Cyclic changes of granulocyte colony-stimulating factor (G-CSF) mRNA in the human follicle during the normal menstrual cycle and immunolocalization of G-CSF protein

K. Yanagi1, S. Makinoda1,4, R. Fujii1, S. Miyazaki1, S. Fujita1, H. Tomizawa1, K. Yoshida1, T. Iura1, T. Takegami2 and T. Nojima3

1 Department of Obstetrics and Gynecology, 2 Medical Research Institute and 3 Department of Pathology, Kanazawa Medical University, Uchinada, 920-0293, Japan


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
BACKGROUND: Ovulation has several similarities with inflammation and is closely connected to the activity of leukocytes and inflammatory cytokines. Since granulocytes are one of the major leukocytes, we focused our attention on the presence and local production of granulocyte colony-stimulating factor (G-CSF) in the human ovary. METHODS: The presence of G-CSF protein in the follicular fluid and perifollicular tissues was examined by Western blot analysis (n = 5) and immunohistochemical staining (n = 10). The relative expression levels of G-CSF mRNA in relation to GAPDH in granulosa, theca and luteal cells during the menstrual cycle were measured by quantitative RT–PCR using TaqMan technology (n = 15). RESULTS: G-CSF protein was detected in all follicular fluid and located mainly in granulosa cells of the follicle and luteal cells. The expression level of G-CSF mRNA in the late follicular phase was 137.6 ± 18.5, which was ~10-fold greater than other phases during the menstrual cycle (P < 0.05). CONCLUSIONS: These results demonstrate that G-CSF is produced in the human follicle shortly before the ovulatory phase and may play an important role in the mechanism of ovulation.

Key words: cytokine/G-CSF/menstrual cycle/ovulation/PCR


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
The process of ovulation has been compared with an inflammatory reaction, since both involve components such as neutrophils, histamine, bradykinin, enzymes and cytokines (Espey, 1980Go, 1994Go; Brännström and Janson, 1992Go; Büscher et al., 1999Go). Among various cytokines, granulocyte colony-stimulating factor (G-CSF), a member of the CSF family, has been shown to stimulate the production, differentiation and functional activation of neutrophilic granulocytes both in vivo and in vitro (Nagata et al., 1986Go; Nomura et al., 1986Go; Metcalf, 1989Go). G-CSF is produced primarily by haemopoietic cells, although several non-haemopoietic cell types, such as osteoblasts, smooth muscle, endothelial and epithelial cells, as well as reproductive tissue cells have also been shown to produce G-CSF (Morstyn and Burgess, 1988Go; Duan, 1990Go; Brännström et al., 1994aGo; Giacomini et al., 1995Go). Women who received ovarian stimulation showed a significant increase of serum G-CSF (Hock et al., 1997Go). We have already reported (Makinoda et al., 1995Go, 1996Go) that serum G-CSF concentration significantly increases during the ovulatory phase compared with all other phases, suggesting that G-CSF may play an important role in ovulation. However, the mechanism of the increase of G-CSF and the location of G-CSF production have not been elucidated yet. To help clarify the mechanism of ovulation, the present study attempts not only to demonstrate the presence of G-CSF and its mRNA in the human follicle, but also to measure the changes of expression of G-CSF mRNA during various menstrual phases using quantitative PCR.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
Subjects and sample collection
All subjects were patients who had received treatment at Kanazawa Medical University Hospital between March 1998 and July 2002. All subjects gave their informed consent to participate in the study. The study protocol was conducted in accordance with the guidelines of the Declaration of Helsinki, as revised in 1983. The follicular fluid was collected from two subjects (aged 35 and 41 years) who had undergone hysterectomy due to uterine fibroids at late follicular phase. The follicular fluid was also collected at the time of oocyte retrieval from three subjects (aged 28, 29 and 33 years) who had received IVF and embryo transfer. Each follicular fluid was centrifuged at 3000 g for 10 min, and the supernatants were stored at –80°C until the analysis.

