Hum. Reprod. Advance Access originally published online on January 4, 2007
Human Reproduction 2007 22(4):989-994; doi:10.1093/humrep/del491
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Characterization and depletion of leukocytes from cells isolated from the pre-ovulatory ovarian follicle
1 Department of Obstetrics and Gynecology 2 Institute of Immunology, Rikshospitalet-Radiumhospitalet Medical Center, Oslo, Norway
3 To whom correspondence should be addressed at: Department of Obstetrics and Gynecology, Rikshospitalet-Radiumhospitalet Medical Center, 0027 Oslo, Norway. Tel: +47 2307 0219; Fax: +47 2307 2940; E-mail: peter.fedorcsak{at}klinmed.uio.no
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Abstract |
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BACKGROUND: Cells isolated from the periovulatory ovarian follicle are often used as a model of ovarian steroidogenesis and corpus luteum formation. The follicular fluid-derived cell (FFDC) population is, however, heterogeneous and in addition to granulosa–lutein cells, non-steroidogenic cells are also present. These non-steroidogenic cells, especially the immune cells, may have important biological functions in this model. Here, we describe a method to isolate FFDC, characterize the phenotype of the immune cells and deplete immune cells from FFDC.
METHODS AND RESULTS: Follicular fluid aspirated transvaginally during IVF was clarified by centrifugation and enzymatic dispersion, labelled for leukocyte-specific markers and analysed by flow cytometry. Leukocytes constituted 22% of FFDC and expressed macrophage/dendritic cell, monocyte and lymphocyte markers. Leukocytes were depleted with anti-CD45-conjugated immunobeads, resulting in an FFDC population with <1.9% leukocytes. Leukocyte-containing FFDC secreted more interleukin-8 in culture than leukocyte-depleted FFDC.
CONCLUSION: Leukocyte-depleted FFDC may serve as a useful model to study the interaction of immune cells and luteinizing cells during corpus luteum formation.
Key words: flow cytometry/follicular fluid/granulosa–lutein cells/leukocytes
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Introduction |
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Cells isolated from the pre-ovulatory ovarian follicle, usually in conjunction with follicle aspiration for assisted reproduction treatment (ART), have been an often-used model in studies on various aspects of ovarian function. Primary cultures of follicular fluid-derived cells (FFDC) have served, among others, in experiments concerning: gonadotrophin-induced gene regulation (Sasson et al., 2004
), release of non-steroid signalling molecules by FFDC (Karström-Encrantz et al., 1998), effects of toxins on steroid synthesis (Barbieri et al., 1986) and the pathophysiology of polycystic ovary syndrome (Fedorcsák et al., 2000). An important, but often overlooked, aspect of this model is that FFDC is a heterogeneous cell population; in addition to luteinizing granulosa cells that constitute the inner lining of the pre-ovulatory follicle, the follicular aspirate also contains immune cells and cells derived from the vaginal epithelium, connective tissue and ovarian stroma. The presence of these non-steroidogenic cells in follicular fluid samples derived during IVF is probably inevitable since the follicles are aspirated transvaginally. When the aspiration needle is being inserted towards the follicle, tissue pieces along the needle's track and cells derived from the accompanying bleeding are also collected in the aspirate.
The presence of immune cells in FFDC is particularly important, since these may directly influence granulosa–lutein(GL) cell function. Leukocytes may constitute, depending on the separation method, between 15 and 75% of the total FFDC (Beckmann et al., 1991;Lachapelle et al., 1996; Enien et al., 1998). Phenotypic characterization based on histochemical staining or leukocyte-specific immunomarkers has revealed the presence of macrophages, monocytes, granulocytes and lymphocytes among these cells (Lachapelle et al., 1996; Smith et al., 2005). Immune cell contamination may arise due to bleeding during follicle aspiration, but according to recent findings bleeding alone cannot entirely account for leukocyte contamination (Smith et al., 2005), probably indicating that tissue-resident cells also contaminate FFDC (Suzuki et al., 1998). The functional significance of leukocytes in FFDC is indicated by experiments finding that leukocyte depletion increases progesterone secretion by FFDC (Beckmann et al., 1991), and that leukocyte-derived cytokines regulate steroidogenesis in cultures of FFDC (Fukuoka et al., 1992) and isolated luteal cells (Castro et al., 1998).
