG- and ET-mediated changes. The results of this study indicate that glucose-induced increased FN production in diabetes may be mediated via ET-dependent NF-κB and AP-1 activation.
- endothelial cells
- activating protein-1
- nuclear factor-κB
increased extracellular matrix (ECM) protein synthesis and capillary basement membrane (BM) thickening are characteristic features of diabetic microangiopathy, which is the most common pathological finding in several chronic diabetic complications, including retinopathy and nephropathy (18, 4). Various pathogenetic mechanisms may be responsible for capillary BM thickening in diabetes. Fibronectin (FN) is one of the most important ECM proteins that mediate a number of functions in BMs. Physiologically, FN plays an important role in cell adhesion, motility, tissue repair, etc. However, its overproduction may decrease motility and replication of many cells, including endothelial cells (ECs) (24). Diabetes increases turnover of vascular ECs in the retina (26). Furthermore, FN synthesis is increased in the retina of diabetic patients with background retinopathy (34). Vasoactive factors like endothelins (ETs), by virtue of their extensive tissue distribution and widespread biological actions, are important mediators of pathogenic changes in several diseases affecting microvasculature, including diabetes. We have previously demonstrated that ET-1 and ET-3 expressions are upregulated in the retina of both diabetic and galactose-fed rats (9, 11). We have further demonstrated that ET receptor blockade prevents hyperhexosemia-induced vasoconstriction, increased ECM production, and BM thickening in the retina and glomeruli of diabetic rats (7, 9, 10, 11). We have shown that diabetes-induced myocardial focal scarring and increased ECM protein production may be blocked by ET antagonism (8). The major ET isoforms having biological significance with respect to diabetic complications are ET-1 and ET-3 (21,22). Several pathways involved in diabetes may also result in increased ET synthesis. Protein kinase C (PKC) activation, secondary to hyperglycemia, may lead to ET-1 mRNA upregulation (18, 21,22). We have demonstrated that vascular endothelial growth factor (VEGF), which is upregulated in diabetes via a PKC-dependent mechanism, as well as reduced nitric oxide (NO), may lead to ET-1 upregulation in ECs (5). Various pathways of tissue injury caused by hyperglycemia in vivo or high glucose (HG) concentration in culture may activate transcription factors such as nuclear factor (NF)-κB and activating protein (AP)-1 (27, 30, 33). NF-κB and AP-1 regulate expression of several genes important in the pathogenesis of diabetic complications (27, 30). NF-κB is present in several cell types, including ECs (38). Normally, NF-κB exists in an inactive form in the cytoplasm bound to an inhibitory protein, IκB. Upon stimulation, IκB is hydrolyzed and the p50/p65 dimer translocates to the nucleus and initiates transcription (1, 6, 15, 27, 30). Resynthesis of IκB, induced by NF-κB, allows sequestration of NF-κB in the cytoplasm, shutting down the NF-κB response (1, 6, 15). ET-1 has been demonstrated to activate NF-κB in the hepatic stellate cells via ETB receptor (12). Activation of genes in different altered physiological and pathological conditions may involve coordinated participation of NF-κB and another transcription factor, AP-1 (17, 37). AP-1 consists of homodimers ofJun or heterodimers Fos and Jun(17, 37). It is also regulated by cellular stress. Furthermore, angiotensin II-induced end organ damage in hypertension has been shown to be mediated via ET receptor-dependent NF-κB and AP-1 activation (29). Human ECs cultured in the presence of HG (25 mmol/l) upregulated expression of many basement proteins including FN, thus mimicking effects of diabetes (3). Therefore, in the present study, we sought to determine the role of NF-κB and AP-1 in mediating HG-induced FN synthesis in ECs in culture and, more importantly, to determine whether such changes are regulated by ETs and their receptors. We used two different types of ECs for our studies. We used HUVECs (human umbilical vein ECs), because these are well-characterized ECs, and HMECs (human microvascular ECs), because diabetic angiopathy involves both microvessels and macrovessels and, therefore, investigations in both microvascular and macrovascular ECs are warranted.