The ovarian tissues were taken from patients with normal menstrual cycle (n = 15; aged 37.3 ± 2.0 years, mean ± SEM) undergoing gynaecological operation with no ovarian diseases. We macroscopically distinguished follicles and the corpus luteum from the ovarian stromal tissues and carefully collected granulosa, theca and luteal cells—removing perifollicular stromal tissues with tweezers and scissors. Granulosa, theca and luteal cells were immediately frozen in liquid nitrogen and stored in –80°C until further processing for isolation of total RNA. The basal body temperature (BBT), estradiol, progesterone, LH and FSH were measured for each patient who was scheduled for the gynaecological operations. Estradiol, progesterone, LH and FSH concentrations in serum were determined by a microparticle enzyme immunoassay [EIA, Biodata S.p.A., Guidonia Montecelio (Roma), Rome, Italy] in accordance with the manufacturer's instructions. The measurable ranges of estradiol, progesterone, LH and FSH were 5–3000 pg/ml, 0.2–40.0 ng/ml, 0.5–200 mIU/ml and 0.5–150 mIU/ml respectively. The menstrual cycle was divided into early follicular (EF), late follicular (LF), ovulatory (OV) and luteal (LU) phases based on BBT, hormonal levels (Table IGo) and histological examinations.


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Table I. G-CSF mRNA by TaqMan PCR and hormonal definition of each phase
 
Ovarian tissues, including the largest follicle or corpus luteum, were collected from other patients (n = 10; aged 36.9 ± 2.5 years, mean ± SEM) and used for immunohistochemical study of G-CSF. The menstrual cycle of these patients was monitored as above and had undergone gynaecological operations without ovarian diseases.

Western blot analysis
Immunoprecipitation and Western Blot were performed to detect the presence of G-CSF protein in the follicular fluid collected from two normal subjects at late follicular phase and three IVF patients. For immunoprecipitation, 100 µl of follicular fluid was added to 15 µl of washed protein A–Sepharose beads and the appropriate goat anti-human G-CSF-specific antibody (G-CSF Ab; kindly donated by Chugai Diagnostics Science Co. Ltd, Tokyo, Japan) at 1:200 (Kiriyama et al., 1993Go). The samples were left at 4°C for 1 h and centrifuged at 2000 g for 5 min. The beads were washed with RIPA buffer (10 mmol/l Tris–HCl, pH 7.4, 1% Nonidet P40, 0.1% sodium deoxycholate, 0.1% sodium dodecyl sulphate (SDS), 0.15 mol/l NaCl, 1 mmol/l EDTA, 10 µg/ml aprotinin) five times and resuspended in sample buffer (0.125 mol/l Tris–HCl, pH 6.8, 2% SDS, 10% glycerol, 5% 2-mercaptoethanol) and boiled at 95°C for 3 min. After centrifuging for 2 min to pellet the beads, the supernatant was immediately electrophoresed using a 12.5% gradient polyacrylamide gel. After electrophoresis, proteins were transferred onto a polyvinylamide-difluoride (PVDF) membrane. The blotted membrane was incubated with the G-CSF antibody at 1:100 for 1 h at 37°C. The membrane was then treated with horse-radish peroxidase-conjugated anti-goat Ig Ab (Code No. P 0449: Dako, Glostrup, Denmark) for 1h at 37°C. Antibody binding was visualized with a 3,3'-diaminobenzidine tetrahydrochloride solution (DAB: Dojin, Kumamoto, Japan) as previously described (Takegami et al., 1994Go). We used 100 ng of recombinant human G-CSF (Pepro Tech EC Ltd, London, UK) and distilled water as positive and negative controls respectively.

