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Original Research Article

Effects of testosterone and 2-hydroxyflutamide onprogesterone receptor expression in porcine ovarianfollicles in vitro

Malgorzata Duda a,*, Malgorzata Durlej-Grzesiak a, Zbigniew Tabarowski b,Maria Slomczynska a

aDepartment of Endocrinology, Institute of Zoology, Jagiellonian University, Cracow, PolandbDepartment of Experimental Hematology, Institute of Zoology, Jagiellonian University, Cracow, Poland

a r t i c l e i n f o

Article history:

Received 12 May 2012

Accepted 10 September 2012

Keywords:

2-Hydroxyflutamide

Progesterone receptor

Granulosa cells

Ovarian follicles

Pig

a b s t r a c t

The purpose of the study was to test the possible role of the androgen receptor (AR)

agonist (testosterone; T), an AR antagonist (2-hydroxyflutamide; 2-Hf) or combination of both

(T + 2-Hf) on progesterone receptor (PGR) expression in cultured porcine granulosa cells (GCs)

or whole follicles. GCs isolated from mature pig follicles (6–8 mmin diameter) were cultured for

48 h. Experimental cultures were carried out with the addition of T (10�7 M), 2-Hf (1.7 � 10�4 M)

or both T and 2-Hf for the last 24 h of culture. To better imitate in vivoconditions, isolated whole

porcine follicles (6–8 mm in diameter) were cultured for 24 h in an organ culture system, with

the addition of the same factors. The cells or sections obtained from cultured follicles were

processed for PGR immunocytochemical or immuno-histochemical staining. In addition,

expression of PGR protein was determined by Western blot and progesterone (P4) concentra-

tions in the culture media weremeasuredby a radioimmunoassay. Wefound that isoform A of

PGR is expressed in both granulosal and follicular cultures. The 2-Hf in the presence of

T increased PGR protein expression in porcine GCs and whole follicles. In both granulosal

and follicular cultures, 2-Hf or T alone inhibited P4 secretion, but simultaneous addition of 2-Hf

and T increased P4 secretion.Our results indicatethatandrogensmay be involved in the control

of PGR expression in porcine GCs in vitro. Moreover, we suggest a potential auto/paracrine

regulation of the follicular function by androgen-dependent signaling pathway.

# 2012 Society for Biology of Reproduction & the Institute of Animal Reproduction and

lish Academy of Sciences in Olsztyn. Published by Elsevier Urban &

Partner Sp. z o.o. All rights reserved.

Food Research of Po

1. Introduction

Reproduction in mammals is controlled by the hypothalamic–

pituitary–gonadal axis through a cascade of hormones whose

* Corresponding author at: Department of Endocrinology, Institute ofPoland.

E-mail address: duda.m@op.pl (M. Duda).

1642-431X/$ – see front matter # 2012 Society for Biology of ReproducPolish Academy of Sciences in Olsztyn. Published by Elsevier Urban &http://dx.doi.org/10.1016/j.repbio.2012.10.006

concentrations are tightly regulated by feedback control

mechanisms [1]. Disruption of normal reproductive function,

either transient or permanent, has been linked to endocrine

disrupting compounds (EDCs) which interact directly to

activate or antagonize the sex hormone receptors such as

Zoology, Jagiellonian University, Gronostajowa 9, 30-387 Cracow,

tion & the Institute of Animal Reproduction and Food Research ofPartner Sp. z o.o. All rights reserved.

r e p r o d u c t i v e b i o l o g y 1 2 ( 2 0 1 2 ) 3 3 3 – 3 4 0334

the androgen receptor (AR; [2]). These compounds constitute a

useful tool to study the involvement of signaling pathways

they block or alter in the physiology of cells equipped with

appropriate receptors. Porcine granulosa cells (GCs) are

considered to be the main site of AR- mediated androgen

action in the follicle because of their strongest expression,

which was demonstrated in whole follicles [3] and GCs in vitro

[4]. The presence of AR within GCs provides a basis for the

interactions with AR agonists and antagonists. The agonist- or

antagonist-bound steroid hormone receptors regulate gene

expression by binding to the regulatory elements in promoters

of susceptible genes, in a tissue-dependent manner. In this

work we focused on 2-hydroxyflutamide (2-Hf) which is

known to interact with the AR. Flutamide (FLU) was the first

AR blocker to achieve a wide-spread use. It is metabolized into

2-Hf, the biologically active form of the drug. FLU was used to

treat prostate cancer, where it competes with testosterone (T)

for binding to ARs, thereby reducing the growth of cancer cells

[5]. It has shown its ability to reduce fecundity in the fathead

minnow by decreasing oocyte maturation in the ovaries and

causing spermatocyte degeneration and necrosis in the testis

[6]. The anti-androgenic activity of 2-Hf and FLU has been

well established [7,8]; it also was demonstrated that 2-Hf

is approximately 1–5 times more potent than its parental

compound [9].

