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BOR - Papers in Press, published online ahead of print January 22, 2003.
Biol Reprod 2003, 10.1095/biolreprod.102.009894
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BIOLOGY OF REPRODUCTION 68, 1982–1988 (2003)
DOI: 10.1095/biolreprod.102.009894
© 2003 by the Society for the Study of Reproduction, Inc.

Environment

Lead Reduces Messenger RNA and Protein Levels of Cytochrome P450 Aromatase and Estrogen Receptor ß in Human Ovarian Granulosa Cells1

Crystel Taupeau3, Joël Poupon5, Dominique Treton4, Aurélie Brosse3, Yolande Richard4, and Véronique Machelon2,3

INSERM Institut Paris-Sud Sur les Cytokines, Unité 355 "Maturation Gamétique et Fécondation,"3 Unité 131 "Cytokines et Immunorégulation,"4 92140 Clamart, France Laboratoire de Biochimie-Toxicologie,5 Hôpital Fernand-Widal, Paris, France


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Exposure to lead causes decreased fertility in women. In the present study, we examined the in vitro effects of lead on cytochrome P450 aromatase (P450 arom) and on estrogen receptor ß (ERß), two key proteins for the human ovary. Aromatase is required for the bioconversion of androgen to estradiol; ERß mediates estrogen effects in granulosa cells. Granulosa cells were collected from women undergoing in vitro fertilization and then cultured with 10 µM lead acetate. Using atomic absorption spectrometry, we showed that lead accumulated in cells. Aromatase activity as measured by a tritiated water production assay was significantly reduced. Using semiquantitative reverse transcription-polymerase chain reaction and Western blotting procedures, we showed that P450 arom and ERß mRNA and protein content were both significantly reduced. Adding 10 µg/ml of cycloheximide, a protein inhibitor, did not eliminate the effects of lead. The present results support the hypothesis that the action of lead on fertility in women may result, in part, from the down-regulation of P450 arom and ERß gene transcription in ovarian granulosa.

environment, estradiol receptor, follicular development, granulosa cells


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Lead is used in many industrial activities, including mining, refining, and producing lead-acid batteries. Although this heavy metal is less widely used today, it remains a significant public health problem. Lead has been widely dispersed in the environment, and it remains in the biotope. People may be exposed to lead via contaminated food or water and fuel additives [1]. The main targets of lead toxicity are the red blood cells, nervous system, and kidney [2]. It may cause changes in the reproductive system as well. A large body of literature indicates that high lead concentrations are associated with reproductive toxicity in men, including testicular tissue disruption [3, 4] as well as altered spermatogenesis and increased sperm pathologies [5–9]. Although few studies have been performed in women, severe cases of lead poisoning have been associated with sterility, miscarriage, abortion, premature delivery, and infant mortality [10–12]. Experimental studies carried out in primates showed that female rhesus monkeys treated with lead exhibited longer and more variable menstrual cycles [13]. Female rhesus monkeys that received lead acetate in drinking water for 75 mo showed a significant decrease in serum progesterone levels, indicating that lead blocks luteal function [14]. In female cynomolgus monkeys chronically exposed for up to 10 yr, blood lead concentrations were approximately 35 µg/dl, which lead to significant decrease in serum levels of LH and FSH, indicating that lead inhibits ovarian function [15].

The objective of the present study was to investigate whether lead affects cytochrome P450 aromatase (P450 arom) and estrogen receptor (ER) ß expression. The P450 arom and ERß are two key proteins that control the production of estradiol and its autocrine effects in the ovary. The product of the CYP19 gene, P450 arom catalyzes three consecutive hydroxylation reactions, converting C19 androgens to aromatic C18 estrogenic steroids [16]. It is a key enzyme for estradiol biosynthesis by ovarian granulosa cells. Follicular aromatase is detectable late in the follicular phase and is at high levels in the corpus luteum in women [17]. The P450 arom is essential for follicular maturation, oogenesis, ovulation, and normal luteal functions in females [18, 19]. The aromatase knockout (ArKO) mouse cannot synthesize endogenous estrogen [20]. These ArKO mice are infertile because of folliculogenic disruption and a failure to ovulate [21]. This phenotype is attributed to the absence of aromatase and the failure of androgen conversion into estradiol. Both ER{alpha} and ERß are present in the human ovary [22]. Although these two receptors are different gene products, they are highly similar in the ligand-binding and DNA-binding domains [23, 24]. The ERß is predominantly expressed in granulosa cells [25]. Estradiol, via ERß, regulates follicular growth and oocyte development. The ERß knockout (BERKO) female mice have a poor reproductive capacity because of folliculogenesis blockade [26]. Therefore, all these data demonstrate that P450 arom and ERß are crucial for folliculogenesis and reproductive functions.

