Mol. Hum. Reprod. Advance Access originally published online on August 8, 2006
Molecular Human Reproduction 2006 12(10):601-609; doi:10.1093/molehr/gal066
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Tumour necrosis factor-
impairs chorionic gonadotrophin ß-subunit expression and cell fusion of human villous cytotrophoblast
Department of Obstetrics and Gynecology, Medical University of Vienna, Vienna, Austria
1 To whom correspondence should be addressed at: Department of Obstetrics and Gynecology, Medical University of Vienna, AKH, Waehringer Guertel 18-20, A-1090 Vienna, Austria. E-mail: martin.knoefler{at}meduniwien.ac.at
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Abstract |
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Growth factors expressed at the fetal–maternal interface modulate hormone expression of placental trophoblasts. The aim of this study was to investigate the effects of different cytokines on hCG subunit mRNA expression in differentiating villous cytotrophoblasts. Quantitative real-time PCR revealed a 1.8- and 6.9-fold increase of hCG-
and hCG-ß mRNA levels, respectively, between 36 and 60 h of term trophoblast syncytialization. Compared with controls, neither interleukin (IL)-1ß, IL-2, IL-4, IL-6, IL-10, IL-13 and IL-15 nor tumour necrosis factor (TNF)- significantly altered hCG- mRNA expression. Similarly, the ILs did not affect hCG-ß transcript levels. In contrast, TNF- suppressed hCG-ß mRNA 3.8- and 1.8-fold at 36 and 60 h of term trophoblast differentiation. Accordingly, hCG secretion was impaired by TNF- but not by the different ILs. Moreover, TNF- reduced luciferase expression of reporter plasmids harbouring the proximal hCG-ß5 promoter to 35 and 77%, respectively, in primary term trophoblasts and trophoblastic SHGPL-5 cells. In addition, counting of nuclei in syncytialized, desmoplakin-negative areas revealed a 1.9-fold reduction of term trophoblast fusion in the presence of TNF-. Similarly, floating explant cultures prepared from first trimester-denuded villi recovered the syncytium 2.8-fold less efficiently during 72 h of cytokine treatment. Concomitantly, TNF- impaired induction of endogenous and secreted hCG-ß protein levels in these cultures. The data suggest that TNF- decreases hCG-ß mRNA and protein expression by reducing gene transcription and trophoblast cell fusion. Suppression of these processes by TNF- could partly explain the adverse effects of the cytokine on placental function and pregnancy outcome.
Key words:
cell fusion/chorionic gonadotrophin/human trophoblast/TNF-
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Introduction |
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Placental hormone expression is critical for the maintenance of gestation and successful pregnancy outcome. In particular, hCG produced by the trophoblast epithelium of the placental villus plays a major role. The hormone consists of two different subunits, hCG-
, which is common to other glycoprotein hormones such as thyroid-stimulating hormone (TSH), FSH and LH, and hCG-ß (a unique subunit) (Pierce and Parsons, 1981
). The classical function of hCG is to maintain production of steroid hormones in the corpus luteum of early pregnancy until the luteo-placental shift in progesterone production occurs (Tulchinsky and Hobel, 1973). However, expression of the LH/hCG receptor in different gestational tissues suggests that the hormone has a pivotal role during pregnancy, including regulation of trophoblast invasiveness and decidualization of endometrial stromal cells (Tao et al., 1995; Han et al., 1999). In vivo, hCG is released by the syncytium, the multinuclear epithelium of the villous placenta which is generated by continuous cell fusion of underlying mononuclear cytotrophoblasts. The hormone has a direct function in this process because it was shown to stimulate in vitro syncytialization involving a protein kinase A-dependent mechanism (Shi et al., 1993). Moreover, growth factors such as transforming growth factor (TGF)-, leukaemia-inhibitory factor (LIF) or epidermal growth factor (EGF) induce hCG and require LH/hCG receptor expression to promote cell fusion, suggesting that the hormone may have a central role in trophoblast differentiation (Yang et al., 2003).