Progesterone (P4) is an essential regulator of female

reproductive functions, such as the control of ovulation,

regulation of the function of corpus luteum or initiation of

decidualization [10,11]. These abundant physiological effects

of P4 are mediated through the intracellular progesterone

receptor (PGR; [12]). PGR is a ligand-activated transcription

factor whose biological activity is affected by hormone-

dependent phosphorylation [13]. In the absence of P4, PGR is

maintained in an inactive complex that contains heat shock

proteins in the nuclei of target cells. After P4 binding, PGR

starts to change distinctly, including dissociation of the heat

shock proteins, phosphorylation, and dimerization. Binding of

PGR to progesterone response elements (PGREs) promotes the

formation of a stable initiation complex, resulting in gene

transcription [14]. PGR is expressed as two isoforms termed

PGRA and PGRB [15] arising from a single gene, each under the

control of a separate promoter [16]. The biological effects of P4

are dependent on the activation of both PGR isoforms [17].

PGRs are widespread in an animal organism. In the reproduc-

tive system they have been found in the ovary, fallopian tube,

muscle and mucous membrane of the uterus, its cervix, vagina

and the nipple [18,19]. Immunolocalization and immunoex-

pression of PGRs in female ovaries still arouses great interest

of researchers. This is mainly because of the need of diagnosis

and treatment of infertility, safety of hormone therapy in the

perimenopausal period, as well as the influence on oncogene-

sis in a nipple, endometrium and ovary. PGR expression in GCs

is induced mainly by luteinizing hormone (LH) but other

signaling pathways can contribute to its regulation [20,21].

On the basis of findings that the anti-androgen 2-Hf

suppresses PGR expression in the uterus, in a manner akin

to down-regulation [22], the aim of the present study was to

test the possible role of the AR agonist: T, an antagonist: 2-Hf

or combination of both on PGR expression in cultured porcine

GCs or whole follicles. We hypothesize that, the exposure of

porcine GCs or whole follicles to these substances would alter

PGR expression and that a mixture of these two compounds

would abrogate their individual effects.

2. Materials and methods

2.1. Sample collection

Porcine ovaries were obtained from Polish Landrace sows at a

local slaughterhouse and placed in cold phosphate-buffered

saline (PBS; pH 7.4, PAA The Cell Culture Company, Dart-

mouth, MA, USA) containing Antibiotic/Antimycotic Solution

(AAS 10 ml/ml; PAA The Cell Culture Company, Dartmouth,

MA, USA). Ovaries were transported to the laboratory within

30 min and rinsed twice with sterile PBS supplemented with

antibiotics. In each experiment, ten ovaries from five animals

were selected for cell isolation. Each ovary yielded 3–5 follicles,

thus the total number of follicles varied from 30 to 50. The

phase of the estrous cycle was determined according to the

established morphological criteria [23]. Medium follicles

(6–8 mm in diameter), classified by morphometric criteria as

healthy [24], were selected for cell and organ cultures. Briefly,

follicles were dissected free from the ovarian stroma and

separately classified under a microscope. Healthy follicles

were characterized by a well-vascularized follicular wall and

the clarity of the follicular fluid. Early atretic and atretic

follicles were traversed by few or no blood vessels and the

surface of the follicles was opaque with the progression of

atresia. This procedure has been chosen to minimize the

variability between tissues and animals.

2.2. Granulosa cell isolation and culture

Granulosa cells were scraped from the follicular wall with

round-tip ophthalmologic tweezers. After collection, GCs were

washed several times in PBS and recovered by low speed

centrifugation (90 � g for 10 min; [25]). Cell viability was tested

by the trypan blue exclusion test (mean � SEM: 92% � 1%). The

cells were seeded in 24-well culture plates equipped with a

round coverslip (Nunc, Kalmstrup, Denmark) at an initial

density of 3 � 105 cells/ml (for immunocytochemical analysis)

or in 6-well culture plates (Nunc) at an initial density of

6 � 105 cells/ml (for Western blot). Control cultures were

carried in McCoy’s 5A medium (HyClone Laboratories Inc.,

W Logan, UT, USA) supplemented with 10% fetal bovine serum

(FBS, PAA The Cell Culture Company) for 48 h. Experimental

cultures (48 h) were carried out in McCoy’s 5A medium with

the addition of T (10�7 M; Sigma–Aldrich, St. Louis, MO, USA),

2-Hf (1.7 � 10�4 M; Sigma–Aldrich) or T and 2-Hf for the last

24 h of culture. After termination of culture, all media were

collected and stored at �20 8C for P4 radioimmunoassay. All

experiments were performed in quadruplicate in five separate

experiments (n = 5).

2.3. Follicle culture and preparation forimmunohistochemical analysis

Whole follicles (n = 36, 6–8 mm in diameter) isolated from

porcine ovaries were cultured on a filter disk on a triangular

r e p r o d u c t i v e b i o l o g y 1 2 ( 2 0 1 2 ) 3 3 3 – 3 4 0 335

stainless steel grid over a well of McCoy’s 5A medium

supplemented with 10% FBS and AAS (5 m/ml; [26]). Follicles

during organ culture were randomly assigned to treatment

groups with the addition of T (10�7 M), 2-Hf (1.7 � 10�4 M) or T

and 2-Hf (T + 2-Hf). All culture media were collected after 24 h

and stored at �20 8C for further P4 analysis, whereas follicles

(n = 3/each group) were fixed in 4% paraformaldehyde,

subsequently dehydrated in an increasing gradient of ethanol

and embedded in paraplast (Sigma–Aldrich). Sections of 5 mm

in thickness were mounted on slides coated with 3-amino-

propyltriethoxysilane (Sigma–Aldrich), deparaffinized, and

rehydrated through a series of decreasing alcoholic solutions.

