Do arachidonic acid metabolites affect apoptosis in bovine endometrial cells with silenced PPAR genes?

Prostaglandins and Other Lipid Mediators 143 (2019) 106336

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Do arachidonic acid metabolites affect apoptosis in bovine endometrial cells with silenced PPAR genes? A.A. Szczepańska, M. Łupicka, A.J. Korzekwa

T

⁎

Department of Biodiversity Protection, Institute of Animal Reproduction and Food Research of Polish Academy of Sciences in Olsztyn, Tuwima St. 10, 10-747 Olsztyn, Poland

A R T I C LE I N FO

A B S T R A C T

Keywords: PPAR Uterus Apoptosis Arachidonic acid metabolites Stromal cells Cow

Peroxisome proliferator-activated receptors (PPARs) are expressed in bovine uterus, and their agonists are arachidonic acid (AA) metabolites. We hypothesised that silencing of PPAR genes in bovine endometrial stromal cells (ESC) would change the intracellular signalling through PPAR and affect apoptosis after cell treatment with different AA metabolites. The study’s aims are detection of apoptosis and examining the influence of prostaglandins and leukotrienes on apoptosis occurring in physiological ESC and cells with silenced PPAR (α, δ, and γ) genes. Silencing the PPARα and PPARδ genes in cells resulted in increased DNA fragmentation and mRNA and protein expression of caspase (CASP) -3 and -8 (P < 0.05). Neither DNA fragmentation nor the mRNA and protein expression of CASP3 and -8 in cells with silenced PPARγ gene were changed compared to physiological cells (P > 0.05). Among PPARs, PPARα and PPARδ appear to inhibit apoptosis, and AA metabolites, as PPAR agonists, modify this process in bovine ESC.

1. Introduction Endometrial cells change morphologically and functionally throughout the oestrous cycle to establish pregnancy [1,2]. In case of a lack of fertilisation, the functional endometrial layer is shed by menstruation in primates [3]. Since menstruation does not occur in nonprimate species, the morphological change of the endometrium seems to be less important than in primates. Cyclic cell proliferation and cell death by apoptosis have been observed in murine [4], rat [5], canine [6], equine [7] and sow [8] uteri. Cell proliferation and apoptosis undergoing cyclic patterns was also observed in the bovine endometrium [9,10]. Arai et al. [9] localized and quantified KI-67 (cell proliferation marker) and cleaved caspase-3 (CCP3) in luminal and glandular epithelia and the stroma throughout the oestrous cycle. The percentage of KI-67 positive cells in stroma was detected the highest in the follicular phase of the oestrous cycle, percentage of CCP3 positive cells was the highest at the early luteal phase. Peroxisome proliferator-activated receptors (PPARs) are transcription factors that belong to the hormone nuclear receptor superfamily. Three major isoforms of PPAR have been identified: PPARα (NR1C1), PPARδ (NR1C2) and PPARγ (NR1C3). Each isoform is encoded by a separate gene [11]. PPARs possess the classic domain structure of other nuclear receptors. They bind to the peroxisome proliferator response element (PPRE) as obligate heterodimers with 9-cis-retinoic acid ⁎

receptor (RXR: RXRα, RXRβ, RXRγ) [12]. The PPAR/RXR complex can be activated by the ligand of either receptor, and the simultaneous binding of both ligands is more efficient [13]. Peroxisome proliferator activated receptors are activated by natural ligands such as fatty acids, eicosanoids and oxidized fatty acids [14,15]. Eicosanoids are a class of fatty acids mainly derived from arachidonic acid (AA), either via the lipoxygenase pathway leading to the formation of leukotrienes (LTs) and hydrooxyeicosatetraenoic acids (HETEs) or via the cyclooxygenase pathway producing prostaglandins (PGs) [14]. Our previous study [16] showed the influence of PGs and LTs on PPAR mRNA expression in bovine uterine cells. Prostaglandins and leukotrienes are synthesised and secreted in the uterus with the intensity dependent on the phase of the oestrous cycle [17]. Prostaglandins affect ovulation, luteal regression, the implantation and maintenance of pregnancy, parturition and postpartum physiology [18]. Endometrial PGF2α and PGE2 are crucial in the regulation of the corpus luteum (CL) activity respective luteolytic or luteotropic actions in various species [19–21]. Leukotrienes are commonly known as potential inflammatory factors [22]. They are also active in the reproductive tract, LTB4 plays luteotropic role in the CL, stimulating P4 and PGE2 secretions, LTC4 stimulates the secretion of luteolytic PGF2α and may enhance luteolytic cascade within the bovine CL steroidogenic cells [23]. Programmed cell death (apoptosis) is a highly conserved mechanism that has evolved to maintain constant cell numbers and

