Abstract
It has been established that estrogen can alter circadian rhythms in behavior and endocrine physiology in rodents. The uterus is a reproductive organ that is critically dependent on regulation by ovarian steroids. Here, we examined the expression of Per1 in different compartments of the uterus, and explored whether the ovarian steroids could regulate Per1 expression employing ovariectomized rat uterus. RT-PCR analysis showed that Per1 was cyclically expressed in the uterus. As revealed by in situ hybridization, the staining intensity of Per1 mRNA was stronger at ZT 8 than at ZT 0 in the uterine luminal epithelium (LE), stroma (S), and myometrium (M) compartments, but was not changed in the glandular epithelium (GE). Both in situ hybridization and immunofluorescence analyses revealed that estradiol (E2) administration induced high expression of Per1 in the LE, GE, and M, and less expression in the S compartment. Progesterone (P4) treatment resulted in an obvious enhancement of Per1 expression in the LE, GE, and S, but unchanged in the M compartment. Furthermore, the E2- and P4-activated Per1 expression was significantly repressed by their respective antagonists, ICI182 780 and RU486. These findings were further supported by RT-PCR analysis of Per1 expression in cultured uterine stromal cells. Collectively, the present data indicate that E2 and P4 might be involved in modification of circadian rhythm via direct regulation of the expression of clock genes.
Introduction
Circadian rhythm is generated by genetically determined biological clock, and is prominently entrained by cues from the 24-h light:darkness cycle (Dunlap 1999, Reppert & Weaver 2001). In mammals, the central clock is located in the suprachiasmatic nucleus of the hypothalamus (Reppert & Weaver 2001). Circadian rhythms also exist in a variety of peripheral tissues, including the heart, liver, kidney, lung, spleen, skeletal muscles (Yamamoto et al. 2004), and the uterus (Dolatshad et al. 2006). At the molecular level, the clock system is composed of interlocked transcriptional and translational feedback loops. The CLOCK and brain and muscle aryl hydrocarbon receptor nuclear translocator-like protein 1 (BMAL1), associated as heterodimers, bind to the E-box enhancer and positively drive the expression of Period genes (Per1, Per2, and Per3) and Cryptochrome genes (Cry1 and Cry2), whose proteins, in turn, form multimeric complexes and feed back to repress the transactivation by CLOCK/BMAL1 in the nucleus (Shearman et al. 2000, Ueda et al. 2005).
The peripheral clocks are synchronized by the central pacemaker through complex interaction of neural, humoral, and behavioral cues (Damiola et al. 2000, Schibler & Sassone-Corsi 2002). So far, the identification of resetting cues and their regulatory effects on clockwork are mostly based on molecular biological studies in cultured cell lines in vitro. A variety of factors have been unraveled having the potential to reset circadian rhythms in peripheral cells, such as cAMP analog, forskolin, phorbol-12-myristate-13-acetate, endothelin, prostaglandin estradiol (E2), fibroblast growth factor, calcium ionophores, retinoic acid, glucocorticoids, and parathyroid hormone (Akashi & Nishida 2000, Balsalobre et al. 2000a,b, Tsuchiya et al. 2005, Hinoi et al. 2006, Nakahata et al. 2006, Shirai et al. 2006). All these factors could induce an acute activation of Per1 expression, and thus synchronize or reset the circadian rhythms in cultured cells. For instance, glucocorticoids could efficiently activate Per1 expression and synchronizes circadian oscillations of clock genes in cultured cells, and could also phase shift the circadian rhythms in peripheral tissues in vivo (Balsalobre et al. 2000b).
Interestingly, it has long been established that ovarian steroids have significant modulatory effects on the circadian rhythms and sleep patterns. It has been demonstrated that estrogen administration alters the circadian rhythms in behavior and endocrine physiology in rodents (Morin et al. 1977, Thomas & Armstrong 1989). Recently, it was found that the phase of estrous cycle has a profound effect on the oscillation of PER2 protein within the brain (Perrin et al. 2006). It was further reported that chronic treatment with E2 drastically alters the circadian rhythms of Per1 and Per2 expression in the liver, kidney, and uterus (Nakamura et al. 2005). In contrast, progesterone (P4) was found to reduce the locomotor activity, but not the period of rhythms (Axelson et al. 1986, Labyak & Lee 1995). Despite the previous evidence, very little is currently known about the molecular mechanism underlying the sex steroidal effects on circadian clockwork. We hypothesize that the ovarian steroids probably modulate circadian rhythms through direct regulation of the expression of clock genes like Per1.