2.4. Immunocytochemstry and immunohistochemistry

The cells were fixed in 2% paraformaldehyde and permeabi-

lized with 0.1% TritonX-100 (Sigma–Aldrich) in Tris-buffered

saline (TBS; 0.05 M Tris–HCl plus 0.15 M NaCl, pH 7.6). To

quench endogenous peroxidase activity, cells were treated for

30 min with 0.1% H2O2. Non-specific binding was blocked by

incubation for 30 min with 5% normal horse serum (Sigma–

Aldrich). Then, cells were incubated overnight at 4 8C with a

mouse monoclonal antibody anti-PGR (NCL-L-PGR-312, clone

16, Novocastra, Leica Biosystems Newcastle Ltd., UK) at a

dilution 1:100. Afterwards, the cells were intensively washed

in TBST (TBS plus 0.1% Tween 20) and incubated for 1.5 h at

room temperature (RT) with biotynylated horse anti-mouse

antibody (1:300; Vector Laboratories, Burlingame CA, USA)

followed by washing with TBST and incubated at RT for 1 h

with avidin–biotin–peroxidase complex (1:1:100; Strept ABC

complex/HRP, DAKO/AS, Glostrup, Denmark). The color

reaction was performed using Stable DAB solution (Research

Genetics, Inc., Huntsville AL, USA) for 4 min. For negative

control the primary antibody was omitted.

Sections of follicles were subjected to a microwave oven

3� for 4 min, in 0.01 M citric acid buffer (pH 6.0), to retrieve

antigens. Endogenous peroxidase activity was blocked by

incubation with 0.3% H2O2 in TBS (Tris-buffered saline, pH

7.4), and non-specific binding was blocked by incubation with

5% normal horse serum (Sigma–Aldrich). Immunohisto-

chemical reactions were performed using the antibody used

in immunocytochemistry and incubation was carried out at

4 8C overnight. Control sections were incubated with 5%

normal horse serum instead of the primary antibody. The

antigens were visualized using biotynylated secondary

antibody – horse anti-mouse antibody (1:300, 1.5 h at RT;

Vector Laboratories, Burlingame, CA, USA), avidin–biotin–

peroxidase complex (1:100, 40 min at RT; StreptABComplex-

HRP, DAKO/AS), and 3,30-diaminobenzidine as the substrate.

Slides were dehydrated and mounted in DPX (Sigma–Aldrich).

The intensity of immunoreaction was analyzed on each

coverslip and section.

2.5. Quantitative evaluation of staining intensity andstatistical analysis

The cells and sections were photographed using the Nikon

Eclipse E200 microscope with the Coolpix 5400 digital camera

(Nikon, Tokyo, Japan) and corresponding software. To esti-

mate quantitatively the intensity of immunoreaction, image

processing and analyses were performed on each coverslip

where at least 500 cells and six different sections from each

examined follicle were analyzed using a public domain ImageJ

software (National Institutes of Health, Bethesda, MD, USA).

The source images were converted to 8-bit grayscale images

and the intensity of PGR staining was measured detachedly

using point selection tool. The background was also analyzed.

Results from each sample were saved as individual means and

interpolated to the following formula:

ROD ¼ODspecimen

ODbackground¼

logðGLblank=GLspecimenÞlogðGLblank=GLbackgroundÞ

!

where GL means gray level for stained area (specimen) and

unstained area (background), and blank is described as a gray

level measured after removing the slide from the light path.

The intensity of immunoreaction was expressed as relative

optical density (ROD).

2.6. Western blot

After termination of culture, GCs were washed twice with ice-

cold saline, and then proteins were extracted with 50 ml of

radioimmunoprecipitation assay buffer (RIPA; Thermo Sci-

entific, Inc., Rockford IL, USA) in the presence of protease

inhibitor cocktail (Sigma–Aldrich). Cultured porcine follicles

were homogenized on ice with a cold Tris/EDTA buffer

(50 mM Tris, 1 mM EDTA, pH 7.5), sonicated and centrifuged at

10,000 � g for 20 min at 4 8C. Supernatant was collected and

stored at �20 8C. Protein concentration was determined with

Bradford reagent (Bio-Rad Protein Assay; Bio-Rad Laborato-

ries GmbH, Munchen, Germany) using bovine serum albumin

(BSA) as a standard. Aliquots (50 mg protein) of cell lysates and

follicle homogenates containing 20 mg of protein were

solubilized in a sample buffer consisting of 62.5 mM Tris–

HCl pH 6.8, 2% SDS, 25% glycerol, 0.01% bromophenol blue, 5%

b-mercaptoethanol (Bio-Rad Laboratories) and heated for

3 min at 99.9 8C. After denaturation, samples were separated

via 10% SDS-polyacrylamide gel electrophoresis under re-

ducing conditions [27]. Separated proteins were transferred

onto a nitrocellulose membrane using a wet blotter in the

Genie Transfer Buffer (20 mM Tris, 150 mM glycine in 20%

methanol, pH 8.4) for 90 min at a constant voltage of 135 V.

