HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

This Article Abstract Full Text (PDF) All Versions of this Article: 291/1/E59    most recent 00582.2005v1 Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in ISI Web of Science Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Citing Articles Citing Articles via HighWire Citing Articles via ISI Web of Science (6) Citing Articles via Google Scholar Google Scholar Articles by Shao, R. Articles by Billig, H. Search for Related Content PubMed PubMed Citation Articles by Shao, R. Articles by Billig, H.

Nuclear progesterone receptor A and B isoforms in mouse fallopian tube and uterus: implications for expression, regulation, and cellular function

¹Section of Endocrinology, Department of Physiology and Pharmacology, The Sahlgrenska Academy; ²Department of Obstetrics and Gynecology, Sahlgrenska University Hospital; ³Perinatal Center, Department of Physiology and Pharmacology, The Sahlgrenska Academy; and ⁴Swegene Centre for Cellular Imaging, Göteborg University, Gothenburg, Sweden

Submitted 28 November 2005 ; accepted in final form 25 January 2006

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Progesterone and its interaction with nuclear progesterone receptors (PR) PR-A and PR-B play a critical role in the regulation of female reproductive function in all mammals. However, our knowledge of the regulation and possible cellular function of PR protein isoforms in the fallopian tube and uterus in vivo is still very limited. In the present study, we revealed that equine chorionic gonadotropin (eCG) treatment resulted in a time-dependent increase in expression of both isoforms, reaching a maximal level at 48 h in the fallopian tube. Regulation of PR-A protein expression paralleled that of PR-B protein expression. However, in the uterus PR-B protein levels increased and peaked earlier than PR-A protein levels after eCG treatment. With prolonged exposure to eCG, PR-B protein levels decreased, whereas PR-A protein levels continued to increase. Furthermore, subsequent treatment with human (h)CG decreased the levels of PR protein isoforms in both tissues in parallel with increased endogenous serum progesterone levels. To further elucidate whether progesterone regulates PR protein isoforms, we demonstrated that a time-dependent treatment with progesterone (P₄) decreased the expression of PR protein isoforms in both tissues, whereas decreases in p27, cyclin D₂, and proliferating cell nuclear antigen protein levels were observed only in the uterus. To define the potential PR-mediated effects on apoptosis, we demonstrated that the PR antagonist treatment increased the levels of PR protein isoforms, induced mitochondrial-associated apoptosis, and decreased in epidermal growth factor (EGF) and EGF receptor protein expression in both tissues. Interestingly, immunohistochemistry indicated that the induction of apoptosis by PR antagonists was predominant in the epithelium, whereas increase in PR protein expression was observed in stromal cells of both tissues. Taken together, these observations suggest that 1) the tissue-specific and hormonal regulation of PR isoform expression in mouse fallopian tube and uterus, where they are potentially involved in regulation of mitochondrial-mediated apoptosis depending on the cellular compartment; and 2) a possible interaction between functional PR protein and growth factor signaling may have a coordinated role for regulating apoptotic process in both tissues in vivo.

progesterone receptor protein isoforms

PR-A and PR-B are coexpressed in most target tissues. However, the regulation of PR proteins has been found to be tissue and cell type specific in female reproductive tissues in vivo (20). Our laboratory has demonstrated that treatment with human chorionic gonadotropin (hCG), mimicking endogenous preovulatory luteinizing hormone surge, induces the levels of PR protein isoforms in periovulatory granulosa cells in mouse ovary (62). Similar observations have been made in ovaries from intact adult mice during the estrous cycle (19) and in humans (22). In addition, the developmental time course of PR mRNA and protein expression has been extensively studied in the uterus by use of either ovariectomized animals (a background devoid of endogenous steroid hormones) treated with selected steroid hormones (27, 30, 31, 70) or intact adult rats during the estrous cycle (46). Furthermore, it has previously been reported that the PR protein isoforms are expressed and regulated in human endometrium during the menstrual cycle (39, 43). Relative to ovary and uterus, earlier studies have demonstrated that specific binding sites for P₄ can be observed using the ligand-binding characteristic assay in human and rodent fallopian tubes, (37, 45, 52, 53, 73). Later reports have demonstrated that PR mRNA and protein expression can be detected by RT-PCR (6, 50, 66), Western blot analysis (6, 66), and immunohistochemical localization (1, 6, 18, 19, 50, 66). Although PR proteins display cyclic changes in human fallopian tubes during the menstrual cycle (1, 73), PR protein isoforms expressed in the fallopian tubes, their regulation, and their role in tubular function are much less well understood.

Apoptosis is an active mode of cell death that is triggered by a variety of physiological and pathological stimuli (23). Inappropriate activation or inhibition of apoptosis may cause or contribute to a variety of diseases (69). Within the complex network of proteins regulating apoptosis, proteins for caspase family are universal effectors in the apoptotic process in mammalian cells (15). Members within this wide family have been divided into initiator caspases (e.g., caspase-9) and executioner caspases (e.g., caspase-3). It is known that functional caspase-9 is the critical player of the apoptotic stimuli, acting through mitochondrial-mediated death pathway (29). When release of cytochrome c (a proapoptotic factor) from mitochondria into cytosol is enhanced in response to apoptotic stimuli, procaspase-9 interacts with cytochrome c, Apaf-1, and ATP to liberate active caspase-9 (34), which in turn generates activation of caspase-3 (21). One of the biochemical hallmarks of apoptosis is the degradation of DNA that is induced by the DNA fragmentation factor (DFF). Caspase-activated deoxyribonuclease (CAD), a subunit of DFF, has been shown to cleave genomic DNA into internuclease fragments activated mainly by caspase-3 in humans and mice (16, 59), resulting in characteristic changes associated with the apoptotic nuclear morphology, such as membrane blebbing, nuclear condensation, and DNA fragmentation (24). In addition to physiological evidence for nuclear PR actions by effect of P₄ in female reproduction [e.g., the establishment and maintenance of pregnancy (20)], several studies from our laboratory and others’ have previously demonstrated that nuclear PRs as intraovarian regulators act in an autocrine manner to influence granulose cell survival in humans and rodents (40, 62). However, the intracellular mechanisms involved in the activation of apoptotic pathways by PR signaling through regulation of PR protein levels in the fallopian tube and uterus are largely unknown.

The present study was designed to investigate the time-dependent expression and regulation of PR protein isoforms in response to gonadotropins in mouse fallopian tube and uterus in vivo using quantitative Western blotting and immunohistochemistry. To understand the impact of PR on the fallopian tube and uterus functions, we have demonstrated the effect of exogenous progesterone on expression of PR isoforms in parallel with expression of cell cycle and proliferation proteins. Furthermore, it assesses the effect of the PR antagonists on the PR protein expression, apoptosis induction and regulation of epidermal growth factor (EGF), and EGF receptor protein expression in both tissues.