In women, these two proteins are highly expressed in granulosa cells from preovulatory follicles [25–27]. In the present study, we used these cells, collected from patients undergoing in vitro fertilization (IVF), as a cellular model to examine the effects of lead on ovarian P450 arom and ERß expression.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Preparation

Cells were obtained from patients (age, 20–42 yr) undergoing IVF-embryo transfer therapy. The present study required no modification of routine IVF protocols. Briefly, the patients were treated for induction of multiple follicles with human recombinant hormone FSH (Gonal-F; Serono, Boulogne, France; and Puregon; Organon, Puteaux, France) or with the human menopausal hormone (75 IU of FSH and 75 IU of LH; Menogon; Ferring GMBH, Kiel, Germany) under medical hypophysectomy induced by gonadotropin-releasing hormone agonist LHRHa (Decapeptyl; Beaufour, Paris, France). Follicular fluid was aspirated from individual hyperstimulated preovulatory follicles under ultrasonographic control (diameter, >18 mm) approximately 36 h after a single injection of 10 000 IU of hCG (Organon). Informed consent for the experimental protocol and use of the cells was obtained from the patient. Cells from aspirates were pooled and centrifuged through a Percoll cushion (1:1 PBS:Percoll) to remove red blood cells. Then, cells were plated at 5 x 105 cells per 60-mm plate in Ham F12-Dulbecco modified Eagle medium (Life Technologies, Inc., Cergy Pontoise, France) supplemented with antibiotic-antimycotic mixture (Life Technologies) and 2 mM L-glutamin (Life Technologies). Cells were cultured at 37°C in an atmosphere of 5% CO2 in air. To allow cell attachment, 10% fetal bovine serum (FBS; Life Technologies) was added to basal medium for 24 h; thereafter, FBS was removed from culture media.

Cell Treatments

Controls were run in parallel in basal medium and in basal medium supplemented with 10 µM sodium acetate (Sigma-Aldrich, Saint Quentin, France). Treatments were run in basal medium supplemented with 10 µM lead acetate (Sigma-Aldrich). Using the Trypan blue exclusion test, we measured the percentage of dead cells. Dead cells were less than 1% in cultures run in the presence of 10 µM lead acetate up to 72 h. Treatments were done for 5 and 24 h after 24- and 72-h culture periods, respectively. To determine whether lead effects require de novo protein synthesis, cells were put in either the presence or absence of 10 µg/ml of cycloheximide (CHX; Sigma-Aldrich) 30 min before the beginning of the lead treatments.

Protein Assay

Protein concentration was determined by the Bradford assay [28] using the Bio-Rad Protein Assay kit (Bio-Rad, Munich, Germany) according to the manufacturer's recommendations. This assay can accurately detect 20 µg/ml in solution and a basal linear range of 20–800 µg/ml.

Lead Quantification

Cells were harvested with trypsin-EDTA 1x (Life Technologies). All glassware was washed three times with triple-distilled water and sonicated. Cellular lysates were stored at -20°C in lead-free Eppendorf microtubes. All materials used for sample preparation and lead determination were previously soaked overnight in nitric acid (Suprapur; Merck, Darmstadt, Germany) and rinsed three times with ultrapure water (resistivity, >18.2 M{Omega}; Milli-Q; Millipore, Saint-Quentin-en-Yvelines, France). Lead in lysates was measured by electrothermal atomic absorption spectrometry on a PE 5100 PC spectrometer (Perkin-Elmer, Les Ulis, France) equipped with a Zeeman background correction system, an HGA 600 graphite atomizer, an AS 60 autosampler, and a lead hollow cathode lamp. Samples (200 µl) were acidified with 15 µl of 14 M HNO3. Three deposits of 40 µl of nontreated lysate samples or one deposit of 40 µl for treated samples were placed in a graphite tube with an integrated platform (part no. B3001264; Perkin-Elmer) and supplemented with 5 µl of palladium modifier (0.05 g/L). Calibration was done using the addition-calibration method, and peak areas were used for calculations. Blank values were determined separately using the same addition technique and were subtracted from the sample results [29]. Atomization temperature was set at 1600°C. When necessary, samples were diluted in HNO3. Results (mean ± SD) were expressed as µg/g protein. Differences between groups were calculated using the Student t-test at P <= 0.01.

