Acidic and Basic Fibroblast Growth Factors Suppress Transcriptional Activation of Tissue Factor and Other Inflammatory Genes in Endothelial Cells
Abstract Tissue factor (TF) is a transmembrane receptor that serves as a cofactor for factor VIIa and initiates the extrinsic pathway of blood coagulation. Under normal physiological conditions, TF is expressed in extravascular and perivascular cells but not in vascular endothelial cells and monocytes. TF can be induced in these cells by inflammatory regulators and other stimulators, such as LPS, thrombin, oxidized lipoproteins, and certain growth factors. An earlier study showed that growing primary cultures of human umbilical vein endothelial cells (HUVECs) with endothelial cell growth supplement (ECGS) and heparin had impaired the ability of monolayers to express surface membrane TF activity after perturbation. The mechanism by which ECGS suppressed TF activity was not known. In the present study, we investigated the effect of recombinant acidic and basic fibroblast growth factors (aFGF and bFGF) on the induction of TF in a HUVEC cell line and a fibroblast cell line. Both aFGF and bFGF suppressed the phorbol myristate acetate–induced expression of TF in endothelial cells but not the serum-induced expression of TF in fibroblast cells. Diminished expression of the cell surface TF activity observed in endothelial cells grown with aFGF or bFGF was due to the accumulation of a lower number of TF mRNA transcripts. TF mRNA stability was not altered in HUVECs grown with aFGF or bFGF. Nuclear run-on experiments revealed that the transcription of TF and several other genes that play an important role in inflammation and angiogenesis was reduced in the endothelial cells that were cultured with aFGF or bFGF. The diminished expression of TF may be part of a generalized response of endothelial cells to FGF that facilitates migration of endothelial cells during angiogenesis.
- Received August 23, 1996.
- Accepted October 15, 1996.
Tissue factor, a cell surface glycoprotein, serves as the cellular receptor (cofactor) for coagulation factor VIIa, and the TF-VIIa complex initiates the coagulation cascade.1 Proper regulation of TF expression is critical for the hemostatic system to maintain a balance between promotion and inhibition of blood clotting. TF is expressed by many cell types that constitute the hemostatic “envelope,” but it is not normally expressed in cells that contact blood, such as monocytes and endothelial cells.2 3 TF can be induced in these cells, to variable degrees, when they are exposed to perturbing stimuli such as endotoxin, thrombin, TNF, IL-1, activated complement factors, and immune complexes.4
Although much is known about the mechanism of TF induction in monocytes and endothelial cells,5 little is known about the downregulation of TF induction in these cells. It was shown that LPS induction of TF expression in monocytes was inhibited by agents that elevate intracellular levels of cAMP, including the prostacyclin analogues6 dibutyryl cAMP and pentoxifylline7 and also by the immunosuppressive cytokines IL-4 and IL-10.8 9 Since the TF mRNA stability was not altered by these agents, it was assumed that these agents suppressed TF mRNA by reducing the rate of TF gene transcription.7 8 9 Ishii et al10 demonstrated that treatment with retinoic acid decreased the expression of TF activity and TF mRNA in HUVECs subsequently exposed to TNF-α, but the mechanism of the suppression was not investigated.
In an earlier study it was shown that HUVECs cultured with ECGS and heparin had impaired ability to express TF activity when stimulated with thrombin and other agonists.11 Further studies revealed that TF mRNA concentration in thrombin-treated cells grown with ECGS/heparin was about 7.5-fold less than those grown without ECGS/heparin.12 Since aFGF was the primary growth factor component in ECGS, it was believed that aFGF was responsible for the suppression of TF induction observed in these studies.12 However, this assumption was not tested in the earlier study. Furthermore, no information was available on the mechanism involved in downregulation of TF induction in HUVECs grown with ECGS.
The present study was the first to test whether recombinant aFGF and/or bFGF could suppress the induction of TF in endothelial cells and delineate the mechanism of the suppression. In addition, we have also analyzed the effect of aFGF and bFGF on the transcription rates of several early genes involved in the inflammatory process. The results show that both aFGF and bFGF suppress the induction of TF and that this suppression stems from the reduced rate of TF gene transcription. The suppressive effect of aFGF and bFGF extends to the transcription of several other genes that may play a role in inflammation, angiogenesis, and atherosclerosis.
