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BOR - Papers in Press, published online ahead of print June 25, 2003.
Biol Reprod 2003, 10.1095/biolreprod.103.015669
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BIOLOGY OF REPRODUCTION 69, 1515–1524 (2003)
DOI: 10.1095/biolreprod.103.015669
© 2003 by the Society for the Study of Reproduction, Inc.

Pregnancy

Induction of p38 Mitogen-Activated Protein Kinase-Mediated Apoptosis Is Involved in Outgrowth of Trophoblast Cells on Endometrial Epithelial Cells in a Model of Human Trophoblast-Endometrial Interactions1

Hsin-Yang Li3,5, Sheng-Ping Chang5, Chiou-Chung Yuan5, Hsiang-Tai Chao5, Heung-Tat Ng5, and Yen-Jen Sung2,4

Institute of Clinical Medicine3 Institute of Anatomy and Cell Biology,4 School of Medicine, University System of Taiwan-National Yang-Ming University, Taipei, Taiwan 112, Republic of China Department of Obstetrics and Gynecology,5 Veterans General Hospital, Taipei, Taiwan 112, Republic of China


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
During embryo implantation in species with hemochorial placentation, such as the mouse and human, trophoblast cells of the attached blastocyst penetrate the luminal epithelium of the endometrium before invasion into the endometrial stroma. Signs of apoptosis were demonstrated in luminal endometrial epithelial cells (EEC) adjacent to the trophoblast cells; however, the signaling mechanisms leading to apoptosis in EEC remain unclear. Because mitogen-activated protein kinases (MAPK) were shown to mediate apoptosis in several model systems and found to be activated in the uterus during decidualization, the possible involvement of MAPK during trophoblast-EEC interactions was studied. By coculturing BeWo human trophoblast spheroids with RL95-2 human EEC monolayers to mimic the blastocyst-endometrial interaction, we found that most spheroids rapidly attached to EEC monolayers and then progressively expanded, with marked dislodgment of EEC adjacent to the spreading trophoblast cells. Immunoblotting analysis showed that both p38 MAPK and extracellular signal-regulated kinase (ERK) were activated in EEC after coculture. However, only SB203580 (a p38 MAPK inhibitor), but not PD98059 (an ERK inhibitor), inhibited trophoblast outgrowth on EEC monolayers through the suppression of p38 MAPK activation in EEC. Furthermore, trophoblast expansion caused prominent EEC apoptosis at the spheroid-EEC interface, as detected by annexin V labeling and valyl-alanyl-aspartyl-[O-methyl]-fluoromethylketone (which binds activated caspases) staining, and SB203580 significantly decreased the percentage of apoptotic cells. Our results, based on a model of human trophoblast-EEC interactions, establish that trophoblast cells cause activation of p38 MAPK in EEC and, consequently, induce apoptosis and displacement of EEC, a process that may facilitate implantation.

apoptosis, implantation, signal transduction, trophoblast, uterus


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Embryo implantation is a crucial event in the establishment of a successful pregnancy [1]. The process of implantation begins with apposition and attachment of the hatched blastocyst to the luminal epithelium of the uterus [2]. In species with hemochorial placentation, including the mouse and human, trophoblast cells of the attached blastocyst penetrate the luminal epithelium and invade the endometrial stroma [3, 4]. Although trophoblast cells were shown to produce metalloproteinases that degrade extracellular matrix in the endometrial stroma [5, 6], it remains unclear how these cells cross the epithelial barrier of the endometrium before reaching the endometrial stroma. In rodent models, ultrastructural signs of apoptosis were observed in uterine epithelial cells surrounding the embryos, suggesting the importance of programmed cell death in implantation [7, 8]. In vitro culture of mouse blastocysts on endometrial epithelial cell (EEC) monolayers also displayed trophoblast outgrowth that was accompanied by apoptosis and dislodgment of EEC around the expanding trophoblast cells [9], indicating a basis of cell-cell interaction for the induction of apoptosis. Evidence of EEC apoptosis was also demonstrated at implantation sites of primates [10], and human blastocysts were found to induce apoptotic reaction in EEC during in vitro coculture [11]. Several secretory and membrane-bound factors, such as transforming growth factor ß and Fas ligand (FasL), were found to be involved in the trophoblast-EEC interaction [9, 11], but the intracellular signaling pathways leading to trophoblast-induced EEC apoptosis remain to be determined.

Members of the highly conserved mitogen-activated protein kinase (MAPK) superfamily, including extracellular signal-regulated kinase (ERK), p38 kinase, and c-Jun NH2-terminal kinase, regulate diverse cellular processes in response to a plethora of extracellular stimuli [12]. Recent studies showed that intraluminal application of a deciduogenic stimulus could activate ERK and p38 MAPK in the mouse uterus [13]. Several factors secreted by trophoblast cells, such as nitric oxide, progesterone, and interleukin-1ß, were also shown to activate p38 MAPK in uterine cells [14, 15]. These findings suggest an important role of MAPK activation in embryo implantation; however, the mechanisms through which MAPK participates in the process of implantation have yet to be defined. Recent studies established that MAPK pathways are critical regulators of apoptosis [16, 17]. For example, p38 MAPK activated by the high concentrations of nitric oxide and progesterone normally present at the trophoblast-decidual interface could induce apoptosis in EEC [14]. In addition, the Fas/FasL death system was found to be active at the embryo-endometrial interface [11, 18, 19], and in various cell models, activation of members of the MAPK superfamily was demonstrated to either mediate Fas-induced apoptosis or enhance FasL expression [20–22]. Therefore, it is likely that trophoblast cells may cause apoptosis in EEC through the activation of members of the MAPK superfamily, thereby enabling successful penetration through the luminal epithelium of the uterus.