RNA Extraction and Polymerase Chain Reaction Procedures

Total RNA was extracted from 106 cells using the SV total RNA isolation kit (Promega France, 69260 Charbonnières, France) according to the manufacturer's instructions. From 2 µg of total RNA, mRNA was transcribed into cDNA using 200 IU of Moloney Murine Leukemia Virus Reverse Transcriptase (RT; Life Technologies) primed with 750 pmol of random hexamers at 42°C for 1 h in the presence of 0.5 mM dNTP, 10 mM dithioreitol, and 5 IU of RNase Inhibitor (Life Technologies). Controls were run with the RT omitted from the RT mix. Samples of cDNA were standardized by measuring ß-actin cDNA by competitive polymerase chain reaction (PCR), which was previously described and validated by Zou et al. [30] in our laboratory. A constant amount of cDNA ß-actin from granulosa cells was amplified in the presence of graded concentrations from 0 to 50 pg of competitor, a recombinant pQB2 plasmid containing an unrelated DNA fragment [31]. The competitive reaction was performed in a final volume of 50 µl containing 1 µl of cDNA, 1 µl of competitor dilution, 10 pM of each primers, PCR reaction buffer, 1.5 µl of 50 mM MgCl2, 2 µl of 6.25 mM dNTP, and 0.5 µl of Taq polymerase (2.5 IU; Life Technologies). Agarose gel electrophoresis showed two characteristic bands corresponding to the competitor products (390 base pairs [bp]) and of the granulosa ß-actin cDNA (237 bp) (Fig. 1). The ratio of the intensity of the bands was set to 1 when the amount of plasmid pQB2 was equal to the amount of cDNA from granulosa cells. Knowing that 1 pg of 371-kilobase plasmid pQB2 contains 2.5 x 109 molecules, we determined the number of actin cDNA molecules in RT samples. We verified that lead treatment did not modify the amount of ß-actin cDNA, and then we compared the quantity of RT-PCR products for P450 arom and ERß in RT samples containing the same amount of ß-actin cDNA molecules.



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  		FIG. 1. Standardization of RT-PCR products by ß-actin competitive RT-PCR. The 237- and 390-bp fragments indicate the amplification products of cells and plasmid competitor, respectively. The ratio of the intensity of the bands was set at 1 for 0.5 pg of plasmid for both samples (control and lead-treated sample), indicating that they contained the same amount of ß-actin cDNA

The PCR for P450 arom and ERß were performed using appropriate PCR protocols on preparations containing the same amount of ß-actin cDNA. The number of cycles was determined during the amplification phase of DNA. Primer sequences and positions are shown in Table 1. Negative controls were run using water and cDNA from human B-lymphocytes. The PCR products were visualized on 2% agarose gel electrophoresis under ultraviolet light. The intensity of the bands was measured using the Gel analyst software (Clara Vision, Orsay, France). The PCR products were quantified using a colorimetric PCR-ELISA assay (Boehringer Mannheim GmbH, Mannheim, Germany). Aliquots of the amplified products were subjected to an additional elongation cycle in the presence of biotinylated (biot) oligonucleotides and digoxigenin-labeled dUTP. Labeled products were quantified by ELISA in streptavidin-coated microtiter plates using peroxidase-conjugated anti-digoxigenin antibody. Absorbance was measured in an ELISA reader plate at 405 nm.


