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Original Studies |
Department of Obstetrics and Gynecology, British Columbia Women’s Hospital, University of British Columbia, Vancouver, British Columbia, Canada V6H 3V5
Address all correspondence and requests for reprints to: Peter C. K. Leung, Ph.D., Department of Obstetrics and Gynecology, University of British Columbia, 2H30-4490 Oak Street, British Columbia Women’s Hospital, Vancouver, British Columbia, Canada V6H 3V5. E-mail: peleung{at}interchange.ubc.ca
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Activin and transforming growth factor-ß (TGFß) are members of the
TGFß superfamily. Activin is a dimeric protein composed of two
ß-subunits, ßA-ßA (activin A), ßB-ßB (activin B), or
ßA-ßB (activin AB) (15). Inhibin, another member of
the TGFß superfamily, is composed of an 
- and one of two
ß-subunits, -ßA (inhibin A) or -ßB (inhibin B). The main
function of these gonadal peptides is to regulate FSH secretion from
the anterior pituitary gland (16, 17). However, as activin
and inhibin are produced locally in the ovary (18), it has
been hypothesized that they may act via an autocrine/paracrine
mechanism to regulate ovarian function (19, 20). Activin
mediates its cellular effects through heterodimeric complexes of type I
and II activin serine/threonine kinase receptors (20). The
importance of the activin in regulating cell proliferation and possibly
ovarian tumor development has been suggested (21, 22, 23). In
the previous studies we have shown that activin stimulated the growth
of OVCAR-3, but not of normal OSE, and altered the expression of the
activin/inhibin subunits (22 23A ). Therefore, activin
appears to act as an autocrine/paracrine factor in epithelial ovarian
tumors, but its role in tumorigenesis has yet to be defined
(23).
TGFß is a growth regulator that affects multiple cellular functions through the TGFß type I and II receptors (TGFßRI and TGFßRII) serine/threonine kinases (reviewed in Refs. 24 and 25). A number of mammalian cells produce TGFß1, -2, and -3, and a significant amount of TGFß is also produced by normal OSE cells (26). In normal and neoplastic ovarian epithelial cells, TGFß has been demonstrated to inhibit cell growth (26, 27, 28, 29). These results indicate that several elements of potential autocrine loops involving TGFßs are present within ovarian cancer cells (30). It has been demonstrated that TGFß inhibits proliferation, but does not induce apoptosis in normal OSE cells. In contrast, TGFß induces apoptosis in some ovarian cancers that are growth inhibited by TGFß (27). In breast, ovarian, and prostate cancer cell lines, relatively low levels of TGFß receptor messenger ribonucleic acid (mRNA) and protein have been demonstrated, and TGFß treatment resulted in an inhibition of growth (31). Enhanced expression of TGFß1 and TGFß3 as well as the loss of expression of TGFßRI contribute to ovarian carcinogenesis and/or tumor progression (32).
The present study was performed to investigate the roles of activin and TGFß in normal, early neoplastic, and tumorigenic OSE cells. The expression of the activin/inhibin subunits and activin receptors was determined. In addition, the effects on cell number and induction of apoptosis by activin and TGFß were examined. Finally, the regulation of proapoptotic Bax and anti-apoptotic Bcl-2 was investigated after treatments with activin and TGFß.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Recombinant human activin A and follistatin were provided by Dr. A. F. Parlow, National Hormone and Pituitary Program of Harbor-University of California-Los Angeles Medical Center (Torrance, CA). Recombinant TGFß1 was purchased from Sigma-Aldrich Corp. (Oakville, Canada).
Cell culture
Normal OSE. OSE cells were scraped from the ovarian surface during laparoscopy for nonmalignant disorders and cultured as previously described (33) in medium 199/MCDB 105 (Life Technologies, Inc., Burlington, Canada; and Sigma-Aldrich Corp., respectively) containing 10% FBS, 100 U/mL penicillin G, and 100 µg/mL streptomycin (Life Technologies, Inc.) in a humidified atmosphere of 5% CO2-95% air and passaged with 0.06% trypsin (1:250)/0.01% ethylenediamine tetraacetate in Mg2+/Ca2+-free Hanks’ Balanced Salt Solution when confluent.