	MATERIALS AND METHODS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Hormones and reagents. Equine chorionic gonadotropin (eCG), P₄, mouse monoclonal anti--actin, mouse monoclonal anti--tubulin, alkaline phosphatase-conjugated goat anti-mouse immunoglobulin G (IgG), and alkaline phosphatase-conjugated mouse anti-goat/sheep IgG were obtained from Sigma Chemical (St. Louis, MO). Human (h)CG and the PR antagonist Org 31710 (26) were obtained from N.V. Organon (Oss, Holland). The PR antagonist RU-486 (4) was obtained from Exelgyn (Paris, France). The antibodies used to detect PR in this study were raised against human PRs and recognized the ligand-binding domain (PR c-19) or DNA-binding domain (PR c-20) of PR in rodent tissues by Western blotting and immunohistochemical analyses. Rabbit polyclonal anti-PR (PR c-19 and PR c-20) and their respective blocking peptides rabbit polyclonal anti-cyclin D₂, rabbit polyclonal anti-p27^Kip1 (p27), mouse monoclonal anti-apoptosis-inducing factor (AIF), rabbit polyclonal anti-caspase-3, rabbit polyclonal anti-caspase-9, rabbit polyclonal anti-CAD, goat polyclonal anti-EGF, rabbit polyclonal anti-EGF receptor, and normal mouse IgG were obtained from Santa Cruz Biotechnologies (Santa Cruz, CA). Mouse monoclonal antiproliferating cell nuclear antigen (PCNA) was obtained from Novocastra Laboratories (Newcastle upon Tyne, UK). Mouse monoclonal anti-cytochrome c and rabbit polyclonal anti-activated caspase-3 were obtained from BD PharMingen (San Diego, CA). Normal rabbit serum and goat serum were obtained from DAKO (Carpinteria, CA). Alkaline phosphatase-conjugated goat anti-rabbit IgG was obtained from Tropix (Bedford, MA). Cy3-conjugated donkey anti-mouse IgG was purchased from Jackson ImmunoResearch Laboratories (West Grove, PA). Other reagents not mentioned in the text were purchased from Sigma or Merck (Darmstadt, Germany) and were of the highest purity grade available.

Experimental animals. The animal studies were reviewed and approved by the ethics committee at Göteborg University, Sweden. All murine analyses were performed in female C57BL/6 mice. Mice (21 days old) were purchased from Taconic M&B, Ejby, Denmark. Mice were initially allowed to acclimate to the facility for 5 days before the beginning of any study and were provided standard chow and tap water ad libitum. All animals were individually housed under defined conditions with temperature control, constant humidity, and a regular light-dark cycle. Mice at 26 days of age with a body weight ranging from 13–15 g were used in this study.

Hormone treatment, tissue preparation, and blood collection. To determine the response to gonadotropins and induction of ovulation (62), immature female mice (30 mice in one experiment) were given a single injection of 5 IU eCG or saline (hormone vehicle). eCG was administered intraperitoneally for 24 or 48 h. In addition, some of the animals (40 mice in one experiment) were injected intraperitoneally with 5 IU eCG followed 48 h later with a single injection of 5 IU hCG (ip) for the indicated time (6, 12, 24, and 48 h). The fallopian tubes and uteri were grossly dissected free of contaminating tissue (e.g., adipose tissue) after mice were killed by decapitation. One-half of the fallopian tube and uterine horn in each animal was either immediately frozen in liquid nitrogen and stored at –70°C for subsequent Western blot analysis or fixed in 4% formaldehyde neutral buffered solution for 24 h at 4°C and embedded in paraffin for histochemical analysis. Protein was harvested from a matched portion of the organ in all instances to ensure that results did not differ because of organ composition. In all cases, trunk blood was obtained from heart puncture before tissue collection. Sera were prepared by clotting, centrifugation, and then collection of supernatants. Sera were stored at –20°C for subsequent analysis.

Validation of eCG and/or hCG mouse model. Ovarian weight of immature female mice remained relatively constant between 26 and 30 days of age. An increase in ovarian weight was noted in mice treated with eCG (day 26). They were increased further within 6 h after subsequent treatment with hCG, remained unchanged through 12 h, and then continued to decrease at 48 h (Fig. 1A). Treatment of mice with eCG for 48 h mimicked ovaries at the preovulatory phase of a reproductive cycle, such as increase in follicular size, cumulus expansion, and oocyte maturation as well as significant changes a number of gonadotropin-dependent gene expression (54). Subsequent hCG treatment (from 6 to 48 h) encompassed the interval preceding ovulation (Table 1) until the luteal phase of the ovarian cycle. Ovarian morphology (47, 48) and numbers of different stage of follicles (5) were assessed in mice with a regimen of 5 IU eCG and/or 5 IU hCG by examining histological step-sections of one ovary (data not shown). Serum estradiol concentration increased by 24 h and peaked 48 h after eCG injection (Fig. 1B), whereas serum P₄ concentration showed a tendency to significantly increase after post-hCG treatment and peaked at 48 h (Fig. 1B). The production of estradiol and progesterone in immature mice treated with eCG and/or hCG in this study is similar to that in adult cycling mice (74). This in vivo model has been used to stimulate follicular development, induce ovulation, and cause significant changes in the expression of PR protein isoforms in mouse ovarian granulose cells (62).

View larger version (14K): [in this window] [in a new window]   Fig. 1. Effect of gonadotropins on ovary weights (A) as well as serum estradiol and progesterone (B) concentrations in immature mice. The number of mice per group is indicated. eCG and hCG, equine and human chorionic gonadotropin, respectively. Trunk blood was obtained from heart puncture. Sera were collected after clotting and centrifugation. Serum estradiol and progesterone concentrations were measured in duplicate aliquots of each sample. Values represent means ± SE. ** and ***Significantly different from untreated animals (time, 0 h), P < 0.01 and P < 0.001, respectively.

