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(Arteriosclerosis, Thrombosis, and Vascular Biology. 1995;15:903-911.)
© 1995 American Heart Association, Inc.

Articles

-Thrombin Stimulates Urokinase Production and DNA Synthesis in Cultured Human Cerebral Microvascular Endothelial Cells

Marie A. Shatos; Thomas Orfeo; Jacqueline M. Doherty; Paul L. Penar; Desire Collen; Kenneth G. Mann

From the Departments of Surgery (M.A.S., T.O., J.M.D., P.L.P.) and Biochemistry (M.A.S., D.C., K.G.M.), University of Vermont College of Medicine, Burlington, Vt, and the Center for Molecular and Vascular Biology (D.C.), Leuven, Belgium.

Correspondence to Marie A. Shatos, Department of Biochemistry, Given Bldg, Rm C-401, Burlington, VT 05405.

	Abstract

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Abstract -Thrombin regulation of endothelial cell (EC) fibrinolysis has been documented by using endothelia derived from a number of anatomic locations but not with those derived from the human cerebral vasculature. In the present study, the fibrinolytic properties of human cerebral microvascular ECs and their regulation by -thrombin are delineated and contrasted with those of human umbilical vein and foreskin microvascular ECs. In cerebral ECs, -thrombin elicited a unique dose-dependent increase in urokinase production and DNA synthesis. Maximal stimulation, observed with 10 nmol/L -thrombin, resulted in a 30- to 50-fold increase in urokinase production and a concomitant fourfold increase in DNA synthesis; the increase in urokinase was reflected in higher steady-state levels of urokinase mRNA. The major urokinase product secreted is the single-chain form of the enzyme. No effect was observed with the addition of other proteases or catalytically inactive variants of -thrombin. A thrombin receptor agonist peptide upregulated urokinase production but had no effect on DNA synthesis, suggesting that fibrinolysis is mediated by the thrombin receptor but that proliferation is regulated by a different pathway. These findings suggest the possibility that the cerebral microvasculature may be a specialized region of the vascular system in which urokinase-type plasminogen activator, not tissue-type plasminogen activator, is the key catalyst of fibrin lysis when the brain responds to thrombotic events and that -thrombin may regulate repair of the cerebral microvascular system.

Key Words: thrombin • brain • endothelial cells • urokinase

	Introduction

Top Abstract Introduction Methods Results Discussion References

 
The vascular endothelium is a highly specialized tissue with diverse functions that are regulated by many hormones, cytokines, and growth factors. Vascular damage leads to focal activation of the coagulation cascade and to the generation of -thrombin during the formation of a blood clot. In addition to its roles in blood coagulation, -thrombin modulates the large-vessel–derived endothelial cell (EC) expression of numerous components, including the fibrinolytic proteins tissue-type plasminogen activator (TPA), urokinase-type plasminogen activator (UPA), and plasminogen activator inhibitor–1 (PAI-1).^{1 2 3} -Thrombin also causes various cells in culture to migrate and proliferate.^{4 5 6}

Most of our knowledge of EC function has been derived from studies of large-caliber vessels such as the umbilical vein and aorta. Limited knowledge is available regarding the influence of -thrombin on the microvasculature. The small vessels of the microcirculatory system, which constitute approximately 95% of the total EC surface in the human body, have not been extensively studied, nor have studies been reported that describe the interaction of thrombin with cultured human brain microvascular ECs. There is a significant incidence of hemorrhage in the cerebral vasculature after pharmacological intervention with TPA.^{7 8 9} It is not known whether this is a result of the cerebral vascular system architecture or a pharmacological response to thrombosis and fibrinolysis. In the present study, we compare the influence of -thrombin on cultured human cerebral microvascular EC (HCMEC), human umbilical vein EC (HUVEC), and human foreskin microvascular EC (HFMEC) fibrinolysis and proliferation.