Immortalized ovarian surface epithelium cell lines. We recently developed a culture system with the cells representing several stages in the neoplastic progression of OSE. The IOSE-29 cell line (referred to as IOSE-Mar in some previous publications) was generated by transfection with the immortalizing SV40 early genes into normal human OSE at passage 5 (12). The IOSE-29EC cell line was made from IOSE-29 at passage 11 by transfecting the full-length mouse E-cadherin complementary DNA (cDNA) under control of the ß-actin promoter (13). IOSE-29EC/T4 and IOSE-29EC/T5 were established from tumors that arose in IOSE-29EC-inoculated SCID mice, and they formed tumors on mesenteries and omentum, invaded the liver and thigh musculature, and produced ascites (14). For monolayer culture, all cell lines were maintained on tissue culture dishes (Corning, Inc., Corning, NY) in a 1:1 mixture of medium 199/MCDB 105 medium supplemented with 10% FBS, 100 U/mL penicillin G, and 100 µg/mL streptomycin.
RNA extraction and RT-PCR procedures
Total RNA was prepared from cultured cells using the RNaid kit
(Bio/Can Scientific, Mississauga, Canada) according to the
manufacturer’s suggested procedure. RNA integrity was confirmed by
agarose gel electrophoresis and ethidium bromide staining (34, 35). The total RNA concentration was determined from
spectrophotometric analysis at A260/280. cDNA was
synthesized from 2.5 µg total RNA by RT at 37 C for 2 h using a
first strand cDNA synthesis kit (Pharmacia Biotech,
Uppsala, Sweden). The synthesized cDNA was used as template for PCR
amplification. Amplification was achieved using a thermal cycler (DNA
Thermal Cycler, Perkin-Elmer Corp., Norwalk, CT). The
amplification profile involves denaturation at 94 C for 60 s,
primer annealing at 51–60 C for 30 s, and extension at 72 C for
90 s. Synthetic oligonucleotides used for PCR primers and PCR
amplification cycle number were listed in Table 1
based on the published sequences. The
amplification profile for human glyceraldehyde phosphate dehydrogenase
(GAPDH), which is a housekeeping gene, involves denaturation at 94 C
for 60 s, primer annealing at 55 C for 30 s, and extension at
72 C for 90 s (36, 37). To avoid overamplification,
the ranges of exponential increase in product formation with numbers of
amplification cycles were determined. The PCR reactions were performed
in 25 µL PCR mixture containing 1 x PCR buffer, 0.2 mmol/L of
each dNTP, 1.6 mmol/L MgCl2, 50 pmol of specific
primers, each cDNA template, and 0.25 U Taq polymerase. The
PCR reaction was performed for two or three independent cDNA
preparations of each RNA sample. Twelve microliters of PCR products
were analyzed by agarose (1%) gel electrophoresis and visualized by
ethidium bromide staining, and the sizes were estimated by comparison
to DNA molecular weight markers.
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After electrophoresis, PCR products were transferred to nylon
membranes (Hybond-N, Amersham Pharmacia Biotech, Arlington
Heights, IL) and fixed by UV irradiation. The blotted membrane was
prehybridized for 3 h at 42 C in prehybridization solution
containing 50% formamide, 5 x SSC (standard saline citrate,
0.1% N-lauroyl sarcosine, 0.02% SDS, and 2% blocking
solution). The prehybridized membrane was hybridized overnight at 42 C
with digoxigenin (DIG)-labeled probe. All cDNA probes were labeled with
a DIG cDNA labeling kit (Roche Molecular Biochemicals,
Laval, Canada). The cDNA clones for -, ßA-, and ßB-subunits were
subcloned from human granulosa cells and verified by sequencing. The
cDNA clones for activin receptors IA and IB were provided by Dr. C.
