* Institute for Risk Assessment Sciences (IRAS), Utrecht University, 3508 TD, Utrecht, The Netherlands; Department of Environmental Toxicology, and
Department of Chemistry, University of California, Davis, California 95616
Received June 2, 2004; accepted August 9, 2004
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ABSTRACT |
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Key Words: H295R; flavonoids; quinolin-4-ones; catechins; aromatase; endocrine disruption; induction; inhibition; transcription.
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INTRODUCTION |
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Several studies have addressed the ability of flavonoids to interfere with the catalytic activity or expression of aromatase (cytochrome P450 19; CYP19), the enzyme responsible for the conversion of androgens to estrogens (Ibrahim and Abul-Hajj, 1990; Kellis and Vickery, 1984
; Le Bail et al., 1998
, 2000
; Pelissero et al., 1996
; Whitehead and Lacey, 2003
). Most of these studies were performed in human placental microsomes, and they reveal significant differences in the relative inhibition potencies of flavonoids. Although the use of microsomes for the study of enzyme inhibitors has numerous advantages, it also has several drawbacks. One aspect is that factors such as cellular uptake and metabolism of flavonoids and their effects on enzyme regulation are generally not taken into consideration. In addition, flavonoid test concentrations used in some studies exceed solubility or are in a range that would normally be toxic to cells or the organism. The use of cell lines may address some of these limitations, and it obviates the need to resort to in vivo experiments. In the present study the human adrenocortical (H295R) cell line was selected as a model system to examine the activity of a variety of natural and synthetic flavonoids on the catalytic activity and gene expression of aromatase. The H295R cell line expresses numerous steroidogenic enzymes, including aromatase, and is a useful bioassay to screen for interferences with steroidogenesis (Rainey et al., 1994
; Sanderson et al., 2000
; Staels et al., 1993
). This cell line has been used to determine inhibitory potencies of compounds toward several steroidogenic enzymes (Johansson et al., 2002
; Ohno et al., 2002
; Sanderson et al., 2002
) and to study the effects of chemicals on regulation of expression of these enzymes (Harvey and Everett, 2003
; Li et al., 2004
; Sanderson et al., 2001
). Here we use the H295R cell line to determine inhibition and induction potencies of a large number of natural and synthetic flavonoids to delineate structureactivity relationships for modulation of aromatase expression and catalytic activity.
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MATERIALS AND METHODS |
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Cell culture conditions. H295R cells were obtained from the American Type Culture Collection (ATCC #CRL-2128) and grown in 1:1 (v/v) Dulbecco's modified Eagle medium/Ham's F-12 nutrient mix (DMEM/F12) containing 365 mg/ml L-glutamine and 15 mM HEPES (GibcoBRL). The medium was supplemented with 10 mg/l insulin, 6.7 µg/l sodium selenite, and 5.5 mg/l transferrin (ITS-G, GibcoBRL), 1.25 mg/l bovine serum albumin (Sigma-Aldrich, Zwijndrecht, The Netherlands), 100 U/l penicilline/100 µg/l streptomycin (GibcoBRL), and 2% steroid-free replacement serum Ultroser SF (Soprachem, France). For the aromatase experiments, cells were treated as described previously (Sanderson et al., 2000). In brief, cells (between 1 x 105 and 2 x 105 cells/well) in 24-well culture plates containing 1 ml medium per well were exposed to various concentrations of the natural (Sigma-Aldrich) and synthetic flavonoid compounds dissolved in 1 µl of DMSO. Negative control cells received 1 µl of DMSO. As positive control for aromatase inhibition, cells were exposed to 1 µM of the steroidal aromatase inhibitor 4-hydroxyandrostenedione in DMSO; as positive control for induction, 300 µM of 8-bromo-cyclic adenosine monophosphate (8Br-cAMP) or 20 µM of forskolin dissolved in medium containing 0.1% (v/v) DMSO was used. Unexposed cells were included as further controls. All treatments were tested in quadruplicate, unless stated otherwise; each concentration-response experiment was performed three times. For the mRNA isolation experiments, cells in 6-well plates were exposed to 4 µl DMSO or the test chemicals in DMSO; a positive control (300 µM 8-Br-cAMP) was included on each plate. Each treatment was tested in triplicate; each experiment was performed three times. DMSO at 0.1% had no effect on CYP19 expression or catalytic activity relative to unexposed cells. Protein concentrations were determined by the method of Lowry (Lowry et al., 1951
), with bovine serum albumin (Sigma-Aldrich) as standard. All exposures were for 24 h, unless stated otherwise.