Corresponding author. E-mail address: [email protected] (A.J. Korzekwa).

https://doi.org/10.1016/j.prostaglandins.2019.106336 Received 10 July 2018; Received in revised form 29 March 2019; Accepted 10 May 2019 Available online 18 May 2019 1098-8823/ © 2019 Elsevier Inc. All rights reserved.

Prostaglandins and Other Lipid Mediators 143 (2019) 106336

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cellular positioning within tissues comprised of different cell compartments [24]. Apoptosis is a common type of cell death associated with morphological features that had been repeatedly observed in various tissues and cell types [25]. The key molecules required for the execution of apoptosis in most systems are caspases (CASPs), which cleave a variety of cellular substrates and thereby cause the characteristic morphology of apoptotic cells [26]. To date, approximately 14 mammalian CASPs have been identified. Caspases share several common features. They are synthesised as inactive zymogens, which can be cleaved into the p20 and p10 subunits to form active enzymes following the induction of apoptosis [27]. Based on CASPs’ functions, they are classified into three groups: (1) inflammatory CASPs (CASP1, -4, -5, -11, -12, -13 and -14) involved in inflammation instead of apoptosis; (2) apoptotic initiator CASPs containing a death-effector domain (CASP8 and -10), or a CASP activation and recruitment domain (CASP2 and -9); and (3) apoptotic effector CASPs (CASP3, -6, -7) [28]. Caspase3 is recognised as a crucial executioner CASP, and CCP3 is widely used to identify apoptotic cells [24,29]. The cleavage and activation of CASP3 needs initiator CASPs (e.g., CASP8 and CASP9) activated by diverse apoptotic stimuli [24]. Moreover, PPARs are involved in apoptosis. Pharmacological agonists of PPARγ induced apoptosis in human cervical cancer HeLa cells [30], whereas PPARδ activation increased proliferation and decreased apoptosis in embryonic cells [31]. Thus, we hypothesised that PPAR genes might be involved in apoptosis in bovine endometrial stromal cells, and arachidonic acid (AA) metabolites – natural PPAR ligands, can modulate the process of apoptosis. The aims of this study were (1) to detect apoptosis in physiological ESC and in cells with silenced PPAR genes and (2) to examine the influence of PGs (PGE2, PGF2α) and LTs (LTC4, LTB4) on apoptosis occurring in physiological ESC and cells with silenced PPAR genes.

Table 1 siRNA sequences used for silencing PPAR genes. No.

Gene name

siRNA sequence 5’ – 3’

1.

PPARα

2.

PPARδ

3.

PPARγ

F: CAGAUCGUUUCCUCCUUUA R: CCAAGACGUUGUCAUCACA F: CCUGUGUUUUUAAUAUAAA R: CAGACUGAUGGAACUUUAA F: CAAUCAGAUUGAAGCUUAU R: AUAAGCUUCAAUCUGAUUG