Recombinant aFGF, bFGF, and culture media F-12 and M-199 were obtained from GIBCO-BRL, Life Technologies; ECGS, fatty acid–free BSA, LPS, TNF-α, and PMA from Sigma Chemical Co; FBS, trypsin-versene mixture, penicillin-streptomycin, and L-glutamine from Bio-Whittaker; [α-32P]UTP (3000 Ci/mmol) and [α-32P]dCTP (3000 Ci/mmol) from DuPont NEN; nitrocellulose membrane BA85 from Schleicher & Schuell; and TRI reagent from Molecular Research Center Inc. Most of the molecular biology–grade chemicals were obtained from either Boehringer Mannheim or United States Biochemicals.
Human TF cDNA was provided by Dr J.H. Morrissey, Oklahoma Medical Research Foundation, Oklahoma City. Human TFPI cDNA was provided by Dr W. Novotny, University of California, San Diego, and IL-8 cDNA was obtained from Dr E. Miller, University of Texas Health Center at Tyler. cDNA probes for fibronectin, TPA, PAI-2, thrombomodulin, collagen type III α-1, thrombospondin, and aFGF were obtained from American Type Culture Collection.
The HUVEC line CRL-1730 was purchased from American Type Culture Collection and maintained at 37°C under 5% CO2 in T-75 flasks in F-12 medium supplemented with 10% FCS, 1% penicillin-streptomycin, 40 μg/mL ECGS, and 15 U/mL heparin. The cells were subcultured by first detaching the cells with trypsin solution and replating them in 24-well culture dishes or in T-75 flasks in the presence or absence of ECGS, aFGF, and bFGF. Heparin 15 U/mL was included in the medium if the medium was supplemented with ECGS and FGF. For many experiments, the cells were first grown in the presence of ECGS/heparin to confluency and then maintained for 48 hours in medium not containing ECGS to obtain no growth factor control. Culture flasks and dishes were coated with human fibronectin (0.65 μg/cm2). A fibroblast cell line, WI-38, was cultured as described in an earlier publication.13
Induction of TF
Confluent endothelial cell monolayers were washed three times with F-12 medium. For induction of TF with thrombin, human α-thrombin (2.5 U/mL) was added to the medium containing 5 mg/mL BSA. For induction of TF with PMA, PMA was added at a final concentration of 10 ng/mL to the medium containing 10% vol/vol FBS. At specific time periods, the medium was removed, and the monolayers were washed twice with F-12 medium or buffer A (10 mmol/L HEPES, 0.15 mol/L NaCl, 4 mmol/L KCl, and 11 mmol/L glucose, pH 7.5) and processed further without delay (see below).
Measurement of TF Procoagulant Activity
Cell surface TF activity was measured as the ability of monolayers to support activation of factor X with the addition of VIIa and CaCl2. Briefly, after two washes with buffer A, monolayers in a 24-well culture dish were overlaid with 0.25 mL of buffer B (buffer A containing 5 mg/mL BSA and 5 mmol/L CaCl2) to which factor VIIa 0.5 μg/mL and factor X 10 μg/mL were added. At the end of 30 minutes, 25 μL of subsample was taken from the well and added to 50 μL of TBS/BSA buffer (50 mmol/L Tris-HCl, 0.15 mol/L NaCl, pH 7.5, and 1 mg/mL BSA) containing 25 mmol/L EDTA. The amount of factor Xa formed in each well was determined by transferring 50 μL of the above mixture to a 96-well microtiter plate and then adding 50 μL of 1.25 mg/mL Chromozym X to each well. The initial rate of color development in milli–optical density units per minute at 405 nm was recorded continuously with a microplate reader (Molecular Devices). The initial rate was converted to micrograms per milliliter of factor Xa from a standard curve prepared by adding 50 μL of Chromozym X to 50 μL of serial dilutions of a 1-μg/mL sample of purified human factor Xa. In initial experiments in which subsamples were removed at increasing times, it was found that the generation of factor Xa was linear up to a 2-hour reaction period.
