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Division of Reproductive Endocrinology, Departments of Obstetrics and Gynecology (O.G.-K., U.A.K., R.A.-R., A.A.) and Pathology (W.Z.), Yale University School of Medicine, New Haven, Connecticut 06520-8063; and Departments of Medical Biology and Genetics (O.G.-K., G.L.) and Histology and Embryology (U.A.K.), School of Medicine, Akdeniz University, Antalya 07070, Turkey
Address all correspondence and requests for reprints to: Aydin Arici, M.D., Division of Reproductive Endocrinology, Department of Obstetrics and Gynecology, Yale University School of Medicine, New Haven, Connecticut 06520-8063. E-mail: aydin.arici{at}yale.edu.
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Introduction |
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PTEN gene encodes a 403-amino acid protein, which is a member of the tyrosine phosphatase family. There is a strong correlation between its phosphatase and tumor suppressor activity that has been demonstrated previously in many advanced cancers in vitro (7, 8, 9).
PTEN inhibits the phosphatidylinositol 3-kinase/Akt signaling pathway by removing the phosphate in D3-phosphate group of phosphoinositide-3, 4, 5-triphosphate (PIP3). Dephosphorylation of PIP3 is a critical determinant for controlling cell growth, proliferation, and survival. Inhibition of PIP3 causes blocking of Akt signaling, which in turn, ends up with an increased activity on proapoptotic molecules such as Bad and Caspase-9 (10). Therefore, PTEN plays important roles in cell survival and apoptosis because it inhibits cell cycle progression by down-regulating cyclin D1 and activates proapoptotic molecules through Akt-dependent and independent pathways (11, 12).
The critical importance of PTEN during development was recognized in mice with homozygous- and heterozygous-targeted deletions of the PTEN gene. Whereas PTEN knockout mice die during early embryogenesis between d 6.5 and 9.5 because of abnormal patterning of ectodermal and mesodermal germ layers, defective placentation, and overgrowth of the cephalic and caudal regions, heterozygous mice develop normally but exhibit an increased risk for a wide range of tumors including endometrial cancer (13, 14). Moreover, in human endometrial cancer, PTEN mutation rate was reported to be approximately 50–80% (15, 16).
Endometrium is capable of blastocyst implantation, regulation of trophoblast invasion, control of infectious agents, and efficient disposal of blood and desquamated cellular debris with menstruation. All of these endometrial events are carried out by mechanisms of proliferation, differentiation, and apoptosis, which are regulated by ovarian steroids.
Mutter et al. (17) have recently reported the changes in endometrial PTEN expression in both mRNA and protein level throughout the menstrual cycle, showing cyclic-related changes. In this study, we hypothesized that PTEN expression in human endometrium is variable throughout the menstrual cycle and during early pregnancy, and ovarian steroid hormones (estrogen and progesterone) regulate PTEN expression. In addition to expanding the previous study’s results to early pregnancy tissues, the present study also investigates PTEN levels and its phosphorylation in endometrial cells in response to sex steroids. The last 50 amino acid residue (354–403) in C-terminal domain of PTEN was referred as PTEN tail and is necessary to maintain stability and to determine its subcellular localization (18). Recently, Vazquez et al. (18) have shown that phosphorylation of the PTEN tail inhibits PTEN activity and decreases its degradation. Therefore, we also investigated the regulation of PTEN phosphorylation by ovarian steroids in cultured human endometrial cells, in vitro. Regulation of PTEN expression by ovarian steroids may help to protect the balance between proliferative and antiproliferative actions in normal endometrium.
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Materials and Methods |
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Endometrial tissue was obtained from human uteri after hysterectomy conducted for benign diseases other than endometrial disease or from endometrial biopsies (n = 22). Decidual tissues from first trimester (n = 10) were collected from clinically normal pregnancies, which were voluntarily terminated by dilation and curettage. Informed consent in writing was obtained from each patient before surgery; consent forms and protocols were approved by the Human Investigation Committee of Yale University and Akdeniz University. The mean age of patients was 41 (range 32–48) yr. The day of the menstrual cycle was established from the patient’s menstrual history and was verified by histological examination of the endometrium. None of the tissues used in the study were from women using exogenous steroids, and all were from premenopausal women. All the endometrial tissues were from women with regular menstrual cycles. Samples were grouped according to menstrual cycle phases: early proliferative (d 1–5), mid-proliferative (d 6–10), late proliferative (d 11–14), early secretory (d 15–18), mid-secretory (d 19–23), and late secretory (d 24–28). Tissues were embedded in paraffin for immunohistochemistry. Some endometrium samples were placed in Hanks’ balanced salt solution (HBSS) and transported to the laboratory for separation and culture of endometrial stromal and glandular cells.
