Abstract
In mares, prostaglandin F2α (PGF2α) secreted from the endometrium is a major luteolysin. Some domestic animals have an auto-amplification system in which PGF2α can stimulate its own production. Here, we investigated whether this is also the case in mares. In an in vivo study, mares at the mid-luteal phase (days 6–8 of estrous cycle) were injected i.m. with cloprostenol (250 µg) and blood samples were collected at fixed intervals until 72 h after treatment. Progesterone (P4) concentrations started decreasing 45 min after the injection and continued to decrease up to 24 h (P < 0.05). In turn, 13,14-dihydro-15-keto-PGF2α (PGFM) metabolite started to increase 4h after an injection and continued to increase up to 72 h (P < 0.05). PGF receptor (PTGFR) mRNA expression in the endometrium was significantly higher in the late luteal phase than in the early and regressed luteal phases (P < 0.05). In vitro, PGF2α significantly stimulated (P < 0.05) PGF2α production by endometrial tissues and endometrial epithelial and stromal cells and significantly increased (P < 0.05) the mRNA expression of prostaglandin-endoperoxide synthase-2 (PTGS2), an enzyme involved in PGF2α synthesis in endometrial cell. These findings strongly suggest the existence of an endometrial PGF2α auto-amplification system in mares.
Introduction
Prostaglandin F2α (PGF2α) released from the uterus is a potent luteolytic agent in many domestic animals (Knickerbocker et al. 1988, Ginther 1992a, McCracken et al. 1999). Prostaglandin F2α auto-amplification systems have been observed in the corpora lutea (CL) of pigs, sheep, and cattle (Guthrie et al. 1979, Rexroad & Guthrie 1979, Tsai & Wiltbank 1997, Diaz et al. 2000, Kumagai et al. 2014) and in the endometrium of sheep and cattle (Wade & Lewis 1996, Kotwica et al. 1999, Duong et al. 2012); however, it is unknown whether mares have a similar system.
Prostaglandin F2α is synthesized through a series of enzymatic reactions, known as the arachidonic acid (AA) cascade. Arachidonic acid is liberated from phospholipids by phospholipase A2 (PLA2). Prostaglandin-endoperoxide synthase 1 (PTGS1) and PTGS2 convert AA to prostaglandin H2 (PGH2) (Smith et al. 1996), and then PGH2 is converted to PGF2α by prostaglandin F synthase (PGFS). Prostaglandin F2α is metabolized by 15-hydroxyprostaglandin dehydrogenase (HPGD) into the PGF metabolite – 13,14-dihydro-15-keto-PGF2α (PGFM).
Mares are more sensitive to the luteolytic effect of PGF2α than other domestic animals, although the reason is unknown. The minimal effective dose of a single injection (8 μg/kg body weight) of a PGF2α analogue in mares (Ginther 1992a) is five times less than that in cows (40 μg/kg body weight) (Ginther & Beg 2012) and 18 times less than that in sheep (144 μg/kg body weight) (Douglas & Ginther 1973). Mares are highly sensitive to PGF2α because the affinity of the luteal cell membrane for PGF2α is approximately ten times greater in horse than in cattle (Kimball & Wyngarden 1977). Another possibility is that the plasma clearance of PGF takes several times longer in mares than in heifers (Shrestha et al. 2012). Thus, in mares, target tissues are exposed to higher concentrations of PGF for longer periods (Shrestha et al. 2012).
As PGF2α plays an important role in the regression of the CL, it is of interest to know how its production is regulated. In vitro studies have shown that equine endometrial PGF2α production is stimulated by many factors such as oxytocin (OT; Watson et al. 1992, Szóstek et al. 2012), tumor necrosis factor-α (TNFα; Szóstek et al. 2014a), and ovarian steroids (Szóstek et al. 2014b); however in mares, it is not known whether PGF2α affects its own production. Our hypothesis is that endometrial PGF2α auto-amplification system is present and contributes to the induction of luteolysis in mares. To test this hypothesis, we examined the effect of PGF2α on PGF2α production both in vivo and in vitro.
Materials and methods
Animals and endometrial tissue collection
In the in vivo study (Experiment 1), we used non-lactating, cyclic Thoroughbred mares (n = 6, 4–11 years of age) from Iburi, Hokkaido (~N42°), Japan. The in vivo study was conducted in February. The mares were examined daily with a B-mode ultrasonographic scanner (ALOKA SSD-620; Hitachi Medical Corporation, Tokyo, Japan) and their ovaries were monitored. The phase of the estrous cycle was evaluated by ultrasonography, rectal palpation, and blood progesterone (P4) concentrations. The day of ovulation was defined as a day 0 of the estrous cycle. The mares were put out to pasture from 6 AM to 3 PM and housed under controlled artificial light. The light was off from 8 PM to 4 AM. The mares had free access to hay and water. All procedures for animal experiments were approved by the local animal care and use committee.