RNA Isolation and Blotting
Total RNA was prepared from 5×106 to 6×106 cells by the acid phenol method using TRI reagent according to the manufacturer's technical bulletin. Total RNA (10 μg) was size fractionated by gel electrophoresis in 1% agarose/6% formaldehyde gels and transferred onto the nitrocellulose membrane by a capillary blot method. Northern blots were prehybridized at 42°C with a solution containing 50% formamide, 5×SSC, 50 mmol/L Tris-HCl, pH 7.5, 0.1% sodium pyrophosphate, 1% SDS, 1% polyvinylpyrrolidone, 1% Ficoll, 25 mmol/L EDTA, and 1% BSA and hybridized with 32P-labeled cDNA probes (106 cpm/mL) as described earlier.12 The cDNA probes were labeled with a nick translation kit. The filters were exposed to either DuPont NEF or Fuji RX x-ray film. Autoradiograms were analyzed with a Millipore Bio-Image laser scanner. In most experiments, a second set of autoradiograms was developed for a longer time period for better visualization of the bands.
Isolation of Nuclei and Run-on Transcription Assay
Nuclei from 4×106 to 6×106 cells were harvested by scraping, collected by centrifugation at 800g for 5 minutes at 4°C, and resuspended in a final volume of 1 mL of homogenization buffer (10 mmol/L HEPES, pH 8.0, containing 10 mmol/L MgCl2, 2 mmol/L DTT, 0.25 mol/L sucrose, and 0.1% Triton X-100). Cells were transferred to a 1-mL Wheaton homogenizer and disrupted by 20 slow strokes with “A”-type fitting on ice. The nuclei were separated by placing the cell homogenate onto a 3-mL cushion of 1.3 mol/L sucrose in homogenization buffer and centrifuging at 10 000 rpm for 15 minutes. The pellet was dissolved in 200 μL of glycerol buffer (40% glycerol, 50 mmol/L HEPES, pH 8.0, 5 mmol/L MgCl2, 0.1 mmol/L EDTA, and 2 mmol/L DTT) and either used immediately or frozen on dry ice and then stored at −80°C until used. Run-on assays were performed with [α-32P]UTP (3000 Ci/ mmol)–labeled RNA as described previously.16
Effect of aFGF, bFGF, and ECGS on Induction of Cell Surface TF Activity
Human endothelial cells were plated in a 24-well culture dish at a density of 100 000 cells/well and grown in control medium, in the medium supplemented with heparin alone, or in the medium supplemented with heparin plus ECGS, aFGF, or bFGF. After 48 hours, the medium was removed and the monolayers were perturbed for 6 hours with PMA to induce TF activity. Cell surface TF activity was measured as the ability of monolayers to support activation of factor X after the addition of VIIa and calcium ions. As shown in Fig 1⇓, the results confirmed the earlier observation made with primary HUVECs11 that the endothelial cells grown with ECGS/heparin and stimulated with PMA expressed reduced levels of cell surface TF activity compared with the similarly treated cells grown in the absence of ECGS/heparin. Furthermore, the results also demonstrated that growing the endothelial cells with aFGF/heparin or bFGF/heparin also markedly suppressed the induction of cell surface TF activity. Growing the endothelial cells in the medium containing heparin alone failed to suppress the induction of TF. Similar results were obtained in experiments in which thrombin was used as an agonist to induce TF activity in endothelial cells (data not shown).
In further experiments, we investigated the effect of varying concentrations of aFGF and bFGF on induction of TF activity in endothelial cells. The endothelial cells were grown in the presence of 2.5 to 40 ng/mL aFGF or bFGF in the presence of a fixed concentration of heparin (15 U/mL), and the confluent monolayers were perturbed with either thrombin or PMA. The results showed that a concentration of 2.5 ng/mL aFGF or bFGF was sufficient to suppress TF induction and a concentration of 10 ng/mL aFGF or bFGF maximally suppressed the induction of TF (data not shown).
