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Biology of Reproduction 59, 759-767 (1998)
©Copyright 1998 Society for the Study of Reproduction, Inc.

Human Endothelial Cell Migration Is Stimulated by Urokinase Plasminogen Activator:Plasminogen Activator Inhibitor 1 Complex Released from Endometrial Stromal Cells Stimulated with Transforming Growth Factor ß1; Possible Mechanism for Paracrine Stimulation of Endometrial Angiogenesis1

Tove Sandberga, Bertil Casslén2,a, Barbro Gustavssona, , and Theo J. Benraadb

a Department of Obstetrics & Gynecology, University Hospital, S-221 85 Lund, Sweden b Department of Experimental and Chemical Endocrinology, Catholic Hospital, 6500 HB Nijmegen, The Netherlands


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Human endometrial stromal cell cultures, stimulated for two days with recombinant transforming growth factor ß1 (TGFß1; 10 ng/ml), contained conditioned medium concentrations of urokinase plasminogen activator (uPA), plasminogen activator inhibitor 1 (PAI1), and uPA:PAI1 complex. Since a number of cellular effects have been reported to follow a binding of enzymatically inactive uPA to the receptor in different cell types, we studied the influence of uPA:PAI1 complex on human umbilical vein endothelial cells (HUVEC) and human microvascular endothelial cells (HMEC-1).

Increasing concentrations of uPA:PAI1 complex as well as free uPA resulted in a dose-dependent stimulation of endothelial cell migration. Stimulation by the complex was of the same magnitude as that of free uPA on a molar basis and reached its maximum at 1 nM in both cell types. PAI1 by itself, however, had no effect on cell migration. The migratory response to both uPA and the uPA:PAI1 complex was inhibited by antibody adhesion to the cell surface receptor for uPA. In addition, we found that TGFß1 had a direct stimulatory effect on migration in both HUVEC and HMEC-1. This response did not, however, involve the binding of uPA to the uPA receptor.

Since TGFßs are expressed in endometrial tissue and reportedly stimulate angiogenesis in other tissues in vivo, though not endothelial cell proliferation in vitro, they may engage in the regeneration of endometrial vasculature indirectly via perivascular cells. We found that the uPA:PAI1 complex, when released from endometrial stromal cells in response to TGFß1, stimulated endothelial cell migration. This suggests a possible mechanism for paracrine stimulation of endometrial angiogenesis.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The human endometrium demonstrates a remarkable growth each cycle in response to estrogen and a subsequent rapid switch to differentiation in response to progesterone. The mechanisms that regulate endometrial angiogenesis, which accompanies regeneration after menstruation, are still largely unknown and controversial. An early study in this area showed higher mitogenic activity of the endometrial endothelial cells in the proliferative phase than in the secretory phase [1]. A later study, using immunological techniques, could not identify significant peaks of endothelial cell proliferation in endometrial tissue over the cycle [2]. Endothelial cell migration was, however, stimulated to a greater extent by endometrial tissue explants from the proliferative phase, rather than from the secretory phase [3]. Furthermore, progestogenic steroids have antiproliferative effects on microvasculature in vivo and on decidual endothelial cells in vitro [4, 5].

Transforming growth factor ß (TGFß) is a family of 25-kDa homodimeric proteins with multifunctional effects on cellular growth and differentiation [6, 7]. In humans, there are at least three well-characterized members, TGFßs 1–3, which are structurally closely related. The biological activity of TGFßs is mediated through their binding to specific cell surface receptors. TGFßs and their mRNA are present in the endometrial tissue throughout the menstrual cycle [8–10]. The content of TGFß1 mRNA is shown to be significantly higher in endometrial tissue obtained in the mid secretory, late secretory, and menstrual phases as compared to the proliferative and early secretory phases [11]. TGFßs are released as latent proforms, which are activated by proteolytic cleavage, e.g., by plasmin. Activation of plasminogen in this context requires the presence of urokinase plasminogen activator (uPA), bound to its cell surface receptor [12]. The enzymatic activity of uPA is controlled by plasminogen activator inhibitor-1 (PAI1), the levels of which are increased by progesterone in the secretory phase [13]. On the basis of previous results indicating that the endometrial release of plasminogen activator (PA) activity is low in the secretory phase, we have suggested that endometrial TGFßs are mainly present in their latent forms during this period. Activation of latent TGFßs is likely to accelerate premenstrually when endometrial content as well as the release of PAs is dramatically increased [14–16]. Such a pattern of activation would imply a role for TGFß in the process of endometrial regeneration and/or angiogenesis.

