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Endocrinology, doi:10.1210/en.2007-1377
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Endocrinology Vol. 149, No. 6 2866-2876
Copyright © 2008 by The Endocrine Society

Differential Regulation of Prostaglandin Production Mediated by Corticotropin-Releasing Hormone Receptor Type 1 and Type 2 in Cultured Human Placental Trophoblasts

Lu Gao, Chunmei Lu, Chen Xu, Yi Tao, Binhai Cong and Xin Ni

Department of Physiology (L.G., C.L., C.X., B.C., X.N.) and Changhai Hospital (Y.T.), Second Military Medical University, Shanghai 200433, People’s Republic of China; and Department of Physiology (C.L.), Harbin Medical University, Harbin 150086, People’s Republic of China

Address all correspondence and requests for reprints to: Dr. Xin Ni, Department of Physiology, Second Military Medical University, 800 Xiangyin Road, Shanghai 200433, People’s Republic of China. E-mail: nxljq2003{at}yahoo.com.cn.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Prostaglandin (PG) production by intrauterine tissues plays a key part in the control of pregnancy and parturition. The present study was to investigate the role of placenta-derived CRH and CRH-related peptides in the regulation of PG synthesis and metabolism. We found that placental trophoblasts expressed both CRH-R1 and CRH-R2. Treatment of cultured placental cells with either a CRH or urocortin I (UCNI) antibody resulted in a significant decrease in PGE2 release. Both CRH and UCNI antibodies significantly decreased mRNA and protein expression of synthetic enzymes cytosolic phospholipase A2 (cPLA2) and cyclooxygenase (COX)-2 and increased mRNA and protein expression of 15-hydroxyprostaglandin dehydrogenase (PGDH), the key enzyme of PG metabolism. CRH-R1/-R2 antagonist astressin and CRH-R1 antagonist antalarmin significantly inhibited PGE2 release, whereas CRH-R2 antagonist astressin-2b had no effect on PGE2 release. Administration of astressin decreased expression of cPLA2 but had no effect on COX-2 expression. Antalarmin reduced cPLA2 and COX-2 expression, whereas astressin-2b did not alter cPLA2 expression but increased COX-2 expression. PGDH expression was enhanced by these three antagonists. Cells treated with exogenous CRH and UCNI showed an increase in PGE2 release and expression of cPLA2 and COX-2 but a decrease in PGDH expression. UCNII and UCNIII had no effect on PGE2 release but decreased COX-2 and PGDH expression. Our results suggested CRH and CRH-related peptides act on CRH-R1 and CRH-R2 to exert different effects on PG biosynthetic enzymes cPLA2 and COX-2 and thereby modulate output of PGs from placenta, which would be important for controlling pregnancy and parturition.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
PROSTAGLANDINS (PGS) PLAY important roles in the processes of pregnancy and parturition in many mammalian species (1, 2). Specifically, PGs have been linked to induction of fetal organ maturation, up-regulation of the fetal hypothalamic-pituitary-adrenal axis (2, 3), stimulation of myometrial contractility and cervical ripening (4, 5), maintenance of uterine and placental blood flow (6, 7, 8), and inhibition of fetal breathing and movement at the time of labor (3, 9). Therefore, regulating the concentration of PGs in uterus is crucial for controlling pregnancy and parturition.

During human pregnancy, placenta is the major source of PGs within intrauterine tissues (3, 6, 10, 11). It has been demonstrated that human placenta is not only capable of synthesizing PGs but expresses high abundance of the oxidation of nicotinamide adenine dinucleotide-dependent 15-hydroxyprostaglandin dehydrogenase (PGDH), the key enzyme for metabolism of active primary PGs to inactive forms (12, 13). Thus, the output of bioactive PGs from placenta is controlled by both biosynthesis and metabolism of primary PGs within placenta. In the PG biosynthesis pathways, the conversion of arachidonic acid into PGG2 and PGH2 by cyclooxygenase (COX) is currently known to be the rate-limiting step (14). There are two isoenzymes of COX, which are designated as COX-1 and COX-2 (15, 16). The increased expression of COX-2 is believed to be associated with the PG synthesis at term and parturition (17, 18). However, there are other enzymes, such as cytosolic phospholipase A2 (cPLA2), which are also potential regulating steps in PGs synthesis in addition to COX (19). The cPLA2 catalyzes the release of free arachidonic acid, an initial and rate-limiting substrate in PG synthesis, from the sn-2 position of membrane phospholipids (14, 20). Recent studies suggested that cPLA2 is associated with increased production of PGs in the intrauterine tissues (21, 22).

Placenta-derived CRH has been implicated to play a major role in the controlling mechanisms for maintenance of pregnancy and initiation of parturition in humans (23, 24). Although the precise biological functions of placental CRH are not well defined, it appears to exert a number of effects in a complex network within intrauterine tissues by autocrine/paracrine as well as endocrine fashions. It has been demonstrated that CRH produced locally could stimulate estrogen but inhibit progesterone production in placental trophoblasts (25, 26). The addition of CRH to primary placental trophoblast cultures stimulates ACTH and PG secretion (27, 28, 29). It has been demonstrated that CRH belongs to a family of peptides that include urocortin (UCN) I, UCNII, and UCNIII as well as fish urotensin I and frog peptide sauvagine (30, 31, 32). UCNI has been shown to be synthesized and secreted by placental and chorion trophoblasts (33) and has the same biological effects as CRH, stimulating ACTH and PG release from cultured placental cells (34). Recently UCNII and UCNIII have also been reported to be expressed in placental cytotrophoblasts and syncytiotrophoblasts (35, 36), but treatment of cultured placental cells with UCNII and UCNIII failed to stimulate ACTH secretion (36), which suggested that the functions of UCNII and UCNIII in placenta may be different from those of CRH and UCNI. However, the effects of UCNII and UCNIII on PG production have not been investigated. In fetal membrane, CRH was able to increase COX-2 and PGDH expression (37, 38, 39). In placenta, the effects of CRH and UCNI on the enzymes responsible for PG biosynthesis and metabolism, however, remain unknown.