Isolation and amplification of RNA. RNA was isolated using the RNA Insta-Pure System (Eurogentec, Belgium) according to the instructions of the supplier and stored at 70°C. The purity of the RNA preparations was verified by denaturing agarose gel electrophoresis. Reverse-transcriptase polymerase chain reactions (RT-PCR) were performed using the Access RT-PCR System (Promega, Madison, WI). The primer pairs and experimental conditions used for promoter-specific amplification of CYP19 mRNA were reported recently (Heneweer et al., 2004). The sequences of the pII- and 1.3-promoter-specific mRNAs were constructed from National Center for Biology Information (NCBI) accession numbers S52794 and D21241, respectively and reported by Heneweer et al. (2004)
. For both primers, amplification was performed using an annealing temperature of 61°C, 1 min extension, and 35 cycles. As reference, RT-PCR was performed on ß-actin mRNA as described previously (Sanderson et al., 2002
). Beta-actin mRNA expression was not affected by treatment with DMSO, 30 µM quercetin, or 300 µM 8Br-cAMP; it was increased slightly but not statistically significantly (<15%) by 30 µM of genistein). Under these conditions ß-actin could be used as a reliable reference amplification response. Amplification products were detected with agarose gel electrophoresis and ethidium bromide staining. Intensities of the ethidium bromide stains were quantified with a FluorImager (Molecular Dynamics, Sunnyvale, CA).
Aromatase assay. The catalytic activity of aromatase was determined based on the method of Lephart and Simpson (1991) with minor modifications. Cells were exposed to 54 nM 1ß-3H-androstenedione (New England Nuclear Research Products [PerkinElmer] Wellesley, MA) dissolved in serum-free culture medium and incubated for 1.5 h at 37°C in an atmosphere of 5% CO2 and 95% air. All further steps were as reported previously (Sanderson et al., 2000
). Aromatase activity was expressed in picomoles of androstenedione converted per hour per milligram cellular protein. The specificity of the aromatase assay based on the release of tritiated water was verified by measuring the production of estrone, using a 125I-labeled double-antibody radioimmunoassay kit (DSL-8700; Diagnostic Systems Inc, Plantation, FL), and by using the irreversible aromatase inhibitor 4-hydroxyandrostenedione (Brodie et al., 1977
) to block the formation of tritiated water. Under these conditions, unexposed or DMSO-exposed cells had a basal aromatase activity of about 2.4 ± 0.2 pmole/h/mg protein.
MTT reduction assay. Mitochondrial function, as an indicator of cytotoxicity, was assessed by measuring the capacity of H295R cells to reduce MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) to formazan (Denizot and Lang, 1986). MTT is reduced to the blue-colored formazan by the mitochondrial enzyme succinate dehydrogenase, which is considered a reliable and sensitive measure of mitochondrial function. The cells in each well on the 24-well plate were incubated for 30 min at 37°C, with 0.5 ml of MTT (1 mg/ml) dissolved in serum-free medium. Then, the MTT solution was removed, after which the cells were washed twice with phosphate-buffered saline (PBS). The formazan formed in the cells was extracted by adding 1 ml of isopropanol and incubating for 10 min at room temperature. The isopropanol was added directly to a plastic cuvette for spectrophotometric analysis (Shimadzu UV-160A, Shimadzu Benelux, Belgium) at an absorbance wavelength of 560 nm. MTT reduction was linear with time for about 45 min and was not affected by DMSO treatment.
Cyclic AMP measurements. Intracellular cAMP concentrations were determined using a commercial enzyme-linked immunoassay kit (DE0450, R&D systems, Abingdon, UK) according to the instructions provided. Briefly, H295R cells were exposed to the test compounds in 12-well plates for up to 6 h, after which the medium was removed and the cells were washed with 50 mM Tris buffer containing 0.9% NaCl (PBS was not used because phosphate interferes with the immunoassay). The cells were lysed for 20 min in 300 µl of 0.1 N HCl; then the lysate was transferred to a 1.5 ml plastic vial, vortexed, and centrifuged at 12,000 x g for 10 min. The supernatant was diluted 1 in 5 with the assay buffer provided by the kit and underwent all other steps, including an acetylation step according to the instructions of the supplier. Each assay was accompanied by a cAMP standard curve.