	Methods

Top Abstract Introduction Methods Results Discussion References

 
Reagents and Supplies
Collagenase (type 1) was obtained from Worthington; rabbit anti–factor VIII/von Willebrand factor (vWf) and fluorescein-labeled goat anti-rabbit IgG were from Cappel Laboratories; fluorescein-labeled Ulex europaeus agglutinin–1 (UEA-1) lectin was from Vector Laboratories, Inc; antibodies to glial fibrillary acidic protein (GFAP) were from Boehringer Mannheim; medium M-199, calf serum, neomycin, nystatin, basal Eagle's medium, vitamins, amino acids, penicillin/streptomycin, L-glutamine, EDTA, trypsin, Hank's balanced salt solution, and RNA calibration mixture were from GIBCO/BRL; EC growth medium (EGM BulletKit containing basal medium and growth factors) was from Clonetics; fetal bovine serum (FBS) was from the Salzman Corp; [³H]thymidine was from ICN Radiochemicals; Pentex fatty acid-poor, fraction V bovine serum albumin (BSA) was from Miles Laboratories, Inc; BCA protein assay reagent was from Pierce; HEPES was from United States Biochemical Corp; 6-keto-prostaglandin F₁ (6-keto-PGF₁) enzyme immunoassay kits were from Cayman Chemical Co; and cycloheximide and all other reagents, unless otherwise specified, were obtained from Sigma Chemical Co.

Solutions
Modified medium M-199, pH 7.2 to 7.4, consisted of 9.87 g/L medium M-199 powder, 1% basal medium Eagle's vitamin solution (100x), 5 mmol/L glucose, 0.1 g/L neomycin (1120 U), 26.2 mmol/L NaHCO₃, 200 mmol/L L-glutamine, and 20% FBS. EC growth medium consisted of MCDB 131 powder, 10 ng/mL human recombinant epidermal growth factor, 2% FBS, 2.8 µmol/L hydrocortisone, 50 µg/mL gentamicin, 50 ng/mL amphotericin B, and 3.0 mg/mL bovine brain extract containing 10 µg/mL heparin. Serum-free medium consisted of modified medium M-199, 0.35% BSA, and 20 mmol/L HEPES, pH 7.2 to 7.4. Phosphate-buffered saline (PBS)–Tween 80 contained 137 mmol/L NaCl, 6 mmol/L Na₂HPO₄7H₂O, 2.7 mmol/L KCl, 1.5 mmol/L KH₂PO₄, pH 7.4, and 0.1% Tween 80. Lysing buffer contained 1% Triton-X 100 in PBS.

Cell Culture
HCMECs were obtained from fragments of cerebral cortex consisting exclusively of gray matter devoid of large blood vessels and assessed to be free from abnormal pathology. All tissue was obtained from the neurosurgical team at the Medical Center Hospital of Vermont, Burlington. HCMECs were isolated by mincing tissue and subjecting it to repeated cycles of collagenase digestion.¹⁰ The supernate containing the cerebral microvessels was then forced through a 70-µm mesh to separate the cells from large undigested tissue fragments. Cells were pelleted and seeded in medium M-199 supplemented with 20% heat-inactivated FBS, 100 µg/mL neomycin, 10 µg/mL amphotericin B, 20 U/mL nystatin, and 2 mmol/L L-glutamine. As cells began to proliferate, microvascular endothelium was further isolated by scraping contaminating cell types from the surface of the culture dish with a rubber policeman. Cells were identified as endothelial if they fulfilled the following four criteria: positive immunocytochemical staining for factor VIII/vWf antigen¹¹ ; binding of UEA-1 lectin, a specific human EC marker that was maintained throughout cell passage¹² ; negative staining for GFAP, which confirms that the cells were not of astrocytic origin¹³ ; and ability to produce prostacyclin (PGI₂), a normal secretory product of endothelium that contributes to EC thromboresistance.¹⁴ Immunocytochemical studies were conducted by using subconfluent cultures of HCMECs established on 2x2-cm² glass coverslips that were then fixed with methanol for 10 minutes at room temperature. For factor VIII/vWf antigen studies, HCMECs were incubated with anti–factor VIII/vWf antibody (1:50 dilution) in PBS overnight at 4°C. Following washing, the coverslips were incubated with fluorescein-labeled goat anti-rabbit IgG (1:25 dilution) for 1 hour at 37°C. HCMECs were incubated overnight at 4°C with UEA-1 lectin (1:50 dilution) in PBS. Both were examined by using fluorescent microscopy (Zeiss IM35 inverted microscope with fluorescein excitation and emission wavelengths of 340 and 510 nm, respectively). PGI₂ production by confluent HCMECs was assessed in serum-free medium (M-199 containing 0.35% BSA and 20 mmol/L HEPES, pH 7.4) at 2, 4, 8, 16, 24, and 38 hours by using a commercial enzyme immunoassay kit for 6-keto-PGF₁.