To test whether the observed suppression of TF induction after a 6-hour treatment with perturbing stimuli in cells grown with aFGF or bFGF could be due to an altered induction pattern of cell surface TF activity, a time-course induction of cell surface TF activity was measured by treating the endothelial cells with PMA for various lengths of time. The results showed a similar pattern of TF induction in endothelial cells that were grown with or without aFGF. In both cases, only a minimal amount of TF activity was measured after a 2-hour stimulation; the activity peaked at about 8 hours and then declined to very low levels by 24 hours (Fig 2⇓). Similar results were obtained with endothelial cells grown with bFGF. These results indicate that suppression of TF induction in cells grown with aFGF and bFGF was not due to a difference in the time-course induction of TF activity.
Effect of aFGF and bFGF on Induction of TF mRNA
TF mRNA accumulation was measured in confluent endothelial cell monolayers that were grown in the presence of various concentrations of aFGF and bFGF and treated with PMA for 2 hours to induce TF mRNA. As shown in Fig 3⇓, both aFGF and bFGF exhibited a dose-dependent inhibitory effect on the induction of TF mRNA, and a concentration of 10 ng/mL aFGF or bFGF maximally suppressed TF mRNA accumulation. A 10-ng/mL concentration of aFGF and bFGF also inhibited thrombin-, LPS-, and TNF-α–induced expression of TF mRNA (Fig 4⇓). Endothelial cells that were grown in the absence or the presence of aFGF and bFGF and not stimulated expressed no measurable TF mRNA (data not shown). The time course of induction of TF mRNA was similar in the cells grown in the presence or the absence of aFGF and bFGF. The mRNA level for TF began to increase during the first hour of PMA treatment, peaked between 2 and 3 hours of PMA treatment, and then declined rapidly (Fig 5⇓). At all time points, the level of TF mRNA was fivefold to sevenfold lower in the cells grown with bFGF than in the cells grown in its absence, which is concordant with the cell surface expression of TF activity. A similar pattern of results was also obtained with the cells grown in aFGF/heparin (data not shown). In other experiments, it was found that the addition of aFGF or bFGF with PMA to the control endothelial cells failed to suppress the induction of TF mRNA in these cells. A 24- to 48-hour exposure to aFGF or bFGF was required for the maximal suppression of PMA-stimulated induction of TF mRNA in endothelial cells.
To test whether the suppressive effect of aFGF and bFGF on induction of TF is specific to endothelial cells or also applicable to other cell systems, we tested the effect of these growth factors on both the constitutive expression and serum induction of TF in a fibroblast cell line, WI-38. These cells express TF constitutively, but the levels could be depleted by serum starvation and then could be induced by the addition of serum. WI-38 cells were grown in the presence or the absence of aFGF and bFGF to confluency, and the expression of cell surface TF activity was measured. The results showed that both aFGF and bFGF failed to suppress constitutive expression of TF (data not shown). In other experiments, WI-38 cells were grown to confluency and then deprived of serum for 48 hours. During this period, the cells were maintained in the serum-free medium supplemented with aFGF or bFGF 10 ng/mL or none. TF mRNA was induced in these cells by the addition of 10% FCS to the medium for 3 hours. The data from these experiments, relative to basal constitutive TF RNA levels (assigned as 100%), were serum deprived, 45%; serum induced, 423%; serum deprived+aFGF, 56%; serum induced+aFGF, 343%; serum deprived+ bFGF, 93%; and serum induced+bFGF, 315% (mean of three independent experiments).
Stability of TF mRNA
To test whether the suppressive effect of aFGF and bFGF on induction of TF mRNA in endothelial cells is due to enhanced degradation of TF mRNA in the cells grown with these growth factors, we next investigated the effect of bFGF treatment on TF mRNA stability by inhibiting mRNA transcription with actinomycin D. The endothelial cells that were cultured with bFGF or the control medium containing no growth factors were incubated with PMA for 2 hours to induce TF mRNA, and then actinomycin D 5 μg/mL was added to inhibit the transcription. The disappearance of TF mRNA with time was analyzed by Northern blot analysis. As shown in Fig 6⇓, the accumulation of mature TF mRNA was decreased similarly in the endothelial cells that were grown with or without bFGF. The apparent half-life of TF mRNA was 44±14 minutes in the presence of bFGF and 51±16 minutes in the absence of bFGF (mean±SD, n=4).