All the CRH family members exert their effects by binding to specific cell surface G protein-coupled receptors. Two major CRH receptor (CRH-R) subtypes have been identified, termed CRH-R1 and CRH-R2. These receptors share 70% homology at the amino acid level but have different binding properties for the members of CRH family. For instance, CRH and UCNI can bind to both CRH-R1 and CRH-R2 (30). CRH has approximately 10-fold lower affinity than UCNI for CRH-R2, whereas UCNII and UCNIII bind exclusively to CRH-R2 (31, 32). Eight and three splice variants of the mRNA for CRH-R1 and CRH-R2, respectively, have been found (40, 41, 42). Human placenta has been shown to express both subtypes of CRH-Rs (35, 43, 44, 45). CRH-R1{alpha}, -R1c, -R1d, and CRH-R2β have also been identified in placenta (43, 46). However, the specific subtype(s) of CRH-Rs responsible for the actions of CRH and CRH-related peptides remains unknown. Thus, the purposes of the present study were to reveal CRH-R1 and CRH-R2 expression in placental tissues as well as cultured placental trophoblasts and, then examine the effects of CRH and CRH-related peptides on PG output and expression of cPLA2, COX-2 and PGDH, the critical enzymes that are responsible for PG synthesis and metabolism, and determine the CRH-R subtype(s) responsible for the action of CRH and CRH-related peptides.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Placental trophoblast culture
Placental tissues were collected from 19 patients (aged 22–30 yr) who underwent elective caesarean section between the 38th and 40th wk of gestation. Tissue collections were performed with approval of Changhai hospital and Second Military Medical University human ethics committees and informed consent from patients. The placental samples were excised from cotyledons after ablation of the choriodecidual layer and transferred immediately to the laboratory and rinsed thoroughly with PBS.

Placental trophoblasts were isolated and cultured according to slightly modified Kliman’s method as described previously (47, 48). Briefly, about 60 g of villi tissue were obtained from the maternal side of the placenta, minced, and then digested with 0.125% trypsin (Invitrogen Corp., Carlsbad, CA) and 0.02% deoxyribonuclease-I (Sigma, St. Louis, MO) in phenol red-free DMEM (Sigma), three times for 30 min each time. The dispersed cells were filtered with 200-µm nylon gauze and loaded onto a discontinued Percoll (Amersham Biosciences, Uppsala, Sweden) gradient (5–70%) and then centrifuged at 2300 x g for 20 min. Cytotrophoblast cells with densities between 1.049 and 1.062 g/ml were collected and then plated into 12-well plates (Corning, Inc. Costar Corp., Cambridge, MA) at a density of 1 x 106/well and grown in phenol red-free DMEM with 10% charcoal-stripped fetal calf serum (FCS) at 37 C in 5% CO2-95% air. On the third day of culture, the medium was changed to FCS-free DMEM containing one of the following treatments: CRH antibody (Santa Cruz Biotechnology, Santa Cruz, CA), UCNI antibody (Santa Cruz), astressin (0.001–1 µmol/liter; Sigma), antalarmin (0.001–1 µmol/liter; Sigma), astressin-2b (0.001–1 µmol/liter; Sigma), CRH (0.001–1 µmol/liter; Sigma), UCNI (0.001–1 µmol/liter; Sigma), UCN II (0.001–1 µmol/liter; Sigma), or UCN III (0.001–1 µmol/liter; Sigma). Control cultures were maintained without additives, and each treatment was performed in quadruplicate for each preparation of cells for 24 h. A well of cells in each treatment was used to assess the cell viability by a dimethylthiazoldiphenyltetra-zoliumbromide (MTT) assay. The assay depends on the reduction of the tetrazolium salt MTT (3-[4,5-dimethylthiazol-2-yl]2,5-diphenyl tetrazolium bromide; Sigma) by functional mitochondria to formazan (49). After a 2-h incubation at 37 C of the cells with MTT, the cells were lysed with dimethyl sulfoxide in Sorensen’s glycine buffer and the formazan crystals solubilized. Absorbance was read at 550 nm using a spectrophotometric microplate reader (Bio-Rad Laboratories, Hercules, CA). It was found that all the treatments did not affect the cell viability in this study (data not shown).

At the end of each experiment, representative wells of cells were fixed and immunostained for cytokeratin and vimentin using primary antibodies (Dako, Inc., Carpinteria, CA) at a dilution 1:1000 to assess cell purity. The results showed that placental cell cultures were predominantly cytokeratin positive (>90%) and vimentin negative (data not shown), suggesting the presence of mainly trophoblast cells and few fibroblasts and decidual cells.

Immunohistochemistry
The placental biopsies from three patients were fixed in buffered formalin before processing the paraffin sections. Paraffin sections (5 µm) were cut, rehydrated, and microwaved in citric acid buffer to retrieve antigens. After inhibition of endogenous peroxidases with 3% H2O2, unspecific antibody binding was blocked with 10% rabbit serum for 30 min. Serial sections were then incubated with specific antibodies against human CRH-R1 and CRH-R2 (both from Santa Cruz, 1:100) overnight at 4 C. The CRH-R1 antibody was directed against an epitope between amino acid positions 81 and 109 of CRH-R1 of human origin, in which no sequence homology existed to CRH-R2. The CRH-R2 antibody was raised against a peptide mapping near the C terminus of CRH-R2 of human origin. The bound antibodies were detected with the biotin-streptavidin-peroxidase system (UltraSensitive-SP kit; MaiXin Biotechnology, Fuzhou, China) using diaminobenzidine (Sigma) as chromogen. Counterstaining was performed with hemalum. Negative controls were performed by substituting primary antibody with a normal IgG in same dilution. To confirm the specificity of primary antibody, preabsorption of the primary antibody with a 10-fold excess of the blocking peptides sc-12381P or sc-20550P (Santa Cruz) was performed.

