1 Service d'Histologie, 2 Unité Inserm U327, 3 Service de Virologie, 4 Service de GynécologieObstétrique, CHU Bichat-Claude-Bernard, and 5 Service de GynécologieObstétrique, Hôpital Rothschild, Paris, France
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Abstract |
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Key words: apoptosis/hepatitis C virus/human granulosa cell/IVF
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Introduction |
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HCV tropism is controversial and the mechanisms of virus entry into cells remain unknown. It was recently advanced that the family of low-density lipoprotein (LDL) receptors might serve as HCV receptors (Agnello et al., 1999). LDL receptors are present on the plasma membrane of granulosa cells (GC) and they are up-regulated by GnRH agonists, human FSH or HCG (Bramley et al., 1987
; Foster et al., 1993
). In addition, each maturing ovarian follicle is perfused by a follicle artery, thus suggesting intimacy among PBMC, HCV, the oocyte and its cumulus. Primary follicular fluid (FF) is produced by the antral follicle until shortly before ovulation and essentially represents a plasma transudate plus GC secretions (Gosden et al., 1988
). We recently confirmed that isolated human GC were sensitive to apoptosis induced by IFN
and an agonistic anti-Fas antibody (Benifla et al., 2002
), as previously reported by Quirk et al. (Quirk et al., 1995
). Thus, we hypothesize that HCV could induce GC apoptosis, by acting directly on GC or by modifying local concentrations of pro-apoptosis substances synthesized by PBMC.
The aim of this study was to test the hypothesis that HCV+ women, recently included in our centre for assisted reproductive technologies, could present an increased spontaneous GC apoptosis percentage compared with HCV women, because of the presence of HCV in vascularized ovarian follicles.
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Materials and methods |
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GC isolation and flow cytometry
After oocyte identification and isolation from FF under a dissecting microscope, each woman's fluid was pooled (n = 12). Aliquots of FF and serum (1 ml each) from each patient were stored at 80°C for subsequent virological analysis. After FF centrifugation at 500 g for 30 min, red blood cells were removed from follicular aspirates by density gradient centrifugation through 10 ml 50% Percoll solution (Pharmacia Biotech, Uppsala, Sweden) for 30 min at 100 g GC were aspirated from the interface and washed in phosphate-buffered saline solution. PBMC were depleted by treatment with anti-CD45-coated magnetic beads (Dynabeads M-450 CD45®; Dynal, Oslo, Norway) according to the manufacturer's instructions. Luteinized GC were dissociated by trypsinization (GIBCO BRL, Glasgow, UK), which was stopped after 5 min by the addition of minimum essential medium (MEM) with Earle's salts and glutamax (GIBCO BRL), supplemented with 10% fetal calf serum (Valbiotech, Paris, France). Purified GC were assessed for apoptosis by a cell cycle analysis of DNA content, as previously described by Makrigiannakis et al. (Makrigiannakis et al., 2000). Briefly, GC suspensions were fixed in ice-cold 70% ethanol for at least 16 h and centrifuged for 5 min at 500 g. GC pellets were treated for 15 min at room temperature with RNase A (180 µg/ml; Roche, Meylan, France), stained for 30 min with propidium iodide (50 µg/ml; Sigma, Saint Quentin Fallavier, France) and analysed by flow cytometry on a EPICS XL (Beckman Coulter, Fullerton, CA, USA). Hypodiploid GC, containing <2n DNA in the cell cycle analysis profile determined by the EXPO 32 software (Beckman Coulter), were considered to be apoptotic. As a positive control, flow cytometry was applied to GC from HCV women. After purification as described above, GC were incubated for 12 h with the anthracycline anti-tumour drug, doxorubicin (100 nmol/l/ml), known to be pro-apoptotic for ovarian cells (Bellarosa et al., 2001
). The cell cycle analysis detected 85% of hypodiploid GC, permitting us to validate our protocol (data not shown).
HCV RNA extraction
HCV RNA was extracted from serum, FF and embryo incubation medium using the Amplicor HCV Specimen Preparation Kit® (Roche) according to the manufacturer's instructions. Internal controls were added to validate the extraction and amplification steps and to assure the absence of false-negative results.
HCV RNA tests
HCV RNA was detected with Cobas Amplicor HCV 2.0 (Roche), which has a sensitivity of 50 IU/ml. Serum and FF virus loads were quantified using Cobas Amplicor HCV Monitor (Roche), which has a sensitivity of 600 IU/ml. To prevent false-positive results due to cDNA contamination, dUTP-uracil-DNA-glycosylase was added to the samples.