Immunohistochemical staining of G-CSF protein in human ovarian tissues
Immunohistochemical staining was performed to detect the presence of G-CSF protein in the follicle or corpus luteum samples using an Envision kit (Dako, Carpinteria, CA, USA). Surgically resected ovarian tissues were quickly fixed in 10% buffered formaldehyde for 24 h and embedded in paraffin. Paraffin-embedded blocks were then cut to 4 µm thick specimens and deparaffinized by ethanol. After washing in distilled water and autoclaved at 120°C for 15 min, inhibition of endogenous peroxidase activity was accomplished by incubation in 3% H2O2 solution dissolved in absolute methanol at room temperature for 5 min. All specimens were washed in distilled water, rinsed with phosphate-buffered saline (PBS), and then incubated at 4°C overnight with mouse anti-human G-CSF monoclonal antibody (Shimamura et al., 1990Go), diluted to 1:200. Specimens were then rinsed with PBS and allowed to react with the Envision polymer (Code No. K 1491: Dako, Carpinteria, CA, USA) for 30 min at room temperature. After rinsing with PBS, peroxidase colour visualization was carried out with 30 mg of DAB dissolved in 150 ml of PBS and added to 10 µl of a 30% H2O2 solution. Nuclear counter staining was carried out with Harris haematoxylin for 3 min before mounting.

RNA extraction
Total RNA was extracted from human granulosa, theca and luteal cells using the AGPC (acid guanidium–phenol–chloroform) method (Chomczynski and Sacchi, 1987Go). Eighty to 100 mg of frozen tissue was homogenized with a nucleic acid extraction reagent (Isogen: Nippon Gene Inc., Osaka, Japan) and chloroform (0.2 ml/1 ml Isogen) and then centrifuged at 12 000 g for 3 min. The aqueous layer containing RNA was collected and further precipitated with isopropanol by centrifugation at 12 000 g for 20 min at 4°C. Pellets were washed with 70% cold ethanol, air-dried and resuspended in RNase-free water. Extracted RNA was quantified by measuring absorption at 260 nm, and its purity was confirmed by electrophoresis. Two µg of RNA were reverse-transcribed into cDNA by incubation in the mixture including 0.5 µl of 100 µmol/l G-CSF antisense primer (5'-TCATCCCAGTGCCCATTGCAGA-3') or 1 µl of 10 µmol/l GAPDH antisense primer (5'-GAAGATGGTGATGGGATTC-3'), 1 µl of 10 mmol/l dNTP, 0.5 µl of 20 IU/µl HPRI (human placenta ribonuclease inhibitor), 2 µl of 5xRT buffer with MgCl2 and 0.5 µl of 7 IU/µl AMV RTase (TaKaRa Shuzo Co., Ltd., Otsu, Japan). The mixtures were adjusted to 10 µl with RNase-free water. RT reactions were performed at 42°C for 1 h, followed by heating to 95°C for 5 min to inactivate the enzyme, and stored at 4°C until the PCR analysis.

Real-time quantitative PCR
Principal aspects of real-time quantitative PCR were previously described (Lee et al., 1993Go; Livak et al., 1995Go; Heid et al., 1996Go). Real-time quantitative PCR analysis was performed by use of a PE Applied Biosystems 7700 Sequence Detector (PE Applied Biosystems, Inc., Foster, CA, USA), which was essentially a combined thermal cycler/fluorescence detector with the ability to optically monitor the progress of individual PCR reactions. In addition to the two amplification primers used in conventional PCR, this system also included a dual-labelled fluorogenic hybridization probe. For real-time quantitative PCR, TaqMan EZ RT–PCR Kit (PE Applied Biosystems) was used. The G-CSF TaqMan system consisted of the amplification primers: G-CSF F (5'-TCTGAGTTTCATTCTCCTGCCTG-3'), G-CSF R (5'-ATTTACCTATCTACCTCCCAGTCCAG-3') and a dual-labelled fluorescent TaqMan probe [5'-(FAM)AGCAGTGAGAAAAAGCTCCTGTCCTCCC(TAMRA)-3']. The control GAPDH (endogenous control) (Ercolani et al., 1988Go), GAPDH F (5'-GAAGGTGAAGGTCGGAGTC-3'), GAPDH R (5'-GAAGATGGTGATGGGATTC-3') and the probe [5'-(JOE)CAAGCTTCCCGTTCTCAGCC(TAMRA)-3'] were made by PE Applied Biosystems. G-CSF TaqMan probe and G-CSF primers were synthesized by Hokkaido System Science, Inc., Sapporo, Japan. TaqMan amplification reaction was set up in a reaction volume of 50 µl, with each reaction volume containing 10 µl of TaqMan EZ Buffer; 0.5 µl of 200 nmol/l of each amplification primer; 2 µl of 100 nmol/l of the corresponding TaqMan probe; 6 µl of 3 mmol/l manganese acetate, 1.5 µl of 300 µmol/l each dATP, dCTP, dGTP and dUTP; 2 µl of 0.1 IU/µl rTth DNA polymerase; 0.5 µl of 0.01 IU/µl AmpErase UNG; and 8 µl of extracted cDNA. RNase-free water was then added to bring the final volume to 50 µl. Thermal cycling parameters were 2 min at 50°C, 10 min at 95°C followed by 40 cycles of 15 s at 95°C; and 1 min at 60°C. Identical thermal profiles were used for both the G-CSF and GAPDH systems. To prevent PCR contamination, strict precautions and anti-contamination measures were taken during the real-time quantitative PCR (Kwok and Higuchi, 1989Go). As a positive control, a series of diluted G-CSF plasmid DNA was used in PCR.