Immunofluorescent staining for CRH-R1 and CRH-R2
Trophoblast cells were grown for 3 d and then fixed in 4% paraformaldehyde for 1 h after washing with PBS. Fixed cells were washed with PBS and incubated with 10% BSA for 1 h. Then the cells were incubated with antibodies raised against human CRH-R1 or human CRH-R2 at a dilution 1:500. All dilutions were made in 1% BSA in PBS. This incubation was performed overnight at 4 C. Subsequently the specimens were washed with PBS three times and then incubated with fluorescein isothiocyanate-conjugated antigoat IgG (1:100) for CRH-R1 or CRH-R2 at 37 C for 1 h in the dark. For negative controls, the primary antibody was either substituted with a normal IgG in same dilution or preabsorbed with the blocking peptide. The cell nuclei were visualized by applying the DNA-specific dye hoechst at a final concentration of 5 µg/ml. Results were viewed under fluorescent microscope using appropriate filters.

RT-PCR and quantitative real-time PCR
After 24 h treatment of cells with various agents, RNA was extracted from individual samples using the TRIzol reagent (Invitrogen) according to the manufacturer’s guidelines. Quantification of total RNA was performed by measuring absorbance at OD 260. The quality of total RNA was controlled by running 1.5% agarose gels and assessed as acceptable if strong and intact 28S rRNA and 18S rRNA bands were visible under UV light after staining with ethidium bromide. Two micrograms RNA were reverse transcribed in a final volume of 25 µl using the Moloney murine leukemia virus reverse transcriptase (Promega, Madison, WI) and then stored at –20 C.

Specific primers for the amplification of the CRH-R2 variants and CRH-R1{alpha}/CRH-R1β were used as described previously (39). PCR solution consisted of 2.0 µl diluted cDNA, 0.4 µM of each paired primers, 2.5 mM Mg2+, 250 µM deoxynucleotide triphosphates, 2 U Taq DNA polymerase (QIAGEN, Beijing, China), and 1x PCR buffer. PCR was set at 94 C (45 sec), 58 C (45 sec), and 72 C (1 min) in a total 40–80 cycles with a final extension step at 72 C for 10 min.

Primers for nested PCR were previously described (39), and were designed to distinguish the different variants of CRH-R1. Reverse transcription products (cDNA) from cells were used as template for the fist round of PCR. After 40 cycles of amplification, 2 µl of the reaction mixture were used for an additional 40-cycle amplification. The primers used in this second round of amplification were internal to the first set of primers.

Ten microliters of the reaction mixture were subsequently electrophoresed on a 1.5% agarose gel and visualized by ethidium bromide, using a 100-bp DNA ladder (Invitrogen) to estimate the band sizes. As a negative control for all of the reactions, distilled water was used in place of cDNA. The identity of the PCR products was confirmed by sequencing. Sequence data were analyzed using Blast nucleic acid database searches from the National Center for Biotechnology Information (Bethesda, MD).

Quantitative real-time PCR was carried out using Rotor-Gene 3000 (Corbett Research, Sydney, Australia) in a total volume of 25 µl reaction mixture following the manufacturer’s protocol, using the 2x Taq PCR master mix (QIAGEN) and 0.2 µM of each primer listed in Table 1Go. Quantitative real-time PCR conditions were optimized according to preliminary experiments to achieve linear relationships between initial RNA concentration and PCR product. The annealing temperature was set at 60 C, and amplification cycles were set at 40 cycles. The specificity of the primers was verified by examining the melting curve as well as subsequent sequencing of the real-time RT-PCR products. As a negative control for all of the reactions, distilled water was used in place of cDNA. Each sample was normalized on the basis of its β-actin mRNA content. The relative expression of the genes of interest was determined by using comparative threshold cycle (Ct) method (50). Briefly, {Delta}Ct in each group was yielded by subtracting the Ct of the housekeeping gene from the Ct of the target gene yields the {Delta}Ct in each group (control and experimental groups). Then subtracting {Delta}Ct of control group from the experimental group obtains the {Delta}{Delta}Ct, which was entered into the equation 2-{Delta}{Delta}Ct and calculated for the exponential amplification of PCR.


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  		TABLE 1. List of primers used for the amplification of cPLA2, COX-2, PGDH, and β-actin in human placental trophoblast cells