Peng (York University, Toronto, Canada). The cDNA clones for activin
receptors IIA and IIB were provided by Dr. W. Vale (The Salk Institute,
La Jolla, CA) and Dr. C. Peng, respectively. The cDNA clones for Bax
and Bcl-2 were subcloned from ovarian cancer cell line (OVCAR-3) and
verified by sequence analysis. The hybridized membranes were detected
by the luminescence method (Roche Molecular Biochemicals)
and exposed to x-ray film for 1–10 min at room temperature. The
specific bands were scanned and quantified using a computerized visual
light densitometer (model 620, Bio-Rad Laboratories, Inc.,
Richmond, CA).
Immunoblot analysis
The IOSE cells (IOSE-29, IOSE-29EC, IOSE-29EC/T4, and IOSE-29EC/T5) were seeded at a density of 2 x 105 cells in 35-mm culture dishes and cultured in a humidified atmosphere of 5% CO2-95% air at 37 C. Cells were washed twice with ice-cold PBS and lysed in ice-cold RIPA buffer (150 mmol/L NaCl, 1% Nonidet P-40, 0.5% deoxycholate, 0.1% SDS, 50 mmol/L Tris (pH, 7.5), and 1 mmol/L phenylmethylsulfonylfluoride, 10 µg/mL leupeptin, and 100 µg/mL aprotinin). The extracts were placed on ice for 15 min and centrifuged to remove cellular debris. The protein content of the supernatants was determined using a Bradford assay (Bio-Rad Laboratories, Inc.). Twenty-five micrograms of total protein were run on 10% SDS-PAGE gels and electrotransferred to a nitrocellulose membrane (Amersham Pharmacia Biotech) (38). The membrane was immunoblotted using rabbit polyclonal antibodies of activin receptors IA, IB, IIA, and IIB provided by Dr. W. Vale (The Salk Institute) or mouse monoclonal antibodies of Bax (BD PharMingen, Mississauga, Canada) and Bcl-2 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA). The loaded amount of protein was normalized with actin protein in the same membrane. After washing, the signals were detected with horseradish peroxidase-conjugated secondary antibody and visualized using the enhanced chemiluminescence chemiluminescent system (Amersham Pharmacia Biotech).
[3H]Thymidine incorporation assay
[3H]Thymidine incorporation assay was performed to analyze the effect on cell number of activin and TGFß in normal and neoplastic OSE cells. The cells were plated in 24-well plates at 2 x 104 cells/well in 0.5 mL medium 199/MCDB 105 supplemented with 10% FBS and antibiotics for 48 h. For activin treatment, cells were treated with 1, 10, and 100 ng/mL recombinant human activin A daily for 6 days as previously described (21, 39). During last 6 h after treatment, the cells were cultured in the presence of the same concentration of activin A and 1 µCi [3H]thymidine (5.0 Ci/mmol; Amersham Pharmacia Biotech). For TGFß treatment, the cells were incubated with 0.1, 1, or 10 ng/mL TGFß and replaced daily for 72 h. During the last 6 h, 1 µCi [3H]thymidine and the same concentration of TGFß were added to each well as previously described (27). The culture medium was then removed and washed with three times with PBS, followed by precipitation with 0.5 mL 10% trichloroacetic acid for 20 min at 4 C (39). The precipitate was washed in methanol twice and solubilized in 0.5 mL 0.1 N sodium hydroxide, and the incorporated radioactivity was measured in a 1217 Rackbeta liquid scintillation counter (LKB Wallac, Inc., Turku, Finland).
