* Department of Zoology, National Food Safety and Toxicology Center, Center for Integrative Toxicology, Michigan State University, East Lansing, Michigan 48824; ENTRIX Inc., East Lansing, Michigan 48864;
Center for Coastal Pollution and Conservation City University of Hong Kong, Kowloon, Hong Kong, SAR China; and
Institute for Risk Assessment Sciences (IRAS), University of Utrecht, 3508 TD Utrecht, Netherlands
Received May 7, 2004; accepted June 3, 2004
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ABSTRACT |
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Key Words: steroidogenesis; bioassay; xenoestrogens; screening.
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INTRODUCTION |
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One type of endocrine disruption takes place when xenobiotics mimic steroid hormones. Of particular concern have been those compounds that mimic endogenous estrogens, sometimes called xenoestrogens. While some reports indicated that endocrine disruption functioned through this mechanism of action, subsequent studies have found that some compounds have more complex mechanisms of action. It has been observed that some compounds can bind to the androgen receptor and function as either androgen agonists or antagonists. Although the effects of endocrine-disrupting chemicals (EDCs) and methods to screen for them have focused on direct interactions with steroid hormone receptors such as ER, AR, and ThR, EDCs can operate several different ways. Firstly, there are several other receptor-mediated processes that control sexual development and homeostasis. Secondly, there are also some nonreceptor-mediated mechanisms. Finally, there are compounds that can modulate steroid hormone production or breakdown and cause endocrine disruption without acting as hormone mimics. These effects are often exerted indirectly via various effects on common signal transduction pathways or by acting on steroid metabolism pathways.
One such example is the effect of the herbicide atrazine. Atrazine has been observed to cause estrogenic effects both in vitro and in vivo but does not bind to the estrogen receptor (Connor et al., 1996; Sanderson et al., 1999
, 2000
, 2001
). While the effects observed in vitro occurred at relatively great concentrations, these results serve as an example of the types of effects that can be observed with in vitro tests. The family of 2-chloro- s-triazine herbicides had a common ability to induce the catalytic activity and mRNA levels of CYP19 using the H295R cell line as a steroidogenic model system (Sanderson et al., 2000
, 2001
). The H295R (a subpopulation of H295 that forms a monolayer in culture) human adrenocortical carcinoma cell line has been characterized in detail and shown to express most of the key enzymes involved in steroidogenesis (Gazdar et al., 1990
; Rainey et al., 1993
; Staels et al., 1993
). Sanderson and coworkers suggested that the effects they observed in the H295R cells occurred by the inhibition of phosphodiesterase with a concomitant increase in cyclic-AMP. The model compound 8-bromo-c-AMP also resulted in the upregulation of CYP19 (aromatase) mRNA.
While this mechanism may not be operating in vivo at all times in all tissues of all species or at relevant environmental concentrations, it is a plausible explanation for the observation that atrazine induced luciferase activity under the control of the ER in MVLN cells (MCF-7-luc, MVLN; Villeneuve et al., 1998). However, experiments demonstrating the expression of aromatase in this cell line have yielded equivocal results. Thus, in addition to other indirect mechanisms of action, it is possible that natural and synthetic chemicals can modulate the endocrine system by acting as direct or indirect stimulators or inhibitors of the enzymes involved in the production, transformation, and or elimination of steroid hormones. Here we present a procedure for screening for the effects of chemicals on the profile of expression of steroidogenic genes. Specifically, we report methods to simultaneously measure mRNA concentrations for 10 steroidogenic enzymes and two housekeeping genes in cultured H295R cells.
The key genes measured in the current study include CYP11A (cholesterol side-chain cleavage); CYP11B1 (steroid 11ß-hydroxylase); CYP11B2 (aldosterone synthetase); CYP17 (steroid 17-hydroxylase and/or 17,20 lyase); CYP19 (aromatase); 17ßHSD1, 17ßHSD4, CYP21B2 (steroid 21-hydroxylase), and 3ßHSD2 (3ß-hydroxysteroid dehydrogenase); HMGR (hydroxymethylgutaryl CoA reductase); and the cholesterol transfer protein StAR (steroid acute regulatory protein). The H295R cells used have the physiological characteristics of zonally undifferentiated human fetal adrenal cells, with the ability to produce the steroid hormones of each of the three phenotypically distinct zones found in the adult adrenal cortex (Fig. 1; Gazdar et al., 1990
; Staels et al., 1993
). Since the cells maintain the ability to express these genes and produce these enzymes, which might otherwise only be expressed in certain tissues or periods of ontogeny, they are a useful model system for potential effects on steroidogenesis.