Data analysis. All responses are presented as means of triplicate or quadruplicate determinations with standard deviations. Each full concentration-response curve was produced three times. Statistically significant differences were determined by one-way analysis of variance (ANOVA) with a significance level of 0.05. IC50 values were determined using Prism 3.00 (GraphPad Software, Inc, San Diego, CA).
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RESULTS |
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DISCUSSION |
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Rotenone may cause aromatase inhibition via several possible mechanisms. It could interact with the heme group via the oxo-group on the 4-position of the C ring, which is common to the flavonoid structures and appears essential (although not necessarily sufficient) for aromatase inhibition. It is also possible that rotenone inhibits CYP enzymes such as CYP19 by interfering with the electron transfer to the heme-iron, by an interaction with either NADPH-cytochrome P450 reductase or NADH-cytochrome b5. The mechanism of action of CYP inhibition by rotenone is of great interest and warrants further investigation.
Aromatase Inhibition by Other Natural Flavonoids
The ability of various natural flavonoid structures to inhibit aromatase activity in vitro has been documented previously in human placental microsomes (Ibrahim and Abul-Hajj, 1990; Kellis and Vickery, 1984
; Le Bail et al., 1998
), in human preadipocytes (Campbell and Kurzer, 1993
), in JEG-3 human placental choriocarcinoma cells (Saarinen et al., 2001
), and in rainbow trout ovarian microsomes (Pelissero et al., 1996
). However, considerable qualitative and quantitative differences were observed among the different test systems. Comparisons of IC50 values among different test systems is not very useful, because these values are highly dependent on experimental conditions. The observed marked qualitative differences, however, are of greater interest. For example, quercetin was found to inhibit human aromatase activity in placental microsomes (Kellis and Vickery, 1984
), had no effect in JEG-3 cells (Saarinen et al., 2001
), and caused aromatase induction in H295R cells in the present study. Also, the ranking of relative inhibition potencies differed among test systems, although some general trends are apparent. In JEG-3 cells (Saarinen et al., 2001
), apigenin (IC50 values) was more potent (0.18 µM) than 7-hydroxyflavone (0.35 µM), chrysin (0.5 µM), naringenin (1.4 µM), and quercetin (>100 µM). In human placenta (Ibrahim and Abul-Hajj, 1990
; Kellis and Vickery, 1984
; Le Bail et al., 1998
), 7-hydroxyflavone and chrysin (0.20.7 µM) were more potent than apigenin (1.22.9 µM), naringenin (9.2 µM), and quercetin (12 µM). In the present study with H295R cells, 7-hydroxyflavone (4 µM) was the most potent inhibitor, followed closely by chrysin (7 µM), then apigenin (20 µM) and naringenin (85 µM); quercetin was an inducer. In general these studies show that the flavones hydroxylated on the A ring were more potent aromatase inhibitors than those with additional hydroxyl-groups on the B ring; substitution of the flavone base-structure (7-hydroxyflavanone, apigenin) with a flavanone (7-hydroxyflavanone, naringenin) resulted in considerably weaker aromatase inhibition, a finding that is consistent with previous reports (Le Bail et al., 1998
; Saarinen et al., 2001
). In strong contrast to our study, where we found no effect of 7-methoxylated flavone and flavanone, LeBail and co-workers (1998)
found them to be relatively potent inhibitors in placental microsomes with IC50 values of 3.2 and 2.6 µM, respectively.