The purpose of the present study was to examine the roles of MAPK pathways in trophoblast-EEC interactions. The BeWo trophoblast line and the RL95-2 EEC line were selected because they retain many properties of their respective normal tissues (i.e., trophoblast and uterine epithelium) [23–25]. We generated spheroidal cell masses from the BeWo cell line to mimic blastocysts and then cocultured these trophoblast spheroids with EEC monolayers formed by the RL95-2 cell line. Immunoblotting assays were performed to determine whether MAPK pathways were activated in EEC monolayers on spheroid attachment and spreading. Subsequently, the effects of MAPK inhibition on trophoblast outgrowth on EEC monolayers were examined. Our results may have clinical ramifications, because they may clarify the physiological significance of MAPK pathways in the implantation process.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
EEC Culture

The human endometrial carcinoma cell line RL95-2 (CRL 1671) was purchased from the American Type Culture Collection (ATCC, Rockville, MD) through the National Health Research Institute Cell Bank (Hsin-Chu, Taiwan) [24, 26, 27]. Cells were maintained at 5% CO2 and 37°C in a 1:1 mixture of Dulbecco modified Eagle medium (Life Technologies, Grand Island, NY) and Ham F-12 (Life Technologies) supplemented with 10% fetal calf serum (FCS; Biological Industries, Kibbutz Beit Haemek, Israel), 10 mM Hepes (BDH, Poole, U.K.), and 5 µg/ml of bovine insulin (Sigma Chemical Co., St. Louis, MO). The cells were subcultured every 3 days by trypsinization (trypsin-EDTA solution; Life Technologies).

Preparation of Trophoblast Spheroids

The human BeWo choriocarcinoma cells (CCL-98; ATCC) were used to produce trophoblast spheroids, which have been employed to study trophoblast adhesion and invasion mechanisms [28, 29]. Cells were cultured in Ham F-12K nutrient mixture (Life Technologies) supplemented with 15% FCS at 5% CO2 and 37°C and subcultured every 3 days with trypsinization. For the preparation of spheroids, BeWo cell concentration was adjusted to 2 x 105 cells/ml, and 10 ml of the cell suspension were plated on a 100-mm plastic Petri dish (Alpha Plus, Taoyuan, Taiwan) in complete growth medium [30, 31]. Spontaneous cell aggregation resulted in abundant spheroidal cell masses in 24-h culture. Spheroids of 50–100 µm in diameter (sizes close to that of an implanting blastocyst) were selected under a dissecting microscope (Nikon SMZ645; Nikon Corp., Tokyo, Japan) for all subsequent experiments.

Immunoblotting Assay for MAPK

Assessment of the phosphorylation of MAPK in EEC in response to spheroid attachment and spreading was accomplished by immunoblotting. The RL95-2 cells were plated in 60-mm culture dishes at 2 x 107 cells/dish in 5 ml of complete culture medium. After 48-h incubation, the culture medium was replaced by serum-free medium supplemented with 1 mg/ml of bovine serum albumin (USB, Cleveland, OH), and serum starvation was continued for 16–18 h, followed by the addition of 500 spheroids to each dish containing the confluent monolayer of EEC. After the indicated coculture time, the supernatant was discarded, and the cells were washed with PBS. The cells were then lysed by keeping the dish on ice for 5 min after the addition of 40 µl of lysis buffer (20 mM Tris [pH 7.5], 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% Triton, 2.5 mM sodium pyrophosphate, 1 mM ß-glycerophosphate, 1 mM Na3VO4, 1 µg/ml of leupeptin, and 1 mM phenylmethylsulfonyl fluoride). The lysed cells were then scraped off the dish, transferred to microcentrifuge tubes, and sonicated. The cell lysates were centrifuged at 18 000 x g, 4°C for 10 min, to remove insoluble substances, and 30 µg of supernatant protein were used in 10% SDS-polyacrylamide gel electrophoresis. After electrophoresis was completed, the fractionated cellular proteins were transferred to polyvinylidene fluoride membranes (NEN Life Science Products, Inc., Boston, MA), followed by blocking with 5% nonfat milk in TBST (25 mM Tris [pH 7.5], 150 mM NaCl, 2.7 mM KCl, and 0.1% [v/v] Tween 20) and incubated overnight with one of the following primary antibodies: rabbit anti-human phosphorylated ERK (phospho-ERK) antibody, rabbit anti-human ERK antibody, rabbit anti-human phosphorylated p38 MAPK (phospho-p38 MAPK) antibody, or rabbit anti-human p38 MAPK antibody (all from Cell Signaling Technology, Beverly, MA). The polyvinylidene fluoride membranes were then extensively washed with TBST and incubated for 60 min with the secondary antibody (goat anti-rabbit antibody; Transduction Laboratories, Lexington, KY). After extensive washing with TBST, the immune complexes were detected by chemiluminescence using the Western Blotting kit from NEN Life Science Products. Quantitation of MAPK activation was accomplished by measuring the gray-scale density of each band on the immunoblot using the Scion Image software (based on NIH Image; Scion Corporation, Frederick, MD) following the user's manual.