In the in vitro studies (Experiments 2–5), uteri (n = 25) were collected from randomly designated cyclic, non-pregnant mares at an abattoir in Kumamoto (~N32°), Japan, from April until the end of July in accordance with the protocols approved by the local institutional animal care and use committee. We used Anglo-Norman mares of various ages (2–11 years) and weighing approximately 600 kg. Mares possessing ovaries and uterus with a macroscopic abnormality, including anovulatory hemorrhagic follicles, were eliminated. The uteri were classified as being in the early, developing, mid, late, or regressed luteal phase by macroscopic observation of the CLs and follicles as described previously (Ginther 1992b, Ferreira-Dias et al. 2006, Kozai et al. 2014). Following determination of the phases, endometrial tissues ipsilateral to the CL or to the follicle > 35 mm (regressed luteal phase) in diameter were immediately frozen in liquid nitrogen and then stored at −80°C until being processed for RNA extraction. For tissue and cell culture, the uteri were submerged in ice-cold sterile physiological saline supplemented with 100 IU/mL of penicillin (611400D3051; Meiji Seika Pharma, Tokyo, Japan) and 100 μg/mL of streptomycin (6161400D1034; Meiji Seika Pharma) and transported to the laboratory on ice.
Tissue culture
Endometrial tissues obtained in the late luteal phase (n = 3) were used for tissue culture. Endometrial tissues were cultured as described previously (Lee et al. 2007). Briefly, endometrial tissues were washed three times in sterile saline solution containing penicillin (100 IU/mL) and streptomycin (100 μg/mL); cut into small pieces (40–50 mg) with a scalpel; washed another three times in Ca2+- and Mg2+-free Hank’s balanced salt solution (HBSS) supplemented with penicillin (100 IU/mL), streptomycin (100 μg/mL), and 0.1% (w/v) bovine serum albumin (BSA) (10735086001; Roche Diagnostics, Manheim, Germany); hanged with steel needles (8N01B; TOP, Tokyo, Japan); and placed into culture glass tubes (12 mm × 75 mm, 73500-13100; Kimble Chase Life Science and Research Products, Vineland, NJ, USA) containing 2 mL culture medium (DMEM/Ham’s F-12, 1:1 (v/v), 12400-024; Invitrogen, Carlsbad, CA, USA) supplemented with penicillin (100 IU/mL), streptomycin (100 μg/mL), and 0.1% (w/v) BSA under 5% CO2 in air.
Epithelial and stromal cell isolation and culture
Early luteal phase uteri were used for isolation of endometrial epithelial and stromal cells. In our research model, the cells obtained from the early luteal phase were used because characteristic feature of cells from the early luteal phase is their optimal viability and small morphological and physiological variability in the long-term cultures as was confirmed in cattle (Fortier et al. 1988, Thibodeaux et al.1991). The epithelial and stromal cells were isolated as described previously (Szóstek et al. 2012), with some modifications. A polyvinyl catheter was inserted into the uterine lumen from the side of the oviduct and the uterine horn was tied at one-third the distance from the uterotubal junction. The uterine lumen was washed three times with 10 mL of sterile Ca2+- and Mg2+-free HBSS supplemented with 100 IU/mL of penicillin (611400D3051; Meiji Seika Pharma, Tokyo, Japan), 100 μg/mL of streptomycin (6161400D1034; Meiji Seika Pharma), and 0.1% (w/v) BSA. Sterile HBSS (30–40 mL) containing 0.3% (w/v) trypsin (T9201; Sigma-Aldrich) was infused into the uterine lumen through the polyvinyl catheter. The uterine horn was immersed in sterile physiological saline and incubated at 38.0°C for 40 min. A suspension containing epithelial cells and undigested tissue fragments was collected into 50 mL tubes, centrifuged (4°C, 200 g, 10 min), and the supernatant was removed. The undigested tissue fragments were digested by stirring for 30 min in 50 mL of sterile HBSS containing antibiotics, 0.1% (w/v) BSA, 0.05% (w/v) collagenase (#CLS1; Worthington Biochemical, Lakewood, NJ, USA), and 0.005% (w/v) DNase I (DNP2; BBI Enzymes, Cardiff, UK). The cell suspension was filtered through metal meshes (100 µm and 80 μm) to remove the undigested tissue fragments. The filtrate was washed three times by centrifugation (4°C, 200 g, 10 min) with Dulbecco’s Modified Eagle’s Medium (DMEM) (D1152; Sigma-Aldrich) supplemented with antibiotics and 0.1% (w/v) BSA. The final pellet of the epithelial cells was resuspended in culture medium (DMEM/Ham’s F-12; 1:1 (v/v)) supplemented with 10% (v/v) bovine serum (16170-078; Gibco, Life Technologies, Carlsbad, CA, USA), 20 μg/mL of gentamicin, and 2 µg/mL of amphotericin B. The epithelial cells were counted using a hemocytometer. The viability of the epithelial cells was higher than 85% as assessed by the trypan blue exclusion test.