The uterus is one of the principal targets of estrogen and P4. It is composed of heterogeneous cell types including luminal and glandular epithelial cells, stromal cells, and muscle layers. In the adult uterus, E2 stimulates the proliferation of epithelial cells (Quarmby & Korach 1984), whereas P4 inhibits E2-induced hyperplasia of the epithelial compartments (Martin et al. 1973). In addition, P4 acts to regulate the proliferation and differentiation of uterine stromal cells (Rider 2002). Herein, we employed the rat uterus as the model system to testify our hypothesis. The results showed that both ovarian steroids could up-regulate the expression of Per1 gene, which sheds some light on how the steroids are involved in the modulation of circadian rhythms.
Materials and Methods
Reagents
E2, P4, mifepristone (RU486), and ICI182 780 were purchased from Sigma. Dulbecco’s modified Eagle’s medium (DMEM)/F12, fetal bovine serum (FBS), and 100 × penicillin–streptomycin solution (PS) were obtained from GIBCO. Collagenase (type I) was purchased from Invitrogen. pGEM-T Easy Vector, T7, and SP6 RNA Polymerases were purchased from Promega. DIG RNA labeling kit, anti-DIG antibody, nitroblue tetrazolium salt (NBT), and 5-bromo-4-chloro-3-indolyl phosphate (BCIP) were obtained from Roche Diagnostics. Anti-PER1 polyclonal antibody was purchased from Affinity Bioreagents (Golden, CO, USA), and Texas Red-conjugated goat F(ab′)2 anti-rabbit IgG (H + L) was from Leinco Technologies (St Louis, MO, USA). All the other chemicals used were of reagent grade and obtained from commercial sources.
Animals and treatments
Wistar rats were maintained under 12 h light:12 h darkness cycles (light between 0800 and 2000 h) with water and food available ad libitum. The female rats at 7 weeks of age were bilaterally ovariectomized (OVX) under ether anesthesia, and allowed for 1 week to recover from surgery. OVX rats were then subjected to hormonal treatments. In the first treatment, at ZT 0 (zeitgeber time, 0800 h), the rats were injected subcutaneously with a single dose of sesame oil (vehicle), E2 (50 μg/kg), or P4 (10 mg/kg). All hormones were given in 100 μl sesame oil. All OVX rats were killed at ZT 0 on the following day, and uteri were collected for histochemical examinations. All procedures were conducted in accordance with the Guideline for Animal Experiment in the Faculty of Medicine, Kyushu University and The Law (No. 105) and Notification (No. 6) of the Government.
Cell culture
Female rats at 3 months of age were mated with fertile males, and the day of finding a copulatory plug or sperm in the vaginal smear was designated as day 1 of pregnancy. Pregnant rats were killed on day 5 of gestation, and uteri were removed for isolation of uterine stromal cells (USCs). The uterine lumens, after flushing with PBS for several times, were filled with PBS containing 0.1% (w/v) collagenase and 1 × PS. The closed uterine horns were incubated in a shaking water bath at 37 °C for 1 h. The dissociated cell suspensions were pooled, and the uteri were longitudinally cut open and pipetted many times to release more USCs. The harvested cells were washed thrice with fresh DMEM/F12, and seeded onto 35 mm collagen-coated dishes at a density of 2 × 105 cells/dish with 2 ml of culture medium (phenol red-free DMEM/F12 supplemented with 10% charcoal-treated FBS and 1 × PS). The medium was changed 15 min after cell seeding to remove epithelial cells. Cells were cultured in a humidified atmosphere of 95% air and 5% CO2 at 37 °C. The USCs were positively immunostained for vimentin but negatively for cytokeratin. The staining result revealed a purity of ~95%.