 
Western blotting analysis
Cells were scraped off the plate in the presence of lysis buffer consisting of 50 mM Tris-HCl, 150 mM NaCl, 0.1% sodium dodecyl sulfate (SDS), 1% Triton X-100, 1% sodium orthovanadate, 100 mM dithiothreitol, 0.6 mM phenylmethylsulfonyl fluoride (Sigma), 1.5 µg/ml aprotinin, and 50 mM leupeptin (Watson, Shanghai, China). The cell lysates were quickly sonicated and centrifuged at 12,000 x g for 5 min at 4 C. The supernatant was collected and protein concentration was assayed using a modified Bradford assay. The samples were diluted in sample buffer (containing 250 mM Tris-HCl, 2% SDS, 10% glycerol, and 0.002% bromophenol blue) and boiled for 5 min. Samples were separated on an SDS-8% polyacrylamide gel, and the proteins were electrophoretically transferred to a nitrocellulose filter at 300 mA for 1.5 h in a transfer buffer containing 20 mM Tris, 150 mM glycine, and 20% methanol. The filter was then blocked in Tris-buffered saline containing 0.1% Tween 20 (TBST) and 5% dried milk powder (wt/vol) for 2 h at room temperature. After three washes with TBST, the nitrocellulose filters were incubated with primary antibody for human cPLA2 (1:500; Santa Cruz), COX-2 (1:500; Santa Cruz), β-actin (1:1000; Santa Cruz), and PGDH (1:500; Cayman, Ann Arbor, MI) at 4 C overnight. After another three washes with TBST, the filters were incubated with a secondary horseradish peroxidase-conjugated IgG (1:1000) for 1 h at room temperature and further washed for 30 min with TBST. Immunoreactive proteins were visualized using the enhanced chemiluminescence Western blotting detection system (Santa Cruz). The light-emitting bands were detected with x-ray film. The resulting band intensities were quantitated using an image scanning densitometer (Furi Technology, Shanghai, China). To control sampling errors, the ratio of band intensities to β-actin was obtained to quantify the relative protein expression level.

RIA of CRH and UCNI
After placental cells were cultured for 72 h, the culture media were replaced with fresh DMEM without FCS and then incubated for another 3 h. Culture media were collected and stored at –80 C for later assay.

CRH and UCNI RIA were performed as previously described (48). Synthetic Tyr-CRH or Tyr-UCNI was iodinated by the chloramine T method, purified by Sephadex G-25 gel chromatography, and used as a tracer. UCNI antibody and CRH antibody were supplied by Phoenix Biotech Co. Ltd. (Beijing, China). UCNI assay did not cross-react with rat/human CRH, somatostatin, neuropeptide Y, and LHRH. CRH assay did not cross-react with ACTH, sauvagine, LHRH, and prepro-CRH. The sensitivity of the assay for CRH and UCNI was 3.2 and 6.4 pg/ml, respectively.

PGE2 measurement
After cells were treated with various agents, media were collected for the determination of PGE2 with EIA kit (R&D Systems, Inc., Minneapolis, MN) according to the manufacturer’s protocol. The standard curve ranged from 5000 to 39.06 pg/ml. The sensitivity of the assay was 39 pg/ml, and the intraassay and interassay coefficients of variation were less than 10 and 15%, respectively (manufacturer’s data).

Statistics
For illustrative purposes, the results are presented as the mean percent control ± SEM. Control cultures were conducted in the absent of exogenous reagents. Data were analyzed by one-way ANOVA with Student-Newman-Keuls multiple comparison methods, with significance determined at the level of P < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Expression of CRH-R1 and CRH-R2 in human placental tissue
As shown in Fig 1Go, in placenta, strong staining for both CRH-R1 and CRH-R2 on syncytiotrophoblasts and cytotrophoblasts were identified. The immunostaining of CRH-R1 and CRH-R2 were abolished after preabsorption of the CRH-R1 and CRHR-2 antibodies with the corresponding blocking peptides, respectively (Fig. 1Go, B and D).


Figure 1
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  		FIG. 1. Immunohistochemistry analysis of CRH receptors in placental villus. Immunostained with antibody against CRH-R1 (A) and CRH-R2 (C). CRH-R1 antibody was preabsorpted with CRH-R1 blocking peptide (B); CRH-R2 antibody was preabsorpted with CRH-R2 blocking peptide (D); the primary antibody was substituted with either normal goat IgG (E) or PBS (F). Arrows indicate representative positive staining. Original magnifications, x400 (A–F).

 
Immunofluorescence revealed that both CRH-R1 and CRH-R2 was identified in plasma membrane and cytoplasm of the cultured placental trophoblasts (Fig 2Go). Western blot analysis showed that a single protein band of approximately 55 kDa corresponding to CRH-R1 or CRH-R2 was observed in cultured placental cells (Fig. 3Go, D and E).


Figure 2
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  		FIG. 2. Immunofluorescence analysis of CRH receptors in cultured placental trophoblast cells. Immunostained with antibody against CRH-R1 (A) and CRH-R2 (B). C and D, The same cells were immunostained with hoechst to show cell nuclei. E, CRH-R1 and nuclei overlie. F, CRH-R2 and nuclei overlie. Negative controls (G) CRH-R1 antibody was preabsorbed CRH-R1 blocking peptide. H, CRH-R2 antibody was preabsorbed CRH-R2 blocking peptide; primary antibody was replaced with normal goat IgG (I) or PBS (J). Original magnifications, x200 (A–J).

 

Figure 3
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  		FIG. 3. PCR analysis of CRH-R1 and CRH-R2 variants and Western blot analysis of CRH-R1 and -R2 in cultured human placental trophoblasts. A, Nested PCR was conducted to go across exons 2–7 and 9–14 of CRH-R1 as described previously (39 ). Isoforms are distinguished by different molecular weight of amplified bands. Bands indicated by arrow 1 can be {alpha}, d, f, or g isoforms. Bands indicated with arrow 2 are specific for isoform c. Arrow 3 indicates CRHR1e. Bands indicated with arrow 4 can be {alpha}, β, c, and e. Arrow 5 indicates CRHR1f. B, The existence of CRH-R1{alpha} and -R1β were confirmed by a set of primers as described previously (39 ). C, CRH-R2β was detected using specific primers as described previously (39 ). D and E, Western blotting analysis of CRH-R1 and CRH-R2 protein.

 
Eight isoforms of CRH-R1 have been described, which were derived from different intron/extron splicing (40). We used the specific primers for CRH-R1{alpha}/CRH-R1β and a nested RT-PCR protocol to screen for the presence of these variants in cultured placental trophoblasts as described previously (39, 40). CRH-R1{alpha}, CRH-R1β CRH-R1c, CRH-R1e, and CRH-R1f isoforms were identified (Fig. 3Go, A and B).