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  		TABLE 1. Primer pairs and PCR condtions

Aromatase Activity

Aromatase activity was assessed by measuring the release of tritiated water production from 1ß-[3H]androstenedione previously described by Garzo and Dorrington [32] and Al-Gubory and Machelon [33] in our laboratory. Cells were incubated at 37°C for 4 h in medium containing 3 µCi of 1ß-[3HN]androstenedione (Amersham Life Science, Orsay, France) equivalent to a final concentration of 0.2 µM. The reaction was stopped by transferring the media into glass vials containing 2 ml of chloroform. Samples were vortexed and stored overnight at -20°C. After centrifugation (3000 x g for 10 min), 500 µl of the upper aqueous layer were collected and treated for 10 min with 500 µl of an aqueous suspension of 5% charcoal and 5% dextran to remove excess 1ß-[3HN]androstenedione. The charcoal was removed by centrifugation at 5000 x g for 30 min. In a supernatant aliquot (200 µl), the radioactivity from tritiated water (3H2O) was measured in a liquid scintillation counter. The quantified radioactivity was corrected by subtracting the blank-tube radioactivity. To determine the protein content using the Bio-Rad Protein Assay, attached cells were solubilized with 0.5 N NaOH and neutralized with 0.5 N HCl. Aromatase activity was expressed in as picomoles of 1ß-[3HN]androstenedione metabolized per hour per milligram of cell protein. Controls and treated samples were compared using independent Student t-tests. Differences were considered to be statistically significant at P <= 0.05.

Western Blot Analysis

Cells were scraped and then homogenized for 1 h at 4°C into ice-cold buffer containing 10 mM Tris HCl (pH 7.6), 1% Triton x-100, 5 mM NaCl, 30 mM sodium pyrophosphate, 50 mM NaF, 400 µM orthovanadate, and protease-inhibitor mixture, which had been optimized and tested for this use (Sigma). Total homogenized lysates were centrifuged for 30 min at 13 000 x g at 4°C, and the resulting supernatant was collected and stored at -20°C. Proteins (30 µg from each sample) were separated by SDS-PAGE (8% resolving gel). They were then transferred onto nitrocellulose membranes (Hybond-ECL; Amersham). Nonspecific binding was prevented by incubating the membranes in TBS (20 mM Tris-HCl [pH 7.6] and 137 mM NaCl) containing 5% skim milk and 0.1% Tween 20 for 1 h at 20–22°C. The membranes were washed and then incubated overnight at 4°C with polyclonal antibodies against P450 arom (1:500 rabbit polyclonal antiserum; a gift of Pr. Kitawaki, Kyoto, Japan), ERß (2 µg/ml; Santa Cruz Biotechnology, Santa Cruz, CA), or ß-actin (0.2 µg/ml; Santa Cruz Biotechnology). The antibodies bound to the proteins on the nitrocellulose membrane were detected using peroxidase-conjugated anti-rabbit immunoglobulin (Ig) G (1:10 000) or anti-goat IgG (1:3000) antibody for 1 h at room temperature. Membranes were washed again in TBS-Tween 20 (0.1%). Blots were developed using chemiluminescent substrate from the detection system (ECL; Amersham) and exposed on Kodak film (Sigma). Lysate from Jurkat cells (human T-lymphocyte line; Upstate Biotechnology, Charlottesville, VA) was used as a negative control. We used prestained SDS-PAGE standards (Bio-Rad S.A., 94203 Ivry, France) to estimate the apparent molecular size of the blotted proteins. Exposed films were scanned on an Imstar densitometer (Paris, France), and the intensity of signals was measured using FotoLook SA 2.0 software (AGFA-Gevaert, Munich, Germany).

Statistics

Arithmetic means, medians, and SDs were calculated and statistical analyses performed using Statistica software (A. Lange, Polish Academy of Science, Warsaw, Poland). Differences in intracellular lead content and aromatase activity between groups were calculated using the Student t-test. The amounts of PCR products measured by absorbance at 405 nm were related to cDNA serial dilutions. We controlled that all curves correlated at P < 0.05 using the Kendall test. Under these conditions, differences between each couple of points, control and lead, were tested at each cDNA dilution using the Wilcoxon test at P < 0.05. The relative intensities of Western blot signals expressed as a percentage of control values were compared using the chi-square test at P < 0.01 to measure the effect of lead on the amount of proteins.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Lead Accumulated in Human Granulosa Cells

Lead was measured by atomic absorption spectrometry. Data are expressed as µg lead/g protein and represent the mean ± SD (n = 5 experiments). After a 5-h culture period in the presence of 10 µM lead acetate, lead content in granulosa cells was 84.6 ± 52.2 µg/g. After 24 h, it was 392 ± 62 µg/g, and after 72 h, it was 2391 ± 1743 µg/g. Lead content in cells cultured in the absence of lead acetate was undetectable (<=0.001 µg/g).