Aromatase Induction by Natural Flavonoids
The human aromatase gene is known to be under the differential control of several tissue-specific promoters (Bulun et al., 2003; Harada et al., 1993
; Simpson et al., 1993
). For example, aromatase in human gonads is regulated mainly through the proximal promoter pII, and to a lesser extent through promoter I.3, both of which are stimulated by the cAMP-dependent protein kinase A (PKA) second messenger pathway. Healthy breast adipose stromal tissue utilizes promoter 1.4, which is stimulated by the glucocorticoid signaling pathway. However, in malignant breast tumors a promoter-switch appears to occur, resulting in strongly increased pII and 1.3 promoter activity (Agarwal et al., 1996
; Kamat et al., 2002
). In H295R cells, it would appear that several promoters are more or less actively involved in the induction of aromatase activity. The pII-specific and I.3-specific aromatase transcripts have been detected in unstimulated and forskolin/cAMP-stimulated H295R cells in two recent studies (Heneweer et al., 2004
; Watanabe and Nakajin, 2004
). Both studies could not detect 1.4 promoter activity. Clearly, further characterization of the promoter-specific regulation aromatase in H295R cells is required and of great interest. Nevertheless, we can use the cell line to evaluate the effects of various compounds on the pII-specific and I.3-specific regulation of aromatase expression, providing us with information on two promoters that are important in aromatase regulation in gonads and malignant breast tissue.
In the present study, quercetin was found to be a relatively potent and efficacious inducer of aromatase activity in H295R cells, followed by genistein, as also shown previously (Sanderson et al., 2002), and flavone. Both quercetin and genistein increased intracellular cAMP concentrations, and elevated cAMP-mediated pII and 1.3 promoter-specific mRNA levels in H295R cells, indicating that these (iso)flavones act through the PKA second messenger pathway. As quercetin, genistein, and to a lesser extent flavone are known phosphodiesterase inhibitors in several tissues (Kuppusamy and Das, 1992
; Nichols and Morimoto, 2000
), it is likely that the flavonoids cause aromatase induction in H295R cells via this mechanism. The observed potency (in decreasing order) of quercetin, genistein, and flavone to inhibit phosphodiesterase activity corresponded in rank-order with their capacity to stimulate cAMP-mediated lipolysis in rat adipocytes (Kuppusamy and Das, 1992
), as well as with their ability to induce aromatase activity in H295R cells in the present study.
Implications for Human Health
Concentrations of (iso)flavonoids in human blood fall within the nM to lower µM range (Manach et al., 2004). Blood plasma concentrations of quercetin and genistein, for example, generally range from less than 15 nM to about 1.5 µM, although in soy-rich diets peak concentrations of over 4 µM may be reached shortly after ingestion (Manach et al., 2004
). Blood plasma concentrations do not necessarily reflect target tissue concentrations, which may well be higher in intestine and liver than in other tissues. We observed the initial signs of aromatase induction by genistein and quercetin at concentrations above 1 µM in H295R cells. However, in vivo induction of aromatase activity by these flavonoids in experimental animals has not been established so far. It is also unknown whether pII-specific and 1.3-specific aromatase expression can be influenced by xenobiotics in humans. Thus, at present it is not possible to say whether, and to what extent, aromatase expression in vivo would be affected by high dietary intakes of genistein and quercetin. The biological significance and toxicological implications of the observed aromatase induction in vitro requires further in vivo studies.
Little information is available about human blood plasma concentrations of flavonoids such as apigenin, chrysin, and naringenin. The first inhibitory effects on aromatase activity in vitro in H295R cells were observed at concentrations above 1 µM for chrysin and apigenin, and above 10 µM for naringenin. Blood levels of naringenin, for example, are generally around 100 to 150 nM (Erlund et al., 2002), but they may reach peak levels of over 2 µM after consumption of orange or grapefruit juice (Manach et al., 2004
). As a human diet contains several flavonoids with aromatase inhibitory properties in various concentrations, it may be possible for combined tissue concentrations to be reached that result in a certain degree of aromatase inhibition. However, under normal dietary conditions, flavones occur in complex mixtures of which the individual components have various (inhibitory or inductive) effects on enzymes such as aromatase, making the prediction of a net effect difficult. Also, the ultimate beneficial effects usually assigned to flavonoid exposure are the product of a balanced set of interactions; these include not only the effects of these compounds on aromatase activity but also their effects on other enzyme systems and their (often more potent) antioxidant (Bravo, 1998
; Duthie and Crozier, 2000
), anti-inflammatory, and anti-proliferative effects (Kuntz et al., 1999
). Therefore, the effects of flavonoids on aromatase activity cannot be seen in isolation. However, the situation may be different for exposures to single flavonoids (e.g., chrysin) in the form of food supplements. It is likely that consumption of high quantities of such food supplements, often containing more than 100 times the normal dietary intake, results in tissue concentrations of a single flavonoid that are sufficiently high to inhibit aromatase activity. Strong decreases of aromatase activity in females may result in a number of reproductive disturbances, such as disruption of the menstrual cycle (Brodie et al., 1989
), and loss of bone density (Turner et al., 1994
). In men, decreased estrogen synthesis may also result in deleterious effects on bone homeostasis (Vanderschueren et al., 1998
) and disruption of spermatogenesis (Carreau et al., 2003
).