2.3. Transfection of cells For transfection experiments, ESC were trypsinised with trypsinEDTA solution (0.05% trypsin and 0.02% EDTA). The trypsinised cells were seeded in 6-well plates at a density of 1 × 105 living cells/mL and cultured for 24 h in 10% FCS. Then, cells were transfected using Lipofectamine 2000 reagent (Invitrogen, 11668, Carlsbad, CA) in 1% FCS, according to the manufacturer’s protocol. Cells were transfected with 50 nM siRNA constructs targeting PPARα, PPARδ, or PPARγ, or with scrambled siRNA (Sigma, SIC001). The siRNA sequences are listed in Table 1. The medium was replaced with a stimulation medium DMEM (Sigma, D5796) supplemented with 0.1% BSA (Sigma, A2058) and antibiotics, after 24-h incubation. For negative control (NC) ESC were treated with Lipofectamine 2000 and scrambled siRNA for 24 h, then cells were harvested and frozen in −80 °C for further analysis. 2.4. Cell treatment Transfected and non-transfected ESC were treated with PGE2 (10−6 M; Sigma, P0409), PGF2α (10−6 M; Sigma, P0569), LTC4 (10−6 M; Cayman, 20210, Ann Arbor, MI), or LTB4 (10-6 M; Sigma, L0517) for 48 h in stimulation medium supplemented with 0.1%BSA and antibiotics. After 48 h treatment cells were harvested and frozen in −80 °C for further analysis. The AA metabolite concentrations and the duration of cell stimulation were chosen based on previous studies [23,33], and a preliminary study (data not shown). As a positive control of apoptosis (expression of CASPs and DNA fragmentation), cells were stimulated with tumor necrosis factor-α (TNF-α) and interferon γ (IFNγ; 10 ng/ml each) [34].

2. Materials and methods 2.1. Material collection Uteri from Holstein/Polish Black and White heifers (Bos taurus) aged 3–5 years were obtained at a local slaughterhouse (‘Warmia’, Biskupiec, Poland) and were transported to the laboratory on ice within 40 min. A total of ten (n = 10) uteri in mid-luteal phase (days 8–10) of the oestrous cycle were collected. Each animal was examined by a veterinarian via per rectum ultrasound-guided examination before slaughter to confirm the absence of reproductive tract disorders. The oestrous cycle stage was confirmed by macroscopic observation of the ovary and uterus, as described previously [32], and was confirmed by determination of the progesterone (P4) levels in peripheral blood plasma using a radioimmunoassay. The concentration of P4 ranged between 5.5–6.7 ng/ml. Animals were slaughtered for economic and herd renewal purposes. All procedures were approved by the local Animal Care and Use Committee, Olsztyn, Poland (agreement no. 83/2012/N).

2.5. Total RNA isolation Total RNA was extracted from the cells using a method described by Chomczynski and Sacchi [35]. TRI-Reagent (Sigma, T9424) was used according to the manufacturer’s instructions. The concentration and purity of RNA were assessed using a NanoDrop 1000 instrument (Thermo Fisher Scientific, ND-1000, Wilmington, DE). One microgram of each RNA sample was reverse-transcribed using the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems, 4368813, Cheshire, UK), as described in the supplier’s protocol. cDNA was stored at −20 °C until real-time PCR was performed.

2.2. Cell isolation and in vitro culture

2.6. Real – time PCR

Endometrial stromal cells were isolated from the uterine horn ipsilateral to an ovary with active corpus luteum (CL) by enzymatic dissociation, as described by Skarzynski et al. [33]. After isolation, cells were seeded at a density of 1 × 106 living cells/mL in 2 mL of culture medium (Dulbecco’s Modified Eagle’s medium [DMEM]; Sigma, D5796, St. Louis, MO, USA) per well in collagen-coated 6-well plates. The medium was supplemented with 10% foetal calf serum (FCS; Sigma, 12133C) and antibiotics (gentamicin [Sigma, G1272] and neomycin [Sigma, N1142]). Cells were cultured in humidified atmosphere of 5% CO2 and 95% air. Cells were cultured to reach 70% confluency (approximately 5–6 days). The culture medium was changed every 2 days.

The mRNA expression levels of CASP3 and CASP8 in cells were determined by quantitative real-time PCR using an Applied Biosystems 7900 system (Applied Biosystems, Foster City, CA, USA) with the SensiFAST SYBR Hi-ROX Kit (Bioline Reagents, BIO-92002, London, UK), according to the manufacturer’s instructions. The real-time PCR reaction mixture (10 μL) contained 7 μL of SensiFAST SYBR Hi-ROX Master Mix, 0.2 μM of sense and antisense primers, and 3 μL of reversetranscribed cDNA (30 ng). The primer sequences used to determine mRNA expression of CASP3 and CASP8, ACTB, RN18S and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were designed using the Primer Express Software 3 (Applied Biosystems). The primer sequences, 2

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Table 2 Oligonucleotide sequences used for real-time PCR. No.