Cell lines
Ishikawa cells (a well-differentiated endometrial adenocarcinoma cell line) were provided kindly to us by Dr. R. Hochberg (Department of Obstetrics and Gynecology, Yale University School of Medicine) from a frozen stock. Thawed cells were maintained in T75 flasks (BD Biosciences, Franklin Lakes, NJ) until the passage as described previously (19). RL-95 cells were purchased from American Type Culture Collection (Manassas, VA).
Isolation and culture of human endometrial stromal and glandular cells
Endometrial stromal and glandular cells were separated and maintained in monolayer culture, as described previously (20). Endometrial tissue was rinsed in HBSS to remove blood and debris. Briefly, endometrial tissue was digested by incubation of tissue minces in HBSS (Sigma, St. Louis, MO) that contained HEPES (25 mmol, Sigma), penicillin (200 U/ml, Sigma), streptomycin (200 mg/ml, Sigma), collagenase (1 mg/ml, 15 U/mg, Sigma), and deoxyribonuclease (0.1 mg/ml, 1500 U/mg, Sigma) for 30 min at 37 C with agitation. The dispersed endometrial cells were separated by filtration through a wire sieve (73-µm diameter pore, Sigma). The endometrial glands (largely undispersed pieces) were retained by the sieve, whereas the dispersed stromal cells passed through the sieve into the filtrate.
Stromal cells were plated in Ham’s F12/DMEM (1:1 vol/vol; Sigma) containing fetal bovine serum (10% vol/vol, Life Technologies, Inc., Rockville, MD) and antibiotics-antimycotics (1% vol/vol, Life Technologies, Inc.) in T-75 plastic flasks (Falcon, Bedford, MA). Cells were plated in plastic flasks (75 cm2, Falcon), maintained at 37 C in a humidified atmosphere (5% CO2 in air), and allowed to replicate to confluence. Thereafter, the stromal cells were passed by standard methods of trypsinization and plated in culture dishes (100-mm diameter) and were allowed to replicate to confluence, which takes approximately 7–10 d. Endometrial stromal cells after first passage were characterized previously (20), and were found to contain 0–7% epithelial cells, no endothelial cells, and 0.2% macrophages. In present experiments, cultures were confirmed to have similar purity. Experiments were commenced 1–3 d after confluence was attained. The confluent cells were treated with serum-free, phenol red-free media (Sigma) for 24 h before treatment with steroids.
Endometrial glandular cells (largely intact glands and sheets of surface epithelium) were collected by back washing the sieve and plated in six-well plates, previously coated with growth factor-reduced matrigel (Collaborative Research, Boston, MA). Cells were maintained in DMEM containing 10% fetal bovine serum, 1% antibiotics-antimycotics, and D-valine (substituted for L-valine to inhibit stromal cell growth, Life Technologies, Inc.) (21). Glandular cells reach confluence in 5–7 d. Endometrial glandular cells in culture were characterized using cytokeratin-7 antibody and were found to contain 0.1–2% leukocytes, 1–4% stromal cells, and no endothelial cells. If cultures were contaminated with more than 10% other cells than glandular cells, these cultures were excluded from the study. Experiments with glandular cells were conducted 1–3 d after confluence was attained. Only pure glandular cells were treated with sex steroids. Cells were treated with serum-free, phenol red-free media for 24 h before treatment with steroids. Stromal and glandular cells were treated with 17ß-estradiol (1 x 10-8 M) and progesterone (1 x 10-8 M) for various times for Western blot analysis and immunocytochemistry staining.
