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Am J Physiol Heart Circ Physiol 290: H1624-H1634, 2006. First published November 4, 2005; doi:10.1152/ajpheart.01233.2004
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Mechanisms of low-density lipoprotein-induced expression of connective tissue growth factor in human aortic endothelial cells

Mimi Sohn,1 Yan Tan,1 Bing Wang,1 Richard L. Klein,1,3 Maria Trojanowska,2 and Ayad A. Jaffa1

Divisions of 1Endocrinology, Diabetes, and Medical Genetics and of 2Rheumatology and Immunology, Department of Medicine, Medical University of South Carolina, and 3The Ralph H. Johnson Veterans Affairs Medical Center, Charleston, South Carolina

Submitted 7 December 2004 ; accepted in final form 1 November 2005


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Hyperlipidemia is a recognized risk factor for atherosclerotic vascular disease. The underlying mechanisms that link lipoproteins and vascular disease are undefined. Connective tissue growth factor (CTGF) is emerging as a key determinant of progressive fibrotic diseases, and its expression is upregulated by diabetes. To define the mechanisms through which low-density lipoproteins (LDL) promote vascular injury, we evaluated whether LDL can modulate the expression of CTGF and collagen IV in human aortic endothelial cells (HAECs). Treatment of HAECs with LDL (50 µg/ml) for 24 h produced a significant increase in the mRNA and the protein levels of CTGF and collagen IV compared with unstimulated controls. To explore the mechanisms by which LDL regulates CTGF and collagen IV expression in HAECs, we determined first if CTGF and collagen IV are downstream targets for regulation by transforming growth factor-beta (TGF-beta). The results demonstrated that TGF-beta produced a concentration-dependent increase in the protein levels of CTGF. To assess whether the induction of CTGF in response to LDL is mediated via autocrine activation of TGF-beta, HAECs were treated with LDL for 24 h in the presence and absence of anti-TGF-beta neutralizing antibodies (anti-TGF-beta NA). The results demonstrated that the increase in CTGF induced by LDL was significantly inhibited by the anti-TGF-beta NA. To investigate the upstream mediators of TGF-beta on activity of CTGF in response to LDL, HAECs were treated with LDL for 24 h in the presence and absence of cell-permeable MAPK inhibitors. Inhibition of p38mapk activities did not affect LDL-induced TGF-beta1, CTGF, and collagen IV expression. On the other hand, SP-600125, a specific inhibitor of c-Jun NH2-terminal kinase, suppressed LDL-induced TGF-beta, CTGF, and collagen IV expression, and PD-98059, a selective inhibitor of p44/42mapk, suppressed LDL-induced TGF-beta and CTGF expression. These findings are the first to implicate the MAPK pathway and TGF-beta as key players in LDL signaling, leading to CTGF and collagen IV expression in HAECs. The data also point to a potential mechanistic pathway through which lipoproteins may promote vascular injury.

transforming growth factor-beta; mitogen-activated protein kinase; collagen IV

DIABETES MELLITUS is associated with increased morbidity and mortality derived mainly from cardiovascular complications. The progression of vascular lesions is enhanced in the diabetic state, and early atherosclerotic lesions are characterized by endothelial dysfunction, accumulation of inflammatory cells, and intima vascular smooth muscle cell (VSMC) proliferation, migration, and extracellular matrix (ECM) deposition in the vessel wall (9, 10, 17, 34). In addition, endothelial cells tend to lose their physiological role as a semipermeable membrane barrier that separates the blood cells and soluble components, such as proteins or lipoprotein complexes, from the cells of the underlying the endothelium. This stage in the development of atherosclerosis is a complex process involving interactions between lipids and activated cells of the immune system, T cells, monocytes, or macrophages, which then convert into foam cells (40).

Although the association of hyperlipidemia and atherosclerosis is well established, the cellular signaling mechanisms that promote atherosclerosis are still undefined. In this regard, the cytokine transforming growth factor-beta (TGF-beta) has been shown to play a pivotal role in vessel wall remodeling after vascular injury (41). Recent studies have implicated a key role for the cytokine TGF-beta as a mediator of vessel wall remodeling after endothelial injury (3, 28). Recent evidence indicates that TGF-beta may mediate its fibrotic effects via activation of CTGF (13). The expression of CTGF is induced by TGF-beta in VSMC, indicating that CTGF may act along a causal pathway for atherosclerosis. Indirect evidence supporting this notion is provided by the fact that high levels of expression of CTGF mRNA and protein concentrations occur in VSMC and endothelial cells of advanced human atherosclerotic lesions (31). In addition, VSMC expressing CTGF were localized predominantly in areas with extracellular matrix production and especially in areas around the fibrous cap, thus indicating that CTGF may regulate the production of matrix proteins in these cells (31).

CTGF was originally identified as a product of human umbilical vein endothelial cells that were both chemotactic and mitogenic for fibroblasts (5). It is now known that CTGF belongs to a new gene family, CCN (named after prototype members of this family, CTGF, cyr61, and nov) (4). The molecular weight of CTGF-like factors varies between 35 and 40 kDa, and the structure of these molecules consists of four modules: an NH2-terminal insulin-like growth factor (IGF) binding protein-like domain, a von Willebrand factor type C repeat domain, a thrombospondin type 1 repeat domain, and a COOH-terminal dimerization domain (4).

An emerging role of CTGF is that of a prosclerotic factor. CTGF was shown to be expressed in atherosclerotic lesions (35) and to mediate adherence and migration of monocytes, VSMCs, and activated platelets, suggesting a functional role in the development of atherosclerotic lesions and plaque rupture (6, 18). Endothelial cells themselves are targets of CTGF, leading to proliferation, migration, and in vitro and in vivo angiogenesis (1, 37).