Gene name

Sequence 5’ – 3’

Product size (bp)

GenBank accession no.

1.

PPARα

168

NM 001034036.1

2.

PPARδ

162

NM 001083636.1

3.

PPARγ

172

NM 181024.2

4.

GAPDH

103

BC 102589

5.

ACTB

256

K00622

6.

RN18S1

365

AF176811

7.

CASP3

134

NM_001077840.1

8.

CASP8

F: GGTGGAGAGTTTGGCAGAACCAGA R: TCCCACTGCCCAGCTCCGATC F: TCCGAAAGCCCTTCAGTGA R: GGATGGCCTCCACCTGAGACA F: AGGACATTCCGTTCCCAAGAGC R: CCATGAGGGAGTTGGAAGGCT F: CACCCTCAAGATTGTCAGCA R: GGTCATAAGTCCCTCCACGA F: CCAAGGCCAACCGTGAGAAAAT R: CCACATTCCGTGAGGATCTTCA F: AAGTCTTTGGGTTCCGGG R: GGACATCTAAGGGCATCACA F: AAGCCATGGTGAAGAAGGAA R: GGCAGGCCTGAATAATGAAA F: TGTCACAATCGCTTCCAGAG R: CCGAGGTTTGCTTGTCATTC

119

NM_001045970.2

2.8. Terminal deoxynucleotidyl transferase dUTP nick end-labelling (TUNEL) assay

GenBank accession numbers and product lengths are detailed in Table 2. To evaluate the efficiency of reactions, a standard curve was generated using serial dilutions of the appropriate cDNA. Amplification was preceded by an initial enzyme activation step (2 min, 95 °C). The PCR steps were as follows: 40 cycles of denaturation (5 s, 95 °C), annealing and extension (20 s, 60 °C). After amplification, melting curves were acquired by stepwise increases in temperature (50–95 °C) to ensure that a single product was amplified, and no primer-dimer structures were formed. To confirm that only one amplification product was present, the dissociation curve analysis was carried out after each realtime experiment. Control reactions lacking the template or primers were performed to confirm that products were free of primer-dimers and genomic DNA contamination, respectively. The GAPDH reference gene was selected as the most stable gene and was unaffected by the experimental conditions in all samples. The ΔΔCt method was used to process the data. Expression is presented in arbitrary units.

Following the manufacturer’s instructions, DNA damage was evaluated and quantified with a colorimetric apoptosis detection kit (Titer TACS, R&D System, Minneapolis, MN) that uses TUNEL staining in a 96well format. Forty-eight hours after stimulation, cells were trypsinised and suspended in DMEM. Cells were subsequently counted and transferred into a 96-well plate (2 × 105 cells/well). Cells were then fixed with 3.7% buffered formaldehyde for 5 min, followed by washing with PBS. Cells were then subjected to labelling procedure, following the manufacturer’s instructions. The reaction was stopped with 2 N HCl 30 min after substrate addition, and the absorbance was measured at 450 nm with a microplate reader. For a comparison, a positive control (nuclease-treated control) was kept to confirm the permeabilisation and labelling reaction. 2.9. Statistical analyses

2.7. Western blotting The differences in mRNA and protein expression, and the levels of DNA fragmentation were statistically analysed by One-way ANOVA followed by the Bonferroni post hoc test (GraphPAD PRISM Version 6.00, San Diego, CA, USA).