Maternal serum levels of hCG peak at 9th to 10th week of gestation and fall to a plateau at lower concentrations thereafter (Braunstein et al., 1976). Because concentrations of the free -subunit increase until parturition, levels of holo-hCG are probably determined by selective expression of the ß-subunit (Braunstein et al., 1980). Indeed, hCG-ß is exclusively expressed in the syncytium and can only be observed in cytotrophoblasts before the 6th week of gestation (Maruo et al., 1992). Similarly, hCG- mRNA is predominantly found in the syncytiotrophoblast but can also be detected in a few cytotrophoblasts (Hoshina et al., 1982). During in vitro trophoblast cell fusion, hCG secretion increases as a consequence of induced hCG- and hCG-ß mRNA transcription (Feinman et al., 1986; Knöfler et al., 1999; Knöfler et al., 2004). In the differentiating cultures, hCG- transcript levels rise earlier than hCG-ß mRNA suggesting that production of the latter is more strictly dependent on syncytium formation (Kato and Braunstein, 1989; Ringler et al., 1989). Because little hCG is stored intracellularly, secretion of the hormone is thought to be mainly triggered by de novo synthesis of its subunits (Hussa, 1980).
Growth factors expressed at the fetal–maternal interface affect trophoblast differentiation and function including placental hormone production (Morrish et al., 1998). Besides well-known inducers of hCG such as EGF, granulocyte-macrophage colony-stimulating factor (CSF) or CSF (Morrish et al., 1987; Garcia-Lloret et al., 1994), different interleukins (ILs) were suggested to augment expression of the particular hormone. Indeed, numerous cytokine receptors, i.e. IL-1, IL-2, IL-4, IL-6, IL-10, IL-13, IL15, and tumour necrosis factor (TNF) receptor are detectable on placental trophoblasts (Masuhiro et al., 1991; de Moraes-Pinto et al., 1997; Dealtry et al., 1998; Knöfler et al., 1998; Zygmunt et al., 1998a; Hanna et al., 2000). However, with respect to hCG regulation discordant results were published. IL-1 induces hCG production in JAR and JEG-3 cells (Yanushpolsky et al., 1993; Strohmer et al., 1997), whereas expression was not affected in isolated first trimester trophoblasts (Meisser et al., 1999b). The IL-6/IL-6 receptor system was suggested to mediate IL-1, IL-6 and TNF--dependent hCG release in early cytotrophoblasts (Nishino et al., 1990; Masuhiro et al., 1991; Li et al., 1992). However, a more recent work failed to demonstrate IL-6-dependent changes of hCG secretion in these cultures (Meisser et al., 1999a). TNF- was shown to stimulate hCG release from JEG-3 and JAR cells (Pedersen et al., 1995), whereas the cytokine suppressed hCG secretion from first trimester villous cytotrophoblasts (Meisser et al., 1999b).
From the present literature, we concluded that cytokine-dependent hCG expression could be an acquired feature of trophoblast tumour cells. Therefore, the aim of this study was to analyse cytokine-dependent hCG gene expression in differentiating villous trophoblasts purified from term placentae. Among the different cytokines tested, only TNF- significantly modulated hCG-ß mRNA expression in these cultures. Therefore, the effects of this particular cytokine on hCG-ß mRNA expression and gene transcription were studied in detail using different trophoblast cell models of early and late gestation. The fact that hCG production can be regarded as a hallmark of trophoblast syncytialization also prompted us to investigate the effects of TNF- on cell fusion. Our data suggest that the cytokine impairs both hCG-ß5 gene promoter activity and trophoblast syncytialization which could explain diminished hCG secretion and synthesis of ß-subunit mRNA. Under pathological conditions, increased TNF- levels may therefore disturb normal trophoblast function by decreasing cellular fusion and hCG expression. Inhibition of these processes could play a role in gestational diseases such as pre-eclampsia showing elevated, placental TNF- concentrations (Wang and Walsh, 1996) and defects in trophoblast syncytialization (Jones and Fox, 1981).
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Materials and methods |
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Collection of placental tissues
Placental tissue of early (between 8th and 12th week) and late (between 38th and 40th week) pregnancy was obtained from legal abortions and Caesarean sections, respectively. Utilization of tissues was approved by the local ethical committee and required informed consent of women donating their placentae.