Three isoforms of CRH-R2, termed CRH-R2{alpha}, CRH-R2β, and CRH-R2{gamma}, were found previously (41, 42). PCR amplification was conducted by using specific primers for the CRH-R2 isoforms (39). It showed the detection only of CRH-R2β in cultured placental trophoblasts (Fig. 3CGo). None of the other CRH-R2 isoforms (CRH-R2{alpha} and CRH-R2{gamma}) were detected. To confirm the negative outcome for CRH-R2{alpha} and CRH-R2{gamma} were due to a lack of expression but not the failure of the primers to work effectively in the PCR systems, we used cDNA generated from human myometrium and brain tissue biopsies as positive control for CRH-R2{alpha} and CRH-R2{gamma}, respectively. The results suggest that the unique set of primers work effectively in the PCR system and the negative outcome is due to a lack of expression (data not shown).

Effects of CRH and UCNI antibodies on PGE2 release and expression of cPLA2, COX-2, and PGDH
Previous studies demonstrated that the placental trophoblasts secrete CRH and CRH-related peptides UCNI (24, 26, 33). We measured CRH and UCNI content in culture media of placental cells and found that CRH content was 186 ± 59 pg/ml, whereas UCNI content was 93 ± 29.7 pg/ml up to 3 h incubation. To investigate the effects of these endogenous peptides on PG production, cells were treated with either a CRH or UCNI antibody, which resulted in a dose-dependent decrease in PGE2 release after a 24-h treatment period (Table 2Go).


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  		TABLE 2. Effects of CRH and UCNI antibodies on PGE2 content in culture media of placental cells (picograms per milliliter)

 
To elucidate the effect of endogenous CRH and UCNI on PG synthetic and metabolic pathway, we determined mRNA and protein expression of synthetic enzymes cPLA2 and COX-2 as well as PGDH, the key enzyme of PG metabolism. As shown in Fig. 4Go, the CRH antibody and the UCNI antibody dose-dependently decreased mRNA and protein levels of cPLA2 and COX-2 and increased PGDH mRNA and protein expression after a 24-h treatment period. At a concentration of 2 µg/ml, CRH antibody significantly decreased cPLA2 mRNA level to 55 ± 4.7% of control and protein level to 49.6 ± 8.6% of control (Fig. 4AGo), decreased COX-2 mRNA level to 56 ± 1.6% of control and protein level to 58 ± 9.2% of control (Fig. 4BGo), and increased PGDH mRNA level to 388% ± 29.5% of control and protein level to 200 ± 10.3% of control (Fig. 4CGo). UCNI antibody, at a concentration of 2 µg/ml, reduced cPLA2 mRNA level to 70.7 ± 2.0% of control and protein level to 49.6 ± 8.6% of control (Fig. 4DGo), reduced COX-2 mRNA level to 38.5 ± 6.4% of control and protein level to 37.3 ± 10.6% of control (Fig. 4EGo), and increased PGDH mRNA level to 300.9 ± 25.7% of control and protein level to 230.9 ± 23.1% of control (Fig. 4FGo).


Figure 4
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  		FIG. 4. Effects of CRH and UCN I antibodies on expression of cPLA2, COX-2, and PGDH in cultured placental syncytiotrophoblasts. Cells were treated with either a CRH (A–C) or urocortin antibody (D–F) for 24 h. mRNA and protein levels of cPLA2, COX-2, and PGDH were measured by real-time RT-PCR and Western blot analysis, respectively. Representative protein bands are presented on the top of corresponding histogram. Cells treated with control IgG (2 µg/ml, 1:100) had no significantly effect on the expression of cPLA2, COX-2, and PGDH. Data are presented as mean percent control ± SEM for five cultures from five patients (n = 5) performed in triplicate. *, P < 0.05; **, P < 0.01, compared with control.

 
Effects of CRH receptor antagonists on PGE2 release and expression of cPLA2, COX-2, and PGDH
CRH and its related peptides exert their actions through two subclasses of CRH-Rs, CRH-R1 and CRH-R2. To elucidate the specific CRH receptor that is involved in the regulation of PG production, we examined the effects of astressin, the CRH-R1/R2 nonselective antagonist, CRH-R1 antagonist antalarmin, and CRH-R2 antagonist astressin-2b on PG production and the expression of cPLA2, COX-2, and PGDH.

Treatment of cells with increasing concentration of astressin resulted in a dose-dependent decrease in PGE2 release after a 24-h treatment period (Table 3Go). Astressin dose-dependently decreased mRNA and protein levels of cPLA2 and increased the expression of PGDH (Fig. 5Go, A and C). However, it did not affect the mRNA and protein levels of COX-2 at the dosages of 1 nmol/liter to 1 µmol/liter (Fig. 5BGo). At a concentration of 1 µmol/liter, astressin significantly reduced cPLA2 mRNA level to 48.5 ± 5.3% of control and protein level to 45.1 ± 5.8% of control and enhanced PGDH mRNA level to 223.7 ± 40% of control and protein level to 396.9 ± 51.3% of control.


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  		TABLE 3. PGE2 concentration in culture media of placental cells treated with CRH receptor antagonists, CRH, and CRH-related peptides (picograms per milliliter)

 

Figure 5
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  		FIG. 5. Effects of CRH-R1/R2 antagonist astressin on expression of cPLA2, COX-2, and PGDH in cultured placental syncytiotrophoblast cells. Cells were treated with increasing concentration of astressin for 24 h, and then cells were harvested for measurement of cPLA2 (A), COX-2 (B), and PGDH (C) mRNA and protein levels. Representative protein bands were presented on the top of corresponding histogram. Values are presented as mean percent control ± SEM for six cultures from six patients (n = 6) performed in triplicate. *, P < 0.05; **, P < 0.01, compared with control.