Protein expression of CCP3 (17 kDa and 19 kDa), CCP8 (12 kDa and 20 kDa) and GAPDH in cells was determined by western blotting. Proteins were extracted from cultured cells by incubation with lysis buffer containing 50 mM Tris-HCl (pH 8.0), 150 mM NaCl, 5 mM EDTA, 0.1% SDS, 1% Triton X-100, 0.5% sodium deoxycholate, and protease inhibitors (Sigma, P8340). The lysates were stored at −86 °C until further analysis. Protein concentrations were assessed by the Bradford method. Equal amounts of protein (32 μg) were dissolved in SDS gel-loading buffer, heated to 95 °C for 4 min, and separated by 10% SDS-PAGE. Separated proteins were electroblotted onto 0.2-μm nitrocellulose membranes in transfer buffer. After blocking in 5% non-fat dry milk prepared in TBS-T buffer for 1.5 h at room temperature (RT), the membranes were incubated overnight with a 1:100 dilution of an antiCASP3 antibody (Sigma, C8487), 1:100 dilution of an anti-CASP8 antibody (Sigma, SAB3500404), and anti-GAPDH antibody (Sigma, G8795) which was used as a reference. Antibodies were selected based on the recognition of a single immunoreactive protein of an appropriate molecular weight and tested with available blocking peptides. Membranes were incubated with a 1:20,000 dilution of secondary polyclonal anti-rabbit, or anti-mouse alkaline phosphatase-conjugated antibodies (Sigma, S3687, A4187, and S3562) for 1.5 h at RT. Proteins were visualised by incubating the membranes with 0.4 mg/mL nitroblue tetrazolium chloride and 0.2 mg/mL 5-bromo-4-chloro-3-indolylphosphate toluidine salt (Sigma, 72091) suspended in Tris-buffered saline (pH 9.5). The Quantity One 1-D Analysis Software (BioRad, Hercules, CA, USA) was used to quantitate western blots.

3. Results The expression of PPARs mRNA was successfully silenced, PPARα mRNA expression was decreased by 75% compared with ESC treated with scrambled siRNA, PPARδ by 60%, and PPARγ by 64% (Fig. 1). 3.1. Apoptosis detection in bovine physiological ESC and the effect of AA metabolites Treatment with arachidonic acid (AA) metabolites did not affect the mRNA expression of CASP3 (Fig. 2A, P > 0.05), CASP8 (Fig. 2B, P > 0.05), protein expression of cleaved CASP3 (CCP3) (Fig. 2C, D, P > 0.05), and CCP8 (Fig. 2E, F, P > 0.05). TUNEL assay also did not show any changes in DNA fragmentation in bovine physiological ESC after treatment with AA metabolites (Fig. 2G, P > 0.05). 3.2. Apoptosis detection in bovine ESC with silenced PPARα gene and the effect of AA metabolites Silencing PPARα gene caused an increase in mRNA expression levels of CASP3 (Fig. 3A, P < 0.01) and CASP8 (Fig. 3B, P < 0.05), it also upregulated protein expression of CCP3 (Fig. 3C, D, P < 0.001), CCP8 (Fig. 3E, F, P < 0.001) and DNA fragmentation (Fig. 3G, P < 0.001) 3

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Fig. 1. The mRNA expression of PPARα, PPARδ and PPARγ in physiological ESC (NC-negative control, treated with scrambled siRNA) and in ESC with silenced PPARα, PPARδ and PPARγ genes (C-control group). The data were normalised to the expression of the reference gene GAPDH and are presented in arbitrary units. Bars represent mean ± S.E.M. ***, P < 0.001 (One-way ANOVA, followed by the Bonferroni posthoc test).

P < 0.001). Prostaglandin F2α decreased protein expression of CCP3 (Fig. 4C, D, P < 0.001) and CCP8 12 kDa (Fig. 4E, P < 0.01), and DNA fragmentation (Fig. 4G, P < 0.001). Leukotriene C4 decreased protein expression of CCP3 (Fig. 4C, D, P < 0.001) and CCP8 (Fig. 4E, F, P < 0.05), and DNA fragmentation (Fig. 4G, P < 0.01). Leukotriene B4 decreased mRNA expression of CASP3 (Fig. 4A, P < 0.05), protein expression of CCP3 (Fig. 4C, D, P < 0.001) and CCP8 (Fig. 4E, F, P < 0.05), and DNA fragmentation (Fig. 4G, P < 0.01) in bovine ESC with silenced PPARδ gene.