Measurement of Spheroid Expansion on EEC Monolayers

Trophoblast outgrowth on the EEC monolayer occurred approximately 60 min after spheroid-EEC coculture, when most spheroids became attached to the monolayer. After 24 h of coculture, these spheroids flattened and expanded, with the areas of trophoblast outgrowth being several-fold the original spheroid sizes. Hematoxylin-eosin staining was first performed to determine whether EEC were displaced by trophoblast outgrowth. A quantitative method was also developed to further examine the effects of MAPK inhibitors on spheroid expansion on EEC monolayers. The RL95-2 cells were plated at 3 x 105 cells per 100 µl per well in 96-well plates that had been marked earlier with quadrants on the exterior surface under each well to enable localization of the same sets of spheroids at different coculture time points. After 16 h of incubation, EEC grew into confluent monolayers, and fresh complete RL95-2 culture medium containing 10% FCS was changed. Then, EEC monolayers were treated with 1–20 µM SB203580 (a p38 MAPK inhibitor; Calbiochem-Novabiochem, San Diego, CA), 1–20 µM PD98059 (an ERK inhibitor; Calbiochem-Novabiochem), or the corresponding vehicles for 60 min, followed by the addition of trophoblast spheroids to the EEC monolayers at approximately 10 spheroids/well. The MAPK inhibitors were dissolved in dimethyl sulfoxide and diluted appropriately with culture medium to give the desired final concentration in cell culture (final concentration of dimethyl sulfoxide in cell culture was 0.38% for treatment with SB203580 and 0.053% for treatment with PD98059). The spreading of spheroids was observed at different coculture intervals (1, 7, 14, or 24 h) under an inverted microscope (Nikon Diaphot) and photographed using a cooled charge-coupled device camera system (Photometrics CoolSNAP fx; Roper Scientific Inc., Tucson, AZ). To define the margins of trophoblast outgrowth, the spheroid-EEC coculture was labeled with 10 µM 5-chloromethylfluorescein diacetate (CellTracker Green CMFDA; Molecular Probes, Eugene, OR), a nontoxic fluorescent probe, for 30 min before each time of photography. Because individual BeWo trophoblast cells spread far more extensively on the culture plate than the RL95-2 EEC, CellTracker staining appeared relatively dim for the BeWo cells and bright for the RL95-2 cells (see Fig. 1). Such clear contrast in staining feature enabled unambiguous demarcation of the boundary of trophoblast outgrowth on the RL95-2 cell lawn. The areas of spreading spheroids were measured using the Scion Image software system. The spheroid areas at 1 h of coculture were considered as their original sizes, and fold-expansion in spheroid areas was calculated for 7, 14, and 24 h of coculture, respectively.



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  		FIG. 1. Morphological changes in the BeWo trophoblast spheroid and RL95-2 EEC monolayer after indicated intervals of spheroid-EEC coculture. A) Expansion in spheroid areas after indicated intervals of spheroid-EEC coculture. Photographs shown are the results of the same spheroid at 1, 7, and 24 h after coculture; arrows indicate the margins of the spheroid-EEC interface. B) Staining of the spheroid-EEC coculture at 24 h with the fluorescent probe CellTracker Green CMFDA. C) Hematoxylin-eosin staining of the spheroid-EEC coculture at 24 h. Pictures shown are representative of three independent experiments. Bar = 50 µm

Detection of Apoptotic Cells

Apoptotic cells were detected with annexin V labeling [32] for phosphatidylserine, which were translocated from the inner to the outer leaflet of the plasma membrane in early apoptosing cells, and valyl-alanyl-aspartyl-[O-methyl]-fluoromethylketone (VAD-FMK) staining for cells expressing activated caspases [33]. Percentages of apoptotic cells were also calculated. Briefly, monolayers of EEC were prepared on eight-well chamber slides (Nalge Nunc International Corp., Naperville, IL) by plating RL95-2 cells at 1 x 106 cells/well in 300 µl of complete culture medium and then incubating overnight. Next, 10 spheroids were delivered into each well in the presence or absence of the MAPK-inhibitors SB203580 and PD98059. In some wells, EEC monolayers were kept void of spheroids to serve as controls. After 24 h of coculture, annexin V labeling or VAD-FMK staining was performed to detect apoptotic cells. For annexin V labeling, the culture wells were first rinsed twice with PBS and replenished with 100 µl of an annexin-binding buffer (10 mM Hepes, 140 mM NaCl, and 2.5 mM CaCl2, pH 7.4). Then, 5 µl of the fluorescein-5-isothiocyanate-conjugated annexin V (FITC-annexin V; Molecular Probes) were added to each well, and the cells were incubated at room temperature for 15 min. In some experiments, propidium iodide (1 µg/ml) was added to the samples for identifying cells with membrane leakage that may cause a false-positive reaction during annexin V staining. After the incubation, cells were washed with annexin-binding buffer and examined under a fluorescence microscope (Olympus BX50; Olympus, Tokyo, Japan) following removal of the culture-chamber frames from the chamber slide.

For VAD-FMK staining, 10 µM FITC-conjugated VAD-FMK (CaspACE FITC-VAD-FMK in situ marker; Promega, Madison, WI), which irreversibly binds activated caspases [34], was added to cells at the end of coculture. After 20 min of incubation at 37°C and 5% CO2, cells were washed twice with PBS and examined under a fluorescence microscope.

To determine the percentage of apoptotic cells in total cells, all cells were harvested by trypsinization, thoroughly but gently mixed, stained with FITC-annexin V or FITC-VAD-FMK, and then mounted on a slide using a cell centrifuge (Cytospin 3; Shandon, Cheshire, U.K.) for examination under a fluorescence microscope. Briefly, RL95-2 EEC monolayers were prepared in 96-well plates by seeding the cells at 3 x 105 cells per 100 µl per well and then incubating overnight. Next, 10 spheroids were delivered into each well in the presence or absence of SB203580 and PD98059. In each plate, three to six wells of EEC monolayers were kept void of spheroids to serve as controls. After 24 h of coculture, cells that had detached from the plate were collected by aspiration. Cells that had remained attached, on the other hand, were collected by trypsinization and pooled with the detached cells. All collected cells were then stained with FITC-annexin V or FITC-VAD-FMK as described above. Cell concentration was adjusted to 2 x 104 cells/ml, and 100 µl of cell suspension were subjected to cytospin at 45 x g for 5 min. The slides were examined under a fluorescence microscope to count the number of annexin V-positive or activated caspase-positive cells. The total number of cells was determined by staining with 2 µg/ml of DAPI (4',6'-diamidino-2-phenylindole). The percentage of annexin V-positive or activated caspase-positive cells in total cells was then calculated.