After overnight blocking with 5% non-fat milk in TBS, 0.1%

Tween 20 (dilution buffer) at 4 8C with gentle shaking, the

membranes were treated with the primary antibody (for

details see immunohistochemistry subchapter: anti PGR,

dilution 1:1000) for 1.5 h at RT. b-Actin was used as an internal

control (1:2000; Sigma–Aldrich). The membranes were

washed and incubated with a secondary antibody conjugated

with the horseradish–peroxidase labeled horse anti-mouse

IgG (Vector Laboratories; dilution 1:1000) for 1 h at RT. The

signals were detected by chemiluminescence using Western

blotting Luminol Reagent (Santa Cruz Biotechnology). The

blots were visualized using the ChemiDocTM and all bands

were quantified using the Image LabTM 2.0 Software (BioRad

Laboratories). The bands were densitometrically scanned,

and the data obtained for PGR were normalized against

b-actin. The protein level within the control group was

arbitrarily set as 1, against which statistical significance was

analyzed.

[(Fig._1)TD$FIG]

Fig. 1 – The effects of testosterone (T) and antiandrogen (2-hydroxyflutamide, 2-Hf) treatment on progesterone receptor (PGR)

immunostaining in cultures of porcine granulosa cells (A–D) and whole follicles (E–H). (A and E) Control cultures; (B and F)

T-treated cells/follicles; (C and G) 2-Hf-treated cells/follicles; (D) H: T + 2-Hf-treated cells/follicles. Nuclear localization of PGR

in granulosa cells (black arrows), cytoplasmic localization of PGR in granulosa cells (white arrowheads), nuclear staining in

theca cells (white arrows). (I and J) Semiquantitative analysis of PGR staining intensity (means W SEM) in granulosa cell (I)

and follicle (J) cultures exposed to androgen receptor agonist (T), antagonist (2-Hf) or both (T + 2-Hf), expressed as relative

optical density (ROD) of diaminobenzidine brown reaction products. Different letters indicate statistically significant

differences at *p < 0.05 or **p < 0.01; bar = 50 mm.

r e p r o d u c t i v e b i o l o g y 1 2 ( 2 0 1 2 ) 3 3 3 – 3 4 0336

r e p r o d u c t i v e b i o l o g y 1 2 ( 2 0 1 2 ) 3 3 3 – 3 4 0 337

2.7. Radioimmunoassay

Samples of the culture media were analyzed for P4 content

using the radioimmunoassay [28]. [1,2,6,7-3H] progesterone

(specific activity 96 Ci/mmol; Hartmann Analytic GmbH,

Braunschweig) was used as a tracer and an antibody induced

in sheep against 11a-hydroxyprogesterone succinyl: BSA

(a gift from Professor Brian Cook, University of Glasgow,

Glasgow, Scotland). The sensitivity of the assay was 20 pg.

Coefficients of variation within and between assays were

below 5.0% and 9.8%, respectively.

2.8. Statistical analysis

Statistical analysis was performed using Statistica 5.1 program

(StatSoft Inc., Tulsa, OK, USA). Immunocyto- and immunohis-

tochemical data were examined by one-way analysis of

variance followed by Tukey’s test. Radioimmunological and

Western blot data are expressed as means � SEM (n = 3). Each

experiment was performed in quadruplicate. Significant

differences in steroid medium concentration between the

control and experimental cultures were analyzed by Student’s

t-test. Statistical significances were set at *p < 0.05, **p < 0.01

or ***p < 0.001.

3. Results

3.1. PGR immunolocalization and Western blot analysis

The anti-PGR antibody used in this study for immunohis-

tochemistry detects only the isoform A of PGR. The PGR

immunostaining was detected in the nuclei of control and

experimental GCs. Control and T-treated GCs showed similar

PGR staining (Fig. 1A and B), whereas treatment with 2-Hf

resulted in a reduced staining (Fig. 1C). The most intensive

reaction for PGR was observed in GCs treated with T + 2-Hf

(Fig. 1D).

PGR immunostaining was observed in various cell types of

control and experimental follicles (Fig. 1E–H). A weak TC

staining was observed in control follicles (Fig. 1E) and follicles

treated with T or 2-Hf (Fig. 1F and G). In theca cells (TC), PGR

[(Fig._2)TD$FIG]

Fig. 2 – Representative Western blots of PGR protein expression

homogenates (B) from control, testosterone (T), 2-hydroxyflutam

n = 3/per treatment). Different letters indicate statistically signif

was always detected in nuclei and the intensity of TC staining

slightly increased after addition of T + 2-Hf (Fig. 1H). PGR

expression in the GCs was consistently higher than in other

cell types. In follicles cultured with T + 2-Hf, the intensity of

nuclear staining in GCs was the strongest (Fig. 1H), whereas

the intensity of immunoreaction dramatically decreased in

follicles cultured with the addition of 2-Hf (Fig. 1G). A decrease

in the staining intensity of GCs was accompanied with

cytoplasmic localization. Semiquantitative evaluation of the

intensity of the immunocyto- and immunohistochemical

reaction in the GCs and whole follicles expressed as relative

optical density (ROD) confirmed generally the qualitative data

(Fig. 1I and J).