The observation that TGFß stimulates angiogenesis in vivo [17–19], but not endothelial cell proliferation in vitro [20, 21], suggests a paracrine involvement of other cell types. In this study, we explored the possibility that the uPA:PAI1 complex, released from TGFß1-stimulated endometrial stromal cells, stimulates migration in human umbilical vein endothelial cells (HUVEC) and human microvascular endothelial cells (HMEC-1).


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Materials

Earle's minimum essential medium (MEM) without phenol red, Hanks' buffered salt solution (HBSS) without phenol red, molecular cell develomental biology medium (MCDB131), fetal bovine serum (FBS), and glutamine were from Gibco (Paisley, Scotland). Crude collagenase, deoxyribonuclease (DNase), tosyl-L-lysine-chloromethyl-ketone (TLCK), estradiol, progesterone, epidermal growth factor, hydrocortisone, penicillin-streptomycin-fungizone, BSA, lactoperoxidase, gelatine, p-aminobenzamidine, bromphenol blue, ethidium bromide, formaldehyde, formamide, 2-mercaptoethanol, 3-(N-morpholino)propanesulfonic acid (MOPS), and trizma base were obtained from Sigma (St. Louis, MO). Ukidan was from Serono (Geneva, Switzerland). Nitex nylon meshes with pore sizes of 350 µm and 35 µm were obtained from Tetko (Elmsford, NY). 125I, [3H&;, and the Megaprime DNA labeling system were obtained from Amersham (Solna, Sweden), and Gene Screen Plus nylon filters from Dupont (Boston, MA). The methanol was obtained from Lab Scan (Dublin, Ireland); SDS from Bio-Rad (Richmond, CA), Giemsa stain, Titriplex III (EDTA), and boric acid from Merck (Darmstadt, Germany); and scintillation liquid Ultima Gold from Packard Instrument Company (Meriden, CT). Tissue culture plates, IsoTip filter tips, Mµlti Safeseal microcentrifuge tubes, and Spin-X centrifuge filter units were from Costar (Broadway/Cambridge, MA); Bacto agar, tryptone, and yeast extract were from Difco (Detroit, MI); and competent cells, Wizard Minipreps DNA Purification system, and blue/orange loading dye were obtained from Promega (Madison, WI). RNeasy Total RNA preparation kit was obtained from Qiagen (Hilden, Germany), glycogen and restriction enzymes from Boehringer Mannheim Scandinavia AB (Bromma, Sweden), and molecular size standards for DNA and RNA from Gibco BRL (Bethesda, MD). Monoclonal antibodies to human vimentin and cytokeratin, normal rabbit serum, biotinylated rabbit anti-mouse antibodies and avidin-biotin-peroxidase complex were from Dako AS (Glostrup, Denmark); and diaminobenzidine was from Saveen Biotech AB (Malmö, Sweden).

The probe for uPA was generously provided by Dr. F. Blasi (Milano, Italy), the probe for uPA receptor by Dr. L.R. Lund (Copenhagen, Denmark), the probe for PAI1 by Dr. T. Ny (Umeå, Sweden), purified PAI1 and uPA:PAI1 complex by Dr. P. Andreasen (Aarhus, Denmark), and recombinant human TGFß1 by Dr. C.H. Heldin (Uppsala, Sweden). The human ß-actin control probe was acquired from Clontech (Palo Alto, CA).