in ESC (C- control group compared with NC-negative control group). Prostaglandin E2 did not affect CASP3 mRNA expression (Fig. 3A, P > 0.05), but decreased CASP8 mRNA expression (Fig. 3B, P < 0.05), CCP3 protein expression (Fig. 3C, D, P < 0.001), CCP8 protein expression (Fig. 3E, F, P < 0.001) and DNA fragmentation (Fig. 3G, P < 0.001) in ESC with silenced PPARα gene. Prostaglandin F2α decreased CASP3 mRNA expression (Fig. 3A, P < 0.001), CASP8 mRNA expression (Fig. 3B, P < 0.05), CCP3 protein expression (Fig. 3C, D, P < 0.001), CCP8 protein expression (Fig. 3E, F, P < 0.001) and DNA fragmentation (Fig. 3G, P < 0.001) in ESC with silenced PPARα gene. Leukotriene C4 decreased CASP3 mRNA expression (Fig. 3A, P < 0.001), CASP8 mRNA expression (Fig. 3B, P < 0.01), CCP3 protein expression (Fig. 3C, D, P < 0.001), CCP8 protein expression (Fig. 3E, F, P < 0.001) and DNA fragmentation (Fig. 3G, P < 0.001) in ESC with silenced PPARα gene. Leukotriene B4 decreased CASP3 mRNA expression (Fig. 3A, P < 0.05), CCP3 protein expression (Fig. 3C, D, P < 0.001), CCP8 protein expression (Fig. 3E, F, P < 0.001) and DNA fragmentation (Fig. 3G, P < 0.001) in ESC with silenced PPARα gene.

3.4. Apoptosis detection in bovine ESC with silenced PPARγ gene and the effect of AA metabolites Silencing of PPARγ gene did not change the mRNA expression of CASP3 (Fig. 5A, P > 0.05), CASP8 (Fig. 5B, P > 0.05), protein expression of CCP3 (Fig. 5C, D, P > 0.05), CCP8 (Fig. 5E, F, P > 0.05) nor DNA fragmentation level (Fig. 5G, P > 0.05) in bovine ESC. Treatment with AA metabolites did not significantly affect the mRNA expression of CASP3 (Fig. 5A, P > 0.05), CASP8 (Fig. 5B, P > 0.05), protein expression of CCP8 (Fig. 5E, F, P > 0.05), and DNA fragmentation level assessed by TUNEL assay (Fig. 5G, P > 0.05). However PGE2 decreased CCP3 protein level (Fig. 5C, D, P < 0.001), and all other treatments (PGF2α, LTC4, LTB4) decreased CCP3 19 kDa subunit protein expression (Fig. 5D, P < 0.001) in bovine ESC with silenced PPARγ gene.

3.3. Apoptosis detection in bovine ESC with silenced PPARδ gene and the effect of AA metabolites Silencing of PPARδ gene in ESC increased mRNA expression of CASP3 (Fig. 4A, P < 0.05) and CASP8 (Fig. 4B, P < 0.05). Prostaglandin E2 decreased mRNA expression of CASP3 (Fig. 4A, P < 0.05), protein expression of CCP3 (Fig. 4C, D, P < 0.001) and CCP8 12 kDa (Fig. 4E, P < 0.01), and DNA fragmentation (Fig. 4G, 4

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Fig. 2. Effect of PGE2, PGF2α, LTC4 and LTB4 on apoptosis in physiological bovine endometrial stromal cells. (A) mRNA expression of CASP3. (B) mRNA expression of CASP8. The data were normalised to the expression of the reference gene GAPDH and are presented in arbitrary units. (C) protein expression of CCP3 17 kDa. (D) protein expression of CCP3 19 kDa. (E) protein expression of CCP8 12 kDa. (F) protein expression of CCP8 20 kDa. The data were normalised to expression of the reference protein GAPDH and are presented in arbitrary units. (G) TUNEL test for DNA fragmentation, optical density (OD) at 450 nm. Bars represent mean ± S.E.M. All experimental groups are compared with C-control group column. *, P < 0.05; **, P < 0.01; ***, P < 0.001 (One-way ANOVA, followed by the Bonferroni post hoc test). C, non-treated control. NC, negative control, treated with scrambled siRNA. 5