Quantification of apoptotic cells
To quantify the induction of apoptosis by activin and TGFß in IOSE-29EC cells, DNA fragmentation was measured using the cell death detection enzyme-linked immunosorbent assay (ELISA; Roche Molecular Biochemicals) as previously described (40). Briefly, the cells (1 x 104) were plated in each well of 24-well plates. After treatments with activin (1, 10, and 100 ng/mL) for 6 days or TGFß (0.1, 1, and 10 ng/mL) for 72 h, the media were collected and stored during the treatments, the cells were washed with PBS, and 0.1 mL lysis buffer was added. After 15-min incubation on ice, apoptotic cells in cell lysates and medium were assayed for DNA fragments according to the manufacturer’s protocol. The same amount (1 µg) of cell lysate was used for the cell death ELISA. DNA fragmentation was measured at 405 nm against an untreated control.
Statistical analysis
Data are shown as the mean of three individual experiments and are presented as the mean ± SD. In the proliferation study, values are expressed as the percentage of growth compared with the control value and are the mean ± SD of three individual experiments with triplicate samples. For the quantification of apoptotic cells, values are expressed as the percentage of DNA fragmentation compared with the untreated control value and are the mean ± SD of three individual experiments with duplicate samples. The data were analyzed by one-way ANOVA, followed by Tukey’s multiple comparison or Dunnett’s test. P < 0.05 was considered statistically significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The mRNA levels of -, ßA-, and ßB-subunits in IOSE-29
(passages 13–16), IOSE-29EC (passages 15–17), IOSE-29EC/T4 (T4), and
IOSE-29EC/T5 (T5) were investigated by RT-PCR and Southern blot
analysis. The possibility of cross-contamination was ruled out, because
no PCR products were observed or detected in the negative control
[TmA(-); without template in the PCR reaction] by ethidium bromide
(data not shown) and Southern blot analysis (Fig. 1). A linear relationship between PCR
products and amplification cycles was obtained for GAPDH and -,
ßA-, and ßB-subunits in all cell types (data not shown). The
expected PCR products of GAPDH and -, ßA-, and ßB-subunits were
obtained at 373, 382, 377, and 424 bp, respectively, and were confirmed
by Southern blot analysis (Fig. 1) and sequence analysis (data not
shown). Semiquantitative analysis of the present study demonstrated
that all types of activin/inhibin subunits are expressed in IOSE-29,
IOSE-29EC, T4, and T5. Interestingly, ßB-subunit was less expressed
in IOSE cell lines than in OVCAR-3 cells (Fig. 1).
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The mRNA levels of activin receptors IA, IB, IIA, and IIB in
IOSE-29 (passages 13–16), IOSE-29EC (passages 15–17), T4, and T5 were
investigated by RT-PCR and Southern blot analysis (Fig. 2). The expected sizes of PCR products
for activin receptors were obtained as 651, 684, 456, and 699 bp,
respectively, using sense and antisense primers located within the
intracellular kinase domains of each activin receptor. The PCR of GAPDH
was amplified to rule out the possibility of RNA degradation and was
used to control the variation in mRNA concentrations in the RT
reaction. PCR products of the predicted sizes were obtained and
confirmed by Southern blot analysis using DIG-labeled probes (Fig. 2)
and sequence analysis (data not shown). Semiquantitative analysis of
the present study demonstrated that all forms of activin receptors were
observed in IOSE-29, IOSE-29EC, T4, and T5 cells.
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Immunoblot analysis was performed using the rabbit polyclonal
antibodies against activin receptor IA (amino acids 474–494), IB
(amino acids 493–505), IIA (amino acids 482–494), and IIB (amino
acids 524–536) based on COOH-terminal amino acids in IOSE cell lines.
As shown in Fig. 3, activin receptor IA
protein (60 kDa) was observed in all cell types. The OVCAR-3 cell line
was used for positive control of the expression of activin receptors
(22). Similarly, activin receptor IB protein (55 kDa) was
observed in all cell types (Fig. 3). In addition, activin receptors IIA
and IIB were clearly detected at 80 and 60 kDa, respectively, in IOSE
cell lines and OVCAR-3 cells (Fig. 3). Immunoblot analysis of the
present study demonstrated that all forms of activin receptor protein
were observed in IOSE-29, IOSE-29EC, T4, and T5.