After the collection of the epithelial cells, the uterine horn was slit open with scissors to expose the endometrial surface. Endometrial strips were excised from the myometrium layer with a scalpel, washed once with sterile HBSS containing antibiotics, and cut into small pieces (1 mm3) with a scalpel. The minced tissues were digested by stirring for 30 min in 50 mL of sterile HBSS containing antibiotics, 0.1% (w/v) BSA, 0.05% (w/v) collagenase, and 0.005% (w/v) DNase I. The cell suspension was filtered through metal meshes (100 µm and 80 µm) to remove the undigested tissue fragments. The filtrate was washed by centrifugation (4°C, 200 g, 10 min) with Tris-buffered ammonium chloride to lyse the red blood cells, then washed three times by centrifugation (4°C, 200 g, 10 min) with DMEM supplemented with antibiotics, and 0.1% (w/v) BSA. The final pellet of the stromal cells was resuspended in culture medium, as described above, for the epithelial cells. The stromal cells were counted using a hemocytometer. The viability of the stromal cells was higher than 85% as assessed by the trypan blue exclusion test.
The dispersed epithelial and stromal cells were seeded separately at a density of 1 × 105 viable cells/mL in 24-well cluster dishes (662160; Greiner Bio-One, Frickenhausen, Germany) and cultured at 38.0°C in a humidified atmosphere of 5% CO2 in air. To purify the stromal cell population, the medium was changed 3 h after plating, by which time-selective attachment of stromal cells had occurred (Szóstek et al. 2012). Alternatively, when the epithelial cells attached, which occurred between 24 and 48 h after plating, the medium was changed. The medium for both types of cells was changed every 2 days until the cells reached confluency. The homogeneity of epithelial and stromal cells was evaluated using immunofluorescent staining for specific markers of epithelial (cytokeratin) and stromal cells (vimentin) as described previously (Szóstek et al. 2012). The epithelial cell contamination of stromal cells was 1% and the stromal cell contamination of epithelial cells was less than 1%.
Experimental procedures
Experiment 1: Effect of PGF2α analogue on circulating P4 and PGFM concentrations
Mares (n = 6) at the mid-luteal phase (days 6-8) were injected i.m. with 1 mL of a PGF2α analogue (Estrumate®; cloprostenol, 250 μg/mL; Intervet, Tokyo, Japan). Cloprostenol, an analogue of PGF2α, was used to induce functional and morphological regression of CL (luteolysis).
Blood samples were collected through a catheter from the jugular vein at fixed intervals (0, 1/4, 2/4, 3/4, 1, 2, 3, 4, 5, 6, 8, 10, 12, 16, 20, 24, 48, and 72 h). The time of cloprostenol injection was defined as 0 h. Blood samples were collected into two sterile plastic tubes containing 200 μl of the stabilizer solution (0.3 M EDTA, 1% acetylsalicylic acid, pH 7.3). Whole blood and plasma, prepared by centrifuging the tubes at 1700 × g for 10 min, were stored at −30°C for later measurements of blood P4 and plasma PGFM concentrations.
Experiment 2: mRNA expression of PGF2α receptor (PTGFR) in equine endometrium in different luteal phases of the estrous cycle
The aim of this study is to determine stage-dependent PTGFR mRNA expression in the endometrium. The endometrial samples were obtained from mares postmortem at an abattoir at the early (n = 6), developing (n = 5), mid (n = 4), late (n = 3), and regressed (n = 7) luteal phases. The mRNA expression of PTGFR was determined using real-time PCR.