In order to control the effects of leukocyte contamination in experiments on cultured FFDC, several methods have been proposed for leukocyte depletion. On the basis of their distinct physical characteristics, lymphocytes can be removed by exposure to high-salt solutions, which increase cellular density and allow depletion by density gradient centrifugation (Beckmann et al., 1991). Another method takes advantage of a selective adherence of leukocytes to the surface of cell culture plastic vessels; macrophages adhere early, whereas lymphocytes are non-adherent (Beckmann et al., 1991; Sasson et al., 2004). In addition, immunobead separation using leukocyte-specific antibodies conjugated to paramagnetic beads has also been used for leukocyte depletion (Arici et al., 1996; Smith et al., 1997; Enien et al., 1998; Salmassi et al., 2005). On the basis of these precedents, we explored a method for FFDC isolation that allows phenotypic characterization of contaminating leukocytes by flow cytometry and efficient depletion of leukocytes by density gradient centrifugation and anti-CD45-conjugated paramagnetic beads. Since the common leukocyte antigen CD45 is a pan-leukocyte marker expressed by all bone marrow-derived cells except erythrocytes and platelets, this method has the potential to target all leukocytes in the FFDC sample.
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Materials and methods |
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Isolation and processing of FFDC
Follicular fluid was aspirated transvaginally, under ultrasound guidance during follicle puncture, for ART (n = 25). Male factor infertility was the treatment indication for the participating couples. Pituitary down-regulation with GnRH agonist, ovarian stimulation with recombinant FSH and ovulation induction with hCG have been described earlier (Fedorcsak et al., 2004
). Women gave written informed consent, and the protocol of this study was approved by the Regional Committee for Medical Research Ethics, Health Region South (protocol no. 2005/01). After isolation of cumulus–oocyte complexes, follicular fluid was pooled and centrifuged at 300 g for 8 min. The pellet was exposed to hemolysis buffer (150 mM NH4Cl, 10 mM NaHCO3, 0.1 mM EDTA pH 7.2), clarified by centrifugation and washed once in Hank's buffered salt solution (HBSS; Invitrogen, Carlsbad, CA, USA). The pellet was then exposed to a mixture of collagenase (0.28 mg ml–1), hyaluronidase (0.23 mg ml–1), DNase (0.38 mg ml–1) and 0.8 mg ml–1 bovine serum albumin (BSA) in Phosphate-buffered saline (PBS) for 30 min at 37°C (reagents from Sigma–Aldrich, Oslo, Norway). After enzymatic digestion, the cellular pellet was dispersed by tituration through a flame-pulled Pasteur pipette, and filtered through a 40 µm mesh (CellStrainer, BD, Franklin Lakes, NJ, USA).
Density gradient centrifugation
Single cell suspension of FFDC was layered onto a discontinuous density gradient consisting of 90, 45 and 22.5% layers of Percoll (Amersham Biosciences, Uppsala, Sweden) in HBSS, and centrifuged at 1000 g for 20 min. The layer in the interphase between 45 and 22.5% Percoll was collected, washed in HBSS and layered over 100% fetal calf serum (FCS; Invitrogen) and centrifuged at 250 g for 15 min to remove platelets. The final pellet was dissolved in Dynal buffer (2% FCS, 2 mM EDTA in PBS) and subjected to immunobead leukocyte depletion.
Immunobead depletion of leukocytes
FFDC dispersed in Dynal buffer (2 x 106 cells per ml buffer) dispersed in 1 ml Dynal buffer were mixed with 4 x 107 paramagnetic beads conjugated with mouse anti-CD45 IgG2a (Dynal Biotech, Oslo, Norway) and rotated for 1 h at 4°C. Beads were captured according to the manufacturer's instructions and the remaining cells were washed once in Dynal buffer.
Aliquots were removed after the steps of (i) enzymatic dispersion, (ii) density gradient centrifugation and (iii) leukocyte depletion, and each of these cell samples were examined for cell viability (using differential staining with a mixture of acridine orange and ethidium bromide), labelled for flow cytometry or seeded on the culture surface. Each experiment was replicated at least three times.