Total RNA extraction and RT-PCR
Female rats at 9–10 weeks of age at diestrus were killed at 4-h intervals over a daily cycle. Two uterine horns of each rat were collected, with one horn used for total RNA extraction and the other for in situ hybridization. In addition, cultured USCs were harvested at the indicated times and subjected to total RNA isolation. The RNA concentration was determined using 260/280 u.v. spectrophotometry (Pharmacia Biotech), and RNA integrity was checked by agarose gel electrophoresis. One microgram of total RNA was utilized for cDNA synthesis in 20 μl mixture using Moloney–murine leukemia virus (M-MLV) reverse transcriptase (Amersham Biosciences) and oligo-dT primer according to the manufacturer’s protocol. The expression levels of β-actin and Per1 were examined by PCR using the following primers: β-actin (NM_031144), forward: 5′-TTG CGC TCA GGA GGA GCA AT-3′ (445–464), reverse: 5′-ATC ATG TTT GAG ACC TTC AA-3′ (1069–1088); Per1 (XM_340822), forward: 5′-AGTACG CGC TGG CCT GTG TCA A-3′ (689–710), reverse: 5′-GGC GCT TCATAA CCG GAG TGG AT-3′ (1222–1244). PCR was performed in a 10 μl mixture containing 1 × PCR buffer, 0.25 U AmpiTaq-Gold enzyme (Applied Biosystems, Tokyo, Japan), 0.2 mM of each dNTP, 1.5 mM MgCl2, 0.2 μM of each primer, and 1 μl of 1:5 diluted cDNA as a template. Semiquantitative RT-PCR was performed to measure the expression levels of Per1 and β-actin (internal control). The following cycling conditions were applied after an initial denaturation step (95 °C for 10 min), denaturation at 95 °C for 50 s, annealing at 50 °C for 50 s, and extension at 72 °C for 50 s. The 30 cycles of PCR for both Per1 and β-actin were currently used within the linear range of amplification based on the amplification curves. The PCR products were subjected to 2% agarose gel electrophoresis. The bands, stained by ethidium bromide, were visualized by u.v. fluorescence. Densitometric intensity was quantified by Scion Image (NIH software).
In situ hybridization
A 556 bp (689–1244) fragment of PCR product of Per1 (cDNA from USCs) was purified from agarose gel and cloned into pGEM-T Easy Vector. Sense and antisense probes were transcribed in vitro from linearized plasmids containing T7 or SP6 polymerase sites using a DIG labeling kit. Uteri collected from normal and OVX rats were cut into 6–8 mm pieces, and embedded in OCT compound for immediate freezing on a bed over liquid nitrogen. The tissues were cryosectioned at 8 μm thickness, and mounted on silane-coated slides. After fixation (4% paraformaldehyde, pH 7.4) and acetylation (0.25% (v/v) acetic anhydride in 0.1 M triethanolamine (pH 8.0)), each tissue was covered with 200 μl hybridization buffer and prehybridized in a humidified chamber at 42 °C for 2 h. The hybridization buffer consists of 4 × (SSC), 50% (v/v) formamide, 100 mg/ml dextran sulfate, 1 × Denhardt’s solution, 10 mg/ml salmon sperm DNA, and 0.5 mg/ml tRNA. Hybridization was carried out in the hybridization buffer containing 1 μg/ml denatured (80 °C for 10 min) antisense or sense RNA probe at 42 °C for 16 h, with the buffer covered with parafilm protecting from evaporation. After hybridization, the sections were first rinsed in 5 × SSC, and then washed twice in 50% (v/v) formamide/2 × SSC at 42 °C for 30 min, 2 × SSC and 0.2 × SSC at 42 °C each for 30 min. After washing with buffer 1 (0.1 M Tris (pH 7.6) and 0.15 M NaCl) for 10 min at room temperature (RT), the slides were incubated in buffer 2 (1.5% (w/v) blocking reagent in buffer 1) at RT for 2 h. Immunological examination of the hybridized probe was performed in a humidified chamber with anti-DIG antibody conjugated with alkaline phosphatase (1:500 dilution in buffer 2) overnight at 4 °C. Slides were washed twice in buffer 1 for 15 min at RT and equilibrated in buffer 3 (0.1 M Tris (pH 9.5), 0.15 M NaCl, and 50 mM MgCl2) for 5 min. For color development, slides were incubated in a NBT/BCIP solution at RT for 2–4 h. The reaction was stopped with Tris–EDTA (0.1 M Tris–HCl, 0.01 M EDTA (pH 8.0)) for 15 min, and the slides were cover slipped for signal visualization.