MATERIALS AND METHODS
All reagents were obtained from Sigma Chemical (St. Louis, MO) unless otherwise mentioned.
The HUVEC and HMEC lines were obtained from American Type Culture Collection (Rockville, MD) and Clonetics (Clonetics, Walkersville, Maryland), respectively. These cells were plated at 2,500 cells/cm2 in EC growth medium (EGM) (Clonetics). EGM is a modified MCDB 131 formulation and is supplied with 10 μg/l human recombinant epidermal growth factor, 1.0 mg/l hydrocortisone, 50 mg/l gentamicin, 50 μg/l amphotericin B, 12 mg/l bovine brain extract, and 10% fetal bovine serum. Cells were grown in 25-cm2 tissue culture flasks. Appropriate concentrations of glucose were added to the medium when cells were 80% confluent. l-Glucose was used as a control. In all experiments, the specific ETA blocker TBC11251 (courtesy of Dr. R. Tilton, Texas Biotechnology, Houston, TX) and the PKA inhibitor N-tosyl-l-phenylalanine chloromethyl ketone (TPCK) were used at 10 μmol/l (5). Phorbol 12-myristate 13-acetate (PMA) was used at 60 μg/l (5). The selective ETB antagonist BQ788, dual ETA and ETB antagonist bosentan (courtesy of Dr. M. Clozel, Actelion, Allschwill, Switzerland), and the PKC inhibitor chelerythrine were used at 1 μmol/l (5). NF-κB inhibitor SN50 and inactive peptide control SN50M (Calbiochem, La Jolla, CA), as well as dual NF-κB and AP-1 inhibitor curcumin, were used at 20 μmol/l (23, 31). ET-1 and ET-3 (Peninsula Laboratories, Belmont, CA) were used at 5 nmol/l, and NF-κB inhibitor aminopyrrolidine-2,4-dicarboxylate (PDTC) was used at 100 μmol/l (25). All experiments were carried out after 24 h of incubation unless otherwise indicated. Three different batches of cells, each in duplicate, were investigated.
Cell proliferation and cell viability.
Cell viability was measured by trypan blue dye exclusion, and cell proliferation was evaluated by the microculture tetrazolium assay using 2,3-bis(2 methoxy-4-nitro-5-sulfophenyl)-5-[(phenylamino) carbonyl]-2H-tetrazolium hydroxide (XTT) (36). Cells were seeded in 100-ml, 96-well plates (3,200 cells/well, 10,000 cells/cm2). After 24 h, experimental agents were applied (100 ml) and the cultures were incubated for 1, 3, or 5 days at 37°C. XTT (50 μg) and 0.38 mg of phenazine methosulfate were added to each well (50 μl) after cell inoculation. The cells were incubated at 37°C for 4 h, and the plates were mixed on a mechanical plate shaker. Absorbance at 450 nm was measured with the Bio-Rad model 3550 microplate reader (Bio-Rad Laboratories, Hercules, CA). All experiments were performed in triplicate.
Preparation of nuclear protein fractions.
Nuclear extracts of HUVECs and HMECs were prepared as described elsewhere, with some modifications (19, 40). Rapid detection of octamer binding proteins with “mini-extracts” was prepared from a small number of cells. Briefly, the cells were washed, resuspended in phosphate-buffered saline, and centrifuged (7,000g for 15 s). The pellet was resuspended in 0.4 ml of cold buffer A [10 mmol/l HEPES, pH 7.9, 10 mmol/l KCl, 0.1 mmol/l EDTA, 0.1 mmol/l EGTA, 1 mmol/l 1,4-dithiothreitol (DTT),and 0.5 mmol/l PMSF] by gentle pipetting. The cells were allowed to swell on ice for 15 min. Twenty-five microliters of a 10% Igepal CA-630 was added, and cells were vortexed vigorously. The homogenate was centrifuged (10,000 g for 30 s). The nuclear pellet was resuspended in 50 μl of ice-cold buffer C (20 mmol/l HEPES, pH 7.9, 0.4 mol/l NaCl, 1 mmol/l EDTA, 1 mmol/l EGTA, 1 mmol/l DTT, and 1 mmol/l PMSF), and the tube was vigorously rocked at 4°C for 15 min on a shaking platform. The nuclear extract was centrifuged at 4°C (15,000 g for 5 min), and the supernatant was frozen at −70°C. The protein concentrations were measured using the BCA protein assay, with bovine serum albumin as a standard (Pierce, IL).