The experiments described in this article used cells that were isolated from three donors aged 29, 60, and 82 years. Following characterization, cells were seeded into 24-well tissue-culture dishes at a seeding density of 1x10⁵ cells/well and grown to confluence in the media described above but without the addition of amphotericin B. Cells were used at passages 1 through 3.

Primary cultures of HUVECs were established¹⁵ by using a brief collagenase digestion. Cells were initially plated into 75-cm² culture flasks at a density of 1x10⁶ cells and grown to confluence in modified M-199 as described above. In experiments, cultures from passages 1 and 2 were seeded in serum-free medium and treated as described above for HCMECs.

HFMECs (a gift of Clonetics Corp) were isolated from a single neonatal male donor and were provided as a growing culture. Cells were grown in EC growth medium until confluent. For experimentation, HFMECs were seeded in serum-free medium by using the protocol described above.

Assay Techniques
The concentrations of TPA, UPA, and PAI-1 antigens were measured by using specific enzyme-linked immunosorbent assays (ELISAs).^{16 17 18} 6-Keto-PGF₁ concentration was determined by enzyme immunoassay according to the manufacturer's instructions (Cayman). The protein concentration of cell lysates was determined by using the bicinchoninic acid protein assay, using BSA for calibration, according to the manufacturer's instructions. Inhibition of protein synthesis was accomplished with 3.6 µmol/L cycloheximide in serum-free medium. The proliferative response of cells to -thrombin was assessed over a 500-fold range of thrombin concentrations by measuring [³H]thymidine incorporation, cell number, and total cellular protein in control and experimental cultures. Confluent cell monolayers were placed in serum-free medium (with or without -thrombin) for 18 hours, pulsed with [³H]thymidine (1 µCi/mL) for 6 hours, washed twice with PBS, and incubated for 30 minutes in serum-free medium containing 4.1 mmol/L unlabeled thymidine. Monolayers were disrupted into single-cell suspensions by using 0.25% trypsin/EDTA. Cell number was established by counting in a hemocytometer; for measurement of [³H]thymidine incorporation, cells were layered on glass microfiber filters (Whatman International Ltd) and denatured with 10% trichloroacetic acid; the filters were transferred into ChemFluor-1 (ICN), and the radioactivity was quantified by scintillation counting. Protein determinations were performed by using the bicinchoninic acid method on parallel cultures.

Preparation of Reagents
-Thrombin (3200 IU/mg) was prepared by the method of Lundblad et al.¹⁹ Prethrombin-2 was prepared by the selective proteolysis of human prothrombin.²⁰ Human coagulation factor Xa and D-phenylalanyl-L-propyl-L-arginine chloromethylketone (FPRcK)–modified -thrombin were provided by Dr Paul Haley, Hematologic Technologies, Essex Junction, Vt. Human plasmin was prepared by the method of Claeys et al.²¹ A thrombin receptor agonist peptide (TRAP) consisting of the sequence SFLLRNPNDKYEPF was provided by Dr William Church, Department of Biochemistry, University of Vermont.

Experimental Protocol
After the cells had grown to confluence, they were rinsed twice with PBS and refed with serum-free medium containing -thrombin or other experimental agents. The cells were then incubated at 37°C under culture conditions of 95% air and 5% CO₂. At designated intervals, cell-conditioned medium was collected and centrifuged at 10 000g to remove cellular debris, and the medium was made 0.01% in Tween 80 and stored at -20°C. Cell lysates were harvested in lysing buffer and stored at -20°C. ELISAs were performed within 72 hours of harvesting samples.

Electrophoresis and Immunoblotting of UPA
Molecular characterization of secretion products was undertaken in the following manner. Aprotinin (10 U/mL) was added to conditioned medium, after which the medium was concentrated from 10 to 1 mL by ultrafiltration on a Centricon-10 membrane. The concentrate was immunoadsorbed by using a mouse monoclonal anti-UPA IgG coupled to Sepharose 4B (50 µL of a 50% substituted Sepharose slurry was added for 2 hours at room temperature to 1 mL concentrated media). Immunoadsorbed UPA antigen was eluted by incubation in 2% sodium dodecyl sulfate (SDS) sample buffer and brought to 5% ß-mercaptoethanol prior to SDS–polyacrylamide gel electrophoresis on a 5% to 15% gradient gel. Fractionated proteins were electrophoretically transferred to nitrocellulose and immunoblotted by using anti-UPA antibodies.²² Molecular weights were determined by calibration with the following protein standards: phosphorylase b (M_r=106 000), BSA (M_r=80 000), ovalbumin (M_r=49 500), carbonic anhydrase (M_r=32 500), soybean trypsin inhibitor (M_r=27 500), and lysozyme (M_r=18 500).