Immunofluorescence analyses
Uteri were collected from vehicle or hormone-treated OVX rats and frozen as outlined previously. Air-dried tissue sections of 8 μm thickness were fixed for 5 min in acetone at −20 °C. Non-specific binding was blocked using 2% (v/v) goat serum in PBS (blocking buffer) for 30 min. Sections were incubated for 12–18 h at 4 °C with rabbit anti-mPER1 polyclonal antibody diluted in blocking buffer (1:500). Rabbit serum was substituted for the primary antibody as negative control. After washing with PBS, they were incubated with the second antibody for 1 h at RT. Sections were subsequently washed in PBS and mounted with Mount-Quick Aqueous (Daido Sangyo, Japan). Immunostaining was detected under a fluorescence microscope (Nikon, Japan).
Statistical analyses
The quantification of in situ hybridization and immunofluorescent staining intensity were performed as previously reported (Han et al. 2007). A box was drawn through each compartment in a random orientation, and the pixel value of each box was measured by NIH Scion Image. A mean value was obtained from the determination of three boxes for each slide section. The values in all figures were shown as means ± s.d. of three independent sections from different samples. The non-specific staining intensity with sense RNA probe was used to normalize Per1 mRNA staining signal and the fluorescence intensity after incubation with normal rabbit serum to standardize PER1 protein intensity. The RT-PCR results were similarly expressed as means ± s.d. of three independent experiments on different RNA sets. The differences between them were evaluated using Student’s t-test following one-way ANOVA. A P value < 0.05 was considered significant.
Results
Per1 expression in different compartments of the diestrous uterus
The RT-PCR analysis showed rhythmic changes of Per1 mRNA level in the rat uteri at the diestrous stage (Fig. 1). The trough and peak of Per1 expression appeared around ZT 0 and ZT 8 respectively. The transcript level of Per1 was then analyzed by in situ hybridization in different compartments of the uterus at ZT 0 and ZT 8. The expression of Per1 mRNA can be readily detected in both the endometrium and the myometrium (M) at ZT 0 (Fig. 2A–C) and ZT 8 (Fig. 2D–F). Strong signals were observed in the luminal epithelium (LE) and glandular epithelium (GE), as compared with that in the stroma (S). The staining intensity of Per1 mRNA was approximately twofold higher at ZT 8 (Fig. 2D, E and H) than at ZT 0 (Fig. 2A, B and H) in the S, LE, and M compartments. However, no obvious change of Per1 expression was found in the GE compartment (Fig. 2C, F, and H). No specific products were observed when using the sense riboprobes (Fig. 2G).
Effects of E2 and P4 on Per1 expression in different uterine compartments
To examine the effects of E2 and P4 on the expression of Per1 in the uterus, OVX rats were treated with steroid hormones at ZT 0 and uteri were collected 24 h later. In situ hybridization analysis showed that the E2-primed uteri exhibited much stronger staining in the LE (ca 3.1-fold), GE (ca 2.8-fold), and M (ca 3.2-fold), while less intense signal in the S (ca 2.0-fold) compartment (Fig. 3C, D, and G), as compared with the control uteri (Fig. 3A, B and G). P4 administration resulted in intense staining in the S (ca 3.3-fold), LE (ca 2.4-fold), and GE (ca 2.1-fold) compartments, but not in the M (ca 1.2-fold; Fig. 3E–G). Immunofluorescent staining for PER1 protein was also examined in the oil-, E2-, and P4-administrated uteri. In the control uteri, weak staining of PER1 was observed in the S compartment, and relatively stronger staining in the LE and GE (Fig. 4A and B). With E2 treatment (Fig. 4C, D, and G), PER1 protein was more intensively expressed in the S (ca 2.2-fold), LE (ca 3.2-fold), and GE (ca 2.1-fold) as compared with the counterparts of the control uteri. Treatment with P4 also resulted in stronger immunostaining of PER1 in the S (ca 2.6-fold), LE (ca 2.9-fold), and GE (ca 2.9-fold) compartments (Fig. 4E, F and G). No staining was observed in the negative control (data not shown). Taken together, immunohistochemical observations of Per1 expression in response to E2 or P4 were similar to the results revealed by in situ hybridization.