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To evaluate the role of recombinant human activin A (rh-activin A)
in IOSE cell lines, IOSE-29 and IOSE29EC were treated with different
concentrations (1, 10, and 100 ng/mL) of rh-activin A for 6 days. The
proliferative index was measured by thymidine incorporation assay.
Follistatin, which is an activin-binding protein, was used to block the
action of activin in the cell proliferation study. As seen in Fig. 4, a dose-dependent decrease in cell
number was observed after rh-activin A (1, 10, and 100 ng/mL) treatment
in IOSE-29 (Fig. 4A; 100.0 ± 7.63 vs. 83.7 ±
6.06, 67.9 ± 4.10, or 59.9 ± 9.06) and IOSE-29EC (Fig. 4B;
100.0 ± 5.89 vs. 75.9 ± 9.11, 61.4 ± 8.11,
or 52.9 ± 9.70) cells. Cotreatment with follistatin (100 ng/mL)
and activin blocked the growth inhibitory effect of activin in both
cell lines (Fig. 4). However, no significant difference was observed
with follistatin treatment only.
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To examine the role of TGFß, normal OSE, IOSE-29, and IOSE-29EC
cells were treated with different concentrations (0.1, 1, and 10 ng/mL)
of TGFß for 72 h. As shown in Fig. 5A, TGFß (1 and 10 ng/mL) induced a
significant decrease in normal OSE cell proliferation in a
dose-dependent manner (100.0 ± 15.62 vs. 58.6 ±
11.78 or 43.3 ± 12.03). Also, a significant decrease was observed
after TGFß treatments (1–10 ng/mL) in IOSE-29 (Fig. 5B; 100.0
± 5.03 vs. 81.1 ± 7.59 or 69.8 ± 4.08).
Treatment with TGFß resulted in a decrease in the proliferative index
in IOSE-29EC cells (Fig. 5C; 100.0 ± 11.70 vs.
74.2 ± 5.63, 67.1 ± 7.05, or 55.0 ± 6.75).
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To examine the role of activin in the induction of apoptosis, DNA
fragmentation was measured by cell death detection ELISA. To quantify
the induction of apoptosis, IOSE-29EC cells were treated with
rh-activin A for 6 days. As shown in Fig. 6A, treatments with 10 and 100 ng/mL
activin increased DNA fragmentation in a dose-dependent manner
(100.0 ± 8.06 vs. 190.6 ± 13.58 or 221.3 ±
15.72). Cotreatment with follistatin (100 ng/mL) and activin attenuated
the effect of activin. No significant difference was observed with
follistatin treatment only in IOSE-29EC cells. Similarly, IOSE-29EC
cells were treated with different concentrations of TGFß for 72
h. Treatments with TGFß induced a significant increase in DNA
fragmentation in a dose-dependent manner in IOSE-29EC cells (Fig. 6B;
100.0 ± 5.20 vs. 123.7 ± 10.03, 191.3 ±
16.94, or 201.9 ± 25.06).
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The mRNA levels of Bax and Bcl-2 in IOSE-29 (passages 13–18) and
IOSE-29EC (passages 13–18) were investigated by RT-PCR and Southern
blot analysis (Fig. 7). A linear
relationship between PCR products and amplification cycles was obtained
for GAPDH, Bax, and Bcl-2 in all cell types (data not shown). PCR
products of GAPDH, Bax, and Bcl-2 were determined to be 373, 323, and
459 bp, respectively, and this was confirmed by Southern blot analysis
using DIG-labeled probes (Fig. 7) and sequence analysis (data not
shown). No difference was observed in the expression level of Bax mRNA
between IOSE-29 and IOSE-29EC cells. In contrast, the expression level
of Bcl-2 mRNA was higher in IOSE-29EC cells than in IOSE-29 cells (Fig. 7).