Prostaglandins and Other Lipid Mediators 143 (2019) 106336

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Fig. 3. Effect of PGE2, PGF2α, LTC4 and LTB4 on apoptosis in bovine endometrial stromal cells with silenced PPARα gene. (A) mRNA expression of CASP3. (B) mRNA expression of CASP8. The data were normalised to the expression of the reference gene GAPDH and are presented in arbitrary units. (C) protein expression of CCP3 17 kDa. (D) protein expression of CCP3 19 kDa. (E) protein expression of CCP8 12 kDa. (F) protein expression of CCP8 20 kDa. The data were normalised to expression of the reference protein GAPDH and are presented in arbitrary units. (G) TUNEL test for DNA fragmentation, optical density (OD) at 450 nm. Bars represent mean ± S.E.M. All experimental groups are compared with C-control group column. *, P < 0.05; **, P < 0.01; ***, P < 0.001 (One-way ANOVA, followed by the Bonferroni post hoc test). C, non-treated control. NC, negative control, treated with scrambled siRNA. 6

Prostaglandins and Other Lipid Mediators 143 (2019) 106336

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Fig. 4. Effect of PGE2, PGF2α, LTC4 and LTB4 on apoptosis in bovine endometrial stromal cells with silenced PPARδ gene. (A) mRNA expression of CASP3. (B) mRNA expression of CASP8. The data were normalised to the expression of the reference gene GAPDH and are presented in arbitrary units. (C) protein expression of CCP3 17 kDa. (D) protein expression of CCP3 19 kDa. (E) protein expression of CCP8 12 kDa. (F) protein expression of CCP8 20 kDa. The data were normalised to expression of the reference protein GAPDH and are presented in arbitrary units. (G) TUNEL test for DNA fragmentation, optical density (OD) at 450 nm. Bars represent mean ± S.E.M. All experimental groups are compared with C-control group column. *, P < 0.05; **, P < 0.01; ***, P < 0.001 (One-way ANOVA, followed by the Bonferroni post hoc test). C, non-treated control. NC, negative control, treated with scrambled siRNA. 7

Prostaglandins and Other Lipid Mediators 143 (2019) 106336

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Fig. 5. Effect of PGE2, PGF2α, LTC4 and LTB4 on apoptosis in bovine endometrial stromal cells with silenced PPARγ gene. (A) mRNA expression of CASP3. (B) mRNA expression of CASP8. The data were normalised to the expression of the reference gene GAPDH and are presented in arbitrary units. (C) protein expression of CCP3 17 kDa. (D) protein expression of CCP3 19 kDa. (E) protein expression of CCP8 12 kDa. (F) protein expression of CCP8 20 kDa. The data were normalised to expression of the reference protein GAPDH and are presented in arbitrary units. (G) TUNEL test for DNA fragmentation, optical density (OD) at 450 nm. Bars represent mean ± S.E.M. All experimental groups are compared with C-control group column. *, P < 0.05; **, P < 0.01; ***, P < 0.001 (One-way ANOVA, followed by the Bonferroni post hoc test). C, non-treated control. NC, negative control, treated with scrambled siRNA. 8

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4. Discussion

apoptosis has already been confirmed in human endometrial cells [44,45]. Lebovic et al. [45] indicated that in human immortalised endometriotic epithelial and stromal cells, PPARγ activation decreased the expression of CCP3 protein and simultaneously did not decrease PGE2 biosynthesis, but rather inhibited expression of its receptors EP2 and EP4. In addition, the expression of mRNA and protein for CASP3, under the influence of PGE2, in stromal cells with the silenced PPARγ gene was inhibited [45]. These results are consistent with our results regarding the effect of PGE2 on CASP3 expression in cells with the silenced PPARγ gene. Previously, we have demonstrated interactions between the PPAR and RXR subunits in the PPAR-RXR receptor complex in bovine stromal cells that affect the transmission of the cellular signal from an agonist, e.g., the AA metabolites [16]. Thus, the complete effect of the receptor complex in the process of apoptosis requires further research including the RXR subunit. The overall results show that apoptosis occurs in the bovine endometrium, and among PPARs, primarily PPARα and PPARδ are involved in the inhibition of this process. Moreover, AA metabolites, as PPAR agonists, decreased the programmed cell death in cells with silenced PPARα and -δ genes. The mRNA expression of PPARs was modified differently, dependent on the PPAR type in primary ESC after stimulation with bacterial lipopolysaccharide (our unpublished study), which indicates the role of PPARs in the process of inflammation in bovine uterus. It is necessary to continue the research taking into consideration the roles of the RXR subunit in the process of programmed death of uterine cells, both in physiological and pathophysiological conditions, in different stages of the oestrous cycle.