Tissue Processing

Endometrial tissue was obtained from uteri removed because of benign nonendometrial pathology, i.e., cervical dysplasia, dysmenorrhoea, uterine prolapse, and fibromyomas. All patients were parous, 30–45 yr of age, and had regular menstrual cycles. Permission to use part of the endometrium was granted by the University Review Board for studies on human subjects. Endometrial pathology was excluded by histopathologic examination of the formalin-fixed portion of the endometrium. The endometrium was gently scraped off from the upper part of the uterine cavity immediately after the removal of the uterus.

Endometrial tissue was proteolytically disintegrated, and the stromal cells were collected by methods previously described [22]. The stromal cell cultures were 96–99% pure when analyzed by immunocytochemistry using antibodies against vimentin and cytokeratin [13].

HUVECs were prepared from human umbilical cords essentially as earlier described [23]. After being flushed with HBSS to remove coagulated blood, the vein was filled under pressure with the dissociation solution, which is crude collagenase 1.25 g/L in HBSS. The umbilical cord was clamped on both ends to maintain the pressure, and incubated at 37°C for 15 min. After draining the collagenase solution, the vein was flushed with ice-cold HBSS containing 20% FBS (to inactivate the collagenase). Endothelial cells were recovered by centrifugation at 1000 rpm for 10 min, and the pellet was resuspended in MEM containing 20% FBS. HMEC-1 were generously provided by the Center for Disease Control (Atlanta, GA) [24].

Tissue Culture

Stromal cells were plated at 100 000 cells/well. Cultures were rinsed with HBSS after one day, and the experiment was begun when the cultures were confluent. Cells were grown in MEM supplemented with 10% FBS, 2 mM glutamine, 100 000 IU/L penicillin, 100 mg/L streptomycin, and 0.25 mg/L fungizone. Phenol red was omitted from HBSS and MEM to avoid interference with steroid receptors. Cultures were incubated in humidified air with 5% CO2 at 37°C. Confluent cultures were stimulated with either TGFß1 10 ng/ml or HBSS for control. Conditioned media were collected and stored at -20°C until assayed.

HUVEC were grown in 6-well plates coated with 0.1% gelatine in Earle's MEM supplemented with 20% FBS, glutamine 2 mM, penicillin 100 000 IU/L, streptomycin 100 mg/L, and fungizone 0.25 mg/L. The cultures were grown to confluence in 2–3 days.

HMEC-1 were grown on uncoated plastic in medium MCDB131 supplemented with 10 ng/ml epidermal growth factor, 1 µg/ml hydrocortisone, and 15% FBS.

Assays for uPA, PAI1, and uPA:PAI1 Complex

Urokinase PA was measured by an RIA [25]. The assay measured both the high and low molecular weight forms of uPA. It also measured 90% of uPA in complex with PAI1 in the range of 1–10 ng/ml.

PAI1 was measured using a commercial ELISA kit, Imulyse PAI1 (Biopool, Umeå, Sweden), which detected the active and latent forms of PAI1, as well as PAI1 in complex with PAs.

The concentration of the uPA:PAI1 complex was measured in a four-antibody ELISA [26]. Plates were incubated overnight at 4°C with sheep anti-chicken IgG coating antibody (100 µl/well). Next morning the plates were washed, blocked with 1% BSA for 2 h at 37°C, washed again, and subsequently incubated for 2 h at 37°C with the catching antibody chicken anti-analyte (100 µl/well). Samples or reference standards were added and incubated overnight at 4°C. Washing was followed by 100 µl tagging antibody, rabbit anti-analyte, for 2 h at room temperature. After being washed, plates were incubated with 100 µL/well detecting antibody (horseradish peroxidase (HRP)-labeled goat anti-rabbit IgG) for 2 h at room temperature, and after another wash, the o-phenylenediamine substrate was added, and incubation continued for 30 min in darkness. The reaction was stopped by adding H2SO4 1 M, and absorbance was subsequently measured at 492/620 nm.