Lead Decreased the Amount of P450 Arom and ERß mRNA

We compared the quantity of RT-PCR products for P450 arom and ERß in RT samples containing the same amount of ß-actin cDNA molecules. Controls were performed in parallel in basal medium and in basal medium with 10 µM sodium acetate added to it. No significant differences were detected between the control run in sodium acetate and the control run in the basal medium. No signal was detected in negative controls with water and B-lymphocyte cDNA (data not shown).

Figure 2 shows data obtained in a representative experiment using samples of cells cultured for 5 h (after culturing for 72 h) in the presence of 10 µM lead acetate (treated cells) versus cells cultured in the absence of lead (controls). The photograph of the ethidium bromide-stained gel (Fig. 2A) shows a characteristic, 202-bp PCR fragment that decreases in intensity with serial cDNA dilutions from 1:1 to 1:64. The PCR signals for P450 arom were less intense in lead-treated samples versus controls for equal cDNA dilutions. The amount of PCR products was measured by PCR ELISA. Absorbance measured at 405 nm was related to cDNA dilutions. Data revealed a 3-fold decrease in the presence of 10 µM lead acetate (Fig. 2B).



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  		FIG. 2. Lead effect on P450 arom mRNA. A) The photograph of the ethidium bromide-stained gel shows a characteristic, 202-bp PCR fragment, the intensity of which decreases with serial cDNA dilutions. Preparations treated with 10 µM lead acetate gave less-intense PCR signals than controls for equal cDNA dilutions. B) The PCR products were quantified by PCR ELISA. In this representative experiment, the absorbance measured at 405 nm was related to cDNA dilutions and revealed a 3-fold decrease in P450 arom mRNA in samples obtained from cells incubated with 10 µM lead acetate for 5 h after culturing for 72 h versus control in basal medium

Figure 3 shows data obtained in the same representative experiment by quantifying ERß mRNA. The photograph of the gel (Fig. 3A) shows a characteristic, 439-bp PCR fragment that decreased in intensity with serial cDNA dilutions from 1:2 to 1:32. The amount of PCR products measured by PCR ELISA was decreased by 5-fold in cells cultured in the presence of 10 µM lead acetate (Fig. 3B).



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  		FIG. 3. Lead effect on ERß mRNA. A) The photograph of the gel shows a characteristic, 439-bp PCR fragment, the intensity of which decreased with serial cDNA dilutions from 1:2 to 1:32. B) In this representative experiment, the absorbance measured at 405 nm by PCR ELISA was related to cDNA dilutions and revealed a 5-fold decrease in ERß mRNA in cells cultured in the presence of 10 µM lead acetate for 5 h after culturing for 72 h versus control in basal medium

Results from four series of experiments run after a 72-h pretreatment period (5- and 24-h lead exposure) were statistically analyzed. First, we controlled by using the Kendall test that all curves (optic density [OD] related to cDNA dilutions) correlated at P < 0.05. Then, we compared each couple of points (control and lead) for each cDNA dilutions using the Wilcoxon test (Fig. 4). Results are shown in Table 2. Considering that the amount of PCR products reflected the amounts of transcripts, they revealed that lead significantly decreased the amount of P450 arom and ERß transcripts in granulosa cells. Additional experiments were run after a 24-h pretreatment period (5- and 24-h lead exposure). Although these data could not be statistically analyzed, they clearly revealed that lead decreased P450 arom (2- to 4-fold) and ERß (5-fold), which confirms the results obtained after culturing for 72 h.