To provide further evidence that GCs and whole follicles

express PGR protein and to confirm the specificity of the

antibody, Western blot analysis was performed. The antibody

used for Western blot detects both A and B isoform of PGR, but

only a single band of 94 kDa (PGRA) was detected in this study.

Densitometric analysis of relative protein level revealed a

distinct decrease in PGR level, in both granulosal and follicular

cultures after 2-Hf treatment. In addition, an increase in

PGR level was observed in GCs and whole follicles exposed to

T + 2-Hf (Fig. 2A and B).

3.2. Progesterone medium concentration

Administration of T and 2-Hf alone decreased granulosal P4

secretion (1.3-fold and 4.4-fold, respectively; p < 0.05; Fig. 3A),

whereas T + 2-Hf significantly increased P4 secretion (1.4-fold;

p < 0.05; Fig. 3A). Similarly, T and 2-Hf significantly inhibited P4

production by porcine follicles (1.4-fold and 2.2-fold, respec-

tively; p < 0.05; Fig. 3B), while T + 2-Hf significantly stimulated

follicular P4 production (1.6-fold; p < 0.05; Fig. 3B).

4. Discussion

The study was focused on the possible involvement of AR

signaling on the induction and localization of PGR by using

specific AR antagonist, 2-hydroxyflutamide and AR agonist,

testosterone. Our results indicated that the AR antagonist

exerts a negative effect on PGR protein expression in both

(mean W SEM) in granulosa cell lysates (A) and follicular

ide (2-Hf) and T + 2-Hf- treated cells (48 h) or follicles (24 h;

icant difference at *p < 0.05 or ***p < 0.001.

[(Fig._3)TD$FIG]

Fig. 3 – Progesterone secretion (mean W SEM) by granulosa cells (A) and whole follicles (B) cultured for 48 and 24 h,

respectively (n = 3 independent experiments). Different letters indicate statistically significant difference at *p < 0.05 and

**p < 0.01; (C) control, T: testosterone, 2-Hf: 2-hydroxyflutamide, T + 2-Hf: testosterone + 2-hydroxyflutamide.

r e p r o d u c t i v e b i o l o g y 1 2 ( 2 0 1 2 ) 3 3 3 – 3 4 0338

cultured porcine GCs and whole ovarian follicles. Additionally,

2-Hf induced perinuclear rather than nuclear distribution of

PGR in granulosa cells after whole follicle culture.

2-Hydroxyflutamide is considered to be a potent, nonste-

roidal antiandrogen which has, as yet, no other known

agonistic or antagonistic hormonal properties. It binds to

the AR and competitively inhibits the binding of T and

dihydrotestosterone (DHT; [29]). However, similar to the

results of the current study, findings of Chandrasekhar and

Armstrong [22] suggested an anti-progestational activity of

2-Hf, which hitherto has been claimed to lack other agonistic

or antagonistic activities [30]. It has been observed that 2-Hf

blocks ovulation and the preovulatory LH surge in hCG-primed

prepubertal rats [31], and these effects are reversed by an

injection of LHRH or P4 on the day of proestrus [32]. 2-Hf also

delays the initiation of implantation, fetal development, and

parturition in pregnant rats and suppresses decidualization

after artificial stimulation of the sensitized uterus of ovariec-

tomized steroid-treated pseudopregnant rats [33].

In the present study we demonstrated that PGR protein

expression in granulosa cells and whole follicles of pigs is

decreased after the addition of 2-Hf to the culture medium.

Interestingly, simultaneous addition of T and 2-Hf significant-

ly increased the PGR staining intensity. Thorough analysis of

PGR distribution within the cultured follicles revealed quali-

tative differences in the localization and staining intensity. In

control follicular cultures, PGR staining was observed in

granulosa and theca layers, while in follicles treated with

2-Hf, the theca layer had almost no PGR immunoreaction. This

is consistent with the report by Durlej et al. [34] who

demonstrated the differential expression of both PGR isoforms

in the porcine ovary during the estrous cycle, and suggested

cell-specific influence of progesterone.

It was reported that the ability of antiandrogens to

antagonize certain androgenic effects and to agonize others

may result from its unique interactions with the AR [35]. 2-Hf

can bind to the AR and induce conformational changes of the

receptor responsible for conveying the androgen signal on

cytoplasmic transduction pathways, but is unable to induce

the conformational changes necessary for DNA binding and

gene regulation [36]. Furthermore, Hickey et al. [37] indicated

the existence of a non-genomic pathway of androgen action

leading to modulation of the transcriptional activity of target

genes. The authors found that FSH-stimulated P4 secretion

was suppressed by DHT and 2-Hf in porcine granulosa. We

hypothesize that androgens may affect PGR protein expres-

sion via AR during follicular development, however to reveal

the actual mechanism it requires additional examination.

2-Hf is known to inhibit expression of AR-responsive genes

such as 5-a reductase, which metabolizes T into DHT in rats

[38], caspase-8 or FADD-like apoptosis regulator [39]. 2-Hf can

also block androgen-regulated expression by indirect modes

of action, e.g.by inhibition the AR action via the recruitment of

corepressors [40]. Antiandrogens can also act as weak

androgens when acting alone and as antagonists when

acting in combination with androgens [41]. In addition,

2-Hf may activate other receptor pathways such as the aryl

hydrocarbon receptor [42]. It may also induce feedback

regulation at the level of gonadotropins [43]. Thus, it is

possible that 2-Hf employs multiple action modes involved in

receptor activation.