Quantification of UPA mRNA Levels by Northern Blot Analysis
Total cellular RNA was extracted from the cells by lysis with 1 mL/well of 5 mol/L guanidinium isothiocyanate and 25 mmol/L sodium citrate, pH 7.0, containing 0.5% sarkosyl and 8% ß-mercaptoethanol according to the procedure described by Chomczynski and Sacchi²³ ; this procedure was followed by cold phenol extraction. The final RNA pellet was resuspended in 50 µL H₂O, and the concentration was determined by absorbance at 260 nm. RNA was denatured with 6 mol/L glyoxal for 1 hour at 50°C. For Northern blot analysis, glyoxal-treated RNA (10 µg) was subjected to electrophoresis in a 1% agarose gel followed by capillary transfer to a Zetaprobe membrane. After transfer, the membrane was baked for 1 hour at 80°C under vacuum and prehybridized for at least 4 hours at 65°C in the following hybridization mixture: 50% formamide, 5x saline–sodium citrate buffer (SSC), 10x Denhardt's solution, 50 mmol/L phosphate buffer, pH 7.6, 5% dextran sulfate, 1 mmol/L EDTA, and 1% SDS containing a specific [³²P]RNA probe for UPA RNA. Antisense RNA probes were generated by SP6 polymerase transcription of a linearized pSP65 plasmid containing nucleotides spanning 931 through 979 of the cDNA sequence.²⁴ The probe was freshly prepared by using a Promega transcription kit and routinely had a specific activity of 0.5 to 2.0x10⁹ cpm/µg RNA. The probes were heat denatured and added to the hybridization solution at a concentration of 10⁶ cpm/mL. Membrane washing conditions at 65°C were as described by the manufacturer of Zetaprobe, with a final wash of 0.05x SSC at 65°C. Autoradiography was performed by using Reflection NEF autoradiographic film (Du Pont/New England Nuclear) at 72°C. Loading levels of RNA samples were determined by using a pT7 RNA 18S template (Ambion). This template contained an 80-bp antisense fragment of a highly conserved region of the human 18S ribosomal RNA (rRNA) gene. Antisense RNA probes were generated by T7 polymerase transcription of a linearized pT7 RNA 18S plasmid that had been freshly prepared by using a Promega transcription kit as described above. Membrane washing and autoradiography were performed as described above. The molecular sizes of UPA mRNA and 18S rRNA were determined by using an RNA calibration mixture. Total cellular RNA was isolated and subjected to Northern blot analysis, and samples (three per treatment) were probed for UPA mRNA and 18S rRNA as described above and quantified by using densitometric scanning (Technology Resources, Inc). The density of UPA mRNA and 18S rRNA bands was measured and assigned arbitrary units. Each UPA mRNA band was normalized to its respective 18S rRNA band, and these values were multiplied by the mean density of the 18S rRNA for that group. Data are expressed as arbitrary density units and are mean±SD (n=3).

Statistics
The statistical significance of experimental results was determined by using the standard Student's t test for nonpaired comparisons.

	Results

Top Abstract Introduction Methods Results Discussion References

 
Characterization of HCMECs, HUVECs, and HFMECs
HCMECs (Fig 1, top) exhibit an elongated spindle-shape morphology that lacks the characteristic cobblestone morphology typical of cultured endothelium derived from large vessels such as HUVECs (Fig 1, middle) or from ECs derived from the noncerebral microvasculature (HFMECs) (Fig 1, bottom). HCMECs align themselves longitudinally as they grow, and upon reaching confluence, which occurs after 12 to 16 days in culture, they form a contact-inhibited monolayer.

View larger version (97K): [in this window] [in a new window]   Figure 1. Photomicrographs. Top, Human cerebral microvascular endothelial cells in situ have an elongated morphology in the subconfluent phase. As cultures reach confluence, they become contact inhibited and exhibit a more oval-type morphology, as shown (original magnification x160). Middle, Human umbilical vein endothelial cells, which are recognized by their classic cobblestone morphology (original magnification x160). Bottom, Confluent human foreskin microvascular endothelial cells, which demonstrate contact inhibition in vitro, are characterized by both their prominent nucleus and distinct cytoplasm (original magnification x200).