Cell culture of villous cytotrophoblasts of term placentae
Cytotrophoblasts of term placentae were isolated by enzymatic dispersion and density gradient centrifugation as described previously (Kliman et al., 1986; Fisher et al., 1989). Briefly, tissue was digested two times (each 30 min) in Hanks' balanced salt solution containing 25 mM HEPES, 0.125% trypsin and 250 IU/ml DNase I in a shaking water bath (37°C). After each digestion step, the supernatant was neutralized by adding fetal calf serum (FCS) to a final concentration of 10% and centrifuged to remove tissue particles. The supernatants were pooled and filtered over nylon (mesh size 70 µm). Cells were then fractionated on a 5–70% discontinuous Percoll gradient (Pharmacia Biotech, Uppsala, Sweden). Trophoblast cells were isolated from the middle layer of the gradient (density of 1.048–1.062 g/ml). After centrifugation, cells were immunopurified by depleting contaminating human leukocyte antigen (HLA)-I-positive cells with anti-human HLA-avidin–biotin–peroxidase antibody (clone W6/32; Sigma Chemical Co, St. Louis, MO, USA; 0.2 µg/106 cells) conjugated to anti-mouse immunoglobulin G (IgG) magnetic beads (Dynal, Oslo, Norway) as previously detailed (Knöfler et al., 2004). Pure trophoblasts (>98% cytokeratin-7-positive cells) were seeded on plastics at a density of 5 x 105 cells/cm2 and cultivated in Dulbecco’s modified Eagle’s medium (DMEM) containing 2% FCS (Pall, East Hills, NY, USA). Each cell preparation was also routinely seeded in chamber slides and checked by immunocytochemistry at 12 and 60 h of cultivation using cytokeratin-7 antibodies (final concentration: 8.3 µg/ml, clone OV-TL 12/30, DAKO, Glostrup, Denmark) and vimentin antibodies (final concentration: 1.2 µg/ml, clone Vim 3B4, DAKO) to detect trophoblasts and contaminating stromal cells, respectively (Knöfler et al., 2000). For cytokine stimulation, IL-1ß, IL-2, IL-4, IL-6, IL-10, IL-13, IL-15 or TNF- (all purchased from Strathmann, Hamburg, Germany) were added 12 h after trophoblast preparation at a final concentration of 10 ng/ml.
First trimester villous explant culture
Preparation of villous explants of first trimester placentae was performed as described recently (Bauer et al., 2004). For removal of the syncytium, a recent protocol was adopted with minor modifications (Baczyk et al., 2005). Briefly, tissue was washed with ice-cold phosphate-buffered saline (PBS) and transferred into serum-free explant culture medium (HamF12 : DMEM 1:2, supplemented with 50 µg/ml gentamycin). Small pieces (3 x 3 mm) were cut under the microscope and transferred into the digestion solution containing serum-free explant culture medium, 5% trypsin (Invitrogen, Carlsbad, CA, USA) and 0.5% DNase I (Sigma). After 15 min of gentle agitation at 37°C, digestion was stopped with FCS to a final concentration of 10%. Before and after trypsin treatment, some of the explants were immediately fixed and embedded in paraffin. Sections of those samples were screened for the complete removal of the syncytial layer and intactness of the cytotrophoblast epithelium using fluorescence immunohistochemistry (discussed below). For cytokine treatment (24, 48 and 72 h), explants were washed twice with pre-warmed PBS, seeded into 24 wells (each 10 explants per well) and cultivated in culture medium with 10% FCS in the absence or presence of 1 or 10 ng/ml TNF-. After the incubation period, the explants were either fixed in 4% buffered formalin for immunohistochemistry or lysed to obtain cellular protein extracts and total RNA. The supernatants (1 ml per 24 well) were snap-frozen and stored at –80°C for further analyses.
Cultivation of SGHPL-5 cells
Cytotrophoblastic SGHPL-5 cells were cultivated in a 1:1 mixture of DMEM/Ham’s F-12 (GibcoBRL LifeTechnologies, Paisley, UK) as described (Choy and Manyonda, 1998).
Immunodetection of apoptotic cells
Trophoblasts prepared by the Kliman method (n = 3) were seeded on eight-well chamber slides (Nunc, Wiesbaden, Germany) and cultivated in the presence or absence of 10 ng/ml TNF-. For detection of apoptosis, cells were fixed after 60 h with 4% formaldehyde, blocked with 10% FCS in PBS and incubated with the monoclonal M30 CytoDEATH antibody (dilution: 1:20, Roche, Mannheim, Germany). To visualize the apoptotic cytokeratin-18 neo-epitope, we incubated cells with 2 µg/ml fluorescein isothiocyanate (FITC)-conjugated 488 Alexa Flour anti-mouse IgG (A-11017, Molecular Probes, Eugene, OR, USA). Nuclei were counterstained with 4'6-diamidine-2'-phenylindole dihydrochloride (DAPI, 1:1000, Roche), and the slides were mounted with Flouromont G (Southern Biotechnology, Birmingham, UK) and digitally photographed. For each condition, four different fields of 
600–800 cells were counted under the fluorescence microscope and the ratio of M30-positive cells to DAPI-stained nuclei was evaluated. For evaluation of apoptosis in first trimester explant cultures, M30-stained cytotrophoblasts were counted on several sections after 72 h of incubation with TNF-. Between 200 and 300 total nuclei per condition were counted in each three different experiments/explant preparations.