Statistical Analysis

Data regarding the densitometric analyses, spheroid spreading assays, and apoptosis assays were expressed as the mean ± SEM. Statistical significance between groups was determined by one-way ANOVA, followed by the Fisher post hoc least-significant-difference (LSD) test. Concentration-dependent effect was analyzed by simple linear regression of all response data against concentration levels of the treatment. All analyses were performed using the SAS program (SAS Institute Inc., Cary, NC) on a Pentium IV-based personal computer.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Morphodynamics of Trophoblast Spheroid Spreading on EEC Monolayers

The spheroids began to flatten and expand on EEC monolayers by 60 min of coculture, as was also observed in our previous study [35]. Approximately 2-fold (191% ± 12% of the original sizes, data obtained from three independent experiments) and 4-fold (425% ± 54% of the original sizes, data obtained from three independent experiments) expansion in spheroid areas, as measured using the NIH Image-based Scion Image software (see Materials and Methods), were observed at 7 and 24 h, respectively (Fig. 1A). Staining of the cocultured cells with a fluorescent vital dye, CellTracker Green, showed that the brightly stained RL95-2 endometrial cell lawn was displaced by regions of dimly stained BeWo trophoblast cells (Fig. 1B). Hematoxylin-eosin staining further demonstrated that RL95-2 EEC, with nuclei distinctively smaller than those of BeWo trophoblast cells, were largely absent in the trophoblast region, confirming the dislodgment of EEC by spheroid spreading (Fig. 1C).

Effect of Trophoblast Spheroid Attachment and Spreading on MAPK Activation in EEC

The MAPK are activated by phosphorylation of specific tyrosine and threonine residues, and the relative levels of phosphorylated MAPK in total MAPK represent the degree of MAPK activation. To determine whether p38 MAPK and ERK were activated during the physical interaction between trophoblast cells and EEC, monolayers of EEC grown on 60-mm culture dishes were confronted by spheroids, and total lysates (including both trophoblast cells and EEC) were collected after various coculture intervals. For the detection of MAPK activation, protein samples were subjected to immunoblotting assays to measure levels of phosphorylated MAPK relative to total MAPK. Two bands of phospho-p38 MAPK were detected by the anti-human phospho-p38 MAPK antibody used in the present study (Fig. 2A, phospho-p38). The lower band of phospho-p38 MAPK in spheroid-EEC coculture remained at basal levels, but the level of the upper band (Fig. 2A, arrow) of phospho-p38 MAPK increased significantly after 7 h of spheroid-EEC coculture and remained elevated at 24 h as compared with that of EEC samples not yet exposed to spheroids (designated as 0 min). Densitometric analysis of the immunoblots from three independent experiments using the Scion/NIH Image software also showed a significant increase in the levels of the upper band of phospho-p38 MAPK at 7 and 24 h of coculture as compared with control EEC levels at 0 min (P < 0.05 between the 7-h coculture and 0-min groups, P < 0.05 between the 24-h coculture and 0-min groups; n = 3) (Fig. 2B). No significant difference was observed in p38 MAPK phosphorylation between 7- and 24-h coculture (P > 0.05 between the 7- and 24-h coculture groups) (Fig. 2B). Samples from BeWo spheroids alone exhibited basal levels of p38 MAPK activity similar to those of the "naïve" EEC. On the other hand, the exposure of EEC monolayers to spheroids evoked a marked rise in phospho-ERK within 10–60 min of coculture, with the level of phospho-ERK returning to baseline after 7 h (Fig. 2C, phospho-ERK1 and phospho-ERK2). Densitometric analysis of the immunoblots from three independent experiments also confirmed the above statement (Fig. 2D). The degree of ERK activation remained at basal levels for samples from BeWo spheroids alone. Thus, both p38 MAPK and ERK were activated during spheroid-EEC coculture.



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  		FIG. 2. MAPK activation in EEC after coculturing with trophoblast spheroids. Monolayers of RL95-2 EEC were exposed to BeWo spheroids and cocultured for various intervals as indicated, followed by protein extraction. Immunoblotting was performed to detect the degree of MAPK activation (relative levels of phosphorylated MAPK in total MAPK), and the results were subjected to densitometric analysis. The level of MAPK activation in "naïve" EEC without spheroid coculture (0 min) was arbitrarily set as 1.0. Data are expressed as the mean ± SEM. A) An immunoblot representative of three independent experiments showing phosphorylation of p38 MAPK (phospho-p38) in EEC after coculture. Arrow indicates the upper band of phospho-p38 MAPK. Total p38 MAPK (total p38) served to normalize loading differences. B) Densitometric analysis of results from three independent experiments as performed in A. Asterisk indicates significant difference (P < 0.05) as compared with groups other than the 7- and 24-h coculture groups. C) An immunoblot representative of three independent experiments showing phosphorylation of ERK (including phospho-ERK1 and phospho-ERK2) in EEC after coculturing with spheroids. Total ERK (including total ERK1 and total ERK2) served to normalize loading differences. D) Densitometric analysis of results from three independent experiments as performed in C. Asterisk indicates significant difference (P < 0.05) as compared with groups other than the 10-min, 30-min, and 1-h coculture groups. P, Positive control (murine BV2 microglia treated with 3 mM sodium nitroprusside for 60 min); PD, EEC monolayers pretreated with 10 µM PD98059 and then cocultured with spheroids for 30 min; SB, EEC monolayers pretreated with 10 µM SB203580 and then cocultured with spheroids for 7 h; Sph, BeWo spheroids alone