Cells were identified as being of endothelial origin when the following four criteria were fulfilled: positive immunocytochemical staining for factor VIII/vWf antigen (Fig 2, top) (considered to be the most reliable marker for cells of endothelial origin)¹⁰ ; binding of UEA-1 (Fig 2, middle); negative staining for GFAP (data not shown); and positive production of PGI₂ (Fig 2, bottom). Primary and passage-one cultures of HCMECs displayed a strong multifocal perinuclear staining for factor VIII/vWf. Cultured human fibroblasts, used as a negative control in our studies, did not stain for factor VIII/vWf, whereas HUVECs and HFMECs, our positive controls, stained intensely. -L-Fucosyl moieties recognized by UEA-1 are highly specific for and are selectively expressed by membranes and organelles of human ECs.^{12 25} UEA-1 is recognized as being particularly sensitive for the demonstration of human microvascular endothelium. Moreover, cultured human ECs retain their UEA-1 surface markers after multiple passages in long-term culture conditions.²⁶ HCMECs, HFMECs, and HUVECs exhibited a strong positive immunofluorescence for UEA-1. In contrast, human lung fibroblasts used as a negative control did not react with the fluorescein-labeled UEA-1 lectin. Astrocytes, a potential contaminant, exhibited the marker GFAP, but HCMEC cultures exhibited negative immunofluorescence to GFAP. This confirmed that the HCMECs were not of astrocytic derivation. We examined the ability of HCMECs to synthesize and secrete PGI₂, which is known to be secreted by large-vessel endothelium. HCMECs, like other types of human ECs, synthesize PGI₂. Baseline secretion by HCMECs at 24 hours in serum-free medium is similar to that of HUVECs during the same time period. HCMECs secrete 2700±924 pg/1x10⁶ cells, and under identical culture conditions HUVECs secrete 1600±100 pg/1x10⁶ cells.

View larger version (41K): [in this window] [in a new window]   Figure 2. Top and middle, fluorescent photomicrographs of human cerebral microvascular endothelial cells (HCMECs) (original magnification x200). Top, The presence of factor VIII–von Willebrand factor antigen is shown. HCMECs display the perinuclear fluorescence characteristic of cells of endothelial origin. Middle, Binding of Ulex europaeus agglutinin–1 lectin to HCMECs. The cytoplasmic fluorescence exhibited by HCMECs is specific for the interaction of this lectin with endothelial cells. Bottom, Line graph showing time course of prostacyclin production by HCMECs. Prostacyclin production was assessed by measuring 6-keto-prostaglandin F1 (pg 6 keto-PGF1), the stable end product of prostacyclin metabolism in serum-free medium conditioned by HCMECs. Data represent mean±SD (n=4) for each time point.

Comparison of EC Fibrinolytic Properties
In order to establish basal levels of fibrinolytic protein secretion by various endothelia, confluent cultures of HCMECs, HUVECs, and HFMECs were placed in serum-free media for 24 hours, after which cell-conditioned medium and cell lysates were assayed by ELISA for TPA, UPA, and PAI-1. Under baseline conditions, results for HCMEC-conditioned media (mean±SD, n=6) were 2.5±0.4, 1.0±0.3, and 1000±130 ng/1x10⁶ cells for TPA, UPA, and PAI-1, respectively. Corresponding values were 2.0±1.0, <0.2, and 3700±550 ng/1x10⁶ cells for HUVEC-conditioned media (n=9) and 10±2, 11±1, and 2200±500 ng/1x10⁶ cells for HFMEC-conditioned media (n=6). There were no significant levels of TPA, UPA, or PAI-1 associated with the cell lysates in any of the above endothelia.