Our recent studies [44] revealed that androgens play a

critical role in selective apoptosis of granulosa cells during

porcine follicular atresia. It has been shown that 2-Hf, in the

presence of a high T level, attenuates induction of granulosal

cell apoptosis. The response of the ovarian follicular cells to

the combined effects of survival and death promoting signals

determines their ultimate fate: atresia or ovulation. Proges-

terone is one of the factors that are induced by LH, and it has

been reported to act as an antiapoptotic factor in luteinized rat

and human granulosa cells [45,46]. In our experiment,

simultaneous addition of T and 2-Hf to GC culture media of

granulosa cells and intact follicles caused a significant

increase in P4 production. Such increase is typical for cells

undergoing luteinization [47].

We found that intact follicles or GCs treated with T or 2-Hf

alone produced less P4 than controls. This is in agreement with

Henderson et al. [48] who demonstrated that bovine GCs

treated with T secreted lower amounts of P4 than untreated

cells. Moreover, Tanabe et al. [49] reported that P4 production

by porcine granulosa cells, in the presence of gonadotropins, is

modulated in a paracrine or an autocrine fashion by andro-

gens. These authors also found that androgens may exert their

actions partially by altering the activity of cholesterol side

chain cleavage enzymes. In turn, the inhibitory effect of

T antagonist, FLU on P4 secretion was demonstrated by Rangel

et al. [50] who showed that the absence of an increased P4

release resulted in a lack of preovulatory LH surge.

Our medium P4 data corresponded well with the granulosal

PGR expression. The positive P4 effects on the expression of its

r e p r o d u c t i v e b i o l o g y 1 2 ( 2 0 1 2 ) 3 3 3 – 3 4 0 339

own receptors is well known. The presence of PGR in the

granulosa and theca layer of preovulatory follicles marks the

cells as targets for P4 action. Results of studies performed on

PR-A knockout (PRAKO) mice clearly indicated that the

presence of PGRA in granulosa cells of preovulatory follicles

was critical for ovulation [51]. The 2-Hf-induced changes in

PGR expression and P4 level may help to understand the

granulosa cell apoptosis and follicular atresia.

In the present study, we also demonstrated that some

granulosa cells of follicles treated with 2-Hf showed both

nuclear and perinuclear/cytoplasmic PGR immunostainning.

According to D’Haeseleer et al. [52] this type of staining may

reflect the cycle-dependent localization of PGR. A negative

correlation was found between the PGR staining intensity and

the plasma P4 concentration. It is not clear whether in the cells

producing low amounts of P4, the receptor was unbound and

inactive or was ready to be ubiquitinated.

In summary, the present study revealed for the first time

that testosterone antagonist 2-Hf can affect PGR protein

expression and progesterone secretion by porcine granulosa

cells and whole ovarian follicles in culture, indicating that

androgens are involved in normal follicular development. The

down-regulation of PGR expression after exposure to 2-Hf

confirms its antiprogestational activity and demonstrate a key

role of androgens in physiological granulosa cell apoptosis and

porcine follicular atresia.

Acknowledgments

The study was supported by the Ministry of Science and

Higher Education (Grant no. N303 538838). M. Durlej-Grzesiak

is a scholar of the Foundation for Polish Science (START

Programme 2011).

r e f e r e n c e s

[1] Palermo R. Differential actions of FSH and LH duringfolliculogenesis. Reproductive BioMedicine Online2007;15:326–37.

[2] Tabb MM, Blumberg B. New models of action for endocrine-disrupting chemicals. Molecular Endocrinology2006;20:475–82.

[3] Slomczynska M, Tabarowski Z. Localization of androgenreceptor and cytochrome P450 aromatase in the follicle andcorpus luteum of the porcine ovary. Animal ReproductionScience 2001;65:127–34.

[4] Duda M, Slomczynska M. Immunohistochemicallocalization of androgen receptor in two subpopulations ofporcine granulosa cells in vitro. Reproduction in DomesticAnimals 2007;42:22–5.

[5] Singh SM, Gauthier S, Labrie F. Androgen receptorantagonists (antiandrogens): structure–activityrelationships. Current Medicinal Chemistry 2000;7:211–47.

[6] Jensen KM, Kahl MD, Makynen EA, Korte JJ, Leino RI,Butterworth BC, et al. Characterization of responses to theantiandrogen flutamide in a short-term reproductionassay with the fathead minnow. Aquatic Toxicology2004;70:99–110.

[7] Clark CR, Nowell NW. The effect of the non-steroidal anti-androgen flutamide on neural receptor binding of

testosterone and intermale aggressive behaviour of mice.Psychoneuroendocrinology 1980;5:39–45.

[8] Neri RO. Antiandrogens. In: Sighal RL, Thomas JA, editors.Cellular mechanisms modulating gonadal hormone action.Baltimore: University Park Press; 1976. p. 233–62.

[9] Neri RO, Peets E, Watnick A. Antiandrogenicity of flutamideand its metabolite Sch 16423. Biochemical SocietyTransaction 1979;1:565–9.