Because such MAPK activation, as detected by immunoblotting, could arise from either BeWo or RL95-2 cells, we then investigated the origin of protein in the coculture samples. We delivered the same amounts of spheroids (~500 spheroids/dish) as used in the above experiments to 60-mm culture dishes with or without EEC monolayers. After the indicated interval of incubation at 37°C and 5% CO2, total protein was extracted and subjected to quantification. The amount of protein arising from EEC in spheroid-EEC coculture was calculated by subtracting the protein amount of spheroids cultured alone from that of the spheroid-EEC coculture sample. In this way, the percentages of protein originating from EEC in spheroid-EEC coculture were estimated to be 95.8% ± 0.5% and 93.6% ± 0.6% at 1 and 24 h of coculture, respectively (n = 3).

Roles of MAPK Pathways in Trophoblast Spheroid Outgrowth on EEC Monolayers

Trophoblast spheroids flattened and expanded on EEC monolayers as the spheroid-EEC coculture time increased, with marked dislodgment of surrounding EEC (Fig. 1). Because both p38 MAPK and ERK were activated by physical contact between EEC and trophoblast cells as shown above, we examined whether MAPK activation was essential for trophoblast spheroid outgrowth on EEC monolayers. We introduced inhibitors of p38 MAPK (SB203580) and ERK (PD98059) to EEC monolayers 60 min before the delivery of trophoblast spheroids. The concentrations of SB203580 and PD98059 were adjusted between 1 and 20 µM, as commonly used in numerous other studies [36, 37]; concentrations of these two MAPK inhibitors greater than 20 µM were cytotoxic (data not shown) and, hence, were not employed. After various intervals of coculture in the presence or absence of MAPK inhibitors, spheroids were photographed, and areas of spheroid spreading on EEC monolayers were compared. As shown in Figure 3A, PD98059 appeared to have minimal effects on spheroid outgrowth, whereas SB203580 inhibited spheroid expansion on EEC monolayers at 24 h in a concentration-dependent manner (P < 0.001 and R2 = 0.949 as analyzed by simple linear regression of data from three independent experiments; areas of two to three spheroids were measured in each experiment, and the results of fold-expansion in spheroid areas were averaged). Because 10 µM SB203580 reached near-maximum inhibition of spheroid expansion on EEC monolayers in our experiments (P < 0.05 for 10 µM group vs. control and 5 µM groups and P > 0.05 for 10 µM vs. 20 µM groups as analyzed by one-way ANOVA followed by post-hoc LSD test), this concentration of SB203580 was employed unless otherwise indicated. Immunoblotting analysis showed that 10 µM SB203580 and 10 µM PD98059, as used in the present study, effectively inhibited p38 MAPK and ERK activation, respectively (Fig. 2).



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  		FIG. 3. Effects of MAPK inhibitors on spheroid expansion. A) Effect of MAPK inhibitors on spheroid expansion on EEC monolayers. The RL95-2 EEC monolayers grown in the 96-well plate were treated with 1–20 µM SB203580 (SB), 1–20 µM PD98059 (PD), or an equal volume of vehicle control (C) 1 h before the addition of BeWo trophoblast spheroids. Data from three independent experiments are presented as fold-expansion in spheroid areas after 24 h of coculture relative to their original sizes and are expressed as the mean ± SEM. Asterisk indicates significant difference (P < 0.05) as compared with vehicle-treated controls. B) A representative set of photographs from three independent experiments as performed in A. Bar = 100 µm. C) Effects of MAPK inhibitors on spheroid expansion on culture surfaces without EEC. BeWo spheroids were cultured in 96-well plates without EEC in the presence or absence of 10 µM SB203580 (SB) or 10 µM PD98059 (PD). The control well (C) received an equal volume of vehicle. Data from three independent experiments are presented as fold-expansion in spheroid areas after 24 h of culture relative to their original sizes and are expressed as the mean ± SEM