To test the effects of PR antagonists (RU-486 and Org 31710) on the regulation of PR isoform expression and ovulation in response to gonadotropin stimulation (33), animals were randomly given either 1 mg of RU-486 or 2 mg of Org 31710 in combination with 5 IU of hCG after treatment with 48-h eCG (5 IU ip). Both RU-486 and Org 31710 were dissolved in 100% ethanol and resuspended in sesame oil. All intraperitoneal injections were in a volume of 100 µl. An equal volume of 100% ethanol and sesame oil was injected to control mice. After 24 h of cotreatment with hCG and/or PR antagonist, either RU-486 or Org 31710, the fallopian tubes, and uteri were collected in the different groups for either Western blot or immunohistochemical analysis. In this study, the doses of RU-486 and Org 31710 were chosen on the basis of previous studies (36, 56). Additional treatment with dexamethasone [600 µg/kg dissolved in saline and resuspended in 100 µl of sesame oil (58)] was used for PR antagonist specific effect on ovulation. All drug solutions were freshly prepared before each experiment. In addition, the number of ovulations was determined by flushing the fallopian tubes and counting the number of oocytes retrieved under microscope. Mice exhibiting ovulation after PR antagonist treatments were discarded from the study (Table 1).

Protein extraction and Western blot analysis. Whole cell protein extract and Western blot were done essentially as described previously (64). Tissue extracts were homogenized at 4°C and incubated on ice for 30 min for a high-salt lysis buffer that effectively extracts all DNA-binding proteins. Subsequently, insoluble material was removed by centrifugation at 10,000 g for 30 min at 4°C. The protein content of the extracts was determined using a bicinchoninic acid protein assay (Pierce, Rockford, IL) with BSA as the standard. Equal amounts of protein per sample were heated at 95°C for 5 min with reducing agents and loaded onto 4–12% sodium dodecyl sulfate (SDS)-polyacrylamide gels (Novex, San Diego, CA) using a MOPS-SDS running system. The nonspecific protein-binding site on the polyvinylidene difluoride membranes (Amersham International, Buckinghamshire, UK) was blocked by incubation of the membrane with 5% nonfat milk in a Tris-buffered saline (TBS)-Tween 20 buffer (10 mM Tris, 150 mM NaCl, and 0.1% Tween 20, pH 8.0) for 4 h. The membranes were incubated with the indicated primary antibodies in blocking buffer overnight at 4°C and the appropriate alkaline phosphatase-linked secondary antibodies with 1:40,000 or 1:80,000 dilutions for 2 h. Preparation of anti-PR (c-19 + c-20, 1:250 for each), anti-PCNA (1:1,000), and anti-activated caspase-3 (1:500) has been described previously (62). The primary antibodies anti-cyclin D₂ (1:1,000), anti-p27^Kip1 (p27; 1:1,000), anti-AIF (1:250), anti-caspase-9 (1:250), anti-CAD (1:1,000), anti-EGF (1:250), and anti-EGF receptor (1:250) were used. The immunosignal-CDP-Star substrate for alkaline phosphatase system (Tropix, Bedford, MA) was used to visualize protein bands. To reprobe the blot with another antibody, the blot was rehydrated in methanol, rinsed, and incubated with stripping buffer (65 mM Tris·HCl, 2% SDS, and 100 mM -mercaptoethanol, pH 6.8) at 50°C for 30 min. Blots were then washed for 2 h with several changes of TBS-Tween 20 buffer at room temperature. ECL films of the Western blots were captured with an image scanner (EPSON perfection 2450 photo). The intensities of the immunoreactive bands were converted into digitized signals by use of a computerized densitometry system and quantified by a gel documentation program (ImageQuant version 5.0; Molecular Dynamics, Sunnyvale, CA). Signal intensities of the mouse PR and caspase-3 proteins were normalized to those of mouse -actin protein as ratios to produce arbitrary densitometric units (ADU) of relative abundance. Each Western blot was quantified using a standard curve consisting of serial dilutions of one of the samples included in the assay. All samples for a particular experiment were run on the same gel, or set of gels run at the same time, under the same conditions. Because exposure times, transfer efficiency, and other factors may vary between runs, ADUs for relative protein levels were only directly compared within a set of gels run at the same time. All steps were carried out at room temperature unless otherwise stated.

Histological and immunohistochemical analyses. Tissue blocks were sectioned at 5 µm thickness onto poly-L-lysine-coated slides. They were dewaxed by two 5-min washes in xylene and rehydrated through graded alcohol by 5-min washes with a final wash in water. A standard citrate boiling antigen retrieval procedure was used (62, 64). Sections were blocked in 0.3% H₂O₂ in TBS and 10% normal goat serum. The primary antibody was diluted 1:250 (anti-PR c-19 + c-20) or 1:50 (anti-activated caspase-3) in TBS containing 1% BSA, and the sections were incubated overnight at 4°C. The appropriate secondary antibody was added. After a wash with TBS, sections were stained using the avidin-biotinylated peroxidase complex detection system (ABC kit, Vector Laboratories, Burlingame, CA) according to the manufacturer’s instructions. Immunostaining was then visualized using 3,3-diaminobenzidine tetrahydrochloride (0.5 mg/ml in PBS + 0.01% H₂O₂, pH 7.6) for 10 min. Slides were viewed on a Nikon E-1000 microscope under bright-field optics and photomicrographed using Easy Image 1 (Bergström Instrument, Gothenburg, Sweden). Ovary sections from mice treated with eCG and hCG (62) and human breast carcinoma sections (41) were used as positive controls because nuclear PR is highly expressed in these tissues. Three negative-control procedures (exclusion of either primary antibody or secondary antibody in the tissue sections, application of either normal goat serum or normal rabbit serum, and preabsorption of the anti-PR antibody with a 100-fold excess of antigen) were run simultaneously to confirm the specificity of the immunostaining. Serial sections of the fallopian tubes and uteri were examined in a blinded manner under light microscopy.

For fluorescence immunohistochemistry of cytochrome c, sections were incubated with antibody against cytochrome c diluted 1:200 in PBS containing 1% BSA and 3% fat-free milk. Immunodetection was accomplished using Cy3-conjugated anti-mouse IgG antibody. Sections were washed and mounted with Fluorescent Mounting Media (DAKO, Carpinteria, CA). Tissue images were then acquired with Cy3 staining using an Axiovert 200 microscope (Zeiss, Jena, Germany) equipped with a laser-scanning confocal imaging LSM 510 META system (Zeiss). The immunohistochemical findings illustrated are representative of those observed in numerous sections from multiple animals.

Visualization of apoptosis. Immunohistochemistry was used for detection of denatured DNA with a monoclonal antibody against single-strand DNA (ssDNA) to identify specific apoptotic cells, as described previously (17). All steps were processed according to the manufacturer’s instructions. Negative controls were processed in an identical manner in the same experiment except that anti-ssDNA primary antibody was omitted; incubation was performed with either normal mouse IgG (Santa Cruz Biotechnologies) or 100 U/ml S1 nuclease (Sigma-Aldrich) in acetate buffer after a rinse in ice-cold PBS or PBS with 1% nonfat dry milk; and the controls yielded no reaction product in parallel experiments (data not shown).