Immunohistochemistry and immunocytochemistry
Paraffin-embedded tissue samples were cut and mounted on SuperFrost Plus slides (Erie Scientific Co., Portsmouth, NH). Following deparaffinization, slides were rinsed with PBS for 10 min. Endogenous peroxidase activity was quenched by incubation in 3% H2O2 for 20 min and followed by a rinse in PBS. For antigen retrieval, slides were placed in 10 mM of citrate buffer, microwaved twice for 5 min, and rinsed in PBS. Sections were incubated with 5% normal serum to block nonspecific staining for 30 min at room temperature. Thereafter, sections were incubated overnight at 4 C with mouse monoclonal antihuman PTEN antibodies [1/100, Cell Signaling (Beverly, MA) and Santa Cruz Biotechnology, Inc. (Santa Cruz, CA)]. Antibodies’ specificity were tested in PTEN-mutant Ishikawa and RL-95 cells. For negative control slides, normal mouse IgG isotype (Vector Laboratories, Burlingame, CA) was used instead of primary antibodies for PTEN. The sections were washed in Tris-buffered saline (TBS), incubated with biotinylated horse antimouse IgG (Vector Laboratories), and then incubated with streptavidin-peroxidase complex using the Vectastain ABC Elite kit (Vector Laboratories). Subsequently, the chromogenic reaction was carried out with 3-amino 9-ethyl carbazole (Vector Laboratories) and the reaction was terminated with tap water. Slides were counterstained with hematoxylin before permanent mounting and then evaluated under a light microscope.
To investigate whether intracellular localization of PTEN and phospho-PTEN is different, we performed immunofluorescence staining in endometrial stromal cells plated on chamber slides (Falcon). Cells were treated with estradiol (1 x 10-8 M) for 5 and 15 min, and slides were then fixed in 4% paraformaldehyde at 4 C for 20 min and washed three times with TBS for 5 min at room temperature. After blocking step, chamber slides were incubated overnight at 4 C with mouse monoclonal antihuman PTEN (1/100) and rabbit polyclonal antihuman phospho-PTEN (1/100) antibodies (Cell Signaling). Mouse antihuman PTEN antibody detects the total level of endogenous PTEN protein in cells. Specificity of this antibody was tested by Birle et al. (22) and has been shown that it detects endogenous levels of PTEN when only phosphorylated at Ser380 residue. For negative control slides, normal mouse IgG isotype and normal rabbit IgG (Vector Laboratories) were used instead of primary antibodies for PTEN and phospho-PTEN, respectively. Following the overnight incubation of primary antibody, chamber slides were incubated with Cy-3 conjugated antimouse IgG and Cy-2 conjugated antirabbit IgG (Jackson ImmunoResearch Laboratories, Inc., West Grove, PA), counterstained with 4',6-diamidino-2-phenylindole and mounted with fluorescence specific media (Vector Laboratories). For each experiment, all slides were stained in a single batch and were exposed to identical staining.
The intensity for PTEN immunoreactivity in endometrial tissues was semiquantitatively evaluated as positively stained cells according to the following categories: -, no staining; 1+, weak but detectable; 2+, moderate or distinct; 3+, intense. For each tissue, at histologic score (HSCORE) value was derived by summing the percentages of cells that stained at each intensity category and multiplying that by the weighted intensity of the staining, using the formula [HSCORE= Pi (i + l)], where i represents the intensity scores and Pi is the corresponding percentage of the cells. In each slide, five different areas were evaluated under a microscope with x40 original magnification, the percentage of the cells for each intensity within these areas was determined at different times by two investigators who were blinded to treatments, and the average score was used.
Preparation of nuclear extracts
Nuclear extracts from endometrial stromal cells grown to confluence in 60-mm plates were performed using a nuclear extraction kit (Aktiv Motif, Carlsbad, CA). Briefly, cells were washed with ice-cold PBS and phosphatase inhibitors, removed from the dish by scraping with a cell lifter, and transferred to prechilled tubes. Cell suspensions were centrifuged at 4 C for 5 min at 500 rpm. Pellets were resuspended in hypotonic buffer and incubated for 15 min on ice, after adding a detergent centrifuged again at 4 C for 30 sec at 14,000 x g. The pellet was resuspended in a lysis buffer and incubated for 30 min on ice on a rocking platform. The suspension was centrifuged at 4 C for 10 min at 14,000 x g, and the supernatant (nuclear fraction) was aliquoted and frozen at -80 C. An aliquot of each sample was used to quantify the nuclear protein amount via Coomassie protein assay (Pierce, Rockford, IL). Five micrograms of nuclear extract sample were loaded into each well and assayed according to manufacturer’s directions (Aktiv Motif) using a microplate reader.