Under coculture conditions, the inhibitory effect of SB203580 on spheroid outgrowth could be mediated by blocking spheroid expansion, by rendering EEC resistant to trophoblast invasion, or by the combination of both mechanisms. To determine whether spheroids were the targets of p38 MAPK inhibition, we cultured spheroids in the presence or absence of SB203580 in the 96-well culture plates without EEC. Photographs were acquired at 1 h (regarded as the original sizes) and 24 h of culture to measure spheroid areas. As shown in Figure 3C, trophoblast spreading on EEC-free culture surfaces was unaffected by SB203580 treatment. We then examined whether EEC were rendered resistant to spheroid invasion on p38 MAPK inhibition. First, EEC monolayers were incubated in the presence of 10 µM SB203580 for 24 h, followed by washing three times with culture medium. Spheroids were then delivered to these EEC monolayers and photographed at 1, 7, 14, and 24 h of coculture for analysis of expansion. A 58.6% ± 9.0% reduction of spheroid outgrowth [defined as (fold-expansion in spheroid area in the control group - fold-expansion in spheroid area in the drug-treated group)/(fold-expansion in spheroid area in the control group - 1)] was observed at 7 h of coculture in samples containing 10 µM SB203580-pretreated EEC (P < 0.05 between SB203580-pretreated and control groups, n = 3) (Fig. 4A). When 20 µM SB203580 was used for EEC pretreatment, a 73.3% ± 1.6% reduction of spheroid outgrowth was found at 7 h of coculture (fold-expansion in spheroid areas was 1.29 ± 0.03 in 20 µM SB203580-pretreated group vs. 2.08 ± 0.06 in the control group; P < 0.05 between 20 µM SB203580-pretreated and control groups, n = 3). Although both 10 and 20 µM SB203580 pretreatments of EEC significantly inhibited spheroid expansion as compared with the control group, no significant difference was found between 10 and 20 µM SB203580 pretreatments in reduction of spheroid outgrowth (reduction of spheroid outgrowth was 58.6% ± 9.0% and 73.3% ± 1.6% for 10 and 20 µM SB203580 pretreatment groups, respectively; P = 0.1813 between 10 and 20 µM SB203580 pretreatment groups). This inhibitory effect largely disappeared at 24 h of coculture, because only 9.6% ± 8.1% reduction of spheroid outgrowth by pretreatment with SB203580 was observed at 24 h of coculture (Fig. 4A). To further illustrate the role of EEC p38 MAPK, spheroids were treated with SB203580 for 24 h, washed three times, and then delivered to EEC monolayers. Outgrowth of these SB203580-pretreated spheroids on EEC monolayers was not affected throughout the coculture period (Fig. 4B).



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  		FIG. 4. Differential effects of MAPK inhibition of trophoblast cells versus EEC on spheroid outgrowth. A) Spheroid expansion on MAPK inhibitor-pretreated EEC monolayers. The RL95-2 EEC monolayers grown in the 96-well plate were treated with 10 µM SB203580, 10 µM PD98059, or vehicle control for 24 h. After removal of supernatants and washing with culture medium, BeWo spheroids were added for coculture, and spheroid expansion was examined at 1, 7, 14, and 24 h of coculture. Data from three independent experiments represent fold-expansion in spheroid areas after the indicated coculture time and are expressed as the mean ± SEM. Asterisk indicates significant difference (P < 0.05) as compared with vehicle-treated controls. B) Effects of pretreatment of spheroids with MAPK inhibitors on trophoblast outgrowth. BeWo spheroids were incubated in the presence of 10 µM SB203580, 10 µM PD98059, or vehicle for 24 h, followed by three washings with culture medium. The pretreated spheroids were then added to EEC monolayers grown on the 96-well plate. Spheroid expansion assay was performed as in A, and data from three independent experiments are presented as the mean ± SEM

Involvement of the p38 MAPK Pathway in EEC Apoptosis Caused by Trophoblast Spheroid Outgrowth

The findings of the above experiments established a central role for p38 MAPK activation in the dislodgment of EEC by outgrowing trophoblast spheroids. Because p38 MAPK activation could mediate apoptosis in various cell models [16], we postulated that an encounter of EEC with trophoblast cells may activate p38 MAPK and induce apoptosis in EEC, thereby facilitating trophoblast outgrowth. To verify this postulation, we examined two hallmarks of apoptosis: phosphatidylserine translocation from the inner to the outer leaflet of the plasma membrane [38] and caspase activation [39]. For detection of these apoptosing cells, we first stained the samples with FITC-annexin V, which could bind to phosphatidylserine exposed on the plasma membrane [40]. The EEC monolayers that were kept free of spheroids showed only weak staining with annexin V (Fig. 5A, EEC Only). In contrast, after 24 h of coculture, cells surrounding the expanding spheroid were stained intensely with annexin V (Fig. 5A, Sph+EEC). These cells were impermeable to propidium iodide (data not shown), excluding the possibility of membrane leakage. Most of the apoptotic cells were located at the spheroid-EEC interface; only a few apoptotic cells were found within the spheroid region. Furthermore, these apoptotic cells were morphologically distinguished from trophoblast cells of the expanding spheroids; hence, they were most likely EEC. In addition, the presence of SB203580, but not PD98059, in the coculture markedly diminished the intensity of annexin V staining in cells surrounding the spheroid (Fig. 5A, PD+Sph+EEC and SB+Sph+EEC). Because a certain portion of cells might have become detached as a result of apoptosis, these detached cells were collected by aspiration and then pooled with the attached cells that were collected by trypsinization. Annexin V staining of the pooled samples was performed, and the percentage of apoptotic cells was derived by enumerating annexin V-positive cells and total cells visualized by DAPI staining. As shown in Figure 5B, the percentage of apoptotic cells increased significantly in the coculture as compared with the naïve EEC without spheroid coculture (detached plus attached cells; P < 0.05 between the Sph+EEC and EEC Only groups, n = 3). Moreover, SB203580, but not PD98059, could significantly decrease the percentage of annexin V-positive cells in spheroid-EEC coculture (Fig. 5B, detached plus attached cells; P < 0.05 between the Sph+EEC and SB+Sph+EEC groups, n = 3). The percentage of annexin V-positive apoptotic cells in the attached cells also displayed a similar response profile (Fig. 5B, attached cells; P < 0.05 between the Sph+EEC and EEC Only groups as well as between the Sph+EEC and SB+Sph+EEC groups, n = 3).