Measurement of estradiol and progesterone. Concentrations of total serum estradiol and P₄ were quantified by a radioimmunoassay (RIA) (10, 63) using a Delfia assay kit (PerkinElmer Life and Analytical Sciences, Wallac Oy, Turku, Finland) according to a protocol provided by the manufacturer. All samples were tested as duplicates together in one run. When different volumes of serum were assayed, concentrations were parallel to the standard curve. The sensitivity of the assay was typically better than 50 pmol/l, and the intra-assay coefficient of variation was 3.8–10% for measurement of estradiol, whereas the sensitivity of the assay was typically better than 0.8 nmol/l, and the intra-assay coefficient of variation was 3.3–7.3% for measurement of P₄.

Data analysis and statistics. Data from Western blot analysis are presented in the figures as means ± SE. Five mice for each group were used in one experiment except for the RU-486 treatment, where 15 mice were used for each group. To confirm reproducibility of results, three independent experiments for time course study of PR protein isoform expression in mouse fallopian tubes and uteri were performed and analyzed. All data were analyzed using the Analyze-It program (Analyze-It Software, Leeds, UK). Multiple comparisons between groups were made using one-way ANOVA with Tukey’s post hoc test. Statistical significance was set at P < 0.05.

	RESULTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Effect of gonadotropins on expression and regulation of PR protein isoforms in fallopian tubes and uteri. The Western blot analysis revealed the expression pattern of PR protein isoforms during gonadotropin treatment in the fallopian tubes comparable to that in the uteri. Treatment of immature mice with eCG resulted in the induction of proteins for total PR, PR-A, and PR-B in the fallopian tubes, commencing at 24 h and reaching maximum at 48 h. Furthermore, additional treatment with hCG reduced the total PR protein levels, reaching a minimum at 24 h (Fig. 2A). In the uterus, induction of total PR protein expression was seen, with a maximum at 48-h eCG treatment. However, the changes in protein levels of both PR isoforms in the uterus were different from those in the fallopian tubes. PR-B protein was induced earlier than PR-A protein after eCG treatment in the uterus. As with expression of PR proteins in the fallopian tubes, hCG treatment reduced the total PR, PR-A, and PR-B protein levels to baseline levels from 12 to 48 h (Fig. 3A). In all tissues, no induction of PR or its isoforms was seen at 48 h after injection of saline (data not shown).

View larger version (42K): [in this window] [in a new window]   Fig. 2. Regulation of progesterone receptor (PR) protein isoforms A and B after gonadotropin treatment in immature mouse fallopian tubes. A: fresh fallopian tubes from 26-day-old female mice treated with eCG and/or hCG with the indicated times were collected. Top: total protein was isolated and used for Western blot analysis as described in MATERIALS AND METHODS. Samples were run on a SDS-polyacrylamide gel and immunoblotted with antibody against PR. Bottom: densitometric analysis of levels of PR isoform protein expression in 33 independent experiments (n = 5–10 mice/group in each experiment). -Actin was used as internal control. Levels of PR isoform proteins were expressed as ratio of either PR-A or PR-B densitometric value to -actin. Data are expressed as arbitrary densitometric units (ADU) and represent means ± SE of 3 independent experiments in the graph. ** and ***Significantly different from untreated animals (time, 0 h), P < 0.01 and P < 0.001, respectively. B: standard curve showing linear relationship between immunoreactive band intensity and amount of whole tissue extract protein loaded for gonadotropin-treated mice. Equation of line of best fit was generated by least squares linear regression analysis. C: immunohistological localization of PR in sections of mouse fallopian tube from mice untreated (I-1 and I-2), or treated with eCG for 48 h (II-1 and II-2) and additionally treated with hCG for 12 h (III-1 and III-2), as described in MATERIALS AND METHODS. High specific immunostaining of PR was observed in nuclei of luminal epithelial (le), stromal (str), and smooth muscle cells (m) (48-h eCG treatment); is, isthmic segment; as, ampullary segment. IV: ovary from mouse treated with eCG + hCG served as a positive control tissue. Strong immunoreactivity was seen in granulosa cells. V: the same concentration of nonimmune IgG instead of the primary antibody was used as a negative control in ovarian section. Original magnification (I, II, and III)-1, x10; (I, II, and III)-2, IV, and V, x100. These experiments were repeated using 5 mice/group with similar results.

View larger version (43K): [in this window] [in a new window]   Fig. 3. Regulation of PR protein isoforms after gonadotropin treatment in immature mouse uteri. A: fresh uteri from 26-day-old female mice treated with eCG and/or hCG with the indicated times were collected. Top: total protein was isolated and used for Western blot analysis as described in MATERIALS AND METHODS. Samples were run on an SDS-polyacrylamide gel and immunoblotted with antibody against PR. Bottom: densitometric analysis of levels of PR isoform protein expression in 3 independent experiments (n = 5–10 mice/group in each experiment). -Actin was used as internal control. Levels of PR isoform proteins were expressed as ratio of PR-A or PR-B densitometric value to -actin. Data are expressed as ADU and represent means ± SE of 3 independent experiments in graph. ** and ***Significantly different from untreated animals (time, 0 h), P < 0.01 and P < 0.001, respectively. B: immunohistological localization of PR in sections of mouse uterus from mice untreated (I-1 to I-3), or treated with eCG for 48 h (II-1 to II-3) and additionally treated with hCG for 12 h (III-1 to III-3), as described in MATERIALS AND METHODS. High specific immunostaining of PR was observed in nuclei of endometrial stromal (str; 48-h eCG treatment) and endometrial luminal epithelial (le; additional 12-h hCG treatment) cells; myo, myometrium. Some stromal cells were positive for PR immunostaining in untreated mouse uterus. IV: strong immunoreactivity seen in human breast carcinoma served as a positive control tissue. V: the same concentration of nonimmune IgG instead of the primary antibody was used as negative control in human breast carcinoma section. All original magnification, x100. These experiments were repeated using 5 mice/group with similar results.