Experiment 3: Effect of PGF2α on PGF2α production by endometrial tissues
The aim of this study is to establish the effect of PGF2α on PGF2α production by endometrial tissues in the phase of estrous cycle when the expression of PTGFR was up-regulated. This experiment was carried out in the endometrial tissues (n = 3) obtained from mares at the late luteal phase of the estrous cycle, which was selected based on the results from Experiment 2. Endometrial tissues were treated with vehicle (1% (v/v) ethanol or 0.1 µM of PGF2α (no. 16010; Cayman Chemical, Ann Arbor, MI, USA) for 2 h with or without pretreatment with 10 µM indomethacin (Indo, I7378; Sigma-Aldrich, St. Louis, MO, USA) for 1 h at 38.0°C under 5% CO2 in air. Indomethacin is a nonselective inhibitor of PTGS1 and PTGS2 and belongs to a nonsteroidal anti-inflammatory drug (NSAID). After incubation, tissues were washed six times with sterile physiological saline to remove exogenous PGF2α. Then, the tissues were incubated in fresh culture medium (DMEM/Ham’s F-12; 1:1 (v/v) (no. 12400-024; Invitrogen, Carlsbad, CA, USA)) supplemented with 20 μg/mL of gentamicin (G1397; Sigma-Aldrich) and 0.1% (w/v) BSA for 2 h at 38.0°C under 5% CO2 in air. At the end of incubation, 1 mL of the conditioned media were collected into 1.5 mL tubes containing 10 µl of the stabilizer solution and frozen at −30°C until being processed for PGF2α determination. The tissues were blotted on filter paper and weighed to normalize the PGF2α concentration.
Experiment 4: Effect of PGF2α on PGF2α production by endometrial cells
When the cells collected from the uteri (n = 3) reached 100% confluency (5–6 days after the start of the culture), media were replaced with fresh DMEM/Ham’s F-12 supplemented with 20 µg/mL of gentamicin and 0.1% (w/v) BSA. The endometrial cells were pretreated with or without pretreatment either with Indo (10 µM) or an OT antagonist – L368.899 (100 nM; Nacalai, Tesque, Japan) – for 1 h at 38.0°C in a humidified atmosphere of CO2 in air. After that, the cells were treated with vehicle (1% (v/v) ethanol) or PGF2α (0.1 µM) for 2 h. After incubation, the cells were washed two times with PBS (−) to remove exogenous PGF. Then, the cells were incubated in fresh DMEM/Ham’s F-12 supplemented with 20 μg/mL of gentamicin and 0.1% (w/v) BSA for 2 h at 38.0°C in a humidified atmosphere of 5% CO2 in air. At the end of incubation, conditioned media were collected into 1.5 mL tubes containing 10 µl of the stabilizer solution and frozen at −30°C until being processed for PGF determination. The DNA content, estimated by the spectrophotometric method of Labarca & Paigen (1980), was used to normalize PGF2α concentrations.
An OT antagonist (L368.899) was tested in cultured equine endometrial cells (both epithelial and stromal cells). The cells were pretreated with or without the antagonist (100 nM) for 1 h at 38.0°C in a humidified atmosphere of CO2 in air and then treated with OT (10-7 M) for 2 h. The antagonist inhibited the OT-stimulated PGF2α secretion by both types of cells (Supplementary Fig. 1, see section on Supplementary data at the end of the article), indicating that it was effective.
Experiment 5: Effect of PGF2α on PGF synthesis-related enzymes and PTGFR in endometrial epithelial and stromal cells
The aim of this study is to determine which type of endometrial cells responds to PGF2α action. When the cells (n = 4) reached 100% confluency (5–7 days after the start of the culture), media were replaced with fresh DMEM/Ham’s F-12 supplemented with 20 µg/mL of gentamicin and 0.1% (w/v) BSA. The endometrial cells were treated with vehicle (1% (v/v) ethanol) or PGF2α (0.1µM) for 4 h at 38.0°C in a humidified atmosphere of 5% CO2 in air and then disrupted with 1 mL of TRIZOL Reagent (no. 15596-026; Invitrogen) and stored at −80°C until needed for RNA extraction.
Methods
P4 determination
Concentrations of P4 in whole blood were determined with the PATHFAST assay system (PATHFAST) analyzer using the PATHFAST reagent kit for P4 as described previously (Sugie et al. 2011, Toishi et al. 2013). The assay range of PATHFAST was 0.2–40 ng/mL. The intra-assay coefficient of variation (CV) was from 3.6 to 10.2%.