Purification and Radiolabeling of uPA

Urokinase PA was purified from Ukidan by affinity chromatography on a benzamidine-Sepharose column as previously described [13]. The active enzyme fraction was further separated in high molecular weight uPA (Mr 50 000) and low molecular weight uPA (Mr 33 000) by gel filtration on Sephadex G-100. Immunoblotting of the peak low molecular weight uPA fractions revealed trace amounts of high molecular weight uPA, estimated to 1%. The peak high molecular weight uPA fraction, which was 98–99% pure, was used for 125I-labeling with the lactoperoxidase method [27]. Specific radioactivity was in the range of 0.4–0.6 MBq/µg protein.

Assay of 125I-uPA Binding to Stromal Cell Cultures

Confluent cultures were given serum-free medium for 2 h at 37°C before the experiments. Cells were subsequently incubated on ice for 2 h with HBSS containing BSA, 10 g/L, and 125I uPA.

To assay the binding of radiolabeled uPA, cells were washed four times with ice-cold HBSS and subsequently lysed with 1 M NaOH. Radioactivity of the lysate was counted in a 1260 Multigamma counter (Pharmacia AB, Sweden). Nonspecific binding was assayed in the presence of 100-fold molar excess of unlabeled uPA, and specific binding was calculated as the difference between total and nonspecific binding.

To measure the total number of receptors, endogenous uPA was removed by briefly (2–3 min) exposing the cells to 75 mM acetate buffer, pH 3.0, containing 2.5 mM CaCl2, 0.5 mM Mg Cl2, and 0.3 M NaCl, before incubation with radiolabeled uPA. The number of occupied receptors was calculated as the difference between the total number and the number of free receptors.

RNA Analysis

Total RNA was prepared from stromal cell pellets using the RNeasy Total RNA Purification kit, and 4-µg aliquots were size-separated on 1% agarose gels containing 2.2 M formaldehyde. The RNA was then transferred to Gene Screen Plus nylon filters [28]. The probes were radiolabeled with [32P]dCTP using the random labeling method Megaprime DNA labeling system [29]. The filters were hybridized in 0.25 M sodium phosphate, 7% SDS, 1 mM EDTA at 65°C for 12 h. After hybridization, the filters were washed in 0.02 M sodium phosphate, 1% SDS for 3 x 10 min at 65°C. Autoradiography was performed for 1–12 h, and signal intensities were measured by densitometry on a Bio Image computer system. All filters were hybridized to the probes for PAI1, uPA, and the uPA receptor, and also, in order to verify equal loading, to a human ß-actin cDNA probe.

The probe for PAI1 was a 2-kilobase (kb) cDNA fragment of the PAI1 gene, subcloned into the EcoRI site of pGEM-1 [30]. The uPA probe consisted of a 1.5-kb cDNA fragment of the uPA gene, subcloned into the Pst I site of pBluescript SK [31]. Human uPA receptor cDNA, nucleotides 497–1081, was subcloned into pBluescript KS [32].

Wound Assay for Cell Migration

Endothelial cells were grown to confluence in Earle's MEM with 20% FBS in 6-well tissue culture plates. The wounds were made in the confluent monolayers by pressing a razor blade through the cell-sheet and gently scraping the cells essentially as described by Sato and Rifkin [33]. The cultures were rinsed with HBSS to eliminate debris. The medium was removed and replaced by Earle's MEM without serum containing 0.1% gelatine, and different concentrations of uPA, PAI1, uPA:PAI1 complex, or TGFß1. The plates were incubated at 37°C, 5% CO2 in air for 24 h. After this time, the cells were fixed with absolute methanol for 10 min and subsequently stained with Giemsa (12.5%) for 15 min. The number of cells that had migrated across the wound line were counted per high-power field. The mean of at least five high-power fields was given as the result.

Immunocytochemistry

Endometrial stromal cells were cultured on glass chamber slides coated with poly-D-lysine. The cells were briefly fixed in acetone and frozen at -80°C until processed. After thawing, slides were incubated with normal rabbit serum for 10 min, followed by monoclonal antibodies to vimentin or cytokeratin at 4°C overnight. After washing, the slides were incubated with biotinylated rabbit anti-mouse IgG for 30 min and with preformed avidin-biotin-peroxidase complex for 30 min. Immunostaining was subsequently developed with diaminobenzidine for 5 min, and nuclear counterstaining was obtained with hematoxylin.