Electrophoretic mobility shift assay.
NF-κB and AP-1 consensus oligonucleotide (Promega, WI) DNA probes (Table 1) were prepared by end-labeling with [γ-32P]ATP (Amersham, Quebec, QC, Canada) using T4 polynucleotide kinase. The probes were purified by ethanol precipitation and resuspended in 10 mmol/l Tris and 1 mmol/l EDTA (pH 7.6). Nuclear proteins (5 μg) were incubated with 100,000 cpm of 32P-labeled consensus oligonucleotides for 30 min at room temperature. The incubation was carried out in a buffer containing 10 mmol/l Tris (pH 7.5), 50 mmol/l NaCl, 1 mmol/l MgCl2, 5% glycerol, 0.05% NP-40, 0.5 mmol/l EDTA, 0.5 mmol/l DTT, and 0.5 μg of poly(dI-dC). Protein-DNA complexes were resolved on a standard 6% (NF-κB) and 4% (AP-1) nondenaturing polyacrylamide gel in 0.5× TBE running buffer. After 0.5 h of electrophoresis at 350 V, gels were dried under a heated vacuum onto Whatman paper and subjected to autoradiography (19, 40). Anti-NF-κB (p65) monoclonal antibody and anti AP-1 (c-jun) polyclonal antibody (Santa Cruz Biotechnology, CA) were used for supershift assay. The specificity of binding was further confirmed by incubation with 100-fold unlabeled oligonucleotides. The blots were quantified by densitometry. The analyses for comparison of effects of various reagents were carried out after 4 h of incubation.
Total proteins were resolved by 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis and were analyzed by Western blotting using the antibodies described above. The signals from Western blots were obtained using horseradish peroxidase-conjugated secondary anti-mouse or anti-rabbit antibody (Santa Cruz Biotechnology) and developed using the chemiluminescent substrate (Amersham Pharmacia Biotechnology, Amersham, UK).
TRIzol reagent (Canadian Life Technologies, Burlington, ON, Canada) was used to isolate RNA. RNA was extracted with chloroform, followed by centrifugation to separate the solution into aqueous and organic phases. RNA was recovered from the aqueous phase by precipitation with isopropyl alcohol and suspended in diethyl pyrocarbonate-treated water.
First strand cDNA synthesis.
First strand cDNA synthesis was performed using Superscript-II system (Canadian Life Technologies). RNA was added to Oligo (dT) primers (Canadian Life Technologies), denatured at 65°C, and quenched on ice for 10 min. Reverse transcription was carried out by the addition of MMLV-reverse transcriptase and dNTP at 42°C for 50 min in a total reaction volume of 20 μl. The reaction was terminated by incubating at 70°C for 15 min. The resulting RT products were stored at −20°C.
The amplification of RTs was carried out as previously described with some modifications (5, 7, 9, 10, 11). Competitive PCR was performed for both ET-1 and FN. Competitor DNA fragments were generated using TaKaRa competitor kit. Several dilutions of competitor DNA and the target RT product were mixed to standardize the reaction. The primer sequence and the predicted product size are outlined in Table 1. Reactions were performed in 25-μl volumes containing 1 × PCR buffer, 1.5 mmol/l MgCl2, 250 μmol/l dNTP mix, 1 μmol/l each of amplification primer, 2.5 U Taq polymerase, and 1 μl of RT product. The amplification for FN mRNA (25 cycles) was carried out as follows: 45 s at 94°C (denaturation), 45 s at 54°C (annealing), and 1 min at 72°C (extension). Amplification of ET-1 mRNA was carried out at the same temperatures using 30 cycles. The amplification products were analyzed on a 3% agarose gel in 1 × TBE buffer, stained with ethidium bromide, and visualized under ultraviolet light.