RNA extraction and semi-quantitative RT–PCR
Total villous RNA was prepared from each of 10 floating villi cultivated for 72 h in the absence or presence of 10 ng/ml TNF-. Pooled tissues were frozen in liquid nitrogen and homogenized with a microdismembrator (Braun, Biotech International). The disrupted tissue was covered with Tri-reagent, and total RNA was isolated according to the manufacturer’s instructions (Molecular Research Center Inc, OH, USA). RNA of differentiating, cytokine-treated villous trophoblasts was isolated using the same procedure. Quality and quantity of the extracted RNA was checked with the Agilent Bioanalyzer 2100 (Agilent, Palo Alto, CA, USA). Four micrograms of total RNA was reverse transcribed in 20 µl of reaction volume using 1 µl (200 U) SuperScript RT (Invitrogen) and 0.4 µl Hexant Mix (62.5 A260 U/ml; Roche) according to the manufacturer’s instructions. Semi-quantitative PCR amplification (96°C for 45 s, annealing temperature for 60 s, 72°C for 80 s) was performed with PCR Reagent System (Gibco) in a RoboCycler Gradient 96 (Stratagene, Amsterdam, Netherlands) using 0.5 U Taq polymerase (Invitrogen). Cycle numbers were optimized within the linear range of individual PCR reactions. Oligonucleotide primers, annealing temperatures, product sizes and cycle numbers were as follows: ßhCG-1s, 5'ATGGAGATGTTCCAGGGGC3', ßhCG-1a, 5'TTGTTGGAGGATCGGGGT3' (50°C, 494 bp, 30 cycles), ß-actin-1s, 5'GACAGCAGTCGGTTGGAG C3', ß-actin-1a, 5'CAGGTAAGCCCTGGCTGC3' (55°C, 398 bp, 22 cycles). In all experiments, possible DNA contamination was checked by negative control RT–PCR in which reverse transcriptase was omitted in the RT step. The PCR products were analysed on 1.5% agarose gels containing ethidium bromide and photographed under UV radiation. PCR fragments were sequence verified on a 16 capillary sequencer by using the non-radioactive ABI PRISM Terminator Cycle Sequencing Ready Reaction Kit as specified by the supplier (Applied Biosystems).
Quantitative RT–PCR
Quantitative real-time PCR was performed using the ABI PRISM 7700 Sequence Detection System according to the manufacturer’s instructions (Applied Biosystems). PCR reactions contained 200 µM dNTP, 200 nM of each primer, 1.2 IU Taq and SYBRGreenI (x0.25 final concentration) for signal detection. Primer sequences used for amplification of hCG-, hCG-ß and 18S rRNA (housekeeping control) were as follows: ßhCG-1s, 5'ATGGAGATGTTCCAGGGGC3', ßhCG-1a, 5'TTGTTGGAGGATCGGGGT3', hCG-1s, 5'AAGCCCAGAGAAAGGAGC3', hCG-1a, 5'GGATAAGGAGGAAGGCAG3', 18S rRNA-1s, 5'AGGAATT GACGGAAGGGCACC3' and 18S rRNA-1a, 5'GGACATCTAAGGGCATCACAG3'. Calculation of signals was done as suggested in the PE Biosystems Sequence Detector User Bulletin and elsewhere (Winer et al., 1999). Briefly, threshold cycle (Ct) is defined as the first fluorescent signal reaching statistical significance above background. For each individual condition, 
Ct (the difference of Ct/ßhCG and Ct18SrRNA) values are calculated representing normalization to the housekeeping gene. Subsequently, Ct values are built indicating normalization to controls. The amount of target normalized to an endogenous reference and relative to the unstimulated control is then given by 2–Ct.
Western blot analyses
hCG-ß protein was detected using Western blot analyses in supernatants and cellular lysates of floating villi after 24, 48 and 72 h of TNF- treatment. Protein extracts of each 10 villi were prepared from the phenol-ethanol phase after lysis with Tri-reagent as described by the manufacturer (Molecular Research Center Inc). Briefly, 20 µl of supernatant (normalized to concentration of the respective cellular protein extract) or 25 µg of cellular protein lysate was separated on 12.5% polyacryl amide (PAA) gels and blotted onto nitrocellulose membrane (Protran, Schleicher & Schuell, Dassel, D). Subsequently, immunodetection was performed using standard procedures. After blocking with 3% non-fat milk, the membrane was incubated with rabbit anti-human hCG-ß antibodies (A0231, final concentration: 6 µg/ml, DAKO) overnight at 4°C. Detection was performed with Enhanced Chemiluminescence System (Amersham Pharmacia Biotech), and signals were visualized on autoradiography films. Loading was monitored using a rabbit anti-human actin antibody (A2066, final concentration: 1 µg/ml, Sigma). Antibodies against hCG-ß and actin detected specific signals at 30 and 40 kDa, respectively. As a positive control for hCG-specific signals, 5 IU/ml of urinary hCG preparation (Pregnyl, Organon, UK) was utilized (not shown).