Western blot analysis
Total protein from the cells was extracted using T-PER tissue protein extraction reagent (Pierce), supplemented with protease inhibitor cocktail (1 mM Na3VO4, 10 µg/ml leupeptin, 10 µg/ml aprotinin, and 1 mM phenylmethylsulfonylfluoride; Calbiochem, San Diego, CA). The protein concentration was determined by Bradford assay (Pierce). Twenty micrograms of protein were loaded into each lane, separated electrophoretically by SDS-PAGE using 10% Tris-HCl Ready Gels (Bio-Rad Laboratories, Hercules, CA), and electroblotted onto nitrocellulose membrane (Bio-Rad). The membrane was blocked with 5% nonfat dry milk in TBS-T buffer (0.05% Tween-20 in TBS) for 1 h to reduce the nonspecific binding. Then the membrane was incubated with mouse monoclonal antihuman PTEN antibody (Cell Signaling) overnight at 4 C, then washed three times with TBS-T for 20 min. The membrane was incubated for 1 h with peroxidase-labeled antimouse IgG (Vector Laboratories) and subsequently washed with TBS-T three times for 20 min. Immunodetection was developed with chemiluminescent detecting reagents (NEN Life Science Products, Boston, MA), and subsequently the membrane was exposed to BioMax film (Kodak, Rochester, NY).
After the membrane was stripped/stripping solution (Pierce), the same membrane was reprobed with rabbit polyclonal antihuman phospho-PTEN antibody (Cell Signaling), followed by the other steps as described above. Equal loading of proteins in each lane was confirmed by probing the membrane with mouse monoclonal antihuman glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (Santa Cruz Biotechnology, Inc.). Immunoblot bands for PTEN, phospho-PTEN, and GAPDH were quantified using a laser densitometer. Each PTEN and phospho-PTEN band was normalized to the value obtained from the same lane blotting GAPDH.
Statistical analysis
Levels of Western blot densitometries and HSCORE of immunohistochemistry were normally distributed as tested by Kolmogorov-Smirnov test and were analyzed by ANOVA and post hoc Tukey test for pair wise comparisons. Statistical significance was defined as P < 0.05. Statistical calculations were performed using Sigmastat for Windows, version 2.0 (Jandel Scientific Corp., San Rafael, CA). Each experiment was repeated at least three times using cells prepared from three different endometrial tissues.
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Results |
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We first tested the specificity of the antibodies. Total protein from PTEN-mutant Ishikawa and RL-95 cells was extracted and assessed by Western blot analysis. No specific PTEN band was detected in either Ishikawa or RL-95 cells (Fig. 1
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Immunohistochemistry revealed stronger PTEN staining in stromal cells than glandular cells throughout the menstrual cycle (Fig. 2, A and B, and Fig. 3). During the proliferative phase, PTEN immunoreactivity in stromal cells was mainly in the cell nucleus and less in the cytoplasm (Fig. 2A). Compared with proliferative phase, during secretory phase and early pregnancy this immunoreactivity was mainly in the cytoplasm (Fig. 2, A–C). In general, during the proliferative phase PTEN expression was either weak or not detectable in the glandular epithelium, except for a strong immunoreactivity in early proliferative phase samples. On the other hand, during the secretory phase PTEN immunoreactivity was increased in the glandular epithelium (Fig. 2, A and B, and Fig. 3). The staining in glandular cells was heterogenous. Specifically, some glandular segments remained either negative or weakly positive for PTEN expression, whereas other segments were more positive (Fig. 2B).