 
Treatment of cells with antalarmin dose-dependently reduced PGE2 contents in media, to about 60% of control at a concentration of 1 µmol/liter after a 24-h treatment period (Table 3Go). It caused a decrease in cPLA2 and COX-2 mRNA and protein expression but an increase in PGDH expression (Fig. 6Go). At a concentration of 1 µmol/liter, antalarmin significantly decreased cPLA2 mRNA level to 63.5 ± 2.6% of control and protein level to 56.8 ± 10% of control, decreased COX-2 mRNA level to 45.4 ± 10.3% of control and protein level to 43.1 ± 8.3% of control, and increased PGDH mRNA level to 179.2 ± 29.8% of control and protein level to 195.8 ± 35.1% of control (Fig. 6Go).


Figure 6
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  		FIG. 6. Effects of antalarmin, the specific CRH-R1 antagonist, and astressin-2b, the specific CRH-R2 antagonist, on mRNA and protein levels of cPLA2, COX-2, and PGDH in cultured placental syncytiotrophoblasts. Cells were treated with increasing concentrations of antalarmin or astressin-2b for 24 h, and then cells were harvested for measurement of cPLA2 (A and D), COX-2 (B and E), and PGDH (C and F) mRNA and protein levels. Anta, Antalarmin; As2b, astressin2b. Data are presented as mean percent control ± SEM for six cultures (n = 6) from six patients performed in triplicate. *, P < 0.05; **, P < 0.01, compared with control.

 
Cells were treated with increasing concentration of astressin-2b, which did not change the content of PGE2 in culture media after a 24-h treatment period (Table 3Go). Astressin-2b had no effect on mRNA and protein expression of cPLA2 but significantly increased COX-2 and PGDH mRNA and protein expression (Fig. 6Go). At a concentration of 1 µmol/liter, astressin-2b significantly increased COX-2 mRNA level to 233.2 ± 33.1% of control and protein level to 281.3 ± 50.3% of control and increased PGDH mRNA level to 315.9 ± 40.4% of control and protein level to 309.3 ± 64.5% of control.

Effects of exogenous CRH, UCNI, UCNII, and UCNIII on PGE2 release and expression of cPLA2, COX-2, and PGDH
CRH (1 nmol/liter to 1 µmol/liter) treatment for 24 h dose-dependently increased PGE2 release from placental cells, and this effect could be blocked by either astressin or antalarmin but not astressin-2b (Tables 3Go and 4Go). CRH significantly increased cPLA2 mRNA level to 159.5 ± 17.6 and 207.2 ± 51.7% of control at a concentration of 0.1 and 1 µmol/liter, respectively (P < 0.05 and P < 0.01 vs. control, n = 5). It significantly increased cPLA2 protein level, even at a concentration of 1 nmol/liter, to 198.3 ± 40% of control value (P < 0.05 vs. control, n = 5), At a concentration of 0.01, 0.1, and 1 µmol/liter, respectively, it significantly enhanced cPLA2 protein expression to 335.1 ± 38.2, 495 ± 103.1, and 642.2 ± 114.5% of control. CRH treatment, at a concentration of 0.01 and 0.1 µmol/liter, respectively, caused an increase in COX-2 mRNA level, to 202.6 ± 29.7 and 221.6 ± 16.5% of control (P < 0.05 and P < 0.01 vs. control, respectively, n = 5), and protein level, to 189.3 ± 25.2 and 209.3 ± 16% of control (P < 0.05 and P < 0.01 vs. control, respectively, n = 5). At a high concentration of 1 µmol/liter, CRH did not alter COX-2 mRNA and protein expression (data not shown). Treatment of cells with CRH significantly decreased PGDH mRNA level to 71.7 ± 5.2, 62.1 ± 4.2, and 47.5 ± 4.7% of control value at a concentration of 0.01, 0.1, and 1 µmol/liter, respectively, and protein level to 79.9 ± 6.5, 72.8 ± 4.1, and 43.8 ± 4.6% of control value at a concentration of 0.01, 0.1, and 1 µmol/liter, respectively. Astressin and antalarmin but not astressin-2b reversed the effects of CRH on cPLA2, COX-2, and PGDH expression (Fig. 7Go, A and B).


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  		TABLE 4. PGE2 content in culture media of cells treated with CRH and CRH- related peptides in absence or presence of CRH receptor antagonists (picograms per milliliter)

 

Figure 7
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  		FIG. 7. Effects of CRH and CRH-related peptides on cPLA2, COX-2, and PGDH expression in cultured placental syncytiotrophoblasts. A and B, Cells were treated with CRH (10–7 M) in the presence or absence of astressin (10–6 M), antalarmin (10–6 M), and astressin-2b (10–6 M). C and D, Treatment of cells with UCNI (10–7 M) in the presence or absence of astressin (10–6 M), antalarmin (10–6 M), and astressin-2b (10–6 M), E–H, Treatment of cells with either UCNII(10–7 M) or UCNIII (10–7 M) in the presence or absence of astressin (10–6 M) and astressin-2b (10–6 M). Astr, Astressin; Anta, antalarmin; As2b, astressin-2b. Values are presented as mean percent control ± SEM for five cultures from five patients (n = 5) performed in triplicate. *, P < 0.05; **, P < 0.01, compared with control; #, P < 0.05; ##, P < 0.01, compared with CRH 10–7mol/liter (A and B), UCNI 10–7 mol/liter (C and D), UCNII 10–7 mol/liter (E and F), or UCNIII 10–7 mol/liter (G and H).