Statistical Methods

Results were given as mean ± SEM, and the significance of differences between treatments was calculated using Wilcoxon's signed-rank test. All tests were two-sided, and a 5% level of significance was used. Differences between curves were evaluated with Fisher's protected least-significance-difference (PLSD) test.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Endometrial stromal cell cultures treated with TGFß1 contained increased conditioned medium concentrations of uPA and PAI1 (Fig. 1). Consequently, the concentration of the uPA:PAI1 complex was also increased in the medium. Elimination of complexed uPA, i.e., internalization for lysosomal degradation, is mediated by available uPA receptor binding sites on the cell surface [13]. In order to understand the extracellular accumulation of the uPA:PAI1 complex in TGFß1-treated cultures, we assayed the number of uPA receptor binding sites on the membranes of these cells. Neither the number of free available binding sites nor the number of endogenously occupied binding sites was influenced by TGFß1 (Fig. 2). Northern blotting of total RNA extracted from TGFß1-stimulated cells showed an increase of PAI1 mRNA, but not of uPA mRNA, compared to control cells (Fig. 3). Similarly, no increase was found in the uPA receptor mRNA (data not shown).



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  		FIG. 1. Confluent cultures of endometrial stromal cells were treated for two days with TGFß1 (10 ng/ml) or buffer control, and the conditioned media were analyzed for uPA (n = 26), PAI1 (n = 26), and uPA:PAI1 complex (n = 5). The media from TGFß1-treated cultures had higher concentrations of uPA (p = 0.001), PAI1 (p = 0.001), and uPA:PAI1 complex (p = 0.04) than did control cells.



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  		FIG. 2. Binding of 125I uPA (1 nM) to endometrial stromal cell cultures (n = 4) that had been treated for two days with TGFß1 (10 ng/ml) or ethanol as a control. Cells had been exposed to either acetate buffer pH 3.0, which removed endogenous uPA and allowed quantitation of the total number of receptor binding sites, or to a neutral buffer for quantitation of free, nonoccupied receptor sites. Results are expressed as femtomoles bound ligand per milligram of cellular protein. Neither the number of occupied (total-free) receptors (hatched bars) nor the number of free receptor sites (open bars) were affected by treatment with TGFß1.



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  		FIG. 3. The amount of specific mRNA for PAI1 and uPA was calculated relative to that of ß-actin in total RNA extracts from primary cultures of endometrial stromal cells (n = 6) that had been stimulated with either TGFß1 (10 ng/ml) or ethanol as a control. Autoradiograms were densitometrically scanned, and integrated optical density (IOD) data of the specific mRNAs were related to that of ß-actin mRNA. Stimulation with TGFß1 increased cellular content of PAI1 mRNA (p = 0.04), but not the content of uPA mRNA.

Since extracellular accumulation of the uPA:PAI1 complex can potentially influence adjacent cells in vivo, we tested various concentrations of the uPA:PAI1 complex, as well as its constituents uPA and PAI1, on endothelial cell migration and proliferation. Both uPA and uPA:PAI1 complex, but not PAI1 by itself, induced a dose-dependent increase of migration in both HUVEC and HMEC-1. These two cell-types represent both large-vein endothelial cells and microvascular endothelial cells (Figs. 4 and 5). Stimulation by both uPA and the complex was maximal at 1 nM. Similarities between the curves for free and complexed uPA support the conclusion that the proteolytic activity of uPA was not required for this process. However, receptor binding was required since antibodies to the uPA receptor completely blocked the stimulatory effect of both free and complexed uPA (Fig. 6). Proliferation measured as thymidine incorporation was, on the other hand, not stimulated by either by uPA, PAI1, or the uPA:PAI1 complex in HUVEC (not shown).