Repression of E2- and P4-regulated Per1 expression by respective antagonist treatments
The antagonists for estrogen receptor (ER) and progesterone receptor (PR) were utilized to evaluate whether the stimulatory effects of E2 and P4 on Per1 expression are mediated via their respective receptors. The antagonists were injected at ZT 0, and the steroids were given after 2 h. Pretreatment with ICI182 780 (ER antagonist) caused a significant decrease in the staining intensity of Per1 mRNA in all the examined compartments, the LE (ca 70%), GE (ca 60%), S (ca 60%), and M (ca 50%; Fig. 5C, D and G), as compared with the control administrated E2 alone (Fig. 5A, B and G). On the other hand, the uteri treated with P4 alone exhibited strong staining of Per1 in the LE, GE, and S compartments (Fig. 5E), whereas RU486 (PR antagonist) pretreatment remarkably inhibited Per1 expression in the GE and S by ~60 and 70% respectively (Fig. 5F and H). The intensity of Per1 staining in the LE also showed ~50% reduction (Fig. 5F and H). These findings suggest that ER and PR might mediate E2- and P4-regulated Per1 expression in the uterus respectively.
E2 and P4 regulation of Per1 expression in cultured USCs
The regulation of Per1 expression by E2 and P4 was confirmed in cultured USCs. The USCs reached confluence after 3-day culture, was further cultured in the presence of vehicle (ethanol), E2 (10 nM), or P4 (100 nM), and collected for RT-PCR analysis at 6 and 24 h. E2 significantly though slightly activated Per1 expression at 6 h, but not at 24 h (Fig. 6A). In contrast, P4 treatment resulted in enhanced Per1 transcript level at both 6 and 24 h (Fig. 6B). Furthermore, we examined whether RU486 could antagonize the P4-activated Per1 expression in cultured USCs. Cells were treated with vehicle or 10 μM RU486 for 2 h, and then cultured in the presence of P4 or P4 + RU486. After 6 h, the cells were harvested for analysis of Per1 mRNA level. The expression of Per1 was significantly inhibited in the RU486-treated cells relative to the control cells exposed to P4 alone (Fig. 6C).
Discussion
The uterus, a highly dynamic tissue, undergoes profound remodeling, implicating cyclic proliferation, differentiation, and differential gene expression dependent on the levels of circulating sex steroid hormones. Recent studies have demonstrated that circadian clock genes are rhythmically expressed in the uterus (Nakamura et al. 2005, Dolatshad et al. 2006) as well as other peripheral tissues (Yamamoto et al. 2004). However, their findings were based on the mRNA analyses prepared from the entire uterine tissue using RT-PCR or northern blotting methods. In the present study, we investigated the expression of Per1, a core clock gene, in different compartments of the uterus with in situ hybridization. As expected, our results showed that the staining intensity of Per1 mRNA was obviously higher at ZT 8 than at ZT 0 in the LE, S, and M compartments of the diestrous uterus, which was consistent with the rhythm of Per1 as revealed by RT-PCR. Surprisingly, the staining signal was not apparently changed at ZT 0 and ZT 8 in the GE compartment, suggesting the lack of normal clockwork in control of uterine glandular physiology. It has been previously suggested that cellular differentiation process is responsible for the absence of circadian rhythms in the testis and thymus tissues (Alvarez et al. 2003, Alvarez & Sehgal 2005). Whether or not cellular differentiation contributes to the constant expression of Per1 gene in glandular epithelial cells remains to be explored.
More importantly, the present study demonstrated that the expression of Per1 increased in the rat uterus after treatment with E2 and P4. It is well known that, at ZT 0, the transcription of Per1 is at the lowest level in almost all peripheral tissues and is less likely activated by the CLOCK/BMAL1 heterodimers (Yamamoto et al. 2004). Hence, all the uteri were collected at ZT 0 to evaluate the steroidal effects on Per1 expression. The analysis by in situ hybridization indicates that E2 administration causes higher expression of Per1 in the uterine LE, GE, and M, but to a lesser extent in the S compartment. The promotion of Per1 expression by E2 was further confirmed by immunohistochemical studies. In addition, the finding that ICI182 780 suppressed E2-activated Per1 expression indicates that the regulation of Per1 expression is at least partly through the ER-mediated pathway. Therefore, the differences in Per1 expression among the compartments might be due to differential expression of ER. Actually, expression of ER mRNA or protein is higher in the LE, GE, and M compartments than in the S compartment at 24 h after E2 administration (Nephew et al. 2000). This pattern of ER distribution, to some extent, supports our findings of Per1 expression profile in the uterus in response to E2 treatment. RT-PCR analysis revealed that Per1 expression was slightly augmented by E2 in cultured USCs at 6 h, but not at 24 h. The different observations between in vitro and in vivo may attribute to distinct environmental context. On the other hand, P4 efficiently activated the expression of Per1 in the entire endometrium via the mediation of PR. The PR-mediated up-regulation of Per1 expression by P4 was further supported by the findings in cultured USCs in vitro. However, no obvious enhancement of Per1 transcription by P4 was observed in the M compartment, although the myometrial cells of OVX rat uterus are also positive to PR (Tibbetts et al. 1998). The distinct responsiveness of Per1 expression to P4 in the endometrium and myometrium (M) may also be due to their differential microenvironmental context.