PGFM determination
Concentrations of PGFM in the plasma samples were determined directly by an enzyme immunoassay (EIA) as described previously (Skarzynski et al. 2003). The PGFM standard curve ranged from 48.828 to 12,500 pg/mL, and the ED50 of the assay was 315 pg/mL. The intra- and inter-assay CVs were on average 2.4 and 7.2% respectively. Cloprostenol, according to the manufacturer, is metabolized to 9,11-dihydroxy-15-eto prosto-5-enoic and 9,11,15-trihydroxyprosto-5-enoic (which are excreted via the urine in 5–6 h), and not to PGFM. Thus, the metabolites of cloprostenol would not affect measurements of PGFM. Cloprostenol was confirmed not to show a cross-reaction with PGFM in our assay system.
PGF2α determination
Concentrations of PGF2α in the conditioned media were determined directly by the EIA as described previously (Uenoyama et al. 1997). The standard curve ranged from 15.625 to 4000 pg/mL, and the ED50 of the assay was 250 pg/mL. The intra- and inter-assay CVs were on average 4.8 and 7.5% respectively.
RNA isolation and cDNA synthesis
Total RNA was prepared from endometrial tissues and cells using TRIZOL reagent according to the manufacturer’s directions. Total RNA (1 μg) was reverse transcribed using a ThermoScript RT-PCR System according to the manufacturer’s directions (no. 11146-016; Invitrogen).
Real-time PCR
Gene expression was determined by real-time PCR using a MyiQ system (no. 170-9740; Bio-Rad Laboratories, Melville, NY, USA) and iQTM SYBR® Green Supermix (no. 170-8880; Bio-Rad Laboratories) starting with 2 ng of cDNA. Briefly, for quantification of the mRNA expression, the primer length (20–22 bp) and GC contents of each primer (40–60%) were designed referring to the previous studies (Atli et al. 2010, Kozai et al. 2014) (Table 1). The primers for PTGFR, PTGS1, PTGS2, PGFS, HPGD, and β-actin generated specific 192-bp, 198-bp, 148-bp, 120-bp, 240-bp, and 113-bp products respectively. Each PCR yielded only a single amplification product. The reaction mixture was 10 μl and included 5 μl of iQTM SYBR® Green Supermix, 1 μl of diethylpyrocarbonate-treated (DEPC) water, 2 μl of template cDNA (1 ng/μl), and 1 μl each of forward and reverse primer (final concentration of each = 0.5 μM). As a negative control, DEPC water instead of template cDNA was used. Real-time PCR was performed under the following conditions: 95°C for 3 min, followed by 45 cycles of 94°C for 15 s, 60°C for 20 s, and 72°C for 15 s. Use of iQTM SYBR® Green Supermix at elevated temperatures resulted in reliable and sensitive quantification of the RT-PCR products with high linearity (Pearson correlation coefficient r > 0.99). We used the 2−ΔΔCT method (Livak & Schmittgen 2001) to analyze the relative level of the expression of each mRNA.
Primers used in real-time PCR.
| Gene | Primer | Sequence | Accession No. | Product size | Reference |
|---|---|---|---|---|---|
| PTGFR | Forward | 5′ - CGTGTGCTTGTTTGCTGTTT - 3′ | NM_001081806 | 192 bp | Atli et al. (2010) |
| Reverse | 5′ - ATGGCATTGCACAAGAATGA - 3′ | ||||
| PTGS1 | Forward | 5′ - GCCTGACTCCTTCAGAGTGG - 3′ | NM_001163976 | 192 bp | Atli et al. (2010) |
| Reverse | 5′ - TCTCGGGATTCCTTGATGAC - 3′ | ||||
| PTGS2 | Forward | 5′ - TATCCGCCCACAGTCAAAGAC - 3′ | NM_001081775 | 148 bp | Atli et al. (2010) |
| Reverse | 5′ - TGTTGTGTTCCCGCAGCCAAAT - 3′ | ||||
| PGFS | Forward | 5′ - AAGCCAGGGCTCAAGTACAA - 3′ | NM_001081895 | 120 bp | Atli et al. (2010) |
| Reverse | 5′ - AGCACCGTAGGCAACTAGGA - 3′ | ||||
| HPGD | Forward | 5′ - GTTGCACAGCAGCCTGTTTA - 3′ | DQ385611 | 240 bp | Atli et al. (2010) |
| Reverse | 5′ - CATCGATGGGTCCAAAATTC - 3′ | ||||
| β-actin | Forward | 5′ - ATGGGCCAGAAGGACTCATA - 3′ | NM_001081838 | 113 bp | Kozai et al. (2014) |
| Reverse | 5′ - TTCTCCATGTCGTCCCAGTT - 3′ |
Statistical analysis
The results are expressed as the mean ± s.e.m. values. In in vivo studies, the statistical significance of difference was assessed by using curves and regression analysis (GraphPad Software version 6, San Diego, USA).