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To investigate the mechanism of activin and TGFß in the
induction of apoptosis, the regulation of apoptotic Bax and Bcl-2 was
examined by immunoblot analysis. The IOSE-29EC cells were treated with
increasing doses of rh-activin A and TGFß, respectively, for 24
h, and immunoblot analysis was performed as described in
Materials and Methods. Bax and Bcl-2 protein were detected
at 21 and 26 kDa, respectively. As shown in Fig. 8A, treatment with 10 and 100 ng/mL
activin had no effect on Bax and Bcl-2 proteins in these cells. No
significant difference in Bax protein was observed after TGFß
treatment (Fig. 8B). In contrast, treatment with 1 and 10 ng/mL TGFß
induced a significant decrease in Bcl-2 protein up to 50% (Fig. 8, B
and C; 100.0 ± 5.17 vs. 58.2 ± 7.35 or 54.0
± 5.39). The amount of loaded proteins in the treatment groups was
normalized by actin protein (41 kDa).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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It is well known that activin and TGFß play a role in growth-promoting, growth-inhibiting, or both activities depending on the particular cell type (42, 43). In particular, these factors are prime candidates as regulators of cell proliferation during morphological changes in the ovary as well as in abnormal proliferation and transformation of these tissues (44). The action of activin can be regulated by follistatin, which binds activin with high affinity and neutralizes its actions (45). Treatment with activin stimulated the growth of OVCAR-3 cells, but not that of normal OSE (23A ). Thus, the differential expression and production of activin/inhibin subunits, activin receptors, and follistatin suggest that activin may be involved in neoplastic OSE cell proliferation (21). Continuous treatments with activin (1–100 ng/mL) for 6 days resulted in a significant decrease in cell proliferation in both IOSE-29 and IOSE-29EC cell lines. These growth inhibitory effects of activin were attenuated after cotreatment with follistatin (100 ng/mL), which is a specific binding protein of activin, in these cells. These findings are unexpected, because activin has been thought to be a growth stimulatory factor in some ovarian cancer cell lines (21, 22 23A ). Di Simone et al. demonstrated that treatment with activin (1–100 ng/mL) resulted in an increase in the growth of four ovarian cancer cell lines, which are not synthesizing follistatin, whereas follistatin treatment (1–100 ng/mL) decreased cellular proliferation in these cell lines (21). Most primary epithelial ovarian tumors (96%) synthesize and secrete activin in vitro, and serum levels of activin are frequently elevated in women with epithelial ovarian cancer, but the majority of tumors in culture do not respond to activin or follistatin treatment in cell proliferation (23). The cause of the difference between previous studies and present results in terms of activin’s effect on cell proliferation is not known. A response to exogenous activin appears to be dependent on the production of endogenous activin and follistatin in epithelial ovarian tumors (21, 23). Thus, the amount of secreted activin or follistatin proteins in IOSE cell lines remains to be elucidated. Interestingly, treatment or overexpression of activin resulted in a decrease in cell proliferation, which was blocked by follistatin, in the human ovarian teratocarcinoma-derived cell line (46, 47). Similarly, it has been demonstrated that activin induced growth inhibition in prostate cancer cell lines (39, 48). However, no difference was observed in the growth of normal OSE, even though all forms of activin receptors are expressed in these cells (21). The mechanism of the growth inhibitory effect of activin in IOSE cell lines remains uncertain. It has been demonstrated that the mRNA level of Smad-2, a specific signaling protein for TGFß family, was increased after activin treatment, whereas no difference was observed in Smad-4 mRNA in OVCAR-3 cells (49).