Quantitation was performed by densitometric analysis of the bands using Mocha software (SPSS, Chicago, IL). The densitometric values were expressed as gene to competitor ratio per microgram of total RNA.
Confocal microscopy for NF-κB and FN.
Cells were plated on eight chamber-tissue culture slides and incubated for 24 h. Glucose (25 mmol/l) was added 24 h before different stimulators and inhibitors were added. After 24 h of stimulation or inhibition, cells were fixed with 1:1 methanol:acetone. The cells were then stained using polyclonal rabbit antihuman FN antibody (1/50; DAKO Diagnostics Canada, Mississauga, ON, Canada) or anti-NF-κB mouse monoclonal antibody (1/40; Santa Cruz) antibody. Goat anti-rabbit/anti-mouse IgG labeled with Texas red or FITC (1/100; Vector Laboratories, Burlington, ON, Canada) was used as the secondary antibody. Slides were mounted in Vectashield fluorescence mounting medium with 4′,6′-diamino-2-phenylindole (DAPI; Vector Laboratories) for nuclear staining. Microscopy was performed by an examiner unaware of the identity of the sample using a Zeiss LSM 410 inverted laser scan microscope equipped with fluorescein, rhodamine, and DAPI filters (Carl Zeiss Canada, North York, ON, Canada).
Data are expressed as means ± SE and were analyzed by ANOVA followed by Student's t-test with Bonferroni corrections. Differences were considered significant at values of P< 0.05.
Cell proliferation and viability.
No difference in viability was noted in cells treated with low (5 mmol/l) or high glucose (25 mmol/l) up to 96 h. The cells in HG, however, showed a significantly (P < 0.001) lower proliferation rate compared with cells in low glucose (data not shown).
ET-1 expression by HG.
We then reproduced our previous findings (5) of upregulation of ET-1 mRNA levels in both HUVECs and HMECs exposed to HG in culture medium (data not shown). We also confirmed the inhibitory effects of PKC inhibitor chelerythrine on HG-induced ET-1 gene expression (data not shown).
Modulation of HG-induced FN expression by ET receptor antagonists and inhibitors of PKC, NF-κB and AP-1.
We have previously demonstrated that hyperhexosemia-induced increased expression of ET is an important mediator of increased ECM production in retinas, glomeruli, and the hearts of diabetic rats (9, 11,24, 26, 34). One of the most important ECM proteins that is overexpressed in these organs in diabetes is FN (3, 9, 35 ). As reported by Cagliero et al. (3), HG concentrations increase a number of ECM-related genes, including FN. In the present study, incubation of the HUVECs in HG (25 mmol/l) caused a time-dependent increase in FN mRNA expression, reaching a maximal increase after 24 h, and thereafter maintained a similar level at least up to 72 h (Fig.1 A). Therefore, subsequent experiments were performed with HG treatment for 24 h in both HUVECs and HMECs. FN mRNA expression was demonstrated as three bands at 852 base pairs (bp) (EIIIA+), 582 bp (EIIIA−), and an intermediate band of ∼650 bp. The intermediate band represents a heteroduplex DNA consisting of one strand of each product mentioned above (35). The competitor band was present at 340 bp. A similar increase was also observed when both the ECs in 5 mmol/l glucose were incubated with ET-1 and ET-3 (Fig. 1,B–D). Failure of mimicry by an equal concentration of l-glucose suggests a lack of involvement of hyperosmolality in HG-induced FN overexpression (Fig. 1,B–D). HG-induced increased FN mRNA accumulation was prevented in both cell types by the nonselective ET receptor antagonist bosentan, as well as the selective ETAantagonist TBC11251 (Fig. 1, B–D). The ETB-selective antagonist BQ788 was, however, only effective in blocking FN mRNA expression in HMECs but not in HUVECs (Fig. 1,B–D).