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  		FIG. 4. Statistical analysis of results from four series of experiments (24-h lead exposure after culturing for 72 h). The P450 arom RT-PCR products were quantified by PCR ELISA. Absorbance was measured at 405 nm, and OD values were related to cDNA dilutions from 1:1 to 1:64. Using the Kendall test, we controlled that all curves in each group (control and lead-treated) correlated at P < 0.05. Then, we used the nonparametric Wilcoxon test to compare each data couple, expressed as the mean ± SD


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  		TABLE 2. Lead effect on P450 arom and ERß mRNA after culturing for 72 h

Also, we tested the effect of the protein-inhibitor CHX on the production of P450 arom and ERß mRNA. We compared cells treated with CHX versus cells treated with CHX and lead. Data showed that the intensity of PCR signals was decreased in cells treated with CHX and lead versus cells treated with CHX alone (Fig. 5). Thus, CHX treatment did not eliminate the inhibitory effect of lead on P450 arom and ERß mRNA production.



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  		FIG. 5. Effect of the protein-inhibitor CHX. The intensity of the P450 arom and ERß RT-PCR signals was lower in preparations treated with CHX and lead than in preparations treated with CHX alone

Lead Decreased P450 Arom Activity

Aromatase activity was 24.0 ± 5.8 pmol h-1 mg-1 in cells cultured in the presence of 10 µM lead acetate for 24 h (n = 12 experiments) versus 35.5 ± 3.5 8 pmol h-1 mg-1 in controls (n = 7 experiments). These results show that lead significantly decreased (35% decrease from control) aromatase activity (P <= 0.05, independent Student t-test). No significant difference was observed between cells cultured in basal medium and cells cultured in the presence of 10 µM sodium acetate (controls).

Lead Decreased P450 Arom and ERß Protein Expression

The P450 arom and ERß protein levels were measured using Western blotting procedures in cells cultured in the presence of 10 µM lead acetate for 24 h after culturing for 72 h (n = 3 experiments). Western blot analysis was carried out with cell homogenates (30 µg of proteins). A 55-kDa band was detected by blotting with the P450 arom antibody, a 54-kDa band was detected with the ERß antibody, and a 43-kDa was detected with the ß-actin antibody. In the representative experiment shown in Figure 6, the intensity of the blotting signal detected with the P450 arom antibody in treated cells decreased to 65% of the control value (100%). The intensity of the blotting signal detected with the ERß antibody in treated cells decreased to 25% of the control value (100%). The intensity of ß-actin signals was similar in control and treated cells. Results from three experiments were statistically analyzed using the chi-square test (Table 3). In each experiment, lead significantly decreased P450 arom and ERß protein levels. In contrast, no significant effects were revealed between the control run in sodium acetate and the control run in basal medium.



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  		FIG. 6. Effect of lead on P450 arom and ERß protein content. Cells were incubated with 10 µM lead acetate for 24 h after culturing for 72 h. Immunoblots revealed a 55-kDa band for P450 arom, a 54-kDa band for ERß, and a 43-kDa band for actin used as a standard. The intensity of the bands was measured by scan-analysis software. In this representative diagram, the intensity of the signal, which was 100% in controls without lead acetate, dropped to 65% for P450 arom and to 25% for ERß in treated cells. No decrease was detected in ß-actin blots


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  		TABLE 3. Lead effect on P450 arom and ERß protein content (24-h lead exposure after culturing for 72 h)


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Our data clearly demonstrate that lead entered and accumulated in cultured cells. Cellular content of lead increased after 5, 24, and 72 h, reaching very high values after 72 h without reducing cell viability. Thus, human ovarian granulosa cells may accumulate large amounts of lead. We found that lead decreased basal aromatase activity, P450 arom protein, and transcript production. The mRNA and ERß protein levels were also reduced. By contrast, ß-actin transcript and protein content in granulosa cells were not modified by lead exposure, indicating that the variations in mRNA and protein levels were not caused by variations in the overall amount of mRNA and protein. The CHX did not eliminate the inhibitory effect of lead on P450 arom and ERß mRNA production. That suggests that the observed inhibition of gene expression by lead does not require de novo protein synthesis.