E2 and P4 could bind to their respective ER and PR, which, in turn, bind to specific response elements (ERE and PRE) in the promoters of target genes, thereby regulating gene transcription (Bagchi et al. 1988, Anderson & Gorski 2000, Edwards 2005). Therefore, our present findings suggest that the activated expression of Per1 by ovarian steroids might result from the direct binding of ER or PR to some potential element(s) in Per1 promoter. Indeed, as stated in the previous report (Nakamura et al. 2005), although there is no consensus ERE (5′-GGTCANNNTGACC-3′), some ERE-half sites (5′-GGTCA-3′ or 5′-TGACC-3′) are located in the 5′ flanking region of Per1 promoter (GenBank accession number, AB030818). Additionally, a PRE-half element (5′-TGTTACT-3′) is located in the first intron of Per1 gene within + 868/+ 874 (+ 1, the transcription start site). It remains to be elucidated whether ER and PR promote Per1 transcription by direct binding to their respective putative element(s) in Per1 promoter, or via yet unknown indirect pathways.
The present finding of the pattern of Per1 expression under the regulation by the steroids is attractive. As is well known, the E-box element in Per1 promoter is essential for binding by CLOCK/BMAL1 and generation of circadian rhythms (Darlington et al. 1998, Travnickova-Bendova et al. 2002). In addition, other two important up-regulatory elements, the cAMP response element (CRE) and glucocorticoid response element (GRE), have been clearly defined by in vitro studies. In response to various resetting cues (Balsalobre et al. 2000b, Tsuchiya et al. 2005, Yamamoto et al. 2005, Hinoi et al. 2006), the CRE-binding protein (CREB) or glucocorticoid receptor (GR) was activated, which, in turn, transactivates the transcription of Per1 in the nucleus by binding to the CRE or GRE respectively (Travnickova-Bendova et al. 2002, Yamamoto et al. 2005, He et al. 2007). It is important to note that both CREB and GR act to induce an acute and robust enhancement of Per1 transcription with a peak at 1 h. In contrast, estrogen and P4 tend to induce a chronic up-regulation of Per1 expression. This finding suggests that there may exist other pathway(s) involved in regulating clock gene expression and resetting or modulating the circadian rhythms in vivo by the humoral cues. In vitro studies using clock gene transgenic cells (He et al. 2007,) will help to provide direct evidence of whether E2 or P4 could serve to reset the circadian clockwork.
In conclusion, the present study demonstrates the existence of circadian rhythms in the uterine LE, S, and M compartments, but not obviously in the GE. The finding of up-regulation of Per1 expression by E2 may contribute to understand how ER functions to phase shift the circadian rhythms of clock gene expression and to alter the behavior and endocrine rhythms (Morin et al. 1977, Thomas & Armstrong 1989, Nakamura et al. 2005, Perrin et al. 2006). We also find that P4 efficiently activates the expression of Per1 gene in the uterus, which indicates that P4 might also have profound effects on circadian clockwork. However, although Per1 gene is subjected to the regulation by both E2 and P4, it is less likely that Per1 directly exerts physiological functions but rather indirectly regulates uterine physiology via modulation of circadian clockwork. Furthermore, comprehensive studies on clock gene expression in P4-responsive tissues are required to further understand the potential actions of P4 on the modulation of circadian rhythms.
Cyclic expression of Per1 mRNA in the uterus. Adult female rats at diestrus were killed at 4-h intervals over a daily cycle (ZT 0, 0800 h). Total RNA was extracted and subjected to RT-PCR analysis of Per1 transcript levels, which were normalized to β-actin values (the maximal, set as 100). Data are presented as mean ± of three independent experiments.