Analyses of re-syncytialization by immunohistochemistry
Each 10 fixed villous explant tissues were dehydrated and embedded in paraffin (Merck) as described elsewhere (Bauer et al., 2004). Serial sections (2–3 µm) were prepared, deparaffinized and finally heated in a microwave oven (x2 5 min, 850 W). After incubation in blocking solution (NEN, Boston, MA, USA), slides were incubated overnight with primary antibodies. Sections were double-stained with mouse anti-human E-cadherin (final concentration: 1.25 µg/ml, clone 36, BD Biosciences) and rabbit anti-human Kip2/p57 (final concentration: 1 µg/ml, C-20, Santa Cruz) followed by 1 h treatment with FITC-conjugated goat anti-mouse (final concentration: 5 µg/ml, Molecular Probes) and rhodamine-conjugated goat anti-rabbit antibody (final concentration: 5 µg/ml, Molecular Probes). As a negative control, the primary antibody was replaced by buffer or isotype IgG. Nuclei were counterstained with DAPI. Cytotrophoblasts were identified by positive E-cadherin and Kip2/p57 staining as described recently (Knöfler et al., 2004), whereas syncytiotrophoblasts were negative for both Kip2/p57 and E-cadherin. For evaluation of the extent of re-syncytialization of denuded villi, 20 sections of each explant were prepared. Distribution of nuclei in the cytotrophoblast and syncytial layer of TNF--treated and TNF--untreated villous explants was determined on several sections, respectively. Between 500 and 600 total nuclei per condition were counted in each three different experiments/placentae.
Fusion of villous cytotrophoblasts
Syncytium formation of differentiating term cytotrophoblasts was investigated by immunocytochemistry using desmoplakin antibodies. Briefly, trophoblasts were seeded in chamber slides (200000 cells per chamber). After 12 h, cells were further cultivated in DMEM + 10% FCS either in the absence or in the presence of 10 ng/ml TNF-. Cultures were fixed and immunostained with anti-desmosomal protein (final concentration: 0.44 mg/ml, ZK-31, Sigma) and DAPI after an additional 48 h. Subsequently, chambers were digitally photographed. Per condition, three areas were randomly selected and the number of syncytial nuclei was determined by counting DAPI-positive cells in desmoplakin-negative structures. Additionally, the number of syncytia in each field was counted. Mean values of the three areas were calculated in three independent experiments.
Transfection of reporters and luciferase assay
For reporter studies, term trophoblasts and SHGPL-5 cells were seeded in 24-well plates and transiently transfected using ProFection Mammalian Transfection System as described by the supplier (Promega, Madison, WI, USA). Twelve hours after isolation, villous term trophoblasts were co-transfected with 2.5 µg of a construct harbouring the proximal (–345 to +114) hCG-ß5 promoter cloned into pGL-3 basic (Johnson and Jameson, 1999; Knöfler et al., 2004) and 0.5 µg pCMV-ß-Gal (Clontech, Palo Alto, CA, USA). Subconfluent SGHPL-5 cells were transfected using the same conditions. After an additional 12 h, medium was changed and cells were incubated in the absence or presence of 10 ng/ml TNF-. After an additional 24 h, supernatants were aspirated and cellular protein lysates were prepared in 100 µl reporter lysis buffer (Promega) and stored at –80°C. Two parallel transfections per condition were performed.
Luciferase activity was determined on a luminometer (Lumat LB 9507, EG&G Berthold, Bad Wildbad, Germany) using Luciferase Assay System (Promega) and 10 µl protein extract. Activity of ß-Gal was quantitated on a photometer by determining the conversion of the chromogenic substrate chlorophenol red-ß-D-galactopyranoside (CPRG, Roche Diagnostics, Vienna, Austria) at 570 nm as described (Eustice et al., 1991). ß-Gal values were determined in 10 µl of protein lysate incubated for 15 min at 37°C. Luciferase- and ß-Gal assays were performed in duplicate, and mean values were calculated. Concentration of protein lysates was determined using Biorad Assay Reagent according to the manufacturer's instructions (Biorad, Hercules, CA, USA).