Decreased PR protein isoform abundance after P₄ treatment in fallopian tubes and uteri. The data presented above indicate that the decrease in PR protein isoforms after additional hCG treatment presumably arises from changes in endogenous P₄ levels in vivo. Therefore, we demonstrated the effect of an exogenous P₄ exposure on the levels of PR protein isoforms in the fallopian tubes and uteri determined by Western blot analysis. A time-dependent reduction of PR-A and PR-B protein expression was seen in the fallopian tubes and uteri from immature mice treated with a single injection of P₄ (Fig. 4, A and B). To analyze the molecular mechanisms involved in the effects of P₄, we also compared the expression of the cell cycle proteins (cyclin D₂ and p27) and PCNA (as an indicator of proliferation-related gene expression). In the fallopian tubes, the levels of expression of cyclin D₂, PCNA, and p27 remained relatively constant throughout the various time points during P₄ manipulations (Fig. 4A). However, all of these proteins displayed a decrease in uteri following P₄ treatment (Fig. 4B), suggesting that the P₄ effect on the expression of cyclin D₂, PCNA, and p27 proteins appears to be tissue specific.

View larger version (42K): [in this window] [in a new window]   Fig. 4. Time-dependent regulation of PR isoforms, cyclin D2, antiproliferating cell nuclear antigen (PCNA), and p27 protein expression after progesterone (P4) treatment in immature mouse fallopian tubes and uteri. Western blot analysis was performed on whole tissue protein preparation as described in MATERIALS AND METHODS. Fallopian tubes (A) and uteri (B) were obtained from immature mice treated for varying hours. Samples were run on an SDS-polyacrylamide gel and immunoblotted with antibody against PR, cyclin D2, PCNA, or p27. -Actin was used as control to monitor loading of protein. These experiments were performed twice using different tissue preparations with similar results.

View larger version (123K): [in this window] [in a new window]   Fig. 5. Regulation of PR protein isoforms after PR antagonist treatment in eCG/hCG-primed mouse fallopian tubes and uteri. Top: results from Western blot analysis performed on whole tissue protein preparation as described in MATERIALS AND METHODS. Fallopian tubes (A) and uteri (B) were obtained from immature mice treated with hCG for 24 h only (lanes 1–3) or combined with RU-486 (lanes 4–6) for 24 h followed by eCG for 48 h. Lanes 1–6 represented the same tissue from different mice. Samples were run on an SDS-polyacrylamide gel and immunoblotted with antibody against PR. -Actin was used as control to monitor loading of protein. Bottom: relative expression levels of PR, PR-A, and PR-B proteins after RU-486 treatment. Densitometic values were normalized to that of -actin protein expression. Data are from 3 mice/group of 3 independent performed experiments and are presented as means ± SE. **P < 0.01, ***P < 0.001 compared with mice not treated with RU-486. NC, no change. C: effect of RU-486 on serum estradiol and serum P4 in female mice. No. of mice per group is indicated. Trunk blood was obtained from heart puncture. Sera were subsequently collected after clotting and centrifugation. Serum estradiol and P4 concentrations were measured in duplicate aliquots of each sample. Values represent means ± SE. ***Significantly different from animals treated with eCG for 48 h, P < 0.001. D: immunohistological localization of PR in sections of mouse fallopian tube and uterus from eCG- and hCG-primed mice treated with oil, RU-486, and Org 31710, as described in MATERIALS AND METHODS. All original magnification, x100. These experiments were repeated using 5 mice/group with similar results.

View larger version (51K): [in this window] [in a new window]   Fig. 6. Effect of PR antagonists on apoptosis in eCG/hCG-primed mouse fallopian tubes. A: Western blot analysis was performed on whole tissue protein preparation as described in MATERIALS AND METHODS. Fallopian tubes were obtained from immature mice treated with hCG for 24 h only (lanes 1–3) or combined with RU-486 (lanes 4–6) for 24 h followed by eCG for 48 h. Lanes 1–6 represented the same tissue from different mice. Samples were run on an SDS-polyacrylamide gel and immunoblotted with antibody against apoptosis-inducing factor (AIF), caspase-3, caspase-9, or caspase-activated deoxyribonuclease (CAD). -Actin was used as a control to monitor the loading of protein. B: immunohistological localization of cytochrome c (a–c), activated caspase-3 (d–f and d1–f1), and DNA fragmentation (g–i) in sections of mouse fallopian tubes from eCG- and hCG-primed mice treated with control oil (a, d, d1, and g), RU-486 (b, e, e1, and h), and Org 31710 (c, f, f1, and i) as described in MATERIALS AND METHODS. Images d1–f1 are higher magnifications of d–f. Original magnification: a–c, x63; d–f, x10; d1–f1 and g–i, x100. These experiments were repeated using 5 mice/group with similar results.

View larger version (58K): [in this window] [in a new window]   Fig. 7. Effect of PR antagonists on apoptosis in eCG/hCG-primed mouse uteri. A: Western blot analysis was performed on whole tissue protein preparation as described in MATERIALS AND METHODS. Uteri were obtained from immature mice treated with hCG for 24 h only (lanes 1–3) or combined with RU-486 for 24 h (lanes 4–6) followed by eCG for 48 h. Lanes 1–6 represent the same tissue from different mice. Samples were run on an SDS-polyacrylamide gel and immunoblotted with antibody against AIF, caspase-3, caspase-9, or CAD. -Actin was used as control to monitor loading of protein. B: immunohistological localization of cytochrome c (a–c), activated caspase-3 (d–f), and DNA fragmentation (g–i) in sections of mouse uteri from eCG- and hCG-primed mice treated with control oil (a, d, g, and j), RU-486 (b, e, h, and k), and Org 31710 (c, f, I, and l) as described in MATERIALS AND METHODS. Original magnification: a–c, x63; d–f, x10; g–l, x100. ge, Endometrial glandular epithelia. These experiments were repeated using 5 mice/group with similar results.

View larger version (42K): [in this window] [in a new window]   Fig. 8. Effect of PR antagonists on expression of epidermal growth factor (EGF) and EGF receptor (EGFr) proteins in eCG/hCG-primed mouse fallopian tubes and uteri. Top: Western blot analysis performed on whole tissue protein preparation as described in MATERIALS AND METHODS. Fallopian tubes (A) and uteri (B) were obtained from immature mice treated with eCG for 48 h (lanes 1–3), additional hCG for 24 h (lanes 4–6), or combined hCG with RU-486 for 24 h (lanes 6–9) followed by 48-h eCG. Lanes 1–9 represented the same tissue from different mice. Samples were run on an SDS-polyacrylamide gel and immunoblotted with antibody against EGF or EGFr. -Actin was used as control to monitor loading of protein. Bottom: relative expression levels of EGF and EGFr proteins after RU-486 treatment. Densitometic values were normalized to that of -actin protein expression. Data are from 3 mice/group and are presented as means ± SE. **P < 0.01, ***P < 0.001 compared with 48-h eCG.