Immunoflow cytometry
Aliquots of FFDC were mixed with fluorochrome-conjugated antibodies raised against various leukocyte markers and incubated for 30 min on ice. Antibodies were purchased from BD, Diatec (Oslo, Norway), RnD Systems (Minneapolis, USA) and Dako (Glostrup, Denmark), and were used in the following combinations: (1) IgG1-FITC/IgG1-PE/IgG1-APC (all from BD); (2) CD4-FITC (BD)/CD3-PE (Diatec) /CD45-APC (Dako); (3) CD15-FITC (BD)/CD20-PE (Diatec)/CD45-APC; (4) DC-SIGN-FITC (RnD)/MHCII-PE (Diatec)/CD45-APC; (5) HLA-DR-FITC (BD)/CD11c-PE (BD)/CD45-APC; and (6) CD45-FITC (BD)/CD14-APC (Diatec). The final dilution of antibodies from Dako and Diatec were 1:10, whereas those from BD and RnD were 1:20. The cells were washed and exposed to 2 µg ml–1 propidium iodide immediately before analysis with a FACS calibur instrument (BD) and the CellQuest software (BD). Necrotic and apoptotic cells that stained with propidium iodide were excluded from the data set.
In order to examine whether enzymatic dispersion interfered with detection of cell surface-associated markers, peripheral blood mononuclear cells were exposed to dispersion enzymes under identical conditions as the FFDC, labelled with the leukocyte markers, and analysed with flow cytometry. Detection of leukocyte markers was not influenced by the presence of hyaluronidase, DNase and collagenase (data not shown).
Some aliquots of FFDC were fixed in 4% paraformaldehyde for 5 min, neutralized with 0.1 M glycine in PBS and cytospun onto glass slides. The slides were blocked with 1% BSA in PBS and exposed to mouse monoclonal anti-CD45 IgG1 (1:100 dilution, Dako) or negative control IgG1 (1:100 dilution, Dako), followed by Cy3-conjugated anti-mouse secondary antibody (1:600 dilution, The Jackson Laboratory, Bar Harbor, Maine, USA). Finally, slides were stained with Nile Red (Sigma) and nuclei were counterstained with bisbenzimide. For a positive control of anti-CD45 labelling, the human myelomonocytic leukemia cell line THP-1 (DSMZ, Braunschweig, Germany) was cultured on glass coverslips in the presence of phorbol myristate acetate (PMA, 50 ng ml–1) and was processed as above. Labelled cells were viewed under a Zeiss epifluorescent microscope using a wide band path filter (WBPF, Vysis, Des Plaines, IL, USA), where Nile Red-specific flow cytometry appeared orange, whereas Cy3-specific flow cytometry appeared red. In order to illustrate the findings, serial images were captured using a monochrome digital camera (SPOT, Diagnostic Instruments, Sterling Heights, MI, USA) and a filter set (red, blue and WBPF, Vysis). The images were reconstituted in Photoshop (version 7.0, Adobe, San Jose, CA, USA), where the WBPF channel was pseudo-coloured orange, the red channel was subtracted from the orange channel and the three channels were merged.
Cell culture
FFDC were seeded on gelatin-coated 24-well plates or on 35 mm Petri dishes (Nunc, Roskilde, Denmark) in Dulbecco's modified Earl's medium with Ham's F12 (DMEM/F12, Invitrogen) with 10% FCS. After an overnight incubation (at 37°C in 5% CO2 in air) that allowed attachment and spreading, cells were extensively washed with serum-free DMEM/F12 and incubated for additional 48 h in serum-free DMEM/F12. The conditioned media was removed and stored at –80°C until further assays. The attached monolayer was washed twice with PBS, dissolved in 150 mM NaCl, 50 mM HEPES, 1.5 mM MgCl2, 10% glycerol, 1% Triton-X, 100 mM NaF, 10 mM Na4P2O7, 1 mM PMSF, 1 mM Na-ortho-vanadate and 10 µg ml–1 aprotinin on ice and the cellular protein content was estimated with the BCA assay (Sigma).
Western blot
Conditioned media of leukocyte-depleted and leukocyte-rich FFDC were concentrated 10-fold by lyophilization and re-dissolution in distilled water. The samples were further diluted with water in order to normalize the total cellular protein content of the parallel samples. Ten microliter of this solution was mixed with 2 x reducing sample buffer, boiled, resolved on a 4–20% continuous gradient gel (Bio-Rad, Hercules, CA, USA) and blotted onto a PVDF membrane (Bio-Rad). The membrane was incubated with mouse monoclonal anti-human interleukin (IL)-8 antibody (1:1000 dilution; Sigma) overnight, followed by incubation with anti-mouse peroxidase-conjugated IgG (1:2000 dilution; GE Healthcare, Uppsala, Sweden) for 90 min. The immunocomplexes were detected using enhanced chemiluminescence (GE).