Enzyme-linked immunosorbent assay
Secreted intact hCG was quantitated in supernatants of 60 h villous trophoblast cultures using enzyme-linked immunosorbent assay (ELISA) (Diagnostic Systems Laboratories, Webster, TX, USA). The assay does not cross-react with free hCG- or free hCG-ß subunit or FSH. Minor cross-reactivity to LH (0.3%) and TSH (0.02%) was detected by the manufacturer.
Statistical analyses
Statistical analyses were performed with Student’s t-test using SPSS 12 (SPSS Inc, Chicago, IL, USA). A P-value <0.05 was considered statistically significant.
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Results |
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Term cytotrophoblasts were treated with different cytokines to evaluate their potential influence on hCG expression (Figure 1). Quantitative real-time PCR revealed an induction of hCG mRNA subunit expression between 36 and 60 h of in vitro differentiation (Figure 1A). Transcript levels of hCG-
and hCG-ß increased 1.8- and 6.9-fold, respectively. Compared with controls, treatment with IL-1ß, IL-2, IL-4, IL-6, IL-10, IL-13 or IL-15 (each 10 ng/ml) had no effects on hCG- and hCG-ß mRNA expression. Similarly, none of the ILs altered hCG subunit mRNA expression at a concentration of 1 ng/ml (data not shown). However, TNF- decreased hCG-ß mRNA 3.8- and 1.8-fold at 36 and 60 h of culturing, respectively. The cytokine also slightly affected hCG- mRNA expression at 36 h of cultivation (1.4-fold reduction compared with controls). However, this could not be observed at 60 h of differentiation.
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To assess whether TNF- may also modulate hCG protein levels, we measured soluble concentrations of the hormone in supernatants after 60 h of cultivation (Figure 1B). ELISA revealed that TNF- decreased secretion of hCG to 35% compared with controls (100%). None of the ILs significantly altered the amount of soluble hCG.
The results suggested that down-regulation of hCG secretion by TNF- could be mediated through impaired expression of hCG-ß mRNA. The hCG-ß gene cluster consists of six different genes, of which the promoter of the hCG-ß5 gene is most actively transcribed in choriocarcinoma cells and trophoblasts of early and late gestation (Talmadge et al., 1984; Jameson and Lindell, 1988; Bo and Boime, 1992; Miller-Lindholm et al., 1997). To gain insights into the mechanism of TNF--dependent hCG-ß mRNA repression, we transfected trophoblastic SGHPL-5 cells derived from first trimester placentae and purified term cytotrophoblasts with luciferase reporters harbouring the proximal (–345 to +114) hCG-ß5 5' flanking sequence (Johnson and Jameson, 1999; Knöfler et al., 2004). Luciferase assays revealed that the activity of the promoter was repressed to 77 and 35%, respectively, in SGHPL-5 cells and differentiating villous term trophoblasts after 24 h of treatment with TNF- (Figure 2).
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The fact that the hCG-ß5 promoter was less efficiently repressed in SGHPL-5 cells that are unable to syncytialize compared with the differentiating primary cells prompted us to investigate the influence of TNF- on cell fusion. Counting of nuclei in desmoplakin-negative areas of differentiating term trophoblasts revealed that the extent of syncytialization was decreased upon TNF- treatment (Figure 3). At 60 h of culturing, mean numbers of nuclei per syncytium were 1.9-fold diminished in the presence of the cytokine. In addition, apoptosis was assessed by counting the percentage of M30-positive cell numbers using immunocytochemistry as described (Huber et al., 2006). No statistical differences could be detected after 60 h of culturing between controls (12.9% ± 2.5 SD) and TNF--treated samples (13.1% ± 1.1 SD). Moreover, we investigated the effects of TNF- on first trimester cytotrophoblast cell fusion (Figure 4) and hCG-ß expression (Figure 5) using villous explant cultures. In these organ cultures, the syncytium had been removed by trypsin treatment. The denuded explants recovered the syncytium within 72 h of culturing in medium supplemented with 10% FCS. Representative immunohistochemically stained tissue sections of freshly denuded and recovered placental villi in the absence or presence of TNF- are shown (Figure 4A). To evaluate the extent of re-syncytialization, we determined the ratio of cytotrophoblast nuclei to syncytial nuclei in controls and TNF--treated samples, respectively (Figure 4B). After the recovery period, the ratio of cytotrophoblasts to syncytiotrophoblasts was 2.3 and 6.5 in untreated and TNF-treated explant cultures, respectively, suggesting that syncytialization was 2.8-fold impaired by the cytokine. Concomitantly, up-regulation of endogenous (Figure 5A) and secreted (Figure 5B) hCG-ß protein, as detected by Western blot analyses, was impaired when denuded explants were treated with TNF- during the recovery period. The cytokine also strongly decreased hCG-ß mRNA expression in cultures that had not been treated with trypsin in advance (Figure 5C). Immunohistochemistry with M30 antibodies did not reveal any differences in the number of apoptotic cytotrophoblasts between controls (9.3% ± 2.7 SD) and TNF--treated (8.4% ± 2.6 SD) explant cultures at 72 h of cultivation.