Although hyperlipidemia is now considered a risk factor for the progression of vascular disease, the relationship between increased plasma lipoproteins and endothelial cell dysfunction is poorly defined. Hyperlipidemia can directly or indirectly stimulate the synthesis and release of factors from resident vascular cells, which in turn can influence endothelial cell structure and function in an autocrine or paracrine manner. Therefore, the present study was designed to explore the potential role of LDL in modulating the expression of CTGF in HAECs and to delineate the cellular signaling mechanisms through which this regulation may occur.


   METHODS
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 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Endothelial cell culture. Human aortic endothelial cells (HAECs) were obtained from Cambrex (Walkersville, MD) and cultured using EBM-2 basal serum-free culture medium and EGM-2 SingleQuots growth supplement (2% FBS, 0.1% human epidermal growth factor, 0.04% hydrocortisone, 0.1% vascular endothelial growth factor, 0.4% human fibroblast growth factor-B, 0.1% R3-IGF-1, 0.1% ascorbic acid, 0.1% GA-1000, and 0.1% heparin), which were obtained from Clonetics (Walkersville, MD). HAECs between passages 4 and 9 were used when 70–90% confluent.

LDL preparation and characterization. LDL was prepared as previously described (19). Briefly, blood was taken from fasting healthy nondiabetic volunteers into a lipoprotein preservative-antioxidant cocktail (LPPC) containing EDTA (0.1% wt/vol), chloramphenicol (20 µg/ml), gentamicin sulfate (50 µg/ml), {epsilon}-amino-n-caproic acid (0.13%, wt/vol), aprotinin (1 µg/ml) and diethylenetriaminepentaacetic acid (1 mM) (final concentrations). Phenylmethylsulfonyl fluoride (PMSF, 20 µg/ml final concentration) was added to plasma to retard proteolysis. All samples were processed at low temperature and in the absence of white light to minimize oxidation. All density solutions were supplemented with LPPC, degassed, and purged with NB2B. Plasma density (d) was increased to d = 1.21 g/ml using dried nitrogen and 11 ml layered under d = 1.019 g/ml saline/LPPC. After ultracentrifugation (Beckman VTi50 rotor, 2.5 h, 50,000 rpm, 7°C with slow acceleration and deceleration), the LDL band was harvested by piercing the tube and aspirating into a syringe. LDL isolated by this procedure was free from contamination with apoA-I and albumin. Each LDL preparation was characterized by electrophoresis on 1% agarose gels (Paragon gels, Beckman, Brea, CA). The LDL pools were tested for endotoxin contamination by the Limulus amebocyte lysate (BioWhittaker, Walkersville, MD) according to the manufacturer’s suggestion. These studies were approved by Institutional Review Board of the Medical University of South Carolina, and all subjects signed an informed consent.

Harvesting of cells and collecting of conditioned media. Conditioned media samples were collected, centrifuged, and stored at –80°C until required. Endothelial cells treated with LDL were washed twice in ice-cold PBS, scraped in PBS containing 2 mM sodium vanadate, and centrifuged at 3,000 g for 5 min. Pellets were resuspended in 100 µl of lysis buffer (50 mmol/l Tris·HCl, pH 8.0, 150 mmol/l NaCl, 0.02% sodium azide, 0.1% SDS, 1% NP40, 0.5% sodium deoxycholate, 1 mmol/l PMSF, 1.5 µl/ml protease inhibitor cocktail), incubated on ice for 30 min, and centrifuged at maximum speed for 5 min. The supernatant of cell pellet was used as the protein source, and its concentration was determined by a BCA protein assay kit (Pierce, Rockford, IL) by using BSA as a standard protein.

Western blotting of CTGF and collagen IV. HAECs at 80% confluence were grown in 1% FBS media without growth factor for 24 h. Endothelial cells were then stimulated with LDL (50 µg/ml) for 24 h in the presence and absence of either a p42/p44mapk inhibitor (PD-98059, 40 µM, Calbiochem, La Jolla, CA), a p38mapk inhibitor (SB-203580, 10 µM, Calbiochem, La Jolla, CA), and/or the c-Jun NH2-terminal kinase (JNK) inhibitor (SP-600125, 30 µM, A. G. Scientific, San Diego, CA). Soluble protein (20–25 µg) obtained as described above was separated by SDS-PAGE (12% CTGF, 7.5% collagen IV) under reducing conditions and transferred onto nitrocellulose membrane (Bio-Rad, Hercules, CA) with semidry transfer method (Bio-Rad) at 20 V for 50 min. The membranes were immunoblotted with anti-CTGF polyclonal antibody (1:1,000 dilution, Fibrogen, South San Francisco, CA), anti-collagen IV polyclonal antibody (1:1,000 dilution, Southern Biotechnology, Birmingham, AL), and anti-actin antibody (1:1,000 dilution, Sigma, St. Louis, MO) overnight at 4°C followed by incubating the membranes in a secondary antibody conjugated to horseradish peroxidase (HRP). The immunoreactive bands were visualized using the chemiluminescence reagent Renaissance (NEN Life Science Products, Boston, MA) according to the procedure described by the supplier. Membranes were exposed to Kodak LS film, and bands were measured by densitometry and quantified by Scion Image program (Scion, Frederick, MA).

MAPK assays. HAECs at 80% confluence were serum starved by growing them in serum-free media for 24 h. Quiescent endothelial cells were stimulated with LDL (50 µg/ml) for 5 min. Twenty-five to thirty micrograms of protein were analyzed by SDS-PAGE, and the separated proteins were transferred to nitrocellulose membrane with the semitransfer method and immunoblotted with anti-phospho-p42/p44mapk polyclonal antibody that detects p42 and p44 MAPK only when activated by phosphorylation at Thr202 and Tyr204 (1:4,000 dilution, Cell Signaling Technology, Beverly, MA), anti-phospho-p38mapk antibody that detects p38 only when activated by phosphorylation at Thr180 and Tyr182 (1:1,000 dilution, Cell Signaling Technology), and anti-JNK antibody that detects JNK only when activated by phosphorylation at Thr183 and Tyr185 (1:1,000 dilution, Cell Signaling Technology). Immunoreactive bands were visualized using the chemiluminescence reagent Renaissance, according to the procedure described by the supplier. Membranes were exposed to Kodak LS film, and bands were measured by densitometry and quantified by Scion Image program.