Prostaglandin F2α production was expressed as a percentage of vehicle. The statistical significance of differences in PTGFR mRNA in endometrial tissues and in the concentrations of PGF2α in culture media was assessed by one-way analysis of variance (ANOVA) followed by the Tukey—Kramer multiple comparison test. The statistical significance of differences in PTGS1, PTGS2, PGFS, HPGD, and PTGFR mRNA expressions in endometrial cells was assessed by Student’s t-test. P values < 0.05 were considered significant.
Results
Experiment 1. Effect of PGF2α analogue on circulating P4 and PGFM concentrations
The average circulating P4 and PGFM concentrations at the time of cloprostenol injection (0 h) were 13.6 ± 1.84 ng/mL and 206.5 ± 34.38 pg/mL respectively. Progesterone (P4) concentrations started to decrease 45 min after an injection and continued to decrease up to 24 h (P < 0.05; Fig. 1A). In turn, PGFM started to increase 4 h after an injection and continued to increase up to 72 h (P < 0.05; Fig. 1B).
Mean ± s.e.m. (n = 6) for concentrations of P4 (A) in the whole blood and PGFM (B) in blood plasma from jugular vein at the fixed intervals (0, 0.25, 0.5, 0.75, 1, 2, 3, 4, 5, 6, 8, 10, 12, 16, 20, 24, 48, and 72 h) in mares after intramuscular injection with 1 mL of a PGF2α analogue (Estrumate®; cloprostenol, 250 μg/mL). Asterisks indicate significant differences between experimental data and data from 0 h (P < 0.05).
Citation: Reproduction 151, 5; 10.1530/REP-15-0617
Experiment 2. mRNA expression of PGF2α receptor (PTGFR) in equine endometrium in different luteal phases of the estrous cycle
The level of PTGFR mRNA was higher in the late luteal phase than in the early and regressed luteal phases (P < 0.05; Fig. 2).
PGF2α receptor (PTGFR) mRNA expression in the equine endometrium at different luteal phases. All experimental data are shown as mean ± s.e.m. β-actin was used as a reference gene. Different letters indicate significant differences (P < 0.05) as determined by one-way ANOVA followed by the Tukey—Kramer multiple comparison test.
Citation: Reproduction 151, 5; 10.1530/REP-15-0617
Experiment 3. Effect of PGF2α on PGF2α production in the endometrial tissues
In tissue culture, PGF2α increased PGF2α production compared with control group and a group treated with PGF2α incombination with Indo (P < 0.05; Fig. 3). Prostaglandin F2α in combination with Indo did not affect PGF2α secretion compared with the control group (P > 0.05; Fig. 3). In turn, Indo alone decreased the production of PGF2α compared with control group and a group treated with PGF2α in combination with Indo (P < 0.05, Fig. 3).
Effects of PGF2α on PGF2α production in the equine endometrial tissues. All experimental data are shown as percentage of vehicle. Different letters indicate significant differences (P < 0.05) as determined by one-way ANOVA followed by the Tukey—Kramer multiple comparison test.
Citation: Reproduction 151, 5; 10.1530/REP-15-0617
Experiment 4. Effect of PGF2α on PGF2α production by endometrial cells
In epithelial cells, PGF2α increased PGF2α production compared with control group and a group treated with PGF2α in combination with Indo (P < 0.05; Fig. 4A). Prostaglandin F2α in combination with Indo did not affect PGF2α secretion compared with control group (P > 0.05; Fig. 4A). Indomethacin did not affect PGF2α secretion compared with control group (P > 0.05; Fig. 4A). Oxytocin increased PGF2α secretion compared with control group and a group pretreated with an OT antagonist (P < 0.05; Fig. 5A). Prostaglandin F2α in combination with an OT antagonist increased PGF2α secretion compared with control group (P < 0.05; Fig. 5A). Pretreatment with an OT antagonist did not influence on PGF2α secretion compared with control group (P > 0.05; Fig. 5A).
Effect of PGF2α and PGF2α in combination with Indomethacin on PGF2α production in the equine endometrial epithelial (A) and stromal (B) cells. All experimental data are shown as percentage of vehicle. Different letters indicate significant differences (P < 0.05) as determined by one-way ANOVA followed by the Tukey—Kramer multiple comparison test.
Citation: Reproduction 151, 5; 10.1530/REP-15-0617
Effect of PGF2α and PGF2α in combination with an OT antagonist (L368.899) on PGF2α production in the equine endometrial epithelial (A) and stromal (B) cells. All experimental data are shown as percentage of vehicle. Different letters indicate significant differences (P < 0.05) as determined by one-way ANOVA followed by the Tukey—Kramer multiple comparison test.