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Discussion |
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Here, we investigated cytokine-dependent regulation of hCG mRNA subunit expression as well as the secretion of the intact hormone from primary villous trophoblasts of term placenta. Quantitative real-time PCR suggested that the ILs, IL-1ß, IL-2, IL-4 IL-6, IL-10, IL-13 and IL-15, did not affect hCG-
or hCG-ß transcript levels. Accordingly, the ILs tested did not modulate the release of hCG. Therefore, our data are in agreement with previous studies suggesting that IL-1 and IL-6 do not alter hCG expression in primary cytotrophoblast cultures (Meisser et al., 1999a,b; Monzon-Bordonaba et al., 2002). However, results of others could not be confirmed (Nishino et al., 1990; Masuhiro et al., 1991). In contrast, IL-1 and IL-6 were shown to stimulate hCG release from JEG-3 and JAR cells (Silen et al., 1989; Taniguchi et al., 1992; Yanushpolsky et al., 1993; Strohmer et al., 1997). From these data, we conclude that IL-dependent induction of hCG is a unique feature of choriocarcinoma cell lines. Therefore, tumour cells might not be the ideal trophoblast system to study physiological hormone expression. Interleukin-mediated induction of hCG in JEG-3 and JAR cells might be advantageous for distinct properties of the tumour cells such as hormone-dependent elevation of invasion and migration (Zygmunt et al., 1998b).
Similarly, the effects of TNF- on hCG expression depend on the cellular model system. The cytokine was shown to stimulate hCG secretion from JEG-3 and JAR cells (Pedersen et al., 1995; Jiang et al., 2005). In contrast, previous publications using first or third trimester villous trophoblast cultures demonstrated repression of hCG release in the presence of TNF- (Meisser et al., 1999b; Monzon-Bordonaba et al., 2002). In accordance with these observations, we also detected reduced hCG secretion from differentiating term trophoblast upon addition of the cytokine.
To gain insights into the repressive effect of TNF-, we investigated mRNA expression of individual hCG subunits during cell fusion. Whereas weak effects of TNF- on hCG- mRNA expression were noticed, transcript levels of hCG-ß were significantly decreased by the cytokine at 36 and 60 h of term trophoblast differentiation. Because hCG production is predominantly regulated at the level of mRNA expression (Hussa, 1980), we assume that down-regulation of hCG secretion by TNF- is mainly triggered by repressed synthesis of its ß-subunit. Indeed, transfection experiments using luciferase plasmids harbouring the proximal 5' flanking region of the hCG-ß5 gene promoter demonstrated that the cytokine significantly reduced transcriptional activity in SGHPL-5 cells and term trophoblasts. As SGHPL-5 cells do not fuse in culture, direct repression of hCG-ß5 promoter activity is suggested. This might be accomplished by enhancer elements of the proximal hCG-ß5 promoter such as two different AP-1 recognition sequences which repress transcription of hCG-ß mRNA in choriocarcinoma cells (Pestell et al., 1994). Indeed, constitutive and TNF--inducible AP-1-binding activity could be detected in SGHPL-5 cells and term villous trophoblasts using electrophoretic mobility shift assays (EMSA) and a radiolabelled AP-1 consensus sequence; however, interaction with the putative cognate sequences of the hCG-ß5 promoter could not be observed (Saleh and Knöfler, unpublished). Therefore, the role of potential AP-1-binding sites and other enhancer elements in TNF--mediated repression of the hCG-ß5 promoter requires further analyses.