Promoter activity analysis. The CTGF promoter containing –823 to +74 region linked to luciferase reporter gene was provided by Dr. Gary Grotendorst. CTGF promoter plasmid DNA was extracted by the alkaline lysis procedure and twice purified on cesium chloride-ethidium bromide gradients. For transient transfections, HAECs were seeded in 12-well plates (750,000 cells/well) in medium containing 2% FBS. The following day, cells at 50–80% confluence were switched to serum-free medium and transiently transfected with 0.5 µg plasmid DNA using FuGENE6 transfection reagent (Roche, Indianapolis, IN) according to the manufacturer’s recommendations. After transfection (16–19 h), HAECs were stimulated with LDL (50 µg/ml) and/or TGF-beta (5 ng/ml) for 24 h in the presence and absence of anti-TGF-beta neutralizing antibodies (anti-TGF-beta NA) (R&D Systems, Minneapolis, MN). Cells were then harvested in 100 µl lysis buffer/well (Promega, Madison, WI), and the protein concentration of each sample was determined using the BCA protein assay kit with BSA as a standard protein. Luciferase activity was measured in different protein concentrations (2.5–30 µg) to determine the linear range of luciferase activity.

RT-PCR. RNA was extracted from cells with the use of Qiagen kit (Valencia, CA) according to manufacturer’s protocol. The RNA was converted to cDNA by using MLV-RT (Promega, Madison, WI) according to the manufacturer’s protocol at 37°C for 1 h. The PCR reaction was carried out in 25-µl total volume containing 1x PCR buffer, 200 µM dNTP, 2 ng/µl of each primer, 5 µl of first-strand cDNA, and 1 U of Taq (Qiagen, Valencia, CA). The primers used for amplification of the human TGF-beta1 were 5'-tacctgaacccgtgttgctctc-3' and 5'-aacccgttgatgtccacttgc-3'; those used for human CTGF were 5'-cgggttaccaatgacaacgc-3' and 5'-taatggcaggcacaggtcttg-3'; those used for human collagen type IV were 5'-ggggttacaaggtgtcattggg-3' and 5'-tttccagggtagccagatgctc-3'; and those used for human beta-actin were 5'-gaaccctaaggccaaccgtg-3' and 5'-tggctatagaggtctttacgg-3'. The cycling conditions of human TGF-beta1 were denaturation at 95°C for 5 min followed by 25 cycles of 95°C for 45 s, 60.2°C for 45 s, and 72°C for 45 s; for human CTGF, 25 cycles of 94°C for 45 s, 59°C for 45 s, and 72°C for 45 s; for collagen type IV, 35 cycles of 94°C for 60 s, 60°C for 60 s, and 72°C for 60 s; for beta-actin, 25 cycles of 94°C for 45 s, 55°C for 45 s, and 72°C for 45 s. PCR reactions were visualized on a 1% agarose gel, photographs were taken, and densitometric analysis was performed with the use of the Scion Image program.

Real-time PCR. Real-time PCR was performed according to the recommendations of iCycler iQ Real-Time PCR Detection System (Bio-Rad). The primers used for amplification of the human CTGF were 5'-ttgcgaagctgacctggaagagaa-3' and 5'-agctcggtatgtcttcatgctggt-3', and of beta-actin were 5'-aatgtcgcggaggactttgattgc-3' and 5' aggatggcaagggacttcctgtaa-3'. For each target gene, a standard curve was established. This was achieved by performing a series of threefold dilutions of the gene of interest. Negative control was made using the same volume of RNase-free water instead of sample. The master mix was prepared as 2x SYBR Green Supermix (Bio-Rad) (12.5 µl), forward and reverse primer (0.25 µl, respectively), and distilled, deionized H2O (12 µl). For each well, 22 µl of master mix was loaded first, and then 3 µl of sample was added and mixed well to get total reaction volume of 25 µl. For plate setup, SYBR-490 was chosen as fluorophore. The plate was covered with a sheet of optical sealing film. PCR conditions were 95°C for 3 min, followed by 40 cycles of 95°C for 10 s, 60°C for 1 min for CTGF (for beta-actin, 58°C for 1 min), then 95°C for 1 min, 55°C for 1 min and 100 cycles of 55°C for 10 s. All of the reactions were done in duplicate. The correlation coefficient was between 0.98 and 1, and PCR efficiency was between 80 and 120%. CTGF mRNA levels were expressed relative to beta-actin mRNA.

TGF-beta1 protein level determination. HAECs were cultured in six-well plates (9.6 cm2/well). At 80% confluence, cells were serum starved by the changing of serum-free media within 24 h. Cells were then stimulated for 24 h with LDL (50 µg/ml), in the presence or absence of the p42/p44mapk inhibitor (PD-98059, 40 µM), the p38mapk inhibitor (SB-203580, 10 µM), and/or the JNK inhibitor (SP-600125, 30 µM) in exactly 1.5 ml of medium. TGF-beta1 protein levels were determined by colorimetric enzyme-linked immunosorbent assay kit (ELISA; R&D Systems) in the conditioned media that were activated by 1 N HCl (to measure both active and latent TGF-beta1) according to the manufacturer’s instructions and expressed as picograms per milliliter.