Citation: Reproduction 151, 5; 10.1530/REP-15-0617
In stromal cells, PGF2α increased PGF2α production compared with control group (P < 0.05, Fig. 4B). Prostaglandin F2α in combination with Indo increased PGF2α production compared with control group (P < 0.05; Fig. 4B). The pretreatment with Indo did not affect production of PGF2α compared with control group (P > 0.05; Fig. 4B). Oxytocin increased PGF2α secretion compared with control group (P < 0.05; Fig. 5B). Prostaglandin F2α in combination with an OT antagonist increased PGF2α secretion compared with control group (P < 0.01; Fig. 5B). Pretreatment with an OT antagonist did not influence on PGF2α secretion compared with control group (P > 0.05; Fig. 5B).
Experiment 5. Effect of PGF2α on mRNA expressions of PGF synthesis-related enzymes and PTGFR in the endometrial epithelial and stromal cells
PTGS2 mRNA expression was stimulated by PGF2α in the endometrial epithelial and stromal cells (P < 0.05; Fig. 6). Prostaglandin F2α did not stimulate the mRNA expressions of PTGS1, PGFS, HPGD, and PTGFR in endometrial epithelial and stromal cells (P > 0.05; Fig. 6).
Effects of PGF2α on PTGS1, PTGS2, PGFS, HPGD, and PTGFR mRNA expressions in cultured equine endometrial epithelial (A) and stromal (B) cells. The cells were exposed to PGF2α (0.1 μM) for 4 h. β-actin was used as a reference gene. All experimental data are shown as mean ± s.e.m. Different letters indicate significant differences (P < 0.05) as determined by Student’s t-test.
Citation: Reproduction 151, 5; 10.1530/REP-15-0617
Discussion
Prostaglandin F2α
The preceding results strongly suggest that mares have an endometrial PGF2α auto-amplification system. First, at the time of luteolysis, the expression of PTGFR mRNA is elevated, which suggests that the endometrium is more sensitive to PGF2α at this time. Secondly, both endometrial epithelial and stromal cells expressed PTGFR and secreted PGF2α in response to exogenous PGF2α. Thirdly, injection of a PGF2α analogue caused the plasma PGFM concentration to slowly increase, which suggests that PGF2α is released from the uterus in response to PGF2α. A PGF2α auto-amplification system may partly regulate endometrial PGF2α release in mares at the time of luteolysis. There is a growing body of evidence that the expression of endometrial PTGFR is elevated at the time of luteolysis in mares. Our finding that PTGFR mRNA expression in mares is strongest in the late luteal phase (Fig. 2) agrees with an earlier report that it was highest in late diestrus and early luteolysis (Atli et al. 2010). Ruijter-Villani et al. (2014) confirmed the presence of endometrial PTGFR by immunolocalization but showed that the mRNA expression of PTGFR was similar at days 7, 14, and 21 of estrous cycle.
Prostaglandin F2α is secreted in a pulsatile manner in mares (Neely et al. 1970; Ginther et al. 2008). The route for passage of PGF from the uterus to the CL is a whole-body systemic pathway. As stated in the Introduction section, PGF2α auto-amplification systems have been reported in the CL of pigs and cattle. In in vivo studies, injection of a PGF2α analogue stimulated the secretion of uterine PGF2α in sheep (Wade & Lewis 1996) and cattle (Kotwica et al. 1999, Duong et al. 2012). The injection of cloprostenol on day 10 of the estrous cycle increased PGF2α concentration in uterine venous blood plasma in cows (Duong et al. 2012). In this study, cloprostenol induced luteolysis, as shown by the decrease in circulating P4 concentrations. Luteolysis is generally thought to begin when the P4 concentration precipitously decreases and to end when it falls below 2 ng/mL.
It is unclear from the present in vivo results whether cloprostenol injection directly stimulates PGF2α release from uterus. In view of finding that exogenous PGF2α stimulated PGF2α in vivo studies in other species (Wade & Lewis 1996, Kotwica et al. 1999, Duong et al. 2012), we suggest that PGF2α is a regulator of PGF2α production at the time of luteolysis. However, the regulation of PGF2α release at the time of luteolysis is very complex and includes the interactions of many factors (Ginther & Beg 2009); hence, our in vivo results should be interpreted carefully. Further in vivo studies are needed to test the hypothesis that locally produced PGF2α is involved in the regulation of PGF2α secretion from the endometrium.