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Results |
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Characterization of leukocytes in FFDC
FFDC were subjected to hemolysis, enzymatic and mechanical dispersion and filtration, labelling for leukocyte markers and analysis with flow cytometry (Figure 1 A–D). The mean proportion of CD45+ cells was 22% (range 11–32%, n = 4), which were further characterized as follows: 7.7–25.8% were CD3+ T lymphocytes (herein 4.9–16.3% CD3+CD4+ T helper cells) and 0.6–2.8% were CD20+ B cells. Myeloid cells expressing CD11c (2.3–5.0%) consisted of granulocytes (CD15+, 1.0–2.5%) and monocytes (CD14+, 0.5–2.9%). Macrophages/dendritic cells were identified by co-expression of dendritic cell-specific ICAM-3-grabbing non-integrin (DC-SIGN) and MHC-II (0.3–1.9%).
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To exclude the possibility that GL cells expressed leukocyte markers, FFDC were labelled with Nile Red, which stains intracellular lipid droplets that are characteristic of luteninizing cells. Nile Red positive cells were invariably negative for CD45 (Figure 1E). Furthermore, when FFDC were labelled with anti-CD45 together with other leukocyte markers, CD45 cells were uniformly negative for CD3, CD4, CD11c, CD14, CD15, DC-SIGN, MHC-II and HLA-DR (Figure 1A–D).
Density gradient centrifugation and immunobead depletion
Density gradient centrifugation resulted in three distinct cellular layers in the interphase among 0–22.5, 22.5–45, and 45–90% Percoll layers. The 0–22.5% interphase mainly consisted of anuclear debris and large flattened squamous epithelial cells with picnotic nuclei, whereas the 45–90% interphase was abundant in lymphocytes, therefore these layers were discarded. The 22.5–45% interphase contained the GL cells and an average of 28.7% CD45+ cells (range 3.2–68.7%, n = 24). Compared with the parallel samples that were not exposed to density gradient centrifugation, the proportion of DC-SIGN+MHC-II+ cells was increased 7-fold, CD15+ cells 4.5-fold, CD20+ cells 2.4-fold and CD14+ cells 2.4-fold. The proportion of CD3+ cells was slightly decreased (0.9-fold, n = 2).
In order to deplete immune cells, FFDC purified on Percoll gradient were mixed with anti-CD45-conjugated paramagnetic beads, and CD45+ cells bound to the beads were removed. After immunodepletion, the final sample contained 1.9% CD45+ cells (mean, range 0.4–6.3%; n = 11; Figure 2) and had 87% viability (mean, range 76–97%, n = 9). The total number of viable cells thus obtained was 2.5 x 106 (mean, range 0.4 x 106 –12.9 x 106, n = 16).
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Interleukin-8 secretion by FFDC
In order to demonstrate that leukocyte depletion efficiently reduced the biological effect of leukocytes in cultured FFDC, leukocyte-rich and leukocyte-depleted FFDC were cultured for 48 h and IL-8 secretion was estimated by SDS–PAGE electrophoresis and Western blot of conditioned media. Compared with leukocyte-rich FFDC, IL-8 secretion by leukocyte-depleted FFDC was reduced (Figure 3).
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Discussion |
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In this study we evaluated a stepwise method to efficiently deplete leukocytes from cells isolated from the pre-ovulatory human follicular fluid aspirated during IVF. The method comprised (i) enzymatic and mechanical dispersion of the cells into a single cell suspension, which made the sample accessible for, but did not interfere with, phenotypic characterization of leukocytes by flow cytometry; (ii) density gradient centrifugation that enriched GL cells along with the leukocytes of similar density characteristics and (iii) depletion of leukocytes with anti-CD45-conjugated paramagnetic Dynal beads. After immunodepletion, FFDC had low leukocyte contamination, which was confirmed by flow cytometry.