Our results are the first that have clearly shown an involvement of PPARs in the apoptotic process in bovine ESC. We have demonstrated that silencing the PPARα and PPARδ genes in ESC results in increased DNA fragmentation and expression of CASP3 and -8 at the mRNA and protein levels. Conversely, in ESC with silenced PPARγ gene, no changes in DNA fragmentation and mRNA or protein expression of CASP3 and -8 were observed compared to physiological cells. Although the silencing effect of individual PPAR genes was not complete, the CASPs expression differs statistically significantly in physiological cells and those with silenced PPAR genes. The analysis of literature data suggests that no complete silencing of PPAR genes has been received so far, the achieved silencing effect was 50% compared to control [36]. It is possible that after silencing one type of PPAR, its functions are taken over by a different isoform. Planning the experiment with silencing all types of PPAR could answer the question whether the process of apoptosis in ESC occurs without PPAR's involvement. We have shown that two types of PPAR (α and δ) appear to inhibit the process of apoptosis. In cells with silenced PPARα and PPARδ genes an increase in the expression of CASP3 and -8 occurred, and the DNA fragmentation intensified. In vascular adventitial fibroblasts, PPARα inhibited the process of apoptosis [37]. In contrast it has been reported that PPARα induces apoptosis by affecting the degradation of the Bcl-2 protein in an experiment conducted on hepatocyte cell lines [38]. Hepatocytes are epithelial cells, it might be a reason for inconsistency with the results of our research. No increase was observed in the mRNA and protein expression of CASPs and the level of DNA fragmentation in the cells with silenced PPARγ gene. This PPARγ action contrasts earlier results of Strakowa et al. [39], which showed by the TUNEL assay that the percentage of apoptotic glial brain tumor cells increased under the influence of PPARγ agonists. In addition, the PPARγ agonist stimulated apoptosis in tumor uterine cells [40]. The mRNA and protein expression of PPARγ in bovine endometrium in luteal phase is the lowest among the oestrous cycle [41]. Also our previous study [16] revealed low mRNA and protein expression of PPARγ in bovine ESC in luteal phase. Thus, it is possible that the low expression of PPARγ in luteal phase causes no response to stimulation. Arachidonic acid metabolites in ESC with silenced PPAR genes reduced the mRNA and protein expression of CASPs and the level of DNA fragmentation. It is possible that selected AA metabolites, as PPAR agonists, inhibit apoptosis in the bovine endometrial cells, although it is recognized that there was not complete silencing of the PPARs. The direct influence of AA metabolites on apoptosis in bovine ESC has not been described yet. However, in cow, a positive correlation between the apoptotic rate and heat stress in endometrial cell line was shown by Xiao et al. [42]. In ESC with silenced PPARα and PPARδ genes, most of tested AA metabolites reduced mRNA and protein expression of CASPs and the level of DNA fragmentation. It is possible that selected AA metabolites inhibit apoptosis directly, although it is recognized that there was not complete silencing of the PPARs. In contrast, in cells with silenced PPARγ genes, AA metabolites treatment did not change the mRNA expression of the CASPs, and only PGE2 reduced the protein expression of CCP3. MacLaren et al. [43] showed that synthesis of PGE2 and PGF2α is directly affected by PPARs in bovine epithelial cell line. Stimulating those cells with various PPARs agonists caused PGE2 and PGF2α accumulation. Prostaglandin E2 is associated with reduction of apoptosis and inflammation in female reproductive system [44]. Thus, if PGE2 does directly inhibit apoptosis, silencing of PPARs might effectively reduce PGE2 level and consequently increase apoptosis. In this experiment, when ESC were treated with PGE2 this effect was reversed. Although there is no available literature on the direct involvement of PGs and LTs in the apoptosis of bovine uterine cells under physiological conditions during the oestrous cycle, the contribution of PPARs to

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