Interestingly, we noticed that the hCG-ß5 promoter was down-regulated to larger extents in differentiating primary trophoblasts compared with the cytotrophoblast cell line which cannot form syncytia. Although cell-type-specific differences cannot be excluded, we assume that impaired cell fusion could additionally decrease hCG-ß transcription and expression. Activity of the hCG-ß5 gene promoter is low in freshly isolated term cytotrophoblasts and increases during in vitro syncytialization (Knöfler et al., 2004). Indeed, TNF- reduced fusion of term cytotrophoblasts as well as the reappearance of syncytial nuclei in trypsin-treated, villous explant cultures of early gestation. Therefore, the data suggest that the repressive mechanism is operational at several stages of gestation. Potential TNF--mediated programmed cell death cannot explain the decrease in cell fusion because in both cultures systems similar levels of apoptotic cytotrophoblasts were detected in the absence or presence of the cytokine. This is in agreement with previous observations, suggesting that TNF- alone does not provoke substantial apoptosis in trophoblasts (Knöfler et al., 2000; Bauer et al., 2004; Renaud et al., 2005). The fact that the cytokine inhibited fusion more strongly in the first trimester organ cultures compared with the primary term trophoblasts might be explained by the isolation procedure. The Kliman method results in the generation of disordered, single cells, whereas removal of the syncytial layer in explant cultures maintains epithelial structure and morphology of the underlying cytotrophoblasts.
Inhibition of differentiation and cell fusion by TNF- has already been noticed in other cellular systems such as myogenic cells (Langen et al., 2002). In analogy with other systems, TNF- may directly repress trophoblast cell fusion involving activation of nuclear factor (NF)
B or generation of reactive oxygen species (Langen et al., 2001; Langen et al., 2002).
On the contrary, diminished syncytialization might be indirectly caused by the TNF--induced drop of hCG expression and secretion. Indeed, hCG was shown to induce cell fusion and markers of trophoblast differentiation (Shi et al., 1993). Moreover, hCG and its receptor probably play a pivotal role in trophoblast cell fusion because factors such as EGF, LIF or TGF- were shown to promote syncytium formation through elevation of hCG secretion (Yang et al., 2003). Therefore, further studies such as fusion experiments in the presence of TNF- and exogenous hCG are necessary to more precisely delineate the underlying molecular mechanism.
Whereas the role of TNF- in normal human gestation is still a matter of debate, increased concentrations of the cytokine are thought to be involved in pathological processes of pregnancy (Hunt, 1993). High levels of the cytokine are present in the amniotic fluid of women suffering from chorioamnionitis, and a role of TNF- in spontaneous abortions was suggested (Clark et al., 1999). Inflammatory reactions may also affect placental viability because TNF- was shown to provoke trophoblast apoptosis in combination with Th1 cytokines such as interferon (INF)-
(Yui et al., 1994). Furthermore, several studies indicate an association between a nucleotide polymorphism of the TNF- gene promoter and preterm rupture of membranes and recurrent pregnancy loss (Roberts et al., 1999; Reid et al., 2001).
An adverse role of TNF- is also anticipated in gestational disease in which placental failures are thought to play a critical role. For example, elevated concentrations of the cytokine were detected in sera, placental villi and some decidual stromal cells of pre-eclamptic women (Kupferminc et al., 1994; Wang and Walsh, 1996; Pijnenborg et al., 1998). Indeed, TNF- was shown to inhibit trophoblast migration and motility in vitro (Bauer et al., 2004; Renaud et al., 2005; Huber et al., 2006), suggesting that elevated concentrations of the cytokine could be one cause of restricted trophoblast invasion observed in pre-eclampsia (Pijnenborg et al., 1983). Interestingly, defects in trophoblast syncytialization were also detected in pre-eclamptic placentae (Jones and Fox, 1981), which might be partly caused by TNF-. We speculate that elevated placental concentrations of the cytokine could impair hCG expression and trophoblast cell fusion in these patients.
Recently, recurrent miscarriage was shown to be associated with a constant ratio of cytotrophoblasts to syncytiotrophoblasts, whereas this ratio decreases during normal placental development (Bose et al., 2005). Elevated proliferative capacity of cytotrophoblasts in the pathological placentae was suggested as an underlying cause. However, because aberrant bioactive TNF- levels were noticed in recurrent abortion (Arslan et al., 2004; Chernyshov et al., 2005), it may well be that impaired cell fusion plays an additional role.
In conclusion, our data show that elevated concentrations of TNF- negatively affect trophoblast function by decreasing hCG expression and syncytialization. Besides the suggested harmful effect on trophoblast viability, TNF- may thus exert an adverse role on placental development through inhibition of trophoblast differentiation. Further studies are needed to delineate the diverse effects of the cytokine on placental function of normal and pathological pregnancies.
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We are grateful to G. Whitley for providing SGHPL-5 cells. We thank G. Puller for preparation of graphics.
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