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  		FIG. 5. Annexin V staining for the detection of apoptotic cells in spheroid-EEC coculture. A) Monolayers of EEC were cocultured with BeWo spheroids for 24 h in the absence of MAPK inhibitors (Sph+EEC) or in the presence of 10 µM SB203580 (SB+Sph+EEC) or 10 µM PD98059 (PD+Sph+EEC). To serve as controls, EEC monolayers were also cultured alone without spheroids (EEC Only). At the end of cell culture, cells were stained with FITC-annexin V and photographed. Phase-contrast and fluorescent views of the same field are shown. Annexin V-positive cells with intense fluorescence on their plasma membranes represent cells undergoing apoptosis. Note apoptotic cells at the spheroid-EEC interface and within the spheroid region. Pictures shown are representative of three independent experiments. Bar = 50 µm. B) Quantitative analysis of the percentage of annexin V-positive cells in the four groups as described in A. In one set of experiments, detached cells (collected by aspiration) and attached cells (collected by trypsinization) were pooled and stained with FITC-annexin V (solid bars). Annexin V-positive cells were counted, and data are expressed as the percentage (mean ± SEM) of the total number of cells visualized by DAPI staining (at least 1000 cells were counted). In the other set of experiments, only attached cells were analyzed (open bars). Asterisk indicates significant difference (P < 0.05) as compared with the EEC Only and SB+Sph+EEC groups, n = 3

To further confirm the changes depicted by annexin V labeling, FITC-VAD-FMK staining was performed to detect caspase activation, a crucial event of apoptosis [41]. The cell-permeable FITC-VAD-FMK irreversibly binds to activated caspases and, therefore, fluorescently labels apoptotic cells [33, 34]. As illustrated in Figure 6A, the naïve EEC monolayer without spheroid coculture showed only background staining with FITC-VAD-FMK (Fig. 6A, EEC Only). After 24 h of spheroid-EEC coculture, the percentage of cells with bright FITC-VAD-FMK staining (i.e., activated caspase-positive cells) increased significantly (Fig. 6A, Sph+EEC). Most of these activated caspase-positive cells were situated at the spheroid-EEC interface, with merely a few cells of this kind being detected within the spheroid region. The SB203580 significantly decreased the percentage of activated caspase-positive cells at the spheroid-EEC interface (Fig. 6A, SB+Sph+EEC), whereas PD98059 could not prevent apoptosis in the coculture (Fig. 6A, PD+Sph+EEC). Quantitative analysis demonstrated that the percentage of activated caspase-positive cells in the pooled samples of detached and attached cells increased significantly when EEC were cocultured with spheroids and that SB203580 could effectively block such increase in activated caspase-positive cells (Fig. 6B, detached plus attached cells; P < 0.05 between the Sph+EEC and EEC Only groups as well as between the Sph+EEC and SB+Sph+EEC groups, n = 3). Examination of attached cells also displayed parallel findings (Fig. 6B, attached cells; P < 0.05 between the Sph+EEC and EEC Only groups as well as between the Sph+EEC and SB+Sph+EEC groups, n = 3).



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  		FIG. 6. VAD-FMK staining of activated caspases for the detection of apoptotic cells in spheroid-EEC coculture. A) Monolayers of EEC were cocultured with BeWo spheroids for 24 h in the absence of MAPK inhibitors (Sph+EEC) or in the presence of 10 µM SB203580 (SB+Sph+EEC) or 10 µM PD98059 (PD+Sph+EEC). The EEC monolayers cultured alone without spheroids (EEC Only) served as controls. At the end of cell culture, cells were stained with FITC-VAD-FMK and photographed. Phase-contrast and fluorescent views of the same field are shown. Activated caspase-positive cells with intense fluorescence inside the cells represent cells undergoing apoptosis. Note apoptotic cells at the spheroid-EEC interface and within the spheroid region. Pictures shown are representative of three independent experiments. Bar = 50 µm. B) Quantitative analysis of the percentage of activated caspase-positive cells in the four groups as described in A. In one set of experiments, detached cells (collected by aspiration) and attached cells (collected by trypsinization) were pooled and stained with FITC-VAD-FMK (solid bars). Activated caspase-positive cells were counted, and data are expressed as the percentage (mean ± SEM) of the total number of cells visualized by DAPI staining (at least 1000 cells were counted). In the other set of experiments, only attached cells were analyzed (open bars). Asterisk indicates significant difference (P < 0.05) as compared with the EEC Only and SB+Sph+EEC groups, n = 3


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Embryo implantation in mammals is initiated by attachment of the trophectoderm of the hatched blastocyst to the luminal epithelium of the uterus following close approximation between them [1]. In species with epitheliochorial placentation, such as the pig, no invasion occurs, and trophoblast cells are opposed to the uterine epithelium throughout pregnancy [3]. In the mouse and human, however, the trophoblast cells are invasive; these invasive trophoblast cells traverse the uterine epithelium, invade the decidua, and breach uterine vessels, resulting in a "hemochorial" placenta that is in direct contact with the maternal blood [4, 5, 42]. In both rodent and human models, signs of apoptosis have been shown in uterine epithelial cells that may facilitate passage of trophoblast cells through the epithelial barrier before invasion into the endometrial stroma [7, 8, 43]. Although several factors have been implicated in trophoblast outgrowth on endometrial cells [9, 11, 44], signaling mechanisms leading to apoptosis and displacement of EEC during interaction with trophoblast cells remain elusive. In light of the evidence that MAPK are activated in the mouse uterus during sesame oil-induced decidualization [13], we investigated the roles of MAPK pathways in implantation. We found that both p38 MAPK and ERK were activated in cultured monolayer EEC (RL95-2) after incubation with embryo-like cell masses, the BeWo trophoblast spheroids. Trophoblast spheroid outgrowth on EEC monolayers was significantly inhibited by SB203580 (a p38 MAPK inhibitor), but not by PD98059 (an ERK inhibitor), indicating that p38 MAPK rather than ERK is involved in trophoblast outgrowth. Because spheroid expansion on culture surfaces without EEC was unaffected by inhibition of p38 MAPK, we postulated that SB203580 impeded outgrowth of trophoblast spheroids on EEC by blocking cell death and displacement of EEC. Using annexin V- and VAD-FMK-staining techniques for detecting apoptotic cells, we observed marked EEC apoptosis along the spheroid-EEC interface that could be significantly suppressed by SB203580. These results suggest that trophoblast cells eliminate EEC by inducing apoptosis in EEC through activation of the p38 MAPK pathway, thereby facilitating implantation.