 
UCNI exhibited similar effects as CRH, increasing PGE2 release from placental cells (Table 3Go), and this effect could be blocked by either astressin or antalarmin but not astressin-2b (Table 4Go). Treatment of cells with increasing concentrations of UCNI (1 nmol to 1 µmol/liter) increased cPLA2 and COX-2 expression and decreased PGDH expression. UCNI, at a concentration of 0.1 and 1 µmol/liter, respectively, significantly increased cPLA2 mRNA expression, to 175.2 ± 15.4 and 223.7 ± 39% of control (P < 0.05 vs. control, n = 5), and protein level, to 256.5 ± 35.8, 287.1 ± 42, and 296.9 ± 81.3% of control (P < 0.05 and P < 0.01 vs. control, respectively, n = 5). It significantly enhanced COX-2 protein level, to 173.2 ± 16.2 and 192.6 ± 20.8% of control at a concentration of 0.1 and 1 µmol/liter, respectively (P < 0.01 vs. control, n = 5), but did not influence mRNA expression at the concentrations of 1 nmol to 1 µmol/liter after a 24-h treatment period. UCNI significantly decreased PGDH mRNA level, to 77.7 ± 3.3, 70.9 ± 3.4, and 56.8 ± 3.4% of control at a concentration of 0.01, 0.1, and 1 µmol/liter, respectively, and protein level, to 74 ± 5.5, 71.9 ± 3.5, and 54.9 ± 4.2% of control at a concentration of 0.01, 0.1, and 1 µmol/liter, respectively. These effects could be blocked by astressin and antalarmin but not astressin-2b (Fig. 7Go, C and D).

As expected, treatment of cells with UCNII and UCNIII (0.001–1 µmol/liter) had no effect on PGE2 production (Table 3Go) and were unable to restore the inhibitory effect of astressin on PG secretion (Table 4Go). These two peptides did not influence cPLA2 mRNA and protein expression (data not shown). UCNII and UCNIII caused a decrease in mRNA and protein expression of COX-2 and PGDH at a concentration of 0.1 or 1 µmol/liter. These effects could be reversed by astressin and astressin-2b (Fig. 7Go, E–H). UCNII, at 1 µmol/liter, significantly reduced COX-2 mRNA level, to 54.5 ± 8.3% of control, and protein level, to 42.1 ± 5.8% of control, and reduced PGDH mRNA level, to 74.1 ± 4.1% of control, and protein level, 62.7 ± 4.6% of control (P < 0.05 vs. control, n = 5). Treatment of cells with UCNIII at 1 µmol/liter significantly decreased COX-2 mRNA level, to 47 ± 5.6% of control, and protein level, to 66.8 ± 5.7% of control, and reduced PGDH mRNA level, to 58.2 ± 4.3% of control, and protein level, 53.9 ± 3.3% of control (P < 0.05 vs. control, n = 5).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In placenta, CRH-R1 and -R2 have been described in placental tissues by in situ hybridization and immunocytochemistry (35, 43, 44, 45, 46). In the present study, we showed that CRH-R1 and CRH-R2 were localized in placental syncytiotrophoblasts and cytotrophoblasts as well as cultured placental trophoblasts. Western blot and RT-PCR analysis also revealed these two subtypes of CRH-Rs in cultured placental cells. These data conform that placental trophoblasts express both CRH-R1 and -R2.

It is known that placental trophoblasts synthesize CRH and CRH-related peptides UCNI, UCNII, and UCNIII (24, 33, 36), which can interact with CRH-Rs in these cells. In this study, we demonstrated that, when CRH-Rs were blocked with administration of the specific CRH-R1/-R2 antagonist astressin, there was a significant decrease in PGE2 release. Although we were unable to show secretion of UCNII and UCNIII by placental trophoblasts, we have shown that these cells did secrete CRH and UCNI. Moreover, when endogenous CRH and UCNI were blocked by CRH and UCNI antibodies, respectively, PGE2 release from cultured placental cells was significantly reduced. In addition, it was found that CRH and UCNI antibodies decreased synthetic enzymes cPLA2 and COX-2 expression and increased the expression of PGDH, the key enzyme of PG metabolism. To confirm the effects of CRH and urocortins on PG production, the cultured placental cells were treated with exogenous CRH, UCNI, UCNII, and UCNIII. It was shown that CRH and UCNI increased PG release as well as cPLA2 and COX-2 expression and reduced PGDH expression. UCNII and UCNIII did not affect PG release and expression of cPLA2 but inhibited COX-2 and PGDH expression. Our results suggested that placenta-derived CRH and CRH-related peptides may act in an autocrine/paracrine fashion and play an important role in local regulation of PG production throughout gestation.

To investigate the role of CRH-R1 and CRH-R2 in the modulation of PG production, we examined the effects of selective CRH-R1 and CRH-R2 antagonists, respectively. We have demonstrated for the first time that when CRH-R1 and -R2 were blocked by administration of the specific antagonists, respectively, a differential change in PGE2 production in cultured placental cells was exhibited. CRH-R1 antagonist antalarmin decreased PGE2 release, whereas astressin-2b, the CRH-R2 antagonist, did not alter PGE2 secretion from placental cells, which indicates that CRH and CRH-related peptides produced locally exert divergent effects on PG production through interacting with CRH-R1 and CRH-R2. Our present study also showed that CRH-R1 antagonist decreased synthetic enzymes cPLA2 and COX-2 expression and increased expression of PGDH, the key enzyme of PG metabolism, suggesting that activation of CRH-R1 by endogenous ligands enhances biosynthesis and inhibits metabolism of PG and leads to an increase in output of PG in placenta. CRH-R2 antagonist increased COX-2 and PGDH expression, suggesting that CRH-R2 activation have a tonic inhibitory effect on both PG synthesis and metabolism. Our results also demonstrated that CRH and UCNI, which recognize both CRH-R1 and -R2, stimulated PGE2 release from placental cells. Blocking CRH-R1 and -R2 with astressin resulted in decreased PGE2 secretion. These results were consistent with the effects of CRH-R1 antagonist, indicating that CRH and UCNI modulate PG production preferentially via CRH-R1 in cultured placental cells.