In this in vitro study, PGF2α stimulated PGF2α production by endometrial tissues and by endometrial epithelial and stromal cells. To determine whether the observed PGF2α had exogenous or endogenous origin, the tissues and cells were pretreated with Indo, which is an inhibitor of PTGS1 and PTGS2, i.e., an inhibitor of PGF2α synthesis. Prostaglandin F2α production was significantly lower in the Indo-pretreated groups than in the PGF2α-treated groups of endometrial tissues and cells, indicating that the observed PGF2α is endogenous, not exogenous. Furthermore, PGF2α increased the mRNA expression of PTGS2 in the endometrial epithelial and stromal cells. This finding is in agreement with a previous finding that PGF2α treatment increased PTGS2 mRNA in the equine CL in vivo (Beg et al. 2005). Prostaglandin F2α also increased PTGS2 expression in human endometrial adenocarcinoma cells (Sales et al. 2008) and myometrial cells (Xu et al. 2013) and increased PGF2α synthesis in the CL of pseudopregnant rats by stimulating Ptgs2 expression (Taniguchi et al. 2010).
Oxytocin
Oxytocin is one of the factors regulating PGF2α secretion. In mares, OT is secreted from hypothalamic nerve cells via the posterior pituitary (Gainer et al. 1985) and the endometrium (Stout et al. 2000, Watson et al. 2000). The temporal relationship between PGF and OT secretion is an important aspect of the luteolytic mechanism in ruminants (McCracken et al. 1999) and perhaps in mares (Shand et al. 2000, Utt et al. 2007). The pulsatile PGF2α secretion is generated by a positive feedback loop between luteal and/or hypophyseal OT and uterine PGF2α (McCracken et al. 1999).
Oxytocin increases PGF2α secretion in vitro in the mare endometrium (King & Evans 1987, Franklin et al. 1989, Paccamonti et al. 1999, Utt et al. 2007, Szóstek et al. 2012, Penrod et al. 2013, Szóstek et al. 2013). In mares, OT is secreted into the uterine lumen during late diestrus and early estrus (Stout et al. 2000), so that OT may locally affect PGF2α production by the endometrium.
Oxytocin treatment causes an increase in PGFM concentration in mares (Goff et al. 1984, Betteridge et al. 1985); however, PGF treatment stimulated an increase in OT concentration in only some individuals (Shand et al. 2000, Handler et al. 2004, Utt et al. 2007). In ruminants, endometrial PGF2α stimulates the release of OT from the CL, which subsequently amplifies the pulses of PGF2α (McCracken et al. 1999). It is not clear whether this also happens in mares. A high dose of PGF2α (1.5 mg) increased OT concentrations in mares (Shand et al. 2000). Prostaglandin F2α injection also increased OT concentration in mares; but, the increase was higher at day 8 than at day 13 of the estrous cycle (Utt et al. 2007). Administration of PGF2α elevated the OT concentration in some mares but not in others (Handler et al. 2004). Furthermore, in gilts, experiments with an OT antagonist (CAP-581) showed that OT controls the height and frequency of the PGF2α pulses (Kotwica et al. 1999). Blocking of OT receptors did not prevent luteolysis in sows (Kotwica et al. 1999) but is unclear whether this is also the case in mares.
As mentioned above, OT is produced by the equine endometrium and affects the secretion of PGF2α by equine endometrial tissues and cells. Pretreatment with an OT antagonist showed that PGF2α affected the secretion of PGF2α from epithelial cells (Fig. 5). Unexpectedly, stromal cells pretreated with an OT antagonist and then treated with PGF2α secreted more PGF2α than stromal cells treated only with PGF2α. Although the reason is unclear, it may be because antagonists can sometimes have agonistic actions (Elands et al. 1988, Mouillac et al. 1995, Postina et al. 1996, Barberis et al. 1998, Siemieniuch et al. 2011).
The overall findings strongly suggest that the equine endometrium has a PGF2α auto-amplification system. Further studies are needed to determine whether it has a role in regulating the release of PGF2α at the time of luteolysis.
Supplementary data
This is linked to the online version of the paper at http://dx.doi.org/10.1530/REP-15-0617.
Declaration of interest
The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.
Funding
This study was supported by the Japanese–Polish Joint Research Project under an agreement between the Japan Society for the Promotion of Science (JSPS) and Polish Academy of Sciences and was supported in part by the Equine Research Institute, Japan Racing Association. A Szóstek was supported by Domestic Grants for Young Scientists from Foundation of Polish Science (FNP, Program Start, Poland) and a JSPS post-doctoral fellow (P-14082).
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