Blastocyst adhesion to the uterine epithelium is mediated initially by carbohydrate ligands and then stabilized by integrins or the trophinin-tastin complex [1]. Signaling via integrins of uterine cells evokes Ca2+ influx [45], which has also been shown to be essential for trophoblast adhesion to uterine cells [46]. Because both integrins and Ca2+ have been shown to activate MAPK [47], it is likely that MAPK pathways may play a role in trophoblast-EEC interactions. However, we found that inhibition of ERK and p38 MAPK by PD98059 and SB203580, respectively, had no effect on trophoblast spheroid adhesion to EEC monolayers using a centrifugation-based attachment assay (data not shown and [35]). On the other hand, the present study established that the p38 MAPK pathway is involved in expansion of trophoblast cells on EEC monolayers, because inhibition of p38 MAPK remarkably inhibited trophoblast outgrowth, probably by preventing apoptosis in EEC. We propose that the attachment of hatched blastocysts to the uterine epithelium may activate p38 MAPK to induce apoptosis in luminal epithelial cells, resulting in the dislodgment of these cells to prepare for trophoblast invasion into the endometrial stroma. The glandular epithelial cells that interpose between trophoblast cells and maternal vessels likely may be removed by similar mechanisms associated with p38 MAPK-mediated apoptosis to facilitate the establishment of the hemochorial placenta.

Response time for activation of MAPK following a variety of stimuli may range from minutes to days [48, 49]. In the present study, the level of phospho-ERK and phospho-p38 MAPK (upper band) increased significantly after 10 min and 7 h, respectively, of trophoblast-EEC coculture, as detected by the immunoblotting assay on total cell lysates. Identification of the exact origin of MAPK would be most accurately accomplished by in situ immunochemical techniques to simultaneously localize both MAPK (ERK or p38 MAPK) and specific cellular markers (of either EEC or trophoblast cells). Our immunoblotting assays most likely, though not absolutely, detected MAPK activation in EEC, because the proteins in these samples derived largely from EEC. (The percentages of protein originating from EEC were estimated to be 95.8% ± 0.5% and 93.6% ± 0.6% at 1 and 24 h of coculture, respectively.) This inference was consistent with the finding that spheroid outgrowth was inhibited only when EEC, but not spheroids, were pretreated with SB203580 before coculture, indicating that EEC is the target of SB203580-caused inhibition. One additional approach to prove this concept may be employing transfection techniques to introduce dominant negative p38 MAPK mutants into EEC or trophoblast cells and then examining spheroid expansion on EEC monolayers.

The role of apoptosis in uterine epithelial cells has been implicated in embryonic implantation [7, 50–52]; however, mechanistic studies regarding how apoptosis is induced have been largely lacking. Because p38 MAPK activation was shown to lead to apoptosis [53], it is reasonable to assume that inhibition of spheroid outgrowth by SB203580 was caused, at least in part, by preventing EEC from undergoing apoptosis through impeding p38 MAPK activation. In the present study, we have clearly localized apoptotic cells around the spheroid-EEC interface using annexin V and VAD-FMK staining. These apoptotic cells were more likely to be EEC than trophoblast cells in origin because of their morphological resemblance to neighboring EEC as well as the receding of EEC lawn in response to trophoblast outgrowth. Given that we have identified EEC as the target of SB203580 inhibition of trophoblast outgrowth in the coculture, the finding that SB203580 diminished apoptotic cells at the spheroid-EEC interface further supported the deduction that the apoptotic cells in the coculture originated from EEC. Previous studies have shown that apoptosis in EEC could be induced by transforming growth factor beta; [9], synergy of nitric oxide and progesterone [14], and interleukin-1ß/epidermal growth factor in association with Fas [19, 54]. Some of these factors, being actively secreted by trophoblast cells [55–58], might account for trophoblast-induced p38 MAPK activation and apoptosis in EEC. Characterization of the signaling mechanisms of these factors for the induction of EEC apoptosis may prove to be invaluable for our understanding of embryonic implantation.

In conclusion, the present study demonstrated that trophoblast cells could activate p38 MAPK, and, subsequently, induce apoptosis in EEC. In this way, trophoblast cells removed EEC that reside along the invasion path. We reason that the lack of appropriate p38 MAPK activation for the induction of apoptosis in EEC could result in disorders of inadequate trophoblast invasion, such as implantation failure and preeclampsia. Further investigation is required to establish upstream and downstream events of p38 MAPK-mediated EEC apoptosis and to explore potential clinical applications.


   ACKNOWLEDGMENTS
 
We are grateful to Ya-Hui Chen at Institute of Anatomy and Cell Biology, School of Medicine, University System of Taiwan-National Yang-Ming University for her technical assistance.


   FOOTNOTES
 
1 Supported by a grant from the National Science Council, Taiwan (NSC90-2320-B-010-076). Back

2 Correspondence: Yen-Jen Sung, Institute of Anatomy and Cell Biology, School of Medicine, University System of Taiwan-National Yang-Ming University, 155 Section 2, Li-Nong Street, Taipei, Taiwan 112. FAX: 886 2 28212884; yjsung{at}ym.edu.tw Back

Received: 28 January 2003.

First decision: 22 February 2003.

Accepted: 5 June 2003.


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