Notably, our results demonstrated that the differential effects of CRH-R1 and CRH-R2 on PG production occurred at the synthetic rather than metabolic pathways. In particular, we have demonstrated that CRH-R1 and -R2 antagonists exhibited an opposite action on the expression of COX-2. Moreover, when both CRH-R1 and -R2 were blocked by astressin, no change in COX-2 expression occurred. Treatment of cells with CRH at a high concentration (1 µmol/liter) no longer increased COX-2 expression. Thus, these data supported that CRH-R1 and -R2 exerted an opposite regulatory effect on COX-2 expression in placental trophoblasts. However, we also found that treatment of cells with exogenous UCNI did not change COX-2 mRNA expression but increased COX-2 protein level at a concentration of 0.1 or 1 µmol/liter after a 24-h treatment period. Thus, the mechanisms of exogenous UCNI involved in regulating COX-2 expression need to be further investigated. It is known that placenta is a major source of PGs in the intrauterine tissues (10, 11), whereas chorion tissue serves as a metabolic barrier to control bioactive level of PGs due to its high PGDH expression (12, 51, 52). Interestingly, our previous study has demonstrated that, in chorionic trophoblasts, CRH-R1 and -R2 also exert differential regulatory effects on PG production, which occurs at PGDH, the metabolic pathway (39). In placental and chorionic tissues, CRH-R1 and -R2 exhibit different effects on synthesis and metabolism of PGs, suggesting that effects of CRH and its related peptides on PG output in intrauterus tissues are dependent on cell context.

Both CRHR1 and CRHR2 have multiple known mRNA splice variants. Currently, eight variants of the mRNA for CRH-R1, termed R1{alpha}-h, have been described (40). Several lines of evidence have demonstrated that CRH-R1{alpha} is the main functional CRH-R1 variant (53, 54). Other CRH-R1 variants have distinctive binding and signaling properties. For instance, CRH-R1c has a decreased binding capacity, whereas CRH-R1e appears to attenuate CRH-R1{alpha} signaling in coexpression experiments (55, 56). The CRH-R2 gene has three mRNA splice variants, encoding R2{alpha}, R2β, and R2{gamma} receptor isoforms (41, 42). The CRH-R2β is about 10-fold more potent in second-messenger activation, compared with CRH-R2{alpha} or -R2{gamma}, although their agonist binding and signaling properties of the various CRH-related peptides are not significantly different (42). Studies of the group of Hillhouse and colleagues (43, 57) reported that CRH-R1{alpha}, -R1c, and -R1d but not -R1β were present in placental tissues. However, in cultured placental trophoblasts, we identified not only CRHR1{alpha} and CRHR1c but also CRHR1β, CRHR1e, and CRHR1f variants. This difference may be due to the different samples (placental tissue and cells) and methods that were applied. With regard to CRH-R2, only CRH-R2β, the most potent CRH-R2 variant, was detected in these cells. Interestingly, such expression patterns of the CRH-R1 and CRH-R2 variants were also found in cultured chorion trophoblasts (39). Previous studies demonstrated that, in human pregnant myometrium, the expression of CRH-R1 variants changes as pregnancy progresses toward labor, which may be associated with the changes in the coupling of CRH-Rs to different G proteins and outcome of CRH-R activation (53, 58). Thus, it would be interesting to further investigate the appearance of CRH-R1 variants in placenta during different phases of pregnancy and the roles of different CRH-R1 variants in the regulation of PG production.

Bioactive PGs produced by intrauterus tissues have been implicated as integral uterotonins in the labor process (5). Before parturition, PGs levels within the uterus are greatly increased (59, 60, 61). It has been shown that the biosynthesis and secretion of placental CRH increases exponentially with advancing gestation and reaches the peak at the labor (24), whereas UCNI production in placenta is unchanged during pregnancy (62, 63). Thus, we speculate that the rise of CRH during pregnancy may be associated with increased PG output, which is important for controlling the pregnancy and parturition.

In summary, CRH and CRH-related peptides produced locally may act in an autocrine/paracrine fashion to regulate PG synthetic and metabolic pathways in placenta via two subtypes of CRH receptors, which would be important for the mechanisms controlling pregnancy and parturition.


   Acknowledgments
 
The authors thank the nursing and medical staff of the delivery suites at Changhai Hospital for their cooperation in obtaining placenta.


   Footnotes
 
This work was supported by Natural Science Foundation of China (30170982), Program for Changjiang Scholars, Innovative Research Team in University (IRT0528), and Innovative Foundation for PhD students of Second Military Medical University.

Disclosure Summary: All authors have nothing to disclose.

First Published Online March 6, 2008

Abbreviations: COX, Cyclooxygenase; cPLA2, cytosolic phospholipase A2; CRH-R, CRH receptor; Ct, threshold cycle; FCS, fetal calf serum; MTT, dimethylthiazoldiphenyltetra-zoliumbromide; PG, prostaglandin; PGDH, 15-hydroxyprostaglandin dehydrogenase; SDS, sodium dodecyl sulfate; TBST, Tris-buffered saline containing Tween 20; UCN, urocortin.

Received October 9, 2007.

Accepted for publication February 28, 2008.


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 Introduction
 Materials and Methods
 Results
 Discussion
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