PKC has been known to mediate many effects of HG concentrations. Activation of PKC has previously been shown to activate NF-κB that might activate its target genes (13, 28). To establish a link between PKC and NF-κB activation in HG-induced upregulation of FN expression, we preincubated ECs with two different pharmacological inhibitors of NF-κB, SN50 and PDTC, and a dual inhibitor of NF-κB and AP-1, curcumin. All these agents prevented HG-induced increase of FN mRNA accumulation in both the ECs (Fig. 1,B–D).
In keeping with the RNA data, 25 mmol/l glucose and ETs showed increased immunocytochemically detectable FN protein expression. This increased FN protein expression was prevented by chelerythrine and ET antagonist in both cell types. Similarly SN50, PDTC, and curcumin blocked increased FN immunoreactivity, which was not seen with SN50M or TPCK (Fig. 2).
HG concentration or ET can activate NF-κB and AP-1 in cultured ECs.
As previously reported, HG concentration causes NF-κB activation in bovine aortic ECs (32). In our experiments, incubation of HUVECs and HMECs in HG caused a time-dependent activation of NF-κB. In HUVECs, Western blot analysis of total cellular proteins demonstrated maximum increase at 4–8 h. (data not shown). EMSA, however, demonstrated peak NF-κB activation at 24 h of incubation, which remained at the same level up to 48 h (data not shown). In HMECs, the peak NF-κB activation was also observed after 24 h of incubation (data not shown). Therefore, subsequent experiments were performed with 24-h incubation. Similar NF-κB activation was observed when the cells in 5 mmol/l glucose were incubated with ET-1, ET-3, or PMA (Figs. 3, A andB, and 4,A–C). The specificity of NF-κB activation was further established by supershift assay (Fig. 3, A andB) and also in the experiments in which both HG and ET-induced NF-κB activation were blocked by the NF-κB inhibitor SN50, antioxidant and NF-κB inhibitor PDTC, and dual NF-κB and AP-1 inhibitor curcumin, but not by inactive peptide SN50M (Fig.4). HG-induced NF-κB activation was further blocked by the specific ETA antagonist TBC11251 and by the dual ETA/ETB antagonist bosentan. In HUVECs, ETA antagonist was more effective than ETB antagonist, whereas HMECs were equally responsive to ETA and ETB antagonists (Fig. 4). Similar inhibition of NF-κB activation was seen when the cells in HG were incubated with the PKC inhibitor chelerythrine (Fig. 4,A–C). No effects of PKA inhibitor TPCK were seen in either of the cells (Fig. 4, A–C).
Activation of NF-κB was further confirmed by the nuclear translocation of NF-κB after incubation with HG, as observed by confocal microscopy (Fig. 5). NF-κB p65 was distributed predominantly in the cytosolic and membrane areas in the cells treated with 5 mmol/l glucose (Fig. 5,A–C). However, the cells treated with HG showed intense immunostaining in the nucleus and perinuclear areas (Fig. 5, B and D). A similar result was obtained when the ECs were incubated with ET-1 (data not shown).
Transcription factor AP-1 also showed a similar pattern of glucose-induced changes. In the HUVECs, peak activation was observed at 12 h by EMSA (data not shown). In the HMECs, however, peak activation was observed at 24 h (data not shown). The specificity of AP-1 activation was further established by supershift assay (Figs.6 and 7). In the HUVECs and HMECs, glucose- and ET-induced AP-1 activation were blocked by TBC11251, bosentan, and chelerythrine. No effect of PKA blocking was observed. PDTC and BQ788 were more effective in HUVECs compared with HMECs. However, curcumin was effective in both cell types (Fig. 7).