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, A–C, and Fig. 3). Among all groups, the strongest immunoreactivity was observed in glandular and decidual cells of early pregnancy with a gradual increase as pregnancy advanced (Fig. 2C). Regulation of PTEN and its phosphorylation in endometrial stromal and glandular cells
If endometrial PTEN expression shows the cyclic manner changes throughout menstrual cycles and early pregnancy, its expression and/or its phosphorylation may be regulated by ovarian steroids in a short-term and/or long-term manner. To investigate this hypothesis, endometrial stromal and glandular cells were treated with estradiol (1 x 10-8 M) and progesterone (1 x 10-8 M) for short (5–90 min) and long (3–24 h) time courses. PTEN and phospho-PTEN protein levels were assessed by Western blot analysis. In endometrial stromal cells treated with estradiol, PTEN levels were 28% and 30% higher at 5 and 15 min, respectively, compared with control cells (P < 0.05) (Fig. 4A). Phospho-PTEN levels reached a peak following 15 min of treatment with estradiol. Increased PTEN levels in estradiol-treated cells were associated with elevated levels of PTEN phosphorylation. PTEN phosphorylation in estradiol-treated cells was 29% and 46% higher at 5 and 15 min, respectively, than that observed in control cells (P < 0.05) (Fig. 4B). Long-term estradiol treatment (3–24 h) did not affect phospho-PTEN level, but a nonsignificant decrease was observed in the PTEN protein level after 24 h (Fig. 4C). Short-term treatment (5–30 min) of endometrial stromal cells with progesterone (10-8 M) decreased both PTEN and phospho-PTEN levels when compared with control, which was not statistically significant. PTEN level was 11–26% and phospho-PTEN level was 9–15% lower in cells treated with progesterone compared with control cells at 5 and 30 min, respectively (Fig. 5A). On the other hand, long-term progesterone treatment (10-8 M) induced a 23–76% increase in PTEN level and a 50–37% increase in phospho-PTEN levels at 6–24 h, respectively (P < 0.05) (Fig. 5B).
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). Estradiol treatment induced a marked increase in both PTEN and phospho-PTEN levels, whereas progesterone induced a not significant decrease in PTEN and phospho-PTEN levels.
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To investigate the specific localization of phosphorylated and normal forms of PTEN in the cell, we performed immunofluorescence staining using mouse antihuman-PTEN and rabbit antihuman-phospho-PTEN antibodies in endometrial stromal cells cultured in chamber slides. Immunofluorescence analysis revealed that phospho-PTEN is mostly localized in the nucleus, whereas normal PTEN is mainly in the cytoplasm (Fig. 7, A–C). Endometrial stromal cells plated onto eight-well chamber slides were incubated with estradiol (1 x 10-8 M) for 5–15 min, and cells were analyzed by immunofluorescence staining. Following 15-min estradiol treatment compared with control, there was an increase in nuclear PTEN immunoreactivity and a decrease in cytosolic PTEN immunoreactivity. Double immunofluorescence analysis revealed that nuclear staining is related to phospho-PTEN (Fig. 7, D–F).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Normal endometrium is hormonally regulated in a cyclic manner and undergoes highly dynamic changes throughout the menstrual cycle and during early pregnancy. Estrogen and progesterone play important roles in the regulation of these changes (25). Mutter et al. (17) have recently reported the changes in endometrial PTEN expression throughout the menstrual cycle. Consistent with the data of that study, our immunohistochemical results revealed that PTEN expression in normal and decidualized endometrial tissues is under the cyclic influence of estrogen and progesterone and has a complex and cell-specific distribution and subcellular localization. Stronger PTEN immunoreactivity observed in endometrial stromal cell nuclei compared with its cytosol during the proliferative phase may be related to its increased phosphorylation because our immunofluorescence staining for phospho-PTEN showed a nuclear pattern. The finding that there are no significant changes in PTEN immunoreactivity during mid- and late-proliferative phases suggests that estrogen may not have a stimulatory effect on PTEN expression in vivo in a long-term manner. On the other hand, nuclear localization of PTEN is poorly understood, and may be related to another yet unknown function besides its known cytoplasmic roles. It is well known that the proliferative phase of human endometrium is characterized by a low number of apoptotic cells, and endometrium stromal cells are less sensitive to apoptotic stimuli compared with epithelial cells (26). One of the mechanisms under the less sensitivity against apoptotic stimuli in stromal cells is likely to be related to the regulation of PTEN activity by estrogen.
Higher PTEN immunoreactivity in endometrial stromal and glandular cells during late secretory and early proliferative phases could be related to decreased cell proliferation and increased apoptosis of these cells. Moreover, further increase in PTEN expression in decidual and glandular cells during early pregnancy supports this hypothesis because during the process of implantation and early pregnancy numerous cells undergo apoptosis. Endometrial cell apoptosis seems to play an important role in the regulation of endometrial decidualization and trophoblast invasion (27).