[10] Espey LL, Lipner H. Ovulation. In: Knobil E, Neill JD, editors.The physiology of reproduction. New York: Reven Press;1994. p. 725–80.

[11] Evans AC. Characteristics of ovarian follicle developmentin domestic animals. Reproduction in Domestic Animals2003;38:240–6.

[12] Leonhardt SA, Boonyaratanakornkit V, Edwards DP.Progesterone receptor transcription and non-transcriptionsignaling mechanism. Steroids 2003;68:761–70.

[13] Migliaccio A, Didomenico M, Green S, Defalco A,Kajtaniak EL, Blasi F, et al. Phosphorylation on tyrosine ofin vitro synthesized human estrogen receptor activatesits hormone binding. Molecular Endocrinolology1989;3:1061–9.

[14] Tsai MJ, O’Malley BW. Molecular mechanisms of action ofsteroid/thyroid receptor superfamily members. AnnualReview of Biochemistry 1994;63:451–86.

[15] Conneely OM, Mulac-Jericevic B, Lydon JP. Progesteronedependent regulation of female reproductive activity bytwo distinct progesterone receptor isoforms. Steroids2003;68:771–8.

[16] Kastner P, Krust A, Turcotte B, Stropp U, Tora L,Gronemeyer H, et al. Two distinct estrogen-regulatedpromoters generate transcripts encoding the twofunctionally different human progesterone receptor formsA and B. EMBO Journal 1990;9:1603–14.

[17] Conneely OM, Mulac-Jericevic B, Lydon JP, De Mayo FJ.Reproductive functions of the progesterone receptorisoforms: lessons from knock-out mice. Molecular andCellular Endocrinology 2001;179:97–103.

[18] Slomczynska M, Krok M, Pierscinski A. Localization of theprogesterone receptor in the porcine ovary. ActaHistochemica 2000;102:183–91.

[19] Starczewski A, Brodowska A, Laszczynska M, Piasecka M.The presence and role of progesterone receptor in theovaries of postmenopausal women who have not appliedhormone replacement therapy. Folia Histochemica etCytobiologica 2008;46:277–82.

[20] Clemens JW, Robker RL, Kraus WL, Katzenellenbogen BS,Richards JS. Hormone induction of progesterone receptor(PR) messenger ribonucleic acid and activation of PRpromoter regions in ovarian granulosa cells: evidence for arole of cyclic adenosine 30,50-monophosphate but notestradiol. Molecular Endocrinology 1998;12:1201–14.

[21] Iwai M, Yasuda K, Fukuoka M, Iwai T, Takakura K, Taii S,et al. Luteinizing hormone induces progesterone receptorgene expression in cultured porcine granulosa cells.Endocrinology 1991;129:1621–7.

[22] Chandrasekhar Y, Armstrong DT. Ability of progesterone toreverse anti-androgen (hydroxyflutamide) inducedinterference with preovulatory LH surge and ovulation inPMSG-primed immature rats. Journal of Reproduction andFertility 1989;85:309–16.

[23] Gregoraszczuk E, Wojtusiak A. Histochemical evaluation ofD5,3b-hydroxysteroid dehydrogenase activity in two typesof porcine corpora lutea and granulosa cells in tissueculture. Acta Histochemica 1982;70:22–30.

[24] Lin P, Rui R. Effects of follicular size and FSH ongranulosa cell apoptosis and atresia in porcine antralfollicles. Molecular Reproduction and Development2010;77:670–8.

r e p r o d u c t i v e b i o l o g y 1 2 ( 2 0 1 2 ) 3 3 3 – 3 4 0340

[25] Duda M, Knet M, Tabarowski Z, Slomczynska M. Lutealmacrophage conditioned medium affects steroidogenesisin porcine granulosa cells. Reproductive Biology2011;11:117–43.

[26] Freshney RI. Culture of animal cells. In: Freshney RI, editor.A manual of basic technique. New York: John Wiley andSons; 2005. p. 481–95.

[27] Laemmli UK. Cleavage of structural proteins during theassembly of the head of bacteriophage T4. Nature1970;227:680–5.

[28] Abraham GE, Swerdloff R, Tuchlinsky D, Odell WD.Radioimmunoassay of plasma progesterone. Journal ofClinical Endocrinology and Metabolism 1971;32:619–24.

[29] Brogden R, Chrisp P. Flutamide: a review of itspharmacokinetic properties and therapeutic usein advanced prostatic cancer. Drugs and Aging1991;1:104–15.

[30] Neri R, Florance K, Koziol P, Van Cleave S. A biologicalprofile of a nonsteroidal antiandrogen, SCH 13521 (40-nitro-30-trifluoromethylisobutyoanilide). Endocrinology1972;91:427–37.

[31] Opavsky MA, Chandrasekhar Y, Roe M, Armstrong DT.Interference with the preovulatory luteinizing hormonesurge and blockage of ovulation in immature pregnantmare’s serum gonadotropin-primed rats with anti-androgenic drug, hydroxyflutamide. Biology ofReproduction 1987;36:636–42.

[32] Chandrasekhar Y, Armstrong DT. Human chorionicgonadotropin (hCG) and luteinizing hormone-releasinghormone (LHRH) reverse the blockage of ovulation inpregnant mare’s serum gonadotropin (PMSG)-primedimmature rats by the anti-androgenic drug,hydroxyflutamide. Canadian Journal of Physiology andPharmacology 1988;66:83–787.