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

The time course of PR protein expression in response to gonadotropin stimulation in the fallopian tubes and uteri is different from that in ovary in mice. The current results showed that eCG treatment resulted in an increase in PR protein levels in the fallopian tubes and uteri that appears to be a response to the presence of high levels of serum and ovarian estradiol concentrations in vivo (Fig. 1B and Ljungström K, Weijdegård B, and Shaw R, unpublished data). There is evidence that estrogen peaked from ovulatory follicles induces PR mRNA and protein in rat uterus in vivo (27), and in most target cells in vitro (20) the present results suggest that increase in the expression of PR protein isoforms in the fallopian tubes and uteri may be important for the control of tubular and uterine responsiveness to estrogens in the early events of reproduction (e.g., cell proliferation for tissue development). This effect was attenuated by subsequent treatment with hCG, suggesting that the increase in endogenous P₄ levels (Fig. 1B) inhibits PR protein expression in both tissues. This hypothesis is strongly supported by the finding that exogenous P₄ treatment led to a time-dependent decrease in PR protein expression. After ligand binding, PR protein is rapidly and extensively downregulated in vitro (32). The mechanism for P₄-dependent reduction of PR protein levels has been demonstrated to be due to an alteration in ubiquitination in chicken fallopian tubes in vivo (67). Our results for regulation of PR protein expression are consistent with other reports, which demonstrate cyclic changes in PR proteins in human (52, 53) and rat (18, 50) fallopian tubes as well as human (39, 43, 75) and rat (27) uteri. Together, our animal experiments suggest that the effect of ovarian steroid hormones on regulation of PR protein expression in the fallopian tubes is similar to that in the uteri. It should be pointed out that we observed differences in pattern and expression levels of PR protein isoforms in mouse fallopian tube compared with uterus after gonadotropin stimulus. Previously, work in this laboratory indicated that both PR protein isoforms are regulated by additional treatment with hCG in ovarian granulosa cells (62). The data presented herein demonstrated that regulation of PR-A protein expression always paralleled that of PR-B protein expression in the fallopian tube (Fig. 2A) as well as in ovarian granulosa cells (62). However, PR-B protein levels in uterus were increased and peaked earlier than PR-A protein levels after eCG treatment. Moreover, although the levels of PR-B protein were decreased, PR-A protein expression further increased in the uterus (Fig. 3A). This may reflect that PR-B is the principal ligand-dependent transcriptional activator of P₄-responsive genes, whereas PR-A acts as a dominant repressor mediated by PR-B when the two isoforms are coexpressed in vitro (8, 35). Together, our observations indicate that regulation of relative expression of PR isoforms seems to differ in a stage- and tissue-specific way in female reproductive tissues. The fallopian tube provides an appropriate environment for fertilization and early embryo development (2). P₄ has been reported to decrease epithelial cilia beat frequency in human fallopian tubes (38). We showed that blocking P₄ action by the PR antagonist RU-486 increased the expression of PR protein isoforms in mouse fallopian tubes, in parallel with increased epithelial cilia beat frequency and tubal contractions (38) and accelerated tubal oocyte transport in rodents (65). Although P₄ via nuclear PR functions for remodeling of the uterus during the estrous cycle and pregnancy in mammals (20), our findings imply that nuclear PR protein isoforms may be involved in P₄-mediated physiological processes such as transportation of the oocyte, sperm, and embryo through the fallopian tubes.

Reproductive tissues exist in coordination and balance between the processes of proliferation, differentiation, cell cycle inhibition, and apoptosis (7, 42). In previous studies, P₄ inhibited apoptosis in rat endometrial epithelial cells, which expressed functional nuclear PR in vitro (51), whereas RU-486 induced apoptosis in rabbit uterine epithelium in vivo (57). Our laboratory has also demonstrated that PR antagonists induce apoptosis in mouse ovarian granulosa cells that were expressing nuclear PR in vivo (62) and in vitro (63). These demonstrations suggest that the suppression of apoptosis is likely mediated by existence of nuclear PR in P₄ target tissues. In mouse fallopian tube and uterus, the present study has detailed the molecular events involved in the apoptotic process affected by PR antagonists by using the same experimental models (62) for in vivo study and establish several new points. First, a considerable amount of experimental evidence has conclusively demonstrated a unique role for PR in the regulation of apoptosis by orchestrating mitochondria-mediated activation of caspase-9 and caspase-3, resulting in DNA fragmentation. Little evidence for regulation of AIF protein expression after RU-486 treatment relative to controls was observed, suggesting that AIF may not be involved in PR antagonist-induced caspase-3-dependent apoptosis in both tissues, in agreement with a previous observation showing apoptotic effect on regulation of AIF protein expression in uterus (68). Treatment with either RU-486 (4) or Org 31710, which is devoid of any action on glucocorticoid receptors (26), yielded similar results, indicating that a generalized mechanism for induction of apoptosis is mediated by blockade of PR protein function. Therefore, our results suggest a direct link between PR protein expression and alterations in the mitochondrial-mediated apoptotic pathway in both tissues. Understanding the molecular components of the PR-mediated apoptotic program in female reproductive tissues will be an important step toward the development of novel therapeutic regimens to control cell proliferation and apoptosis during abnormal pregnancy as well as more targeted approaches to female contraception. To the best of our knowledge, this is the first study that shows the molecular details of a linkage between the regulation of PR expression and development of apoptosis in the fallopian tube and uterus of any species. Second, both fallopian tube and uterus are relatively complex tissues composed of many different cell types. For the interpretation of the consequences of an increase in the levels of PR protein isoforms on induction of apoptosis by PR antagonists, the exact localization of apoptotic cells is of great importance. We showed that the induction of apoptosis (i.e., extensive cytochrome c release, caspase-3 activation, and increase in DNA fragmentation) by PR antagonists was predominant in the epithelium, whereas an increase in PR protein expression was observed in stromal cells of both tissues, suggesting that different cell types within a P₄ target tissue can have different cellular fates in response to PR antagonist treatment. Moreover, the inhibition of stromal cells undergoing apoptosis by increase in stromal PR could indicate negative regulation at one or more points within a mitochondrial-dependent apoptotic process. P₄-dependent inhibition of uterine cell proliferation has been demonstrated previously in intact (71) and PR knockout mice (8, 9, 44). Therefore, it is likely that both antiapoptotic (in this study) and antiproliferative effects (7, 42) are mediated by the functional PR protein expression in the fallopian tube and uterus during their development as a result of changing serum steroid levels. However, whether one isoform is more important than the other requires further study. Third, various growth factors, including EGF, play an essential role in regulating cell proliferation and differentiation and in suppressing the cell apoptotic pathway (13). A previous report has shown that treatment with EGF induces PR mRNA expression in adult mouse uterus (11), suggesting that PR-mediated inhibition of apoptosis may be mediated, at least in part, through growth factors. Therefore, we demonstrated that an increase in endogenous P₄ by hCG treatment resulted in an increase in EGF and EGF receptor protein expression, pointing to the pivotal link between P₄-PR and EGF signaling pathways in both tissues. Moreover, knowledge about the specific effects of PR on EGF and its receptor protein expression was supported by treatment with PR antagonists, in accord with previous reports that RU-486 reduces EGF receptor expression in rat uterine stromal cells in vivo (12, 49) and in human endometrial cells in vitro (76). Given the similarity of PR-mediated regulation of EGF and EGF receptor expression in mouse fallopian tube and uterus in the current study, we propose the existence of a possible interaction between PR and growth factor signaling in the regulation of cell proliferation and apoptosis in both tissues. On the other hand, we found that the protective effects of PRs were partial, as epithelial apoptosis still occurred despite an increase in PR protein expression after PR antagonist treatment in the fallopian tubes, suggesting the existence of an additional cell death mechanism that functions against the protective effect of PR. One interpretation of our result is that changes in EGF protein levels could be a main alternative for determination of epithelial cells committed to die in the fallopian tubes.