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  		FIG. 4. Photomicrograph showing the "wound assay" for cell migration. Confluent cultures of HUVEC were wounded with a razor blade. Upper panel shows a freshly wounded culture. Middle panel shows a culture stimulated for 24 h in the presence of PAI1 1 nM, and lower panel demonstrates treatment with uPA:PAI1 1 nM. The mean number of cells migrating across the wound line was calculated per low-power field for each treatment, and expressed as percentage of untreated cultures.



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  		FIG. 5. Cultures of HUVEC (n = 5) and HMEC-1 (n = 6) were exposed to various concentrations of the uPA:PAI1 complex, as well as its constituents uPA and PAI1. The migratory response was assayed as described in Figure 4. Migration was stimulated by free uPA as well as by uPA:PAI1 complex in HUVEC (p < 0.0001 and p < 0.0001 respectively) and HMEC-1 (p < 0.0001 and p = 0.003 respectively).



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  		FIG. 6. HUVEC (n = 3) were stimulated with uPA 1 nM, uPA:PAI1 complex 1 nM, or buffer as a control, in the presence or absence of blocking antibodies to the uPA receptor (Anti uPAR, 10 µg/ml). The stimulatory effect on migration by free as well as complexed uPA was inhibited by the antibodies.

In order to directly elucidate the influence of TGFß1-stimulated stromal cells on endothelial cell migration, confluent stromal cell cultures in 80-cm2 Petri dishes were stimulated with TGFß1 10 ng/ml or HBSS for control for 24 h in serum-free medium. Cultures were subsequently incubated for 24 h in serum-free medium without TGFß1. These conditioned media were collected, and 1.5-ml aliquots were evaluated for stimulatory effect in the HMEC migration assay. Conditioned medium from TGFß1-treated stromal cell cultures increased migration in HMEC by 60% as compared to nonconditioned medium (Fig. 7). This stimulatory effect was significantly inhibited by monoclonal antibodies to the uPA receptor, indicating that the stimulatory response involved this receptor. Conditioned medium from HBSS-treated stromal cell cultures had no significant stimulatory effect on HMEC migration.



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  		FIG. 7. HMEC were incubated with conditioned media (CM) from stromal cell cultures. The stromal cell cultures had been stimulated for 24 h with either TGFß1 (10 ng/ml) or HBSS as a control (n = 7), and subsequently incubated for 24 h in serum-free medium without TGFß1. The stimulatory effect of the conditioned media was expressed as a percentage effect of the nonconditioned medium. Conditioned media from TGFß1-treated cultures increased HMEC migration as compared to that from control cultures (p = 0.008). This increased stimulatory effect was inhibited by monoclonal antibodies to the uPA receptor, auPAR (p = 0.018).

The above results imply a possible paracrine regulatory loop involving the release by TGFß1-stimulated endometrial stromal cells of uPA:PAI1 complex, which subsequently stimulates migration in adjacent endothelial cells. We also considered an alternative model for stromal stimulation of endothelial cell migration; i.e., stromal-derived TGFß1 stimulates endothelial production of uPA, which binds to uPA-receptors on the surface of endothelial cells in an autocrine/paracrine manner, thereby stimulating migration. To test this hypothesis, we studied the migratory response in both HUVEC and HMEC-1 stimulated with TGFß1. The response to TGFß1 in HUVEC showed two clear-cut maxima at 0.001 and 1.0 ng/ml TGFß1, whereas the response in HMEC-1 was different, showing a weak monophasic response between 0.001 and 1.0 ng/ml TGFß1 (Fig. 8). However, antibodies to the uPA receptor did not reduce migration induced by 0.001 and 1.0 ng/ml TGFß1 in HUVEC. This indicates that the effect of TGFß1 was a direct one, and did not involve ligand binding to the uPA receptor (Fig. 9).