Effects of -Thrombin on the EC Fibrinolytic System
Exposure of HCMECs to -thrombin induced a large concentration-dependent increase in UPA secretion, a twofold increase in PAI-1 secretion, and no influence on the secretion of TPA (Table 1). With 10 nmol/L -thrombin, production of UPA was 29±1.6-fold higher than in control cultures; HCMECs treated with 25 and 50 nmol/L -thrombin showed 27±2- and 34±3-fold increases, respectively (data not shown). Half-maximal stimulation was observed with between 1 and 2 nmol/L -thrombin. Cycloheximide (3.5 µmol/L) blocked the increased UPA production in -thrombin–treated HCMECs, indicating that protein synthesis was a requirement for the effect. Western blotting with monoclonal anti-UPA antiserum of the immunoadsorbed UPA antigen from 24-hour conditioned medium from HCMECs revealed a main band, with M_r=54 000 consistent with urokinase (Fig 3, lane 1). Conditioned medium from HCMECs stimulated with 10 nmol/L -thrombin showed an increase in this band corresponding to the increase observed in UPA antigen secretion when assayed by ELISA (Fig 3, lane 4). When immunoadsorbed samples from -thrombin–treated HCMECs were reduced with ß-mercaptoethanol and subjected to electrophoresis and Western blotting, a single band of M_r=54 000 was observed (data not shown). In the basal state, HUVECs produce no detectable urokinase. Thrombin treatment of HUVEC cultures did not result in detectable UPA in the medium. HUVEC cultures showed a 3.5-fold increase in TPA after treatment with -thrombin and no significant change in PAI-1 levels. With the possible exception of TPA, HFMECs were not responsive to -thrombin (Table 1).

View this table: [in this window] [in a new window]   Table 1. Fibrinolytic Responses of Human Cerebral Microvascular (HCMEC), Human Umbilical Vein (HUVEC), and Human Foreskin Microvascular (HFMEC) Endothelial Cells to -Thrombin

View larger version (79K): [in this window] [in a new window]   Figure 3. Immunoprecipitation and Western blotting of urokinase plasminogen activator antigen from 24-hour conditioned medium from cultured human cerebral microvascular endothelial cells (HCMECs). Lane 1, 200 ng single-chain urokinase; lane 2, 100 ng two-chain urokinase; lane 3, control HCMECs; and lane 4, HCMECs treated with 10 nmol/L -thrombin. Samples were not reduced. Molecular-weight standards are indicated at the right (prestained standards were reduced). Both single-chain and two-chain urokinase stocks contain low-molecular-weight urokinase (Mr=33 000).

Time-course studies indicated that increased UPA secretion by HCMECs occurred between 8 and 16 hours after the addition of -thrombin (Fig 4A). In contrast, HCMECs treated with 10 nmol/L -thrombin showed a significant increase in PAI-1 production within 2 hours (105±19 versus 63±3 ng/1x10⁶ cells in control cultures, n=3, P<.03) that was maintained over 48 hours and represented a 1.8-fold increase in the rate of PAI-1 production (Fig 4B).

View larger version (13K): [in this window] [in a new window]   Figure 4. Line graphs showing time courses of (A) urokinase plasminogen activator (UPA) and (B) plasminogen activator inhibitor–1 (PAI-1) secretion by human cerebral microvascular endothelial cells (HCMECs). UPA and PAI-1 antigen (Ag) levels were measured at times ranging from 0 to 48 hours by using enzyme-linked immunosorbent assays in serum-free medium conditioned by control () or -thrombin (10 nmol/L)–treated HCMECs. Data represent mean±SD for n=3 samples. uPA indicates UPA.

Northern blot analysis of total RNA obtained from HCMECs harvested 2, 8, and 24 hours after treatment with 10 nmol/L -thrombin are shown in Fig 5. UPA mRNA (2.4 kb) was significantly increased in treated HCMECs compared with control cells at all time points. Hybridization with the 18S rRNA probe showed that equivalent amounts of RNA had been applied to the gel and that 18S rRNA (2.8 kb) levels were not changed by -thrombin treatment. Quantitation of UPA mRNA by densitometry scanning showed that -thrombin treatment resulted in 2.5±0.9-, 5.1±0.9-, and 2.75±0.4-fold increases (P<.05) in UPA mRNA over control values at 2, 8, and 24 hours, respectively.

View larger version (38K): [in this window] [in a new window]   Figure 5. Autoradiograph showing effect of -thrombin on human cerebral microvascular endothelial cell (HCMEC) urokinase plasminogen activator (UPA) mRNA. HCMECs were treated with 10 nmol/L -thrombin (IIa) for 2, 8, and 24 hours; total cellular RNA was isolated, and Northern blot analysis was performed as described in "Methods." Samples were transferred to a Zetaprobe membrane that was then probed for UPA (2.4 kb) mRNA and 18S ribosomal (2.8 kb) RNA by using specific complementary 32P-labeled RNA probes. The membrane was subjected to autoradiography, and RNA species were distinguished by using an RNA calibration mixture. uPA indicates UPA; C, control.