Statistical analysis. Data are expressed as means ± SE and were analyzed by ANOVA with post hoc corrections (Holm-Sidak method and Dunn’s method) and by Student’s t-test for paired and unpaired analysis. Differences were considered significant if P < 0.05.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Upregulation of CTGF by LDL. To examine if LDL will modulate the expression of CTGF, HAECs were stimulated with a known concentration of LDL (50 µg/ml) for various times to determine the maximal response. The concentration of LDL used is in the subphysiological range and is above those reportedly necessary to saturate the LDL receptor (~30 µg/ml). The protein levels of CTGF were measured by Western blots using specific anti-CTGF antibodies (1:1,000 dilution). The results shown in Fig. 1A demonstrate that LDL stimulation resulted in a significant increase in CTGF protein levels at 3 h with a peak response at 24 h (n = 4, P < 0.05).


Figure 1
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  		Fig. 1. Time response and dose response of low-density lipoprotein (LDL)-induced connective tissue growth factor (CTGF) levels. A: human aortic endothelial cells (HAECs) were stimulated with LDL (50 µg/ml) for various times (3, 6, 24 h). B: HAECs were treated with various concentrations of LDL (0, 10, 25, 50, 75, 100 µg/ml) for 24 h. Cell proteins were separated by SDS-PAGE (12%) and immunoblotted with an antibody against CTGF (1:1,000) and actin (1:1,000). Bar graph represents intensities of CTGF bands relative to actin expressed as percentage above control. Blots shown are representative of 4 experiments. *P < 0.05 vs. control (C).

 
To determine the optimal concentration of LDL for maximal response, HAECs were stimulated with various concentrations of LDL (0, 10, 25, 50, 75, and 100 µg/ml) for 24 h. The protein levels of CTGF were measured by Western blots using specific anti-CTGF antibodies (1:1,000 dilution). The bar graph (Fig. 1B) represents the intensities of the CTGF bands expressed as percent increase above control. The results shown in Fig. 1 indicate that LDL produced a concentration-dependent increase in the expression of CTGF with a peak response at 50 µg/ml (P < 0.01, n = 4). The protein levels of CTGF were expressed relative to actin protein levels measured in the same samples. Actin protein levels measured in the same samples were not different between control and LDL-treated cells.

Control experiments were performed to assess the effects of high-density lipoprotein (HDL) on CTGF levels in HAECs. HAECs were stimulated with HDL (50 µg/ml) for 24 h, and the protein levels of CTGF were measured by Western blots and expressed relative to actin protein levels measured in the same samples. The results demonstrated that there was no significant differences in CTGF protein levels between control and HDL-stimulated HAECs (861 ± 116 vs. 819 ± 123 CTGF/actin protein levels; control vs. HDL, respectively, P = 0.8074, n = 6).

We next examined if LDL will modulate the expression of CTGF mRNA levels. HAECs were treated with LDL at a concentration of 50 µg/ml for 24 h. RNA levels were extracted from the cells, and the CTGF mRNA levels were determined by RT-PCR. beta-Actin mRNA levels were also determined at the same time in the same cell extracts as control. CTGF mRNA levels were expressed relative to beta-actin mRNA, and the data are represented in Fig. 2A. Our findings indicate that LDL significantly increased the mRNA levels of CTGF compared with unstimulated cells (P < 0.002, n = 12).


Figure 2
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  		Fig. 2. LDL increases CTGF mRNA and protein levels in HAECs. A: HAECs were treated with 50 µg/ml of LDL for 24 h. RNA was isolated, and RT-PCR was performed with human CTGF-specific primers. Bar graph represents intensities of CTGF mRNA bands relative to beta-actin mRNA expressed as percentage above control. Blots shown are representative of 12 experiments. *P < 0.005 vs. control. B: HAECs transfected with 0.5 µg plasmid DNA containing the CTGF promoter were treated with 50 µg/ml of LDL for 24 h. Luciferase activity was measured in 5 µg protein, and all values were normalized against expression of plasmid without stimulation. *P < 0.05 vs. control; n = 4 experiments.

 
The RT-PCR study was repeated using real-time PCR. The CTGF mRNA levels were expressed relative to beta-actin mRNA levels. The results demonstrated that LDL stimulation resulted in a significant increase in CTGF mRNA compared with unstimulated cells (54.8 ± 3.69 vs. 45.9 ± 2.88; LDL vs. control, respectively, P < 0.01, n = 6). Thus the data generated from real-time PCR support our data generated using RT-PCR. The optimal time for LDL to illicit maximal response on CTGF mRNA expression was also determined. HAECs were stimulated with LDL (50 µg/ml) for various times (3, 6, 24 h), and CTGF mRNA levels were measured by real-time PCR. The results demonstrate that CTGF mRNA levels were increased 3 h after LDL-stimulation with a peak response at 24 h. CTGF mRNA levels for unstimulated control cells were 4.48 ± 0.51 compared with 6.93 ± 0.78 (LDL-3 h), 7.68 ± 0.67 (LDL-6 h), and 9.36 ± 0.21 (LDL-24 h) (P < 0.01, n = 3).

To determine whether LDL induces CTGF gene expression at the transcriptional level, CTGF promoter containing 805 bp linked to the luciferase reporter gene (0.5 µg pGL2 plasmid DNA) was transiently transfected into HAECs and stimulated with LDL (50 µg/ml) for 24 h. Luciferase activity was measured in cell lysates normalized for protein concentration. The results shown in Fig. 2B demonstrated that LDL significantly increased the CTGF promoter activity compared with basal activity [14,035 ± 1,701 vs. 7,923 ± 857 relative luciferase units (RLU)/s; LDL vs. control, respectively, P < 0.005, n = 4].

We next assessed the effects of LDL on plasmids (pGL2) lacking the CTGF promoter region. HAECs were transiently transfected with plasmids lacking the CTGF promoter followed by stimulation with LDL (50 µg/ml) for 24 h. Luciferase activity was measured in cell lysates normalized for protein concentrations. The results demonstrated that no difference in luciferase activity was detected between control and LDL-stimulated cells (22.2 ± 1.7 vs. 21.8 ± 2.0 RLU/s; control vs. LDL-treated cells, respectively, P = 0.8727, n = 6).