We found that leukocytes constitute a high proportion (17–29%) of dispersed FFDC, confirming earlier findings that leukocyte contamination is an important feature of IVF samples (Lachapelle et al., 1996; Smith et al., 2005). Both circulating leukocytes (lymphocytes, monocytes, and granulocytes) and tissue-resident immune cells (macrophages/dendritic cells) were observed in FFDC, suggesting that these immune cells originate both from a direct bleeding during aspiration and from tissue pieces that are collected in the aspirate. Therefore, it is likely that other non-steroidogenic cells of the follicle wall and ovarian stroma are also present in the dispersed FFDC. Indeed, after a long-term culture (>2 weeks) of FFDC, proliferation of spindle-like cells (putative fibroblasts) and cells forming cobblestone-line colonies (putative endothelial or epithelial cells) are occasionally observed (data not shown). The most probable source of immune cells in the FFDC is the aspiration-induced tissue damage, as the follicular basement membrane apparently inhibits leukocyte transmigration into the folliculor fluid before ovulation, indicated by the absence of leukocytes in the granulosa layer of pre-ovulatory follicles (Takaya et al., 1997; Chang et al., 1998; Suzuki et al., 1998). Nonetheless, it has been claimed that leukocytes, such as CD3+ T lymphocytes and CD68+ macrophages (Piccinni et al., 2001) or CD56+ natural killer cells (Lukassen et al., 2003), occasionally enter the pre-ovulatory follicular fluid. Due to the sampling with transvaginal aspiration, however, the source of these cells could not be ascertained by these studies.
After ovulation, the follicular basal membrane completely breaks down and allows leukocytes, connective tissue and capillaries to invade the collapsing folds of the follicle, and establish a dense vascular network supplying the luteinizing steroidogenic cells. This process presumably requires a close interaction between GL cells and other cell types, in particular luteal leukocytes. Indeed, GL cells may release chemokines that attract leukocytes into the forming corpus luteum (Wong et al., 2002; Kryczek et al., 2005), whereas leukocytes secrete cytokines that regulate steroidogenesis and are probably also involved in vascularization, corpus luteum maintenance and luteolysis (Bukulmez and Arici, 2000; Pate and Landis Keyes, 2001). Interpretation of cell culture experiments on GL cell–leukocyte interactions have been difficult, however, especially when follicular fluid samples derived during IVF are used with insufficient removal of leukocytes. Notably, the technical bias due to leukocyte contamination had not been resolved by density gradient centrifugation during cell separation, since this method does not result in proper separation of leukocyte subsets with density characteristics similar to GL cells.
IL-8 is a member of the CXC chemokine family and attracts leukocytes, predominantly neutrophils, to the inflammation site and presumably contributes to leukocyte accumulation in the corpus luteum (Townson and Liptak, 2003).
The putative major source of IL-8 in the corpus luteum are the GL cells, and their IL-8 secretion is regulated by hCG, TNF-
and IL-1 (Arici et al., 1996; Zeineh et al., 2003). Our findings indicate that IL-8 secretion by the co-cultured GL cells and leukocytes is considerably enhanced when compared with the secretion by GL cells alone, suggesting that either IL-8 is predominantly released by leukocytes, or leukocytes greatly stimulate IL-8 secretion by GL cells. In either case, chemokine release and leukocyte accumulation in the corpus luteum may critically depend on ovarian leukocytes and their interaction with the GL cells.
In conclusion, immunobeads can efficiently deplete immune cells from FFDC and have, indeed, been successfully used to study GL cell-specific expression of IL-8 (Arici et al., 1996), macrophage colony-stimulating factor and its receptor (Salmassi et al., 2005), hCG binding (Enien et al., 1998) and steroidogenic enzyme expression (Smith et al., 1997). We propose that the present method for isolation and processing of FFDC may also be useful when studying GL cell–leukocyte interactions during corpus luteum formation. The dispersed FFDC can be analysed with flow cytometry and the depletion of leukocytes allows the comparison of leukocyte-rich and leukocyte-depleted cultures of FFDC. This method would also be useful in co-culture experiments where leukocyte-depleted FFDC are mixed with specific leukocyte subsets derived from peripheral blood.
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Acknowledgement |
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This study was supported by grants from Norske Kvinners Sanitetsforening and Medinnova AS.
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References |
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Submitted on June 15, 2006; resubmitted on October 30, 2006; resubmitted on November 28, 2006; accepted on December 1, 2006.
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