Effects of -Thrombin on EC Proliferation
-Thrombin increased DNA synthesis in HCMECs as measured by [³H]thymidine incorporation in a dose-dependent manner (Table 2), reaching a maximal fourfold increase with 10 nmol/L -thrombin; levels of [³H]thymidine incorporation at 25 and 50 nmol/L -thrombin were 4.2±0.3- and 4.9±0.3-fold increased, respectively (n=6). Half-maximal stimulation was observed between 1 and 2 nmol/L -thrombin for each cell isolate. Enhanced DNA synthesis was accompanied by an increase in total cellular protein (1.6-fold) at 24 hours. The level of [³H]thymidine incorporation by HCMECs when grown in medium supplemented with 20% FBS was 1.5-fold of the control (cells grown in serum-free medium) value (n=6). No increase in [³H]thymidine incorporation was observed in -thrombin–treated HUVECs or HFMECs.

View this table: [in this window] [in a new window]   Table 2. Proliferative Responses of Human Cerebral Microvascular (HCMEC), Human Umbilical Vein (HUVEC), and Human Foreskin Microvascular (HFMEC) Endothelial Cells to -Thrombin

Specificity of Catalytically Active -Thrombin on Increased UPA Synthesis and Proliferation
Two -thrombin–related proteins without catalytic activity and two related catalytically competent proteases were examined to evaluate the specificity of active -thrombin (Table 3). Prethrombin-2, a catalytically inactive, single-chain molecule, has an amino acid sequence that is identical to that of -thrombin. FPRcK-modified thrombin is catalytically inert but structurally intact -thrombin that is generated by selectively modifying the active site histidine (His₃₆₃) with the tripeptide FPRck. Human factor Xa and human plasmin share with -thrombin the catalytic requirement that a basic amino acid provides the carbonyl group of the scissile bond. None of these proteins affected HCMEC production of UPA and PAI-1, and none increased DNA synthesis. Table 3 presents a representative experiment. Higher concentrations (50 and 100 nmol/L) of FPRcK-modified thrombin, prethrombin-2, and plasmin were also evaluated and found to be ineffective.

View this table: [in this window] [in a new window]   Table 3. Specificity of the HCMEC Proliferative and Fibrinolytic Responses for -Thrombin

Role of the Thrombin Receptor
Many of the effects of -thrombin on different types of cells depend on its binding to and proteolysis of an extracellular region of a transmembrane glycoprotein.^{27 28} TRAPs, peptides with the sequence of the NH₂-terminal region of the proteolyzed binding protein, mimic a number of thrombin effects.^{28 29} In our study, a TRAP with the sequence SFLLRNPNDKYEPF increased UPA production by HCMECs. Table 4 shows a representative experiment in which HCMEC response was evaluated over a range of TRAP concentrations (0.5 to 50 µmol/L). In this experiment, 23% of the UPA level achieved with -thrombin (10 nmol/L) treatment of HCMECs was obtained with 50 µmol/L TRAP. In four additional experiments (n=12), the response involving all three cell isolates to 50 µmol/L TRAP ranged from 33% to 57% of the response to 10 nmol/L -thrombin. In contrast, TRAP was unable to mimic any part of the mitogenic action of -thrombin (Table 4). When data from three additional experiments were combined, the response to 50 µmol/L TRAP was 117±41% (n=12) of untreated HCMECs.

View this table: [in this window] [in a new window]   Table 4. Fibrinolytic and Proliferative Responses of HCMECs to a Thrombin Receptor Agonist Peptide

	Discussion

Top Abstract Introduction Methods Results Discussion References

 
Our studies show that HCMECs, in contrast to endothelium isolated from large vessels (HUVECs) or from the noncerebral microvasculature (HFMECs), respond to catalytically active -thrombin with a selective and dramatic upregulation of UPA synthesis and secretion and that -thrombin is a mitogen for HCMECs. Furthermore, regulation of UPA production is at least partially mediated through the thrombin receptor,²⁷ whereas the proliferative response is not associated with this thrombin receptor.

Although ECs throughout the vasculature share basic structural and functional characteristics, marked differences, dictated largely by their vascular bed of origin, exist in their morphology and differential responsiveness to a variety of mediators.^{30 31} Our studies demonstrate that endothelium derived from the cerebral microvasculature is distinctly different in its interactions with -thrombin than other large- and small-vessel endothelia. -Thrombin, the immediate catalyst of fibrin deposition, is a well-established modulator of TPA and PAI-1 production by cultured endothelium derived from large vessels.³² In our study, we show that HCMECs respond uniquely to catalytically active -thrombin with a selective, enhanced upregulation in UPA production. These observations suggest that UPA, not TPA, is the major profibrinolytic enzyme in the microvasculature of the brain.