The ratio of granulosa, theca and perifollicular stromal cells used in PCR
The samples used in PCR were 80–100 mg, collected macroscopically (see above). The ratio of granulosa, theca and perifollicular stromal cells of each sample was examined microscopically using the samples collected in the same manner (n = 6). Preparing five haematoxylin–eosin specimens from each sample, the numbers of granulosa, theca and stromal cells in the same amount of the specimens for PCR were counted in accordance with the size of follicles.

Statistical analysis
Analysis of variance was used for comparison of the relative expression level of G-CSF mRNA among various ovarian phases. Data are presented as the mean ± SEM. Statistical significance was tested by Fisher's protected least-significant difference test. P < 0.05 was considered statistically significant.


    Results
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
The presence of G-CSF protein in the follicular fluid
Immunoprecipitation and Western blot analysis using anti-G-CSF antibody revealed a specific band of 19 Mrx10–3 kDa in the follicular fluids from all five subjects (Figure 1Go). This band was consistent with G-CSF protein based on the amino acid sequence. No specific band was observed in the negative control (N, distilled water). Bands H and L were IgG-H and -L chains derived from goat immunoglobulin (lanes 1–5, N and P) and human immunoglobulin (lanes 1–5).



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Figure 1. Western blot analysis of G-CSF protein immunoprecipitated from follicular fluids. 100 µl of follicular fluid from each subject (n = 5) (lanes 1 and 2; normal late follicular phase: Lanes 3–5; IVF patients) was immunoprecipitated with anti-G-CSF antibody and separated on 12.5% SDS–PAGE. P = positive control (recombinant human G-CSF without immunoprecipitation). N = negative control (distilled water); M = size marker. As indicated by the arrow, G-CSF protein was detected by Western blot using goat anti-human G-CSF antibody. The bands of H and L were derived from goat and human immunoglobulin in lanes 1–5 and goat immunoglobulin only in Pand N.

 
Immunohistochemical staining of G-CSF protein in human ovarian tissues
Immunohistochemical staining using anti-human G-CSF monoclonal antibody showed staining mainly in granulosa and luteal cells (Figure 2A–DGo). The staining was also observed at surface epithelium (Figure 2EGo). These signals detected with the anti-G-CSF antibodies were specific, since no signals were observed on the negative control (Figure 2FGo).



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Figure 2. Immunohistochemical staining of the largest follicular walls of early follicular (A), late follicular (B), ovulatory (C) phase, corpus luteum (D) and ovarian surface epithelium (E). Negative control (the follicular wall at ovulatory phase) is shown in (F). G = granulosa cells; T = theca cells.