Effect of aFGF and bFGF on the Transcription of TF and Other Inflammatory Responsive Genes
To determine whether the lower number of TF transcripts in PMA-stimulated endothelial cells that were treated with aFGF and bFGF was due to a decrease in transcriptional activation of the TF gene, nuclear run-on experiments were performed. The data demonstrated a minimal transcription of TF in unperturbed endothelial cells and a markedly enhanced transcription of the TF gene after PMA treatment. Furthermore, the data also showed that both aFGF and bFGF suppressed the induction of TF gene transcription (Fig 7⇓). Densitometry of autoradiographs from four experiments revealed that the transcription rate of the TF gene in cells grown with aFGF and bFGF was reduced by threefold and fivefold, respectively, compared with cells grown in their absence (Fig 7B⇓). Transcription of α-tubulin, TF pathway inhibitor, thrombomodulin, fibronectin, collagen, and aFGF were not affected by PMA treatment. Furthermore, the transcription of these genes was not altered whether the cells were grown in aFGF or bFGF. In contrast, the transcription of TPA, PAI-2, and IL-8 genes was markedly enhanced in the endothelial cells upon their stimulation with PMA. aFGF treatment minimally suppressed PMA-induced transcription of TPA (23%) and IL-8 (10%) while suppressing PAI-2 moderately (43%) (n=4). The suppression of transcription of these genes was much more pronounced in cells grown with bFGF. The transcriptions of TPA, PAI-2, and IL-8 were reduced by 52%, 76%, and 24%, respectively (n=4). Transcription of the thrombospondin gene was inhibited by PMA treatment and was further reduced by aFGF (20%) and bFGF (49%).
The present study is the first report to show that bFGF suppresses the induction of cell surface TF activity and accumulation of TF mRNA in HUVECs. The present data obtained with recombinant aFGF not only confirm the assumption made in an earlier study12 that aFGF, a growth component in ECGS, was responsible for the suppression of TF induction in primary cultures of HUVECs grown with ECGS but also extend to delineate the mechanism involved in the suppression. In many cases, growth factors were shown to induce TF expression in endothelial cells and fibroblasts.17 bFGF was shown to induce TF mRNA in 3T3 cells.18 In contrast, our present data show that bFGF and aFGF do not induce any measurable TF mRNA or TF activity in endothelial cells. The inability of aFGF and bFGF to induce TF expression in endothelial cells differs from that of another angiogenic growth factor, vascular endothelial growth factor, which was shown to induce TF expression in endothelial cells.19
A similar pattern of the induction and the decline in cell surface TF activity and TF mRNA accumulation after the PMA treatment in endothelial cells grown in the absence or the presence of aFGF or bFGF suggests that an accelerated degradation of TF mRNA may not be the reason for the suppression of TF activity observed in the cells grown with aFGF or bFGF. This was further supported by the observation that the rate of decay of TF mRNA after the arrest of transcription with actinomycin D was not affected by FGF treatment (Fig 6⇑). The apparent half-life of TF mRNA in PMA-treated endothelial cells that were cultured either in the presence or absence of FGF (44 and 51 minutes, respectively) was similar to the reported value of 48 minutes observed in PMA-perturbed endothelial cells.20 Nuclear run-on assays demonstrated that both aFGF and bFGF reduced the rate of TF gene transcription by threefold and fivefold, respectively. Thus, the present data establish that aFGF and bFGF suppress the expression of cell surface TF activity not by decreasing TF mRNA stability but rather by reducing the rate of TF gene transcription.