Vascular cells are thought to proliferate only in response to injury. -Thrombin initiates proliferation in a variety of cells including vascular fibroblasts and smooth muscle cells^{4 5} and is recognized for its amplification of the proliferation induced by several EC growth factors.³³ Recently, thrombin has been reported to act as a mitogen in cultures of rodent capillary ECs but not in cultured ECs derived from large rodent vessels.³⁴ Our studies highlight the unique response of HCMECs to -thrombin; we report here for the first time that -thrombin is a potent mitogen for these endothelia.

The response of the fibrinolytic system of HCMECs to thrombin has two components: an approximately twofold increase in the rate of PAI-1 production, which occurred within 2 hours of the addition of -thrombin, and an increased production of UPA, which was not observed until after 8 hours of exposure to -thrombin. Thus the fibrinolytic response of HCMECs to -thrombin can be divided into two phases: an early antifibrinolytic phase in which the rate of PAI-1 secretion increases immediately and a late profibrinolytic phase characterized by increased UPA secretion. The delayed increase in plasminogen activator release by HCMECs in response to vessel injury seems logical. The immediate response to vessel injury is plugging the leak, a repair process in which local -thrombin production provides the catalyst required for clot formation. The maintenance of the clot requires suppression of the activity of available plasminogen activators. The primary plasminogen activator inhibitor PAI-1 is released by platelets participating in clot formation³⁵ and is synthesized by ECs.^{36 37} Thus, suppression of plasminogen activator activity is an early requirement, while the demand for increased plasminogen activator activity occurs during the subsequent processes that are aimed at restoring and renovating the vessel.

Based on studies of large-vessel endothelia, mainly those derived from human umbilical vein, a consensus has existed in which the role of the intravascular activator of plasminogen is assigned to TPA, while extravascular catalysis of the conversion of plasminogen to plasmin is postulated to rely on UPA. Complementing these observations supporting TPA as the intravascular plasminogen activator, positive evidence has accumulated for a role of UPA in the extravascular space during wound healing, tissue remodeling, and cell migration.³⁸ Cells involved in these processes synthesize UPA and/or possess specialized receptors for UPA³⁹ that bind UPA but do not interfere with its catalysis of plasminogen activation,⁴⁰ thus localizing plasmin formation at the cell surface. Our studies suggest that in injured regions of the cerebral microvasculature, UPA functions as the primary plasminogen activator, ie, when -thrombin is the agonist.

The HCMEC fibrinolytic and proliferative responses to -thrombin require catalytically active thrombin. Our studies with the TRAP SFLLRNPNDKYEPF indicate that upregulation of PAI-1 and UPA by HCMECs is mediated through the thrombin receptor. TRAP, however, does not promote HCMEC proliferation, suggesting that thrombin induction of cell division may involve additional signaling pathways and/or receptors. In support of our studies, others using identical TRAPs have demonstrated differential regulation of one function but not the other in the same EC.^{41 42} For example, in human saphenous vein endothelium, TRAP stimulates tissue-factor production but has no effect on thrombomodulin expression, whereas thrombin exerts an influence on both tissue factor and thrombomodulin.⁴²

In conclusion, our findings demonstrate that catalytically active -thrombin regulates fibrinolysis and proliferation in human brain microvascular endothelium by different mechanisms. Furthermore, the cerebral microvasculature may be a specialized region of the vascular system in which fibrinolysis is mediated via UPA and not TPA. These data imply that pharmacological intervention with anticoagulant or fibrinolytic agents may produce responses unique to this tissue.

	Acknowledgments

 
These studies were supported by Public Health Service grants NS-3032403 and HL-46703, the Surgical Research Fund, Department of Surgery, University of Vermont, and the Collen Foundation. We wish to thank the Department of Obstetrics and Gynecology, Medical Center Hospital of Vermont, for obtaining umbilical cords. We are grateful to Sabrina Martucci, Clonetics Corp, for arranging the gift of human dermal microvascular endothelium. We thank Dr Paul Haley, Hematologic Technologies, for preparation of FPRck-modified thrombin and factor Xa and Dr William Church and Laurie Ouelette for preparation of the TRAP. We appreciate the excellent secretarial assistance provided by Jeanette Mann.

Received October 18, 1994; accepted April 11, 1995.

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