Treatment with TGFß (0.1–10 ng/mL) induced a significant decrease in the proliferative index of normal and neoplastic OSE cells in a dose-dependent manner. The expressions of TGFß isoforms and its receptors have been demonstrated in ovarian tumors, suggesting an autocrine and/or paracrine role of TGFß (28, 30, 50). TGFß inhibited the proliferation of monolayers of normal human ovarian epithelial cells by 40–70% (26) and by 95% in primary epithelial ovarian cancer cell cultures obtained directly from ascites (51). In contrast, some epithelial ovarian cancer cell lines were found to be relatively resistant to the growth inhibition of exogenous TGFß treatment (26, 52). These data suggest that TGFß may act as a growth inhibitor that prevents inappropriate proliferation of normal OSE cells, whereas loss of this autocrine growth inhibitory pathway may lead to cancer development in vivo and/or immortalization of cell in vitro. The results in this study confirm that TGFß is a prime inhibitory regulator of cell proliferation in both normal and neoplastic ovarian cells and show that it effectively inhibits cell proliferation in early neoplastic and tumorigenic transformation stages.
An increase in proliferation and/or a decrease in apoptosis play critical roles in tumorigenesis. Treatments with increasing concentrations of activin and TGFß resulted in an increase in DNA fragmentation of IOSE-29EC cells in dose-dependent manner. The effect of activin on induction of apoptosis was attenuated after 100 ng/mL follistatin treatment. Previous reports have demonstrated that activin has been shown to induce apoptosis in B cell lymphoma (53, 54), hepatoma (55), and androgen-dependent prostate cancer cells (39). The exact mechanism by which TGFß induced growth inhibition in ovarian tumor cells remains unknown. However, previous studies suggested that binding of TGFß to its receptors initiates a cascade of molecular events that are thought to decrease the activity of cyclin-dependent kinase, resulting in arrest of the cell cycle from G1 into the S phase of DNA synthesis in normal and neoplastic ovarian cells (56). In addition to cell cycle inhibition, TGFß induced apoptosis in epithelial ovarian cancer, but not in normal OSE, suggesting that neoplastic cells are more susceptible to apoptosis than their normal counterparts (27, 57). The present study indicates that both exogenous activin and TGFß induced apoptosis in neoplastic OSE cells that were growth inhibited in vitro. It is hypothesized that growth inhibition by activin or TGFß may be derived from the induction of apoptosis in this model, suggesting that apoptosis may be one of the important phenomena in growth-inhibited ovarian cancer cells. In addition, it can be postulated that IOSE cell lines represent an early transformation stage, because these cell lines are responsive to TGFß treatment in both inhibition of cell growth and induction of apoptosis.
The Bcl-2 gene family has been widely considered to be regulators of cell death (reviewed in Refs. 58 and 59). Among pro- and antiapoptotic genes in the Bcl-2 family, Bax and Bcl-2 genes are dominant regulators for apoptosis. The ratio of Bcl-2 to Bax is important in determining susceptibility to apoptosis (58). It has been shown that steroid hormones and growth factors may regulate pro- or antiapoptotic genes in ovarian and breast cancer cells (40, 57, 60). The present study has demonstrated that mRNAs of Bax and Bcl-2 are expressed in IOSE cell lines. No difference was observed in the expression level of Bax mRNA between IOSE-29 and IOSE-29EC cells. In contrast, the expression level of Bcl-2 mRNA is higher in IOSE-29EC cells than in IOSE-29 cells, suggesting that IOSE-29EC cells may be more resistant to apoptosis. In fact, relatively high expression level of Bcl-2 in IOSE-29EC cells suggests that this cell line is more resistant to serum deprivation than IOSE-29 cells (data not shown). To examine the exact mechanism by which activin and TGFß regulate apoptosis in neoplastic OSE cells, the regulation of proapoptotic Bax and antiapoptotic Bcl-2 protein was investigated after treatment with activin and TGFß, respectively. Treatment with TGFß (1 and 10 ng/mL) resulted in a significant decrease in Bcl-2 protein (up to 50%), whereas no difference was observed in Bax protein level. These findings are in agreement with a previous report that TGFß1 down-regulated the endogenous expression of the antiapoptotic Bcl-2 gene (57). Thus, down-regulated Bcl-2 may elicit apoptosis in IOSE-29EC cells, suggesting that antiapoptotic Bcl-2 is a dominant regulator of apoptosis in these cells. However, no difference was observed in Bax and Bcl-2 protein expression after treatment with increasing doses of activin. It has been reported that the expression of the proapoptotic Bax was unchanged after activin treatment in B cell lymphoma (53); however, overexpression of Bcl-2 suppressed activin-induced apoptosis. Thus, different pro- and/or antiapoptotic genes or another apoptotic pathway may be related to activin-induced apoptosis in this culture system (54, 58).