Sperm preparation, culture and embryo transfer
Discontinuous PureSperm® (Nidacon, Göteborg, Sweden) centrifugation (90 and 45% layers) was used to separate semen samples and the 90% layer was washed in B2 UpGraded INRA medium (CCD, Paris, France) by centrifugation at 500 g for 20 min. Retrieved cumulus oocyte complexes from FF were gently washed twice in B2 UpGraded INRA medium (CCD), as routinely made in our laboratory, especially to discard blood and clots. Oocytes were inseminated with 100 000 motile sperm/ml and fertilization was verified 1820 h later, by the presence of pronuclei and polar bodies, observed under an inverted microscope after complete removal of cumulus cells surrounding the oocytes. ICSI was performed when there was severe oligoasthenozoospermia (3/6 in both groups). Embryos from ICSI or from conventional IVF were cultured for 2 more days, with development monitored daily. We recorded the mature oocyte rate, defined as the percentage of oocytes reaching metaphase II, the fertilization rate, the cleavage rate, the embryo quality (good defined as embryos with <20% cytoplasmic fragmentation and even-sized blastomeres) and the pregnancy rate. The day 2/3 culture medium from each embryo to be transferred or frozen was stored at 80°C until tested for HCV.
Statistical analysis
Values are expressed as mean ± standard deviation (SD). Wilcoxon rank sum test was used with the level of significance set at 5%. The sample size of our study was evaluated on the basis of 3.0 ± 1.5% apoptotic cells in the control group (unpublished data) and at least twice as many apoptotic cells in the HCV+ group, i.e. 6.0%, with the same SD. For a Wilcoxon rank sum test, this led to an inclusion of six subjects in each group.
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Results |
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The percentages of apoptotic GC for HCV+ and HCV groups were 3.08 ± 1.14 and 3.14 ± 1.40% respectively. The two groups did not differ significantly for this or any of the other routine IVF parameters, especially concerning the pregnancy rates, since two pregnancies occurred in each group (Table I).
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Discussion |
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However, it remains unknown if HCV status modifies ovarian follicle development during the IVF stimulation procedure and, consequently, IVF outcome. Because the results of recent studies implicated HCV in apoptosis (Hahn et al., 2000; Izuma et al., 2000
; Piazzolla et al., 2000
; Taya et al., 2000
), and it was previously demonstrated that women who became pregnant after IVF treatment had significantly lower percentages of apoptotic luteinized GC, as assessed by flow cytometry, than those who did not become pregnant despite normal basal serum FSH levels (Oosterhuis et al., 1998
), we thought it important to determine whether HCV could have an impact on the percentage of GC apoptosis. Furthermore, Nakahara et al. have reported that a better oocyte outcome from individual follicles and a higher pregnancy rate in women undergoing IVF were associated with a low GC apoptosis percentage, as estimated by fluorescence microscopy (Nakahara et al., 1997a
,b
). In this preliminary study, on a very limited number of women, we demonstrated that, despite the possible exposure of the follicular micro-environment to HCV and PBMC which might trigger apoptosis via the Fas-mediated pathway (Hahn et al., 2000
; Taya et al., 2000
), GC were not affected, as no statistically significant difference between the GC apoptosis percentages was seen as a function of HCV status. Furthermore, none of the other IVF or ovarian parameters examined differed significantly between groups. The low mean virus load in FF compared with that measured in the sera of our patients seems to indicate that HCV does not replicate in GC from FF, perhaps because the virus does not penetrate into follicular GC. Thus, the presence of HCV in follicular aspirates is probably the consequence of blood contamination during ovarian puncture. However, because GC express LDL receptors on their plasma membrane and it has been postulated that these receptors might enable virus entry into cells (Agnello et al., 1999
), the ability of GC to replicate HCV cannot be excluded and merits investigation.
In conclusion, our preliminary results suggested, for the first time to the best of our knowledge: (i) that in-vivo active chronic HCV infection does not affect follicle development and IVF outcome in women undergoing IVF; and (ii) routine IVF procedures including washing of oocytes and changing the culture media of embryos until transfer seems, based on our RTPCR results, to eliminate HCV. Thus, assisted reproduction might not contribute to increasing the risk of newborns being infected by maternal HCV, a preliminary finding that needs to be confirmed on a larger number of HCV+ women. However, the presence of HCV in FF, as demonstrated in our study, implicates a possible risk of nosocomial contamination, as recently reported by Lesourd et al. in a case of contamination by HCV during assisted reproduction (Lesourd et al., 2000). Thus, in order to prevent this risk, strict security rules (Steyaert et al., 2000
) must be employed in IVF laboratories which cater for infertile couples with HCV+ women.
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Acknowledgements |
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Notes |
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References |
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Submitted on November 10, 2001; resubmitted on December 20, 2001; accepted on March 7, 2002.