Induction of TF gene transcription in endothelial cells is mediated by the functional interaction between Fos-Jun and c-Rel-p65 heterodimers.5 Fos-Jun heterodimers bind to two AP-1 sites, and c-Rel-p65 heterodimers bind to a κB-like site in the TF promoter. Although Fos-Jun heterodimers bind to AP-1 sites in both unstimulated and stimulated endothelial cells, the binding is an absolute requirement in stimulated endothelial cells for c-Rel-p65 heterodimers to facilitate activation of TF gene transcription. Activation of NF-κB (c-Rel-p65) requires phosphorylation, dissociation, and proteolytic degradation of I-κB-α before nuclear translocation of the transcription factors.5 21 Blocking any one of the above steps, including Fos-Jun binding to AP-1, could downregulate the induction of TF expression. Since activation of both NF-κB and AP-1 is mediated through the protein kinase C pathway, an impairment in the protein kinase C pathway could affect both AP-1 binding and NF-κB binding.22 We are currently investigating at what step TF transcription is blocked in endothelial cells that were exposed to FGF.
The suppressive effect of aFGF and bFGF on TF expression appears to be specific to the induction of TF on endothelial cells. Growing a fibroblast cell line with aFGF or bFGF failed to suppress either the constitutive expression of cell surface TF activity or TF mRNA accumulation. Furthermore, both aFGF and bFGF also failed to impair the serum induction of TF in these cells. It was shown that both the basal expression of TF and PMA and serum induction of TF expression in human epithelial cells is controlled by a proximal enhancer containing three overlapping Sp1/Egr-1 binding sites.5 The inability of aFGF and bFGF to suppress either the basal expression or the serum-induced expression of TF in fibroblasts suggests that neither aFGF nor bFGF impairs the interaction of Sp1 and Egr-1 with their binding sites in the TF promoter.
To understand how FGF inhibits the induction of TF in endothelial cells requires an understanding of signal pathways that are involved in the induction of TF as well as FGF signaling pathways in endothelial cells. The binding of FGF to the high-affinity receptor leads to autophosphorylation of FGF receptors, followed by activation of phospholipase C-γ, Raf-1 kinase, and MAP kinases (reviewed in Reference 2323 ). In general, these events are immediate/early cellular events, require only a short exposure time to FGF, and do not require new protein synthesis. However, our data showing that a prolonged exposure (24 hours) to FGF is required for optimal suppression of TF induction suggest that new protein synthesis is required for FGF-mediated suppression of TF. The cellular responses that are elicited upon prolonged exposure to FGF remain largely unknown.
Both aFGF and bFGF are potent angiogenic agents that stimulate proliferation and migration of endothelial cells and other cell types. Evidence shows that exposure of HUVECs to aFGF and bFGF can modulate a number of functional responses, including the synthesis and degradation of several extracellular matrix proteins.24 25 26 Our present data show that aFGF and bFGF inhibited the transcription of TPA, PAI-2, and IL-8. It is speculated that a net decrease in the endothelial cell matrix may facilitate blood vessel migration and development.27 Thus, a decrease in PAI-1,24 PAI-2 (present data), and fibronectin25 by FGF could contribute to the promotion of angiogenesis. Furthermore, recent studies showed that exposure of endothelial cells to bFGF significantly inhibited cytokine-mediated increased expression of cell adhesion molecules, intracellular adhesion molecule (ICAM-1), vascular cell adhesion molecule (VCAM-1), and E-selectin.28 The induction of cell adhesion molecules in vascular endothelium by cytokines released from tumor cells could lead to enhanced interaction between leukocytes and endothelial cells. One could postulate that these cell-cell interactions coupled with cytokine-induced expression of TF in vascular endothelium could limit blood flow in capillaries and subsequently impede angiogenesis. If so, it is reasonable to hypothesize that one mechanism by which FGF promotes angiogenesis and subsequent tumor growth is by the suppression of a specific set of genes whose expression is enhanced in inflammation that could occur in the progression of tumor growth.
Selected Abbreviations and Acronyms
|ECGS||=||endothelial cell growth supplement|
|FGF||=||fibroblast growth factor (a, acidic; b, basic)|
|HUVEC||=||human umbilical vein endothelial cell|
|PAI||=||plasminogen activator inhibitor|
|PMA||=||phorbol 12-myristate 13-acetate|
|TFPI||=||tissue factor pathway inhibitor|
|TNF||=||tumor necrosis factor|
|TPA||=||tissue plasminogen activator|
This study was supported by grant HL-42813 from the National Heart, Lung, and Blood Institute (NHLBI). Dr Rao is a recipient of a Research Career Development Award from the NHLBI.
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