These findings provide the first evidence that LDL can upregulate the expression of CTGF in HAECs.

Induction of TGF-beta by LDL. To investigate whether LDL will stimulate the production of TGF-beta protein levels, HAECs were treated with LDL (50 µg/ml) for 24 h, and TGF-beta protein levels were measured in the conditioned media by ELISA (R&D Systems). The results shown in Fig. 3A demonstrate that LDL treatment resulted in a significant increase in the production of TGF-beta compared with unstimulated cells (P < 0.005, n = 10).


Figure 3
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  		Fig. 3. LDL stimulates transforming growth factor-beta (TGF-beta) mRNA and protein levels in HAECs. A: HAECs were stimulated by LDL (50 µg/ml) for 24 h. TGF-beta1 protein levels in the conditioned media were measured by ELISA kit assay. Values are means ± SE of TGF-beta1 levels expressed in pg/ml. *P < 0.005 vs. control; n = 10 experiments. B: HAECs were treated with 50 µg/ml of LDL for 24 h. RNA was isolated, and RT-PCR was performed with human TGF-beta-specific primers. Bar graph represents intensities of TGF-beta mRNA bands relative to beta-actin mRNA expressed as percentage above control. Blots shown are representative of 12 experiments.*P < 0.005 vs. control.

 
We next assessed whether LDL will stimulate the expression of TGF-beta mRNA levels. HAECs were treated with LDL at a concentration of 50 µg/ml for 24 h, and TGF-beta mRNA levels were determined by RT-PCR and expressed relative to beta-actin mRNA levels measured in the same cell extracts. The results shown in Fig. 3B demonstrated that LDL significantly increased the mRNA levels of TGF-beta compared with unstimulated HAECs (P < 0.005, n = 12).

Induction of collagen IV by LDL. Collagen I and IV are the predominant matrix proteins of the atherosclerotic plaque of diabetic patients, and collagen IV is the main collagen, produced by endothelial cells. Because matrix deposition is the hallmark of atherosclerosis, it is crucial to determine whether LDL will influence the deposition of matrix proteins in cells prone to injury. In this regard, we examined if LDL will modulate the expression of collagen IV mRNA as well as protein levels. HAECs were treated with LDL at a concentration of 50 µg/ml for 24 h. RNA levels were extracted from the cells, and the collagen IV mRNA levels were determined by RT-PCR. beta-Actin mRNA levels were also determined at the same time in the same cell extracts as control. Collagen IV mRNA levels were expressed relative to beta-actin mRNA, and the data are represented in Fig. 4A. Our findings indicate that LDL significantly increased the mRNA levels of collagen IV compared with unstimulated cells (P < 0.0005, n = 12). Collagen IV protein levels in cell extracts were measured by Western blots by using specific anti-collagen IV antibodies (1:1,000 dilution). Collagen IV protein levels were expressed as percentage of control, and the data are represented in Fig. 4B. The results demonstrate that LDL significantly increased the protein levels of collagen IV compared with control (P < 0.001, n = 13). The levels of the structural protein actin were also measured in the same cell extracts as control, to ensure equal loading of the samples into the gel, and were not different between control and LDL-treated cells.


Figure 4
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  		Fig. 4. LDL stimulates collagen IV mRNA and protein levels in HAEC. A: HAECs were treated with 50 µg/ml of LDL for 24 h, and collagen IV mRNA levels were measured by RT-PCR. Bar graph represents intensities of collagen IV mRNA bands relative to beta-actin mRNA expressed as percentage above control. Blots shown are representative of 12 experiments.*P < 0.005 vs. control. B: HAECs were stimulated with 50 µg/ml of LDL for 24 h. Cell proteins were separated by SDS-PAGE (7.5%) and immunoblotted with an antibody against collagen IV (1:1,000) and actin (1:1,000). Bar graph represents intensities of collagen IV bands relative to actin expressed as percentage above control. Blots shown are representative of 13 experiments. *P < 0.0005 vs. control. C: quiescent HAECs were stimulated with 50 µg/ml of LDL for 24 h. Conditioned media from cells were collected. Equal volumes (28 µl/lane) of conditioned media were separated by SDS-PAGE (7.5%) and immunoblotted with an antibody against collagen IV (1:1,000). Bar graph represents intensities of collagen IV bands expressed as percentage above control. Blots shown are representative of 22 experiments. *P < 0.001 vs. control.

 
The levels of collagen IV protein in the conditioned media were also measured. The results shown in the Fig. 4C demonstrate that LDL-treated HAECs produced a significantly higher level of collagen IV in their media than unstimulated cells (2,264.5.6 ± 274.3 vs. 1,310.8 ± 179; LDL vs. control, respectively, P < 0.005, n = 22).

Induction of CTGF by TGF-beta. To explore the cellular mechanisms through which LDL regulates the expression of CTGF in HAECs, we examined whether CTGF is a downstream target for regulation by TGF-beta. The results shown in Fig. 5A demonstrate that HAECs stimulated with TGF-beta (5 ng/ml) for 24 h significantly increased the promoter activity of CTGF compared with unstimulated cells (16,265.13 ± 2,605.7 vs. 7,923.25 ± 856.8 RLU/s; TGF-beta vs. control, respectively, P < 0.005, n = 4). Figure 5B shows the protein levels of CTGF measured in HAECs treated with different concentrations of TGF-beta (0, 10, 20 ng/ml) for 24 h. The levels of the structural protein actin were also measured in the same cell extracts as control to ensure equal loading of the samples into the gel. The results demonstrate that TGF-beta resulted in a concentration-dependent increase in the expression of CTGF (P < 0.005, n = 10). This increase in CTGF protein levels induced by TGF-beta was eliminated in the presence of anti-TGF-beta NA (Fig. 5B). Actin protein levels were not different between control and TGF-beta-treated cells. These findings indicate that TGF-beta is upstream of CTGF and may be involved in modulating the expression of CTGF.