In conclusion, the present study indicates that both activin and TGFß induced growth inhibition and apoptosis in experimentally produced early neoplastic and tumorigenic OSE cells. Furthermore, antiapoptotic Bcl-2 protein was down-regulated by TGFß, whereas no difference was observed in Bax protein by activin or TGFß or in Bcl-2 protein by activin. These results suggest that activin and TGFß may play a role in growth inhibition and induction of apoptosis in early neoplastic and tumorigenic transformation stages of ovarian cancer.
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Acknowledgments |
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Footnotes |
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Received September 26, 2000.
Revised December 12, 2000.
Accepted January 29, 2001.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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- and ßA-subunit and
follistatin mRNAs by cyclic AMP and phorbol ester in cultured human
granulosa-luteal cells. Mol Cell Endocrinol. 121:1–10.[CrossRef][Medline]
and ß1. Biochim Biophys Acta. 1180:130–136.[Medline]
expression in tumor cells correlates with
resistance towards apoptosis. Int J Cancer. 60:854–859.[Medline]
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K.-Y. Kim, K.-C. Choi, S.-H. Park, C.-S. Chou, N. Auersperg, and P. C. K. Leung Type II Gonadotropin-Releasing Hormone Stimulates p38 Mitogen-Activated Protein Kinase and Apoptosis in Ovarian Cancer Cells J. Clin. Endocrinol. Metab., June 1, 2004; 89(6): 3020 - 3026. [Abstract] [Full Text] [PDF]
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C. J. Jorgez, M. Klysik, S. P. Jamin, R. R. Behringer, and M. M. Matzuk Granulosa Cell-Specific Inactivation of Follistatin Causes Female Fertility Defects Mol. Endocrinol., April 1, 2004; 18(4): 953 - 967. [Abstract] [Full Text] [PDF]
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J. L. Carey, L. M. Sasur, H. Kawakubo, V. Gupta, B. Christian, P. M. Bailey, and S. Maheswaran Mutually Antagonistic Effects of Androgen and Activin in the Regulation of Prostate Cancer Cell Growth Mol. Endocrinol., March 1, 2004; 18(3): 696 - 707. [Abstract] [Full Text] [PDF]
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 K.-C. Choi, C.-J. Tai, C.-R. Tzeng, N. Auersperg, and P. C.K. Leung Adenosine Triphosphate Activates Mitogen-Activated Protein Kinase in Pre-Neoplastic and Neoplastic Ovarian Surface Epithelial Cells Biol Reprod, January 1, 2003; 68(1): 309 - 315. [Abstract] [Full Text] [PDF]
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D. C. Danila, X. Zhang, Y. Zhou, J. N. S. Haidar, and A. Klibanski Overexpression of Wild-Type Activin Receptor Alk4-1 Restores Activin Antiproliferative Effects in Human Pituitary Tumor Cells J. Clin. Endocrinol. Metab., October 1, 2002; 87(10): 4741 - 4746. [Abstract] [Full Text] [PDF]
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 P. Y.K. Yong, C. Harlow, K.J. Thong, and S. G. Hillier Regulation of 11{beta}-hydroxysteroid dehydrogenase type 1 gene expression in human ovarian surface epithelial cells by interleukin-1 Hum. Reprod., September 1, 2002; 17(9): 2300 - 2306. [Abstract] [Full Text] [PDF]
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 G. P. Risbridger, J. F. Schmitt, and D. M. Robertson Activins and Inhibins in Endocrine and Other Tumors Endocr. Rev., December 1, 2001; 22(6): 836 - 858. [Abstract] [Full Text] [PDF]
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