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  		FIG. 8. Migratory response of HUVEC (n = 3) and HMEC-1 (n = 5) to various concentrations of TGFß1. HUVEC showed a biphasic response, with a maximal stimulation at 0.001 ng/ml and 1.0 ng/ml. The response of HMEC-1 was weaker and monophasic with a flat peak between 0.001 and 1.0 ng/ml.



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  		FIG. 9. Confluent cultures of HUVEC were wounded and stimulated with TGFß1 0.001 ng/ml and 1.0 ng/ml in the presence or absence of antibodies to the uPA receptor (Anti uPAR). The migratory response to TGFß1 was not affected by the antibodies, indicating that this effect of TGFß1 does not involve ligand binding to the uPA receptor.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Our results show that endometrial stromal cells in culture stimulated with TGFß1 increase their extracellular levels of uPA, PAI1, and uPA:PAI1 complex. The increase of PAI1 is a transcriptional effect, since we have demonstrated that stimulation with TGFß1 of stromal cells that had been transfected with an 800-basepair (bp) sequence of the PAI1 promoter fused to a reporter gene activates a site within the transfected sequence [34]. Increased uPA was not associated with an increase of uPA mRNA but may be secondary to an increased translation or to other posttranslational effects. Increased concentrations of PAI1 in the conditioned medium resulted in an increased formation of the uPA:PAI1 complex. This complex can be endocytosed and degraded by a process that requires free available uPA receptors. Progesterone treatment, like TGFß1 treatment, induces PAI1 production in these cells. However, degradation of uPA is increased and extracellular concentrations of uPA are decreased as a result of an increased number of free uPA receptor sites in progesterone-treated cells [13]. The observation that the number of free receptor sites was not increased and the concentration of uPA was not decreased in TGFß1-treated cells suggests that the number of uPA receptor sites is rate-limiting for the degradation of uPA.

The binding of uPA to its receptor has been shown to induce intracellular phosphorylation processes involving protein tyrosine kinase [35]. Cellular effects that follow the binding of uPA or its amino-terminal fragment to the receptor include proto-oncogene expression [36], mitogenic effects [37], cell adhesion [38, 39], or migration [40, 41], depending on cell type. All these effects are independent of the proteolytic activity of uPA, but require its receptor binding site. Similar binding affinities have been shown for complexed and free uPA in different cells [42] including endometrial stromal cells [43], suggesting that the binding of the complex to uPA receptors can potentially induce the same cellular effects as those described for free uPA. Since, however, the number of uPA receptor binding sites was limited in TGFß1-treated stromal cells, it is possible that a binding of the complex occurs to receptor sites on adjacent cell types. To test this hypothesis, we exposed endothelial cells to various concentrations of uPA:PAI1 complex, free uPA, and PAI1, and found that both free and complexed uPA stimulated endothelial cell migration in a dose-dependent way. The stimulatory curves were almost identical, which agrees with previous observations that the active site of uPA is not needed for the binding to the receptor nor for initiating signal transduction and cellular effects [35–41]. On the other hand, monoclonal antibodies to the uPA receptor inhibited the stimulatory effect of both free and complexed uPA, indicating that the receptor binding site in the A-chain of uPA is required for the functions. Urokinase PA, free or complexed, released from endometrial stromal cells induced by TGFß1, could function as a paracrine signal to stimulate endothelial cell migration. Conditioned medium from TGFß1-treated stromal cell cultures, but not that from unstimulated cultures, significantly increased endothelial cell migration. We furthermore found that this effect was inhibited by the uPA receptor antibodies, suggesting that the increased migration of endothelial cells results from an increased release of uPA from TGFß1-treated stromal cells.

TGFß is reportedly angiogenic in vivo [17–19]. The mechanism by which TGFß induces angiogenesis, however, remains debatable since it has been reported to inhibit endothelial cell proliferation [20, 21] and migration [44, 45]. One possible explanation for the discrepancy between in vivo and in vitro responses to TGFß may be that in vivo effects of TGFß are mediated via other cells in a paracrine way. For example, TGFß is chemotactic for macrophages, which, once attracted to a site of inflammation/wound healing, are capable of releasing additional growth factors such as tumor necrosis factor alpha, itself a potent inducer of angiogenesis [46–48].