The key findings of the present study are the following:1) HG-induced accumulation of FN mRNA and protein in human ECs, requiring activation of NF-κB and AP-1, can be prevented by ET receptor antagonism; 2) these HG-induced effects in human ECs can be mimicked by activation of ET receptor(s); and 3) all the observed effects of HG are very similar in both macrovascular (HUVEC) and microvascular (HMEC) ECs. The only difference in the two cells we observed is that both ETA and ETBmediate the ET response in HUVECs, whereas ETApredominantly mediates the ET response in HMECs. Furthermore, it is well known that HG activates PKC (16). We studied the importance of the PKC pathway in HG-induced activation of human ECs by evaluating the countereffect of PKC inhibitor on HG-induced FN expression and NF-κB and AP-1 activation and by further evaluating the HG-mimicking effects of the PKC activator phorbol ester. Indeed, our results showed the requirement of PKC in HG-induced effects in both HUVECs and HMECs. Activation of PKC is also required for HG-induced upregulation of ET in many cells, including ECs (5). Therefore, HG causing upregulation of ET expression through PKC activation can further activate transcription factors NF-κB and AP-1 and thereby increases FN expression. However, hyperhexosemia-induced ET upregulation can also be controlled by NF-κB and AP-1 (21, 22,33). Furthermore, as ET receptors are predominantly Gq-coupled, their activation may lead to activation of phospholipase C, elevation of [Ca2+]i, and activation of PKC. Therefore, combining the results of our present study with previously published studies (5, 21, 22, 33), we can speculate that PKC, NF-κB, and AP-1 can control both upstream and downstream effector molecules in the HG-induced signaling pathway.
Gel shift and supershift analyses in this study have shown activation of at least p65 subunits of NF-κB and c-Jun subunits of AP-1 in nuclear extracts of both HUVECs and HMECs. The specificity of these bindings has been examined further by competition experiments performed with 100-fold excess unlabeled nucleotides corresponding to NF-κB and AP-1 binding sequences. Elevated nuclear levels of p65 have also been observed in HG-treated ECs by Western blot analysis (data not shown). In hepatoma cells, NF-κB has been shown to play both positive and negative regulators acting on the FN gene (20). Although several potential NF-κB binding sites have been identified in the FN promoter, functional significance of only some of these has been determined. Regulation of FN gene expression has been shown to be offered by −41 NF-κB binding site (20). The DNA sequence between +1 and +136 has been shown to be responsible for part of PKC-mediated activation of FN gene in hepatoma cells (20). A potential NF-κB p65 response element present in the FN promoter has been shown to serve as a positive regulator of FN gene expression (20). Identification and characterization of a NF-κB binding site in the promoter region of FN gene responsible for ET responsiveness in ECs require further investigation. An AP-1 binding site mediating angiotensin II-induced transcriptional activation of FN gene has been shown in its promoter region (39). Whether ET-induced activation of FN gene also utilizes the same region of the gene remains to be investigated.
The effects of HG concentration in various cell types may be mediated by different PKC isoforms. For example, PKCα has been shown to mediate some of the HG-induced effects in porcine aortic ECs (14). Which PKC isoform(s) mediate(s) the HG-induced effect on FN expression in HUVECs and HMECs remains to be determined.
Cyclic AMP response element (CRE) present in FN gene promoter is known to play an important role in FN gene expression (2). However, PKA inhibitor did not reverse HG-induced upregulation of FN expression in the present study. Therefore, CRE in FN gene may not play any important role in HG-induced upregulation of FN in HUVECs and HMECs.
In conclusion, using macrovascular (HUVEC) and microvascular (HMEC) EC lines, we have demonstrated that constitutive expression of functional ET receptor(s) is required for HG-induced upregulation of FN and that ETs can mimic this HG-induced effect. Furthermore, both HG- and ET-induced FN upregulation involve two transcription factors, NF-κB and AP-1. The observations made in the present study are likely to have relevance to the vasculopathy in diabetes. Studies are underway to extrapolate our present findings in vivo using models of diabetic complications.
This study was supported in part by grants from the Canadian Diabetes Association in honor of Margaret Francis and the Canadian Institute of Health Research (MOP 43841).
Address for reprint requests and other correspondence: S. Chakrabarti, Dept. of Pathology, Dental Sciences Bldg., Univ. of Western Ontario, London, Ontario, N6A 5C1, Canada (E-mail:).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
First published September 18, 2002;10.1152/ajpcell.00192.2002
- Copyright © 2003 the American Physiological Society