 
G-CSF mRNA expression in the granulosa, theca and luteal cells during the menstrual cycle
Since we have detected G-CSF mRNA by regular RT–PCR in all follicular tissues (data not shown), we compared the quantity of G-CSF mRNA of granulosa, theca and luteal cells during the normal menstrual cycle by quantitative RT–PCR using real-time TaqMan technology. The relative expression levels of G-CSF mRNA are summarized in Figure 3Go. The relative level of G-CSF mRNA in the early follicular phase was only 13.6 ± 1.7. It increased ~10-fold to 137.6 ± 18.5 in the late follicular phase. The expression levels decreased back to the level of early follicular phase or below in the ovulatory and luteal phase.



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Figure 3. Quantitative analysis of G-CSF mRNA using TaqMan system in granulosa, theca and luteal cells during the various menstrual phases. Each bar shows mean ± SEM of the G-CSF mRNA/GAPDH mRNA ratio. EF = early follicular phase; LF = late follicular phase; OV = ovulatory phase; LU = luteal phase.

 
The ratio of granulosa, theca and perifollicular stromal cells used in PCR
The ratio of granulosa, theca and perifollicular stromal cells used in PCR was shown in Table IIGo. Since each sample used in PCR was 80–100 mg, the ratio of granulosa and theca cells is relatively higher in large follicles than in small and middle-sized follicles (P < 0.05).


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Table II. The ratio of granulosa, theca and perifollicular stromal cells in the macroscopically collected specimens (n = 30 specimens/6 ovaries)
 

    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
The ovary is a site for interaction between the endocrine and immune systems through leukocytes and their secreted products, cytokines. Convincing data showing the intimate relationship between cytokines and leukocytes in follicular development, ovulation and luteal function is available (Norman and Brännström, 1996Go; Bukulmez and Arici, 2000Go).

A variety of leukocyte subtypes has been described in the ovary, including neutrophils, macrophage/monocytes and lymphocytes (Brännström and Norman, 1993Go; Brännström et al., 1993aGo). In the pre-ovulatory or large follicle, the majority of leukocytes is located in the theca, with a predominance of neutrophils and macrophages, as identified by immunostaining with specific antibodies (Brännström et al., 1993aGo; Brännström and Norman, 1993Go). Addition of leukocytes, as compared with a cell-free medium, to a perfused rat ovary model has been shown to significantly enhance the number of ovulations (Hellberg et al., 1991Go). On the other hand, depletion of neutrophils in vivo by specific monoclonal antibodies in the rat has been shown to decrease the number of ovulations (Brännström et al., 1995aGo).

The inflammatory cytokines in serum are elevated during ovulation. Some cytokines are secreted in a cyclic fashion from the ovary (Brännström et al., 1995bGo), while some cytokines reproduce the pro-ovulatory and pro-inflammatory effects seen in the ovary during the final stages of follicular rupture. For instance, interleukin (IL)-1ß and tumour necrosis factor-{alpha} promote ovulation in combination with LH in the perfused ovary (Brännström et al., 1993bGo; Takehara et al., 1994Go) and cytokine antagonists diminish the number of ovulations (Simón et al., 1994Go). Thus, it is now well recognized that cytokines play an important role in many physiological events, including many functions in endocrinology and ovulation (Kennedy and Jones, 1991Go; Adashi, 1992Go).

In regard to the role of CSF families on ovulation, many research studies have focused on M-CSF (Nishimura et al., 1998Go; Shinetugs et al., 1999Go; Kawano et al., 2001Go). Although the importance of macrophages in normal ovary was often reported (Best et al., 1996Go; Suzuki et al., 1998Go), granulocytes were also present in high numbers in the follicular wall, especially in the thecal layer at ovulation (Brännström et al., 1994bGo). Moreover, it was reported that not M-CSF but G-CSF has significant correlation to leukocytosis by gonadotrophin stimulation (Hock et al., 1997Go). Therefore, research on G-CSF on ovulation has practical value. G-CSF stimulates a variety of responses in mature neutrophils, including prolonged survival, phagocytosis and superoxide production (Asano, 1991Go). We have recently demonstrated that serum concentrations of G-CSF are significantly increased during ovulation in women with a normal menstrual cycle (Makinoda et al., 1995Go, 1996Go). In addition, the cultured normal ovarian surface epithelial cells produced more G-CSF than other cytokines (Ziltener et al., 1993Go). However, the precise location and timing of G-CSF production in the human ovary in vivo remained unclear.