There is experimental evidence to indicate that endometrial angiogenesis is regulated by endometrial cells, i.e., endometrial explants as well as separated stromal and epithelial cells influence vascularization in a modified chorioallantoic membrane assay, and cocultures of endothelial cells with endometrial tissue explants stimulate their migration [3, 49]. Endothelial cell mitosis and microvascular density in endometrial biopsies does not, however, vary between the menstrual phases, and it has been suggested that endothelial mitosis is a continuous phenomenon over the cycle, whereas angiogenesis during endometrial repair and growth mainly involves endothelial cell migration [2, 50]. In fact, endometrial tissue obtained in the proliferative phase stimulated endothelial cell migration to a greater extent than tissue obtained in the secretory phase [3]. These authors noted two distinct peaks in the stimulation of endothelial cell migratory activity by proliferative endometria. One occurred postmenstrually and was suggested to coincide with endometrial repair. The other occurred in the mid-late proliferative phase, when endometrial growth is maximal.

Since the amount of TGFß1 mRNA is highest in late secretory and menstrual endometrium [11], and high PA activity, which can initiate activation of latent TGFßs, is released premenstrually [14–16], we have suggested that TGFß1 participates in the postmenstrual repair process. The results in this study show that TGFß1 can stimulate endometrial stromal cells to increase their extracellular concentrations of the uPA:PAI1 complex, which stimulates endothelial cell migration. The actual concentrations we measured in the conditioned media may not accurately reflect in vivo concentrations in the pericellular space, since the volumes in vivo are minute compared to those in our condition media. The in vivo concentration is furthermore subject to variations due to the local binding of the complex to extracellular matrix proteins like vitronectin, heparin, and laminin-nidogen [51–53]. Merely the fact that the complex is accumulated, however, poses a possibility that it has paracrine effects. The uPA:PAI1 complex, which is enzymatically inactive, may travel as a paracrine signal in the pericellular space without being a proteolytic hazard.

We found no effects of free or complexed uPA on thymidine incorporation in the endothelial cells. Such effects, however, may be dependent on tissue culture conditions, since it has been shown that the response of endothelial cells to stimulants like TGFß1 is dependent upon cell shape, proliferative state, and the nature of the substratum [54].

The stimulatory effect of uPA and uPA:PAI1 complex was demonstrated in both umbilical vein endothelial cells and microvascular endothelial cells, and may also be valid for endometrial microvascular endothelial cells. There were, however, differences between the two studied cell types with respect to the migratory response to TGFß1. HUVEC cultures had two distinct maxima, at 0.001 ng/ml and 1 ng/ml, whereas the response of HMEC-1 cultures was monophasic and much weaker. This difference may reflect the different functions of the endothelium of veins and capillaries. Since the migratory response elicited by basic fibroblast growth factor in bovine aortic endothelial cells was secondary to the induction and autocrine binding of uPA to the uPA receptor [40], we examined the possibility that a similar autocrine mechanism might be responsible for the migratory response to TGFß1 in HUVEC and HMEC-1. In contrast to migration in response to uPA and the uPA:PAI1 complex, migration in response to TGFß1 was not inhibited by antibodies to the uPA receptor. This indicates that endothelial cell migration in response to TGFß1 does not involve the uPA receptor. Our finding that TGFß1 stimulates endothelial cell migration stands in contrast to some earlier studies [44, 45]. Different culture conditions, experimental systems, and source of cells can probably explain this discrepancy.


   FOOTNOTES
 
1 This study was supported by grants from the Swedish Cancer Fund (2693-B94–07XCC), the Lund University Medical Faculty, the Lund University Hospital Fund for Cancer Research, and the Gunnar, Arvid, and Elisabeth Nilssons Fund for Cancer Research. Back

2 Correspondence. FAX: 46 46 157868; bertil.casslen{at}gyn.lu.se Back

Accepted: May 11, 1998.

Received: June 12, 1997.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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