The dose most commonly used in the in vitro experiments was 10 µM lead acetate. This also corresponded to the minimal dose that induced the maximal effects in our experiments (data not shown). Although it is quite difficult to compare the doses used in the in vitro experiments to lead levels measured in the blood of humans exposed to lead, health effects associated with lead exposure have been correlated to blood lead levels from 3 to 5 µM for chronic exposure and up to 20 µM for acute exposure. Inhibition of aromatase activity may involve different mechanisms. One mechanism is the direct inhibitory effect of lead. It may be mediated by lead blocking the ferroxy radical involved in all three steps of P450 arom action via iron interaction [34, 35]. Also, it may be mediated by the formation of a complex with the cysteine residues, because lead has a strong affinity for sulfhydryl groups [36]. However, we cannot exclude that the changes in aromatase activity also resulted from concomitant changes in the levels of P450 arom mRNA and protein. This supports the hypothesis that the inhibitory action of lead on aromatase is partly attributable to the down-regulation of gene transcription. It is well known that protein kinase C (PKC) may be implicated as a target for lead [37]. Activation of PKC in cultured granulosa cells from rats [38], sheep [39], and humans [40] down-regulates aromatase gene transcription. This would implicate that lead activated the PKC in human granulosa cells. Further investigations will be necessary to confirm this hypothesis. Decrease in transcript content may be also caused by increased mRNA turnover or specific alteration of the transcription process. Similarly, decrease in ERß proteins was associated with a concomitant decrease in ERß transcripts. The interaction of lead in cells with DNA- or nucleic acid-binding proteins represents a fundamental mechanism underlying lead toxicity [41]. The lead targets cellular proteins that contain the zinc-finger motif protein domain through bonds created with cysteine residues [42]. Thus, lead may interfere with the DNA-binding properties of nuclear transcription factors. Indeed, further investigations will be necessary to demonstrate that lead alters nuclear transcription factors binding to ERß and to P450 arom promoter sequences.

In chronically lead-intoxicated rhesus monkeys, primordial follicles are damaged and follicular development is inhibited [43, 44]. Rats exposed to high lead concentrations develop ovarian follicular cysts [5]. Female mice have a reduced number of small and medium follicles after low lead acetate intoxication [45]. Recently, we showed that lead causes dysfunction of folliculogenesis in mice ovaries with fewer primordial follicles and an increase in atretic antral follicles [46]. All these data indicate that lead may alter folliculogenesis in mammals. The P450 arom is crucial for follicular maturation. Aromatase inhibition may increase the serum androgen:estradiol ratio, which is a cause of follicular atresia and alteration in folliculogenesis [47–49]. Treatments with fadrozole, a specific aromatase inhibitor, significantly reduced the number of healthy antral follicles produced in rats [50]. Autocrine effects of estradiol are mediated in the ovary by ERß; BERKO female mice have a poor reproductive capacity as a consequence of folliculogenetic blockade. All these data indicate that both ERß and aromatase play important roles in the regulation of follicular growth and that lead-induced inhibitory effects on these two factors may be the source of folliculogenetic disorders.

Estradiol is a pleiotropic hormone that plays a key role in mammalian physiological functions outside the female reproductive system, including the cardiovascular system [51] and bone stability [52], by binding to ERß, which is widely expressed in these tissues. Thus, lead might affect numerous other physiological functions in addition to the reproductive system in women by reducing ERß expression. Endocrine disrupters are exogenous chemicals that often block hormonal action. Lead is known as an environmental endocrine disrupter affecting Leydig cell steroidogenesis and the male reproductive system [53, 54] as well as ovarian steroidogenesis in rats [55]. Its action on P450 arom and ERß expression provides further evidence that lead might be considered as a potential endocrine disrupter in women.


   ACKNOWLEDGMENTS
 
We thank the Center for Reproductive Medicine (American Hospital of Paris, Neuilly, France) for providing us with the follicular aspirates and its technicians for their helpful assistance.


   FOOTNOTES
 
1 Supported by Ministère de l'Environnement/Inserm grant EN97D06. Back

2 Correspondence: Véronique Machelon, INSERM Unité 355, 32 rue des Carnets, 92140 Clamart, France. FAX: 33 1 46 32 79 93; veronique.machelon{at}inserm.ipsc.u-psud.fr Back

Received: 21 August 2002.

First decision: 12 September 2002.

Accepted: 26 December 2002.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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