Our study also showed that in the fallopian tube the effect of P₄ treatment on reduction of PR protein isoforms could be dissociated from changes in p27, cyclin D₂, and PCNA protein expression, whereas in uterus this effect was accompanied by decreased levels of p27, cyclin D₂, and PCNA proteins, suggesting that PR may be one of the intracellular molecules involved in cell cycle progression in the uterus (71) and ovary (55) but not in the fallopian tube.

In summary, the present study highlights the developmental regulation of PR protein isoforms by gonadotropins in mouse fallopian tube and uterus in vivo. Our investigation suggests that both isoforms are mediated by estradiol stimulation and P₄ inhibition. Aside from the physiological functions of PR and its isoforms in the female reproductive tract, cellular events leading to cell death or survival are mediated by modulation of PR expression in different cell types in mouse fallopian tube and uterus. Moreover, the decision for cells to undergo apoptosis or to survive may be determined by a potential connection between PR and growth factor functions in these two tissues. Future investigation is likely to uncover the mechanisms by which actions of nuclear PR and EGF are coordinately integrated in vivo.

	GRANTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
This work was supported by Grants 10380 and 13550 from the Swedish Medical Research Council to H. Billig, the Assar Gabrielssons Forsknings Foundation, the Scientific Foundation of Eva och Oscar Ahrens, Wilhelm och Martina Lundgrens, Rudolf och Ellen Maria Mörnes, as well as Hjalmar Svensson and Adlerbertska Research Foundation to R. Shao.

	ACKNOWLEDGMENTS

 
We thank the Swegene Centre for Cellular Imaging at Göteborg University for the use of imaging equipment. A preliminary report was presented at the 38th Annual Meeting of the Society for the Study of Reproduction, Quebec City, Quebec, Canada, July 2005.

	FOOTNOTES

 

Address for reprint requests and other correspondence: R. Shao, Section of Endocrinology, Dept. of Physiology and Pharmacology, Sahlgrenska Academy, Göteborg University, SE-40530, Gothenburg, Sweden (e-mail: ruijin.shao{at}fysiologi.gu.se)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