Figure 5
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  		Fig. 5. Induction of CTGF by TGF-beta. A: HAECs transfected with 0.5 µg plasmid DNA containing the CTGF promoter were treated with 5 ng/ml of TGF-beta for 24 h. Luciferase activity was measured in 5 µg protein, and all values were normalized against expression of plasmid without stimulation. *P < 0.05 vs. control; n = 4 experiments. B: HAECs were treated with 10 and 20 ng/ml of TGF-beta for 24 h in presence or absence of 5 µg/ml of neutralizing anti-TGF-beta antibody (TGF-beta NA). Cell proteins were separate by SDS-PAGE (12%) and immunoblotted with an antibody against CTGF (1:1,000) and actin (1:1,000). Bar graph represents intensities of CTGF bands relative to actin expressed as percentage above control. Blots shown are representative of 10 separate experiments. *P < 0.005 vs. control. #P < 0.001 vs. TGF-beta1.

 
Role of TGF-beta in LDL-induced CTGF expression. To assess whether the induction of CTGF we observed in response to LDL is mediated via autocrine activation of TGF-beta, endothelial cells were treated with LDL (50 µg/ml) for 24 h in the presence and absence of anti-TGF-beta NA (5 and 10 µg/ml), concentrations shown to neutralize TGF-beta action (12). The promoter activity of CTGF gene was measured with the luciferase reporter vector, and the protein levels of CTGF and actin were measured by Western blots. The results shown in Fig. 6A demonstrates that LDL stimulation resulted in a significant increase in the promoter activity of CTGF compared with unstimulated control cells (P < 0.002, n = 6). However, in the presence of anti-TGF-beta NA, the LDL-induced increase in the promoter activity of CTGF was significantly reduced (P < 0.03, n = 6, Fig. 6A). Anti-TGF-beta NA did not significantly influence CTGF promoter basal activity. The results shown in the Fig. 6B demonstrate that LDL once again produced a significant increase in CTGF protein levels (P < 0.005 vs. control, n = 4). This increase in CTGF in response to LDL was significantly inhibited by the anti-TGF-beta NA (P < 0.05 vs. LDL). Anti-TGF-beta NA did not significantly influence the basal protein levels of CTGF. Mouse nonimmune serum had no significant effect on LDL-induced CTGF expression in HAECs (Fig. 6C). These findings suggest that TGF-beta signaling contributes to LDL stimulation of CTGF expression in endothelial cells. Our findings are in support of previous reports showing that LDL stimulates collagen expression in human mesangial cells via activation of TGF-beta (27).


Figure 6
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  		Fig. 6. LDL induces CTGF expression via autocrine activation of TGF-beta. A: HAECs transfected with 0.5 µg plasmid DNA containing the CTGF promoter were treated with 50 µg/ml of LDL for 24 h in the presence and absence of anti-TGF-beta NA (5 and 10 µg/ml). Luciferase activity was measured in 5 µg protein, and all values were normalized against expression of plasmid without stimulation. *P <0.002 vs. control, #P < 0.03 vs. LDL; n = 6 experiments. B: HAECs were stimulated with 50 µg/ml of LDL for 24 h in presence or absence of 5 and 10 µg/ml of anti-TGF-beta NA. Equal amounts of proteins were resolved on SDS-PAGE (12%) and immunoblotted with an antibody against CTGF (1:1,000) and actin (1:1,000). Bar graph represents intensities of CTGF bands relative to actin expressed as percentage above control. Blots shown are representative of 4 experiments. *P < 0.005 vs. control, #P < 0.05 vs. LDL. C: HAECs were stimulated with LDL (50 µg/ml) for 24 h in the presence and absence of mouse nonimmune serum (NS, 5 µg/ml). Bar graph represents intensities of CTGF bands relative to actin expressed as percentage above control. Blots shown are representative of 4 experiments. *P < 0.05 vs. control.

 
Phosphorylation of MAPK pathway by LDL. To determine whether LDL will activate members of the MAPK family, HAECs were treated with LDL (50 µg/ml) for 5 min. Cytosolic proteins were analyzed by SDS-PAGE, transferred to nitrocellulose membranes, and immunoblotted with either anti-phospho-p42/p44mapk antibodies (1:4,000 dilution) or anti-phospho-p38mapk (1:1,000 dilution) and/or anti-phospho-JNK antibodies (1:1,000 dilution). The membranes were also immunoblotted with total nonphosphorylated p42/p44mapk, p38mapk, and JNK antibodies to ensure equal amounts of MAPK loaded into the gels.

The results shown in Fig. 7, A–C, demonstrate that LDL significantly increased the phosphorylation of p42/p44mapk (P < 0.0001, n = 12), p38mapk (P < 0.0001, n = 12), and of JNK (P < 0.0002, n = 12). Total p42/p44mapk, p38mapk, and JNK were not different between control and LDL-treated cells.


Figure 7
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  		Fig. 7. Phosphorylation of MAPK pathway by LDL. Quiescent HAECs were stimulated with 50 µg/ml of LDL for 5 min. Cell proteins were separated by SDS-PAGE (12%) and immunoblotted with a polyclonal antibody against phospho (P)-p44/42 (1:4,000) and total (T) p44/42 (1:4,000) (A), phospho-p38 (1:1,000) and total p38 (1:4,000) (B), and phospho-JNK (1:1,000) and total JNK (1:4,000) (C). Blots shown are representative of 12 experiments. Bar graph represents intensities of phospho-MAPK relative to total MAPK expressed as percent phosphorylation above control. Bars represent means ± SE of 12 experiments. *P < 0.0005 vs. control.