[33] Chandrasekhar Y, Armstrong DT, Kennedy IG.Implantation delay and anti-deciduogenic activity in therat by the anti-androgen, hvdroxyflutamide. Biology ofReproduction 1990;42:120–5.

[34] Durlej M, Tabarowski Z, Slomczynska M.Immunohistochemical study on differential distribution ofprogesterone receptor A and progesterone receptor Bwithin the porcine ovary. Animal Reproduction Science2010;121:167–73.

[35] Zhu X, Li H, Liu JP, Fonder JW. Androgen stimulatesmitogen-activated protein kinase in human breastcancer cells. Molecular and Cellular Endocrinology1999;152:199–206.

[36] Kemppainen JA, Lane MV, Sar M, Wilson EM. Androgenreceptor phosphorylation, turnover, nuclear transport, andtranscriptional activation. Specificity for steroids andantihormones. Journal of Biological Chemistry1992;267:968–74.

[37] Hickey TE, Marrocco DL, Gilchrist RB, Norman RJ,Armstrong DT. Interactions between androgen and growthfactors in granulosa cell subtypes of porcine antral follicles.Biology of Reproduction 2004;71:45–52.

[38] Nguyen TV, Yao M, Pike CJ. Flutamide and cyproteroneacetate exert agonist effects: induction of androgenreceptor-dependent neuroprotection. Endocrinology2007;148:2936–43.

[39] Garcia-Reyero N, Villeneuve DL, Kroll KJ, Liu L, Orlando EF,Watanabe KH, et al. Expression signatures for model

androgen and antiandrogen in the feathead Minnow(Pimephales promelas) ovary. Environmental Science andTechnology 2009;43:2614–9.

[40] Rahman MM, Miyamoto H, Lardy H, Chang C. Inactivationof androgen receptor coregulator ARA55 inhibits androgenreceptor activity and agonist effect of antiandrogens inprostate cancer cells. Proceedings of the NationalAcademy of Sciences of the United States of America2003;100:5124–9.

[41] Wong C, Kelce WR, Sar M, Wilson EM. Androgen receptorantagonist versus agonist activities of the fungicidevinclozolin relative to hydroxyflutamide. Journal ofBiological Chemistry 1995;270:19998–20003.

[42] Coe KJ, Nelson SD, Urlich RG, He Y, Dai X, Cheng O, et al.Profiling the hepatic effects of flutamide in rats: amicroarray comparison with classical aryl hydrocarbonreceptor ligands and atypical CYP1A inducers. DrugMetabolism and Disposition 2006;34:1266–75.

[43] Thackray VG, McGillvray SM, Mellon PL. Androgens,progestins and glucocorticoids induce FSH beta-subunitgene expression at the level of the gonadotrope. MolecularEndocrinology 2006;20:2062–79.

[44] Duda M, Durlej M, Knet M, Knapczyk-Stwora K,Tabarowski Z, Slomczynska M. Does 2-hydroxyflutamideinhibit apoptosis in porcine granulosa cells? An in vitrostudy. Journal of Reproduction and Development2012;58:438–44.

[45] Makrigiannakis A, Coukos G, Christofidou-Solomidou M,Montas S, Coutifaris C. Progesterone is an autocrine/paracrine regulator of human granulosa cell survivalin vitro. Annals of the New York Academy of Sciences2000;900:16–25.

[46] Yacobi K, Tsafriri A, Gross A. Luteinizing hormone-inducedcaspase activation in rat preovulatory follicles is coupled tomitochondrial steroidogenesis. Endocrinology2007;148:1717–26.

[47] Channing CP, Ledwitz-Rigby F. Methods for assessinghormone mediated differentiation of ovarian cells inculture and short-term incubations. In: Hardman JG,O’Malley BW, editors. Methods in enzymology. New York:Academic Press; 1975. p. 183–230.

[48] Henderson KM, McNatty KP, Smith P, Gibb M, O’Keeffe LE,Lun S, et al. Influence of follicular health on thesteroidogenic and morphological characteristics of bovinegranulosa cells in vitro. Journal of Reproduction andFertility 1987;79:185–93.

[49] Tanabe K, Tamura T, Saijo A, Kikuchi M, Sho R, Mashu A,et al. Modulation of progesterone secretion by androgens inporcine granulosa cell cultures. Hormone Research1992;37(1):25–31.

[50] Rangel PL, Sharp PJ, Gutierrez CG. Testosterone antagonist(flutamide) blocks ovulation and preovulatory surges ofprogesterone, luteinizing hormone and oestradiol in layinghens. Reproduction 2006;131:1109–14.

[51] Mulac-Jericevic B, Mullinax RA, DeMayo FJ, Lydon JP,Conneely OM. Subgroup of reproductive functions ofprogesterone mediated by progesterone receptor-Bisoform. Science 2000;289:1751–4.

[52] D’Haeseleer M, Simoens P, Van den Broeck W. Cell-specificlocalization of progesterone receptors in bovine ovary atdifferent stages of the oestrous cycle. Animal ReproductionScience 2007;98:271–81.