	REFERENCES

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 

1. Amso NN, Crow J, and Shaw RW. Comparative immunohistochemical study of oestrogen and progesterone receptors in the fallopian tube and uterus at different stages of the menstrual cycle and the menopause. Hum Reprod 9: 1027–1037, 1994.[Abstract/Free Full Text]
2. Boatman DE. Responses of gametes to the oviductal environment. Hum Reprod 12: 133–149, 1997.[Abstract]
3. Brosens JJ, Tullet J, Varshochi R, and Lam EW. Steroid receptor action. Best Pract Res Clin Obstet Gynaecol 18: 265–283, 2004.[CrossRef][Medline]
4. Cadepond F, Ulmann A, and Baulieu EE. RU486 (mifepristone): mechanisms of action and clinical uses. Annu Rev Med 48: 129–156, 1997.[CrossRef][ISI][Medline]
5. Canning J, Takai Y, and Tilly JL. Evidence for genetic modifiers of ovarian follicular endowment and development from studies of five inbred mouse strains. Endocrinology 144: 9–12, 2003.[Abstract]
6. Christow A, Sun X, and Gemzell-Danielsson K. Effect of mifepristone and levonorgestrel on expression of steroid receptors in the human Fallopian tube. Mol Hum Reprod 8: 333–340, 2002.[Abstract/Free Full Text]
7. Clarke CL and Sutherland RL. Progestin regulation of cellular proliferation. Endocr Rev 11: 266–301, 1990.[ISI][Medline]
8. Conneely OM and Lydon JP. Progesterone receptors in reproduction: functional impact of the A and B isoforms. Steroids 65: 571–577, 2000.[CrossRef][ISI][Medline]
9. Conneely OM, Mulac-Jericevic B, and Lydon JP. Progesterone-dependent regulation of female reproductive activity by two distinct progesterone receptor isoforms. Steroids 68: 771–778, 2003.[CrossRef][ISI][Medline]
10. Couse JF, Yates MM, Deroo BJ, and Korach KS. Estrogen receptor-beta is critical to granulosa cell differentiation and the ovulatory response to gonadotropins. Endocrinology 146: 3247–3262, 2005.[Abstract/Free Full Text]
11. Curtis SW, Washburn T, Sewall C, DiAugustine R, Lindzey J, Couse JF, and Korach KS. Physiological coupling of growth factor and steroid receptor signaling pathways: estrogen receptor knockout mice lack estrogen-like response to epidermal growth factor. Proc Natl Acad Sci USA 93: 12626–12630, 1996.[Abstract/Free Full Text]
12. Dai D and Ogle TF. Progesterone regulation of epidermal growth factor receptor in rat decidua basalis during pregnancy. Biol Reprod 61: 326–332, 1999.[Abstract/Free Full Text]
13. Danielsen AJ and Maihle NJ. The EGF/ErbB receptor family and apoptosis. Growth Factors 20: 1–15, 2002.[CrossRef][ISI][Medline]
14. Donath J, Michna H, and Nishino Y. The antiovulatory effect of the antiprogestin onapristone could be related to down-regulation of intraovarian progesterone (receptors). J Steroid Biochem Mol Biol 62: 107–118, 1997.[CrossRef][ISI][Medline]
15. Earnshaw WC, Martins LM, and Kaufmann SH. Mammalian caspases: structure, activation, substrates, and functions during apoptosis. Annu Rev Biochem 68: 383–424, 1999.[CrossRef][ISI][Medline]
16. Enari M, Sakahira H, Yokoyama H, Okawa K, Iwamatsu A, and Nagata S. A caspase-activated DNase that degrades DNA during apoptosis, and its inhibitor ICAD. Nature 391: 43–50, 1998.[CrossRef][Medline]
17. Frankfurt OS, Robb JA, Sugarbaker EV, and Villa L. Monoclonal antibody to single-stranded DNA is a specific and sensitive cellular marker of apoptosis. Exp Cell Res 226: 387–397, 1996.[CrossRef][ISI][Medline]
18. Fuentealba B, Nieto M, and Croxatto HB. Estrogen and progesterone receptors in the oviduct during egg transport in cyclic and pregnant rats. Biol Reprod 39: 751–757, 1988.[Abstract]
19. Gava N, Clarke CL, Byth K, Arnett-Mansfield RL, and deFazio A. Expression of progesterone receptors A and B in the mouse ovary during the estrous cycle. Endocrinology 145: 3487–3494, 2004.[Abstract/Free Full Text]
20. Graham JD and Clarke CL. Physiological action of progesterone in target tissues. Endocr Rev 18: 502–519, 1997.[Abstract/Free Full Text]
21. Hakem R, Hakem A, Duncan GS, Henderson JT, Woo M, Soengas MS, Elia A, de la Pompa JL, Kagi D, Khoo W, Potter J, Yoshida R, Kaufman SA, Lowe SW, Penninger JM, and Mak TW. Differential requirement for caspase 9 in apoptotic pathways in vivo. Cell 94: 339–352, 1998.[CrossRef][ISI][Medline]
22. Iwai T, Nanbu Y, Iwai M, Taii S, Fujii S, and Mori T. Immunohistochemical localization of oestrogen receptors and progesterone receptors in the human ovary throughout the menstrual cycle. Virchows Arch A Pathol Anat Histopathol 417: 369–375, 1990.[CrossRef][ISI][Medline]
23. Jacobson MD, Weil M, and Raff MC. Programmed cell death in animal development. Cell 88: 347–354, 1997.[CrossRef][ISI][Medline]
24. Janicke RU, Sprengart ML, Wati MR, and Porter AG. Caspase-3 is required for DNA fragmentation and morphological changes associated with apoptosis. J Biol Chem 273: 9357–9360, 1998.[Abstract/Free Full Text]
25. Kastner P, Krust A, Turcotte B, Stropp U, Tora L, Gronemeyer H, and Chambon P. Two distinct estrogen-regulated promoters generate transcripts encoding the two functionally different human progesterone receptor forms A and B. EMBO J 9: 1603–1614, 1990.[ISI][Medline]
26. Kloosterboer HJ, Deckers GH, and Schoonen WG. Pharmacology of two new very selective antiprogestagens: Org 31710 and Org 31806. Hum Reprod 9, Suppl 1: 47–52, 1994.
27. Kraus WL and Katzenellenbogen BS. Regulation of progesterone receptor gene expression and growth in the rat uterus: modulation of estrogen actions by progesterone and sex steroid hormone antagonists. Endocrinology 132: 2371–2379, 1993.[Abstract]
28. Kraus WL, Montano MM, and Katzenellenbogen BS. Cloning of the rat progesterone receptor gene 5'-region and identification of two functionally distinct promoters. Mol Endocrinol 7: 1603–1616, 1993.[Abstract]
29. Kuida K, Haydar TF, Kuan CY, Gu Y, Taya C, Karasuyama H, Su MS, Rakic P, and Flavell RA. Reduced apoptosis and cytochrome c-mediated caspase activation in mice lacking caspase 9. Cell 94: 325–337, 1998.[CrossRef][ISI][Medline]
30. Kurita T, Lee KJ, Cooke PS, Lydon JP, and Cunha GR. Paracrine regulation of epithelial progesterone receptor and lactoferrin by progesterone in the mouse uterus. Biol Reprod 62: 831–838, 2000.[Abstract/Free Full Text]
31. Kurita T, Lee KJ, Cooke PS, Taylor JA, Lubahn DB, and Cunha GR. Paracrine regulation of epithelial progesterone receptor by estradiol in the mouse female reproductive tract. Biol Reprod 62: 821–830, 2000.[Abstract/Free Full Text]
32. Lange CA, Shen T, and Horwitz KB. Phosphorylation of human progesterone receptors at serine-294 by mitogen-activated protein kinase signals their degradation by the 26S proteasome. Proc Natl Acad Sci USA 97: 1032–1037, 2000.[Abstract/Free Full Text]
33. Leonhardt SA and Edwards DP. Mechanism of action of progesterone antagonists. Exp Biol Med (Maywood) 227: 969–980, 2002.[Abstract/Free Full Text]
34. Li P, Nijhawan D, Budihardjo I, Srinivasula SM, Ahmad M, Alnemri ES, and Wang X. Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade. Cell 91: 479–489, 1997.[CrossRef][ISI][Medline]
35. Li X and O’Malley BW. Unfolding the action of progesterone receptors. J Biol Chem 278: 39261–39264, 2003.[Free Full Text]
36. Loutradis D, Bletsa R, Aravantinos L, Kallianidis K, Michalas S, and Psychoyos A. Preovulatory effects of the progesterone antagonist mifepristone (RU486) in mice. Hum Reprod 6: 1238–1240, 1991.[Abstract/Free Full Text]
37. Lukola A and Punnonen R. Estrogen and progesterone receptors in human uterus and oviduct. J Endocrinol Invest 6: 179–183, 1983.[ISI][Medline]
38. Mahmood T, Saridogan E, Smutna S, Habib AM, and Djahanbakhch O. The effect of ovarian steroids on epithelial ciliary beat frequency in the human Fallopian tube. Hum Reprod 13: 2991–2994, 1998.[Abstract/Free Full Text]
39. Mangal RK, Wiehle RD, Poindexter AN III, and Weigel NL. Differential expression of uterine progesterone receptor forms A and B during the menstrual cycle. J Steroid Biochem Mol Biol 63: 195–202, 1997.[CrossRef][ISI][Medline]
40. Markstrom E, Svensson E, Shao R, Svanberg B, and Billig H. Survival factors regulating ovarian apoptosis—dependence on follicle differentiation. Reproduction 123: 23–30, 2002.[Abstract]
41. Masood S, Dee S, and Goldstein JD. Immunocytochemical analysis of progesterone receptors in breast cancer. Am J Clin Pathol 96: 59–63, 1991.[ISI]