 
Role of MAPK pathway in LDL-induced TGF-beta production. To explore whether the MAPK pathway modulates the production of TGF-beta in response to LDL, we measured TGF-beta protein levels in HAECs treated with LDL (50 µg/ml) for 24 h in the presence and absence of membrane-permeable inhibitors of p42/p44mapk (PD-98059, 40 µM), p38mapk (SB-203580, 10 µM), and JNK (SP-600125, 30 µM). The results shown in Fig. 8 demonstrate that TGF-beta protein levels were modestly reduced by the ERK inhibitor PD-98059 (P < 0.005, n = 8). On the other hand, the JNK inhibitor SP-600125 markedly reduced the increased production of TGF-beta protein levels in response to LDL (P < 0.005, n = 7), whereas selective inhibition of p38mapk by SB-203580 did not have a significant effect. These findings demonstrate that LDL utilizes the ERK and JNK pathway to modulate the production of TGF-beta in HAECs.


Figure 8
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  		Fig. 8. Role of MAPK in LDL-induced TGF-beta production. HAECs were stimulated with LDL (50 µg/ml) for 24 h in presence or absence of p44/42mapk inhibitor PD-98059 (PD; 40 µM) (A), p38mapk inhibitor SB-203580 (SB; 10 µM) (B), and JNK inhibitor SP-600125 (SP; 30 µM) (C). Conditioned media were collected and activated to determine the total level of TGF-beta1 protein by using colorimetric ELISA kit assay. Values are means ± SE of TGF-beta1 levels expressed in pg/ml. *P <0.05 vs. LDL; n = 7–8 experiments.

 
Role of MAPK in LDL-induced CTGF production. To explore whether the MAPK pathway modulates the production of CTGF in response to LDL, we measured CTGF protein levels in HAECs treated with LDL (50 µg/ml) for 24 h in the presence and absence of membrane-permeable inhibitors of p42/p44mapk (PD-98059, 40 µM), p38mapk (SB-203580, 10 µM), and JNK (SP-600125, 30 µM). The results shown in Fig. 9 demonstrate that LDL once again produced a significant increase in CTGF protein levels compared with unstimulated cells. However, in the presence of the p44/42mapk inhibitor PD-98059 and/or the JNK inhibitor SP-600125, the increase in CTGF protein levels in response to LDL was significantly suppressed (P < 0.01, n = 7–8). On the other hand, inhibition of the p38mapk by SB-203580 did not alter the expression of CTGF in response to LDL. Actin protein levels measured at the same time and in the same cell extracts were not significantly different among the groups.


Figure 9
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  		Fig. 9. Effect of MAPK inhibitors on LDL-induced CTGF levels. A: HAECs were stimulated with LDL (50 µg/ml) for 24 h in presence or absence of p44/42mapk inhibitor PD-98059 (40 µM) for 45 min; equal amounts of proteins were resolved on SDS-PAGE (12%) and immunoblotted with an antibody against CTGF (1:1,000) and actin (1:1,000). Bar graph represents intensities of CTGF bands relative to actin expressed as percentage above control. Blots shown are representative of 8 experiments. *P < 0.05 vs. control, #P < 0.01 vs. LDL. B: HAECs were stimulated by LDL (50 µg/ml) for 24 h in presence or absence of p38mapk inhibitor SB-203580 (10 µM) for 45 min. Blots shown are representative of 8 experiments. *P < 0.05 vs. control. C: HAECs were stimulated by LDL (50 µg/ml) for 24 h in presence or absence of JNK inhibitor SP-600125 (30 µM) for 45 min. Blots shown are representative of 7 experiments. *P < 0.05 vs. control, #P < 0.01 vs. LDL.

 
This is the first demonstration that the induction of CTGF by LDL in endothelial cells is mediated via activation of the MAPK pathway.

LDL increases collagen IV via JNK-dependent pathway. To evaluate whether activation of the MAPK pathway modulates the increase in collagen IV in response to LDL, endothelial cells were pretreated with the p42/p44mapk (PD-98059, 40 µM), p38mapk (SB-203580, 10 µM), and JNK (SP-600125, 30 µM) inhibitors for 45 min, followed by LDL stimulation for 24 h. The results shown in Fig. 10 demonstrate that LDL produced a significant increase in collagen IV protein levels in the conditioned media compared with unstimulated cells. This increase in collagen IV protein levels was significantly reduced by the JNK inhibitor SP-600125 (P < 0.05, n = 6). However, selective inhibition of p42/p44mapk by PD-98059 and of p38mapk by SB-203580 did not significantly alter the increased production of collagen IV protein levels in response to LDL stimulation. These findings demonstrate that LDL utilizes the JNK pathway to modulate the production of collagen IV in HAECs.


Figure 10
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  		Fig. 10. Role of MAPK pathway in LDL-induced collagen IV expression. A: HAECs were stimulated with LDL (50 µg/ml) for 24 h in presence or absence of p44/42mapk inhibitor PD-98059 (40 µM) for 45 min. Conditioned media were collected. Equal volumes (28 µl/lane) of conditioned media were separated on SDS-PAGE (7.5%) and immunoblotted with an antibody against collagen IV (1:1,000). Bar graph represents intensities of collagen IV bands expressed as percentage above control. Blots shown are representative of 6 experiments. *P < 0.05 vs. control. B: HAECs were stimulated by LDL (50 µg/ml) for 24 h in presence or absence of p38mapk inhibitor SB-203580 (10 µM) for 45 min. Blots shown are representative of 10 experiments. *P < 0.05 vs. control. C: HAECs were stimulated by LDL (50 µg/ml) for 24 h in presence or absence of JNK inhibitor SP-600125 (30 µM) for 45 min. Blots shown are representative of 6 experiments. *P < 0.05 vs. control, #P < 0.05 vs. LDL.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
In the present study we have demonstrated that LDL exerts a significant effect on the expression of CTGF and collagen IV in endothelial cells. We have shown that LDL increases CTGF promoter activity, the mRNA and the protein levels of CTGF, TGF-beta, and collagen IV in endothelial cells. This effect of LDL was mediated via activation of the MAPK pathway.