In this study, the presence of G-CSF protein in the human pre-ovulatory follicle was identified in all samples tested by Western blot analysis using anti-G-CSF antibody. Samples were taken from normal physiological menstrual cycles as well as IVF cycles. Although local production in the human follicle of both IL-1 and IL-6 has been reported (Hurwitz et al., 1992Go; Machelon et al., 1994Go), our previous study (Makinoda et al., 1996Go) revealed that only G-CSF among the cytokines (IL-1ß and IL-6) showed significant increases at the ovulatory phase. For these reasons, we postulated that G-CSF might be produced in the human follicle and attempted to elucidate which cells in the follicle produce G-CSF protein using immunohistochemical methods.

Immunohistochemical staining using anti-human G-CSF monoclonal antibody showed high staining mainly in granulosa cells before ovulation and luteal cells after ovulation. These results indicate that G-CSF protein is located mainly in granulosa cells in the human follicle as well as the normal ovarian surface epithelial cells (Ziltener et al., 1993Go). Since granulocyte was present in high numbers especially in the thecal layer at ovulation (Brännström et al., 1994bGo), G-CSF produced by granulosa cells might induce granulocyte infiltration in the thecal layer.

We then performed quantitative PCR using real-time TaqMan technology to determine the relative abundance of G-CSF mRNA in granulosa, theca and luteal cells at each of the various reproductive stages in the human ovary. Real-time PCR is a rapid, reproducible and highly sensitive method suitable for quantitative analysis of G-CSF mRNA in human ovarian tissue. The quantitative data showed that the expression level of G-CSF mRNA in the late follicular phase was significantly higher than levels in other phases, although the contamination of ovarian stromal cells was observed at the rate of ~20% and the contribution of macrophages in this study was not known. Our immunohistochemical and real-time PCR findings demonstrate that G-CSF is produced mainly at granulosa cells of the human follicle before ovulation.

As to the biological significance of G-CSF in the ovary, it is possible that this cytokine may act to recruit leukocytes into the ovary at the time of ovulation and subsequently regulate their behaviour and their mediators. It is well known that the number of leukocytes, particularly the number of granulocytes, increases during pregnancy (Makinoda et al., 1995Go). Supplementation of leukocytes in the rat ovarian perfusion system has been shown to enhance LH-induced ovulation (Hellberg et al., 1991Go). G-CSF induced elevations in leukocytes might accelerate the ovulatory mechanism. Considering the hormonal backgrounds of the patients, high estrogen before LH surge seems to be the important factor in high levels of G-CSF mRNA. However, the mechanism of regulation of G-CSF production in ovulation is still unclear at present. Further investigations are necessary.

In conclusion, we have demonstrated that G-CSF is produced in the human ovary, and that the expression of G-CSF mRNA is more pronounced during the late follicular phase than during other phases. Our results suggest that G-CSF may play an important paracrine or autocrine role in the human ovulatory process.


    Acknowledgements
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
We thank Prof. Shogo Katsuda of Kanazawa Medical University for his technical advice and Dr Karen Wang, MD, Albert Einstein University for reviewing this manuscript. This work was supported in part by a grant-in-aid for general scientific research (No. 10671574) from the Ministry of Education, Science, Sports and Culture of Japan and Grants for Project Research from the High Technology Research Center of Kanazawa Medical University (P99-3 and H00-3).


    Notes
 
4 To whom correspondence should be addressed. E-mail: mak{at}kanazawa-med.ac.jp Back


    References
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Acknowledgements
 References
 
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Submitted on April 2, 2002; accepted on August 6, 2002.