StructureActivity Relationships for Aromatase Inhibition by Flavonoids and Their Derivatives in H295R Cells
An essential contributor to the aromatase inhibitory effect of flavonoids is the 4-oxo group on the C ring of the flavone base structure (Kao et al., 1998). Hydroxylation of the 7-position on the A ring enhances the inhibitory potency considerably, whereas methoxylation diminishes this effect. As for the effect of additional hydroxylation at the 5-position of the A ring (chrysin), it has been suggested that this reduces aromatase inhibition potency after formation of a hydrogen bond with the 4-oxo group (Kao et al., 1998
). However, we did not observe a dramatic decrease in potency when comparing 7-hydroxyflavone with chrysin. It is also clear that the presence of a 7-hydroxy group on the A ring does not appear to be essential, as various alpha-naphthoflavones (7,8-benzoflavones) are also potent aromatase inhibitors.
Flavones (7-hydroxyflavone, apigenin) were significantly more potent aromatase inhibitors than flavanones (7-hydroxyflavanone and naringenin). It is plausible that the lack of 2,3-double bond in the flavanones results in reduced electronegativity of the 4-oxo group and, subsequently, a weaker interaction of this group with the heme prosthetic group of the aromatase enzyme. Obliteration of aromatase inhibition capacity is seen when flavonoid structures are substituted on the 3-position of the C ring, as was observed for quercetin and genistein. This is consistent with the generally very weak or nonexistent inhibitory effects of these compounds found in other studies (Campbell and Kurzer, 1993; Kao et al., 1998
; Pelissero et al., 1996
; Saarinen et al., 2001
).
Substitutions on the B ring of the flavone base structure generally decrease aromatase inhibitory potency, as can be seen by comparing the natural flavone chrysin to apigenin or 7-OH-flavone to the synthetic flavone Z10. One notable exception is the relatively potent inhibitory effect of 4'-tert-butyl-quinolin-4-one (A4), indicating that a t-butyl substitution on the B ring is an important contributing factor. Whether the influence of the t-butyl group is steric, electronic (by donating electrons), or a consequence of increased lipophilicity is presently unknown. Fluorination of the B ring obliterates aromatase inhibition by alpha-naphthoflavone derivatives (AJ2, AJ6, AJ8, and AJ9) or quinolin-4-one derivatives (A2 and A3). Effects of substitutions on the B ring of the synthetic flavones A7, A8, and A9 could not be discerned in our test system because flavone itself was not an inhibitor.
In summary, the 4-oxo group on the C ring appears to be essential for a flavonoid compound to be able to interact with the heme iron of CYP19 and cause aromatase inhibition. This finding rules out any significant capacity for related compounds that lack the 4-oxo group such as catechins or anthocyanidins, but not the chalcones, to inhibit aromatase activity. Inhibition potency is improved by the presence of a 7-hydroxy or 7,8-benzo group on the A ring, although their presence is not essential. Inhibition potency is also improved by the presence of electron donating groups on the B ring, although again this presence is not essential, whereas an electron withdrawing group results in decreased potency. Decreased aromatase inhibition potency is also caused by groups that interfere (either electronically or sterically) with the 4-oxo group, such as substitutions on the 3 position of the C ring, and large substitutions on the 5- or 6-position of the A ring.
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NOTES |
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1 To whom correspondence should be addressed at Institute for Risk Assessment Sciences (IRAS), University of Utrecht, P.O. Box 80176, 3508 TD Utrecht, The Netherlands. Fax: +31-302535077. E-mail: t.sanderson{at}iras.uu.nl.
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