Interestingly, PTEN expression is not homogenous in glandular cells. Specifically, in the same section, some glands are either negative or weakly positive, whereas others are strongly positive for PTEN immunoreactivity. Recently, similar staining pattern for Fas and proliferating cell nuclear antigen expression in glandular cells were reported by Demir et al. (28). One possibility is that immunohistochemically PTEN-negative glands may have somatic PTEN mutations because endometrium is one of the highest tissues of ratio of somatic PTEN mutations. This may be important for early stage detection of endometrial cancer. A relationship between PTEN-null glands and the incidence of endometrial cancer was recently reported by Mutter et al. (29). Loss of PTEN occurs in approximately 50% of all endometrial carcinomas, increasing to 83% for tumors with adjacent premalignant lesions (29). In the present study, we are reporting for the first time the expression of PTEN during early pregnancy and comparing its expression throughout menstrual cycle. In terms of our in vivo findings, it seems that PTEN expression may be regulated by ovarian steroids throughout the menstrual cycle and early pregnancy, and our in vitro results support this hypothesis. Moreover, increased PTEN expression in secretory phase of endometrium is likely to be regulated by progesterone because in vitro progesterone treatment stimulates PTEN in endometrial stromal cells. Our in vitro results suggest that in endometrial cells progesterone but not estrogen regulates PTEN expression in a long-term manner.
PTEN is comprised of an N-terminal domain known as phosphatase domain, a C-terminal domain that contains the lipid-binding C2 domain, PEST (proline, glutamic acid, serine) domains that regulate protein stability, and the PDZ (pSD-95/Dlg and ZO1) domain, which is important in protein-protein interactions (30). Vazquez et al. (31) have suggested that PTEN is phosphorylated in a specific C-terminal region, referred as PTEN tail, which is rich in serine and threonine residues. PTEN tail is important for PTEN phosphatase activity and is also necessary for maintaining PTEN stability. Two different groups have recently shown that phosphorylated PTEN is less active but more stable, whereas dephosphorylation of PTEN increases its activity and its interaction with PDZ domain-containing binding partners, thus its degradation (22, 31).
Our results demonstrate that, although estrogen does not affect PTEN expression by a long-term manner, it regulates PTEN level by affecting its phosphorylation in a short-term manner. Increased phosphorylation of PTEN tail in the presence of estrogen may regulate both its stability and activity, and may affect its nuclear localization, confirming our in vivo findings. Nuclear PTEN localization has been shown by numerous other studies (32, 33, 34). A balance between PTEN phosphorylation and dephosphorylation regulates its activity. Unfortunately, there are no studies describing a role for phospho-PTEN in the nucleus. Dephosphorylation of PTEN tail would result in an increase in PTEN activity but also in its rapid degradation by Proteosoma. As pointed out above, PTEN phosphorylation affects both its stability and cellular localization of PTEN and/or its ability interacting with other intercellular proteins. One hypothesis is that estrogen may activate a kinase related to the phosphorylation of PTEN. Miller et al. (35) have shown that PTEN is phosphorylated by protein kinase casein kinase II, which also phosphorylates estrogen receptor 
(36). Thus, estrogen-activating casein kinase II may cause a decrease of PTEN activity, but an increase of estrogen receptor activity. Estrogen may lead to a decreased cytoplasmic activity of PTEN by increasing the PTEN phosphorylation, which in turn would cause increased Akt phosphorylation by phosphatidylinositol 3-kinase (37). Therefore, we propose that PTEN is one of the pathways by which estrogen may be acting to affect endometrial stromal cell proliferation and/or apoptosis.
In conclusion, endometrial PTEN expression has temporal and spatial changes throughout the menstrual cycle and during early pregnancy. Changes in the endometrial PTEN pool are likely to be regulated by estrogen and progesterone in both long-term and short-term manner. Estrogen may down-regulate PTEN activity by increasing its phosphorylation, which requires further studies to prove. This in turn may result in increased PIP3, which initiates activation of second messengers involved in cell proliferation and survival. On the other hand, progesterone is likely to regulate the PTEN pool by decreasing its phosphorylation and increasing its protein level in short-term and long-term manners, respectively. We propose that PTEN might be one of the signaling proteins that estrogen and progesterone are acting to affect endometrial cell proliferation and/or apoptosis. It should be interesting to further investigate the molecular mechanisms of PTEN phosphorylation after estrogen treatment in the future therapy for estrogen-dependent diseases such as endometriosis.
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Footnotes |
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Received March 10, 2003.
Accepted July 14, 2003.
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