The risk for atherosclerosis is augmented when plasma LDL-cholesterol levels are increased, and many interventional studies have demonstrated that pharmacological treatment of coronary-prone patients with lipid-lowering agents reduces coronary events and total mortality (14, 36, 38). An important early event in the initiation of atherosclerosis is the increased uptake of monocytes into the intima where they differentiate into macrophages and ingest modified forms of LDL to become foamy macrophages, which give rise to "fatty streaks," the precursor lesion that subsequently leads to development of atherosclerosis (11, 21, 23, 32). Consequently, much attention is now focused on understanding the etiology of the fatty streak and the mechanisms by which LDL can affect components of the atherosclerotic process. However, much less is known about the cellular and molecular mechanisms by which native LDL activates endothelial cells to influence atherogenesis.

Although hyperlipidemia is now considered a risk factor for the progression of atherosclerosis, the initiating and sustaining signaling pathways that link hyperlipidemia to atherosclerosis are not fully realized. Accumulation of LDL within the intima may activate endothelial cells to produce various cytokines and growth factors that may alter endothelial cell behavior by inducing cell proliferation, migration, invasive capacity, and tubulogenesis. In this regard our findings demonstrate that the protein levels of TGF-beta, CTGF, and collagen IV were induced in endothelial cells in response to LDL challenge. Inhibition of TGF-beta activity with neutralizing antibodies partially prevented the rise in CTGF, thus demonstrating that the increase in CTGF protein levels in response to LDL was mediated via activation of TGF-beta-dependent and -independent mechanisms.

Even though TGF-beta has long been regarded as a major driving force in many progressive fibrotic diseases, attention has recently focused on the role of CTGF as a profibrotic factor (25). CTGF, a newly described factor that promotes ECM deposition and fibrosis in many tissues, appears to act downstream of TGF-beta to induce ECM production (33). Several studies demonstrated CTGF is an important mediator in the pathogenesis of atherosclerosis (2, 15, 29, 31). CTGF in endothelial cells is upregulated by lysophosphatidic acid, sphingosine-1-phosphate, and platelets (29). However, in the present study we demonstrate for the first time that LDL can directly and acutely upregulate CTGF protein levels in endothelial cells. Our findings indicate that the upregulation of CTGF by LDL could be mediated through dependent or independent mechanisms involving autocrine activation of TGF-beta.

It is widely accepted that LDL stimulation results in the activation of members of the MAPK family, and activation of this pathway is known to be important in regulating gene expression and endothelial cell growth and function (30). The MAPK are a family of serine-threonine protein kinases that are activated in response to a variety of extracellular stimuli. ERK (p42/44mapk), p38mapk, and JNK constitute three major subfamilies of MAPK that appear to mediate cellular responses, including proliferation, differentiation, and apoptosis (43). ERK plays a major role in cell proliferation and differentiation, as well as in survival mediated by various growth factors (24). On the other hand, p38mapk and JNK are activated by various inflammatory cytokines and environmental stressors, and they play important roles in apoptosis and cytokine production (24). To elucidate which member of the MAPK family may be responsible for the LDL-induced increase in CTGF and collagen IV, we elected to study the role of p42/44mapk, p38mapk, and JNK in HAECs. Our findings indicate that inhibition of JNK by SP-600125 suppressed the expression of TGF-beta, CTGF, and collagen IV levels in response to LDL stimulation, whereas inhibition of the p42/44mapk by PD-98059 resulted in suppression of LDL-induced expression of CTGF. On the other hand, blockade of the p38mapk pathway by SB-203580 did not significantly alter the expression of TGF-beta, CTGF, or collagen IV in response to LDL. This finding implicates JNK as a key player in modulating the signals through which LDL promotes TGF-beta, CTGF, and collagen IV expression in HAECs. Furthermore, it is of interest to note that inhibition of basal JNK activity also reduced the basal production of TGF-beta, thus implicating a key role for JNK kinase in modulating the production of TGF-beta in HAECs. Other studies have implicated JNK in TGF-beta-induced CTGF mRNA expression in human lung fibroblasts, whereas p38mapk and p42/44mapk pathways were implicated in TGF-beta-induced extracellular matrix deposition (8, 16, 41). We also found JNK is a key player in modulating the expression of TGF-beta, CTGF, and collagen I in response to LDL stimulation in rat mesangial cells (38).

Although the initiating and sustaining signals that link LDL to CTGF expression are not yet fully defined, several possibilities may exist. Our data demonstrate that LDL stimulates JNK activation, and JNK has been shown to bind and phosphorylate the DNA binding protein c-Jun and increase its transcriptional activity (24). c-Jun is a component of the activator protein-1 (AP-1) transcription complex, which is an important regulator of gene expression (20). In this regard, the autoinduction of TGF-beta has been shown to be mediated via activation of the AP-1 complex (22). Once activated, TGF-beta can induce CTGF expression via the Smad pathway (7). In fact, a Smad response element is present on the promoter of the CTGF gene (26).

In summary, we demonstrate that the expression of CTGF and collagen IV in endothelial cells are upregulated by LDL and that this regulation is mediated via autocrine activation of TGF-beta. The results also implicate JNK as a key player in modulating the expression of TGF-beta, CTGF, and collagen IV in response to LDL stimulation. The data also point to a potential novel pathway through which lipoproteins may promote atherosclerotic injury.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by National Institutes of Health Grants DK-46543 and HL-55782 (to A. A. Jaffa).


   ACKNOWLEDGMENTS
 
We thank Dr. William Usinger (FibroGen, South San Francisco, CA) for the generous gift of anti-CTGF antibodies.


   FOOTNOTES
 

Address for reprint requests and other correspondence: A. A. Jaffa, Dept. of Medicine, Div. of Endocrinology-Diabetes-Medical Genetics, Medical Univ. of South Carolina, 114 Doughty St., P.O. Box 250776, Charleston, SC 29425 (e-mail: jaffaa{at}musc.edu)

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.


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