Citation: Journal of Endocrinology 194, 3; 10.1677/JOE-07-0172
Representative photomicrographs of in situ hybridization analyses for Per1 mRNA in diestrous uteri. The uterine samples were obtained at ZT 0 (A–C) and ZT 8 (D–F). (C) and (F) show the insert sections of (A) and (D) at higher magnification respectively. Dark shades show the staining of Per1 mRNA. No signal was obtained in the uteri stained with sense probe (G). (H) shows the fold changes of staining intensity of Per1 in different uterine compartments at ZT 8 relative to those at ZT 0 (set at 1). Data are presented as mean ± s.d. of three independent sections. *Statistical significance (P < 0.05), ZT 8 versus ZT 0. Samples from three individual rats were analyzed for each group, and at least three sections were examined by histochemistry. S, stroma; LE, luminal epithelium; GE, glandular epithelium; M, myometrium. Scale bars, 100 μm.
Citation: Journal of Endocrinology 194, 3; 10.1677/JOE-07-0172
Representative photomicrographs of in situ hybridization analyses for Per1 mRNA in the uteri treated with E2 or P4. At ZT 0, the OVX rats were injected subcutaneously with 100 μl oil-containing vehicle (A and B), 50 μg/kg E2 (C and D), or 10 mg/kg P4 (E and F), and uteri were removed after 24 h. (G) shows the fold changes of Per1 staining intensity in different compartments of the E2- and P4-administrated uteri relative to the control (set at 1). Data are presented as mean ± s.d. of three independent sections. *Statistical significance (P < 0.05) compared with the control. Samples from three individual rats were analyzed for each group, and at least three sections were examined by histochemistry. S, stroma; LE, luminal epithelium; GE, glandular epithelium; M, myometrium. Scale bars, 100 μm.
Citation: Journal of Endocrinology 194, 3; 10.1677/JOE-07-0172
Representative photomicrographs of immunofluorescent analyses for PER1 protein in the uteri treated with E2 or P4. At ZT 0, the OVX rats were injected subcutaneously with 100 μl oil-containing vehicle (A and B), 50 μg/kg E2 (C and D), or 10 mg/kg P4 (E and F), and uteri were removed after 24 h. (G) shows the fold changes of Per1 staining intensity in different compartments of the E2- and P4-administrated uteri relative to the control (set at 1). Data are presented as mean ± s.d. of three independent sections. *Statistical significance (P < 0.05) compared with the control. Samples from three individual rats were analyzed for each group, and at least three sections were examined by histochemistry. S, stroma; LE, luminal epithelium; GE, glandular epithelium. Scale bars, 100 μm.
Citation: Journal of Endocrinology 194, 3; 10.1677/JOE-07-0172
Representative photomicrographs of in situ hybridization analyses for Per1 mRNA in the uteri treated with E2, ICI182 780 + E2, P4, or RU486 + P4. At ZT 0, the OVX rats were injected with 100 μl oil-containing vehicle, 3 mg/kg ICI182 780 (ER antagonist), or 30 mg/kg RU486 (PR antagonist), after 2 h, followed by injection with 50 μg/kg E2 or 10 mg/kg P4. The uteri were collected at ZT 0 the following day. (A) and (B) E2, (C) and (D) ICI182 780 + E2, (E) P4, and (F) RU486 + P4. (G) and (H) show the percentage repressions of Per1 staining intensity in different compartments of antagonist-pretreated uteri relative to the control (set at 100%). Data are presented as mean ± s.d. of three independent sections. *Statistical significance (P < 0.05) compared with the control. Samples from three individual rats were analyzed for each group, and at least three sections were examined by histochemistry. S, stroma; LE, luminal epithelium; GE, glandular epithelium; M, Myometrium. Scale bars, 100 μm.
Citation: Journal of Endocrinology 194, 3; 10.1677/JOE-07-0172
Stimulatory effects of E2 and P4 on Per1 expression in cultured USCs. The 3-day cultured USCs were further cultured in fresh medium containing vehicle (A–C), 10 nM E2 (A), 100 nM P4 (B and C), or 100 nM P4 + 10 μM RU486 (RU) (C). The cells were harvested at the indicated times for RT-PCR analysis of Per1 expression. Per1 transcript levels were normalized to β-actin values and expressed as arbitrary units relative to the vehicle control (set as 1). Data are presented as mean ± s.d. of three independent experiments. Different letters are significant (P < 0.05).
Citation: Journal of Endocrinology 194, 3; 10.1677/JOE-07-0172
This research was supported in part by a Grant-in-Aid for Scientific Research from the Japan Society for the Promotion of Sciences (JSPS; 16380200 and 17658131; to M-A H). The authors declare that there is no conflict of interest that would prejudice the impartiality of this scientific work.
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