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MATERIALS AND METHODS |
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The H295R human adrenocortical carcinoma cell lines were obtained from the American Type Culture Collection (ATCC CRL-2128; ATCC, Manassas,VA) and were grown in 75 cm2 flasks with 12.5 ml of supplemented medium at 37°C with a 5% CO2 atmosphere. Supplemented medium was a 1:1 mixture of Dulbecco's modified Eagle's medium with Ham's F-12 Nutrient mixture with 15 mM HEPES buffer. The medium was supplemented with 1.2 g/l Na2CO3, ITS + Premix (1 ml Premix/100 ml medium), and 12.5 ml/500 ml NuSerum (BD Bioscience, San Jose, CA). Final component concentrations in the medium were as follows: 15 mM HEPES, 6.25 µg/ml insulin, 6.25 µg/ml transferrin, 6.25 ng/ml selenium, 1.25 mg/ml bovine serum albumin, 5.35 µg/ml linoleic acid, and 2.5% NuSerum. The medium was changed two to three times per week and cells were detached from flasks for subculturing by use of trypsin/EDTA (Sterile 1x Trypsin-EDTA; Life Technologies Inc., Grand Island, NY). Cells were exposed to chemicals of interest in 6-well Tissue Culture Plates (Nalgene Nunc Inc., Rochester, NY). Cells were dosed with chemicals dissolved in DMSO for 4872 h after plating.
RNA isolation. Before nucleic acid isolation and analysis, cell viability was determined. Cells were visually inspected under a microscope to evaluate viability and cell numbers. Also, cell viability was determined with the Live/Dead cell viability kit (Molecular Probes, Eugene, OR). Cell death was only observed for 17-Ethynylestradiol and lovastatin at concentrations greater than 30 µM; ketoconazole and cyproterone acetate inhibited cell growth at concentrations greater than 30 µM. No adverse effects on cell growth or viability were observed for any of the tested chemicals at maximum concentrations ranging from 30 to 100 µM. Exposures in which either cell death or decreased viability was observed were not used for gene expression analysis.
After removal of the medium, cells were lysed in the culture plate by the addition of 580 µl/well of Lysis Buffer-ß-ME mixture (Stratagene, La Jolla, CA). Cells were mixed and collected by repeated pipetting and transferred to a microcentrifuge tube that was mixed to homogenize and ensure low viscosity of the lysate. After mixing, the homogenate was transferred to a prefilter spin cup seated in a 2-ml tube and was centrifuged in a microcentrifuge for 5 min. The spin cup was removed from the receptacle tube and discarded. For RNA isolation, 700 µl of 70% ethanol was added to the filtrate and the tube was vortexed to mix thoroughly. Half of the mixture was transferred to an RNA binding spin cup seated in a fresh 2-ml tube and this was then centrifuged for 1 min. The spin cup was removed and retained and the filtrate was discarded. This procedure was repeated with the same spin cup using the second half of the sample.
To remove residual DNA prior to reverse transcription, DNase treatment was used; 600 µl of 1x low-salt wash buffer were added to the spin cup containing the RNA, this was centrifuged for 1 min, and the filtrate was discarded. Next, 55 µl of RNase Free-DNase I solution (Stratagene) were added to the fiber matrix inside the spin cup. The sample was incubated at 37°C for 15 min. The sample was then washed with 600 µl of 1x high-salt wash buffer and 600 µl of 1x low-salt wash buffer, centrifuged at maximum speed for 3060 s and discarding the filtrate after each wash. A final wash was done by adding 300 µl of 1x low-salt wash buffer to the spin cup, and the tube was centrifuged for 2 min to dry the fiber matrix. The spin cup was transferred to a fresh 1.5-ml microcentrifuge tube and 80 µl of nuclease-free water was added directly onto the center of the fiber matrix inside the spin cup. The tube was incubated for 2 min at room temperature before centrifugation for 1 min. This elution step was repeated to maximize the yield of RNA. The purified RNA was used immediately for RT-PCR or was stored at 80°C until analysis.
An appropriate dilution of the RNA sample (1:50) was prepared for RNA quantitation. The absorbance of the RNA solution was measured at 260 and 280 nm and the 260/280 ratio was calculated. The concentration of total RNA was estimated using the A260 value and a standard with an A260 of 1 that was equivalent to 40 µg RNA/ml.
cDNA preparation. Total RNA (15 µg) was combined with 50 µM oligo-(dT)20 and 10 mM dNTPs diethylpyrocarbamate- (DEPC-) treated water to a final volume of 12 µl. RNA and primers were denatured at 65°C for 5 min and then incubated on ice for 5 min. Reverse transcription was performed using 8 µl of a master mix containing the following: 5x cDNA synthesis buffer, 0.1 M DTT, RNase OUT 40 U/µl, Cloned AMV Reverse Transcriptase (Invitrogen, Carlsbad, CA), and DEPC-treated water. Reactions were incubated at 50°C for 45 min and were terminated by incubation at 85°C for 5 min. Samples were either used directly for PCR or were stored at 20°C until analysis.
Real-time PCR. Real-time PCR (quantitative PCR) was performed by using a Smart Cycler System (Cepheid, Sunnyvale, CA) in 25-µl sterile tubes using a master mix containing the following: 25 mM MgCl2, 1 U/µl AmpErase (Applied Biosystems, Foster City, CA), 5 U/µl Taq DNA polymerase AmpliTaq Gold, 10X SYBR Green (PE Biosystems, Warrington, UK), nuclease-free water, and between 10 pg and 1 µg of cDNA. The Thermal Cycling program was 94°C for 10 min as follows: 5060°C for 30 s to 1 min; 6872°C for 1 min/kb followed by 3540 cycles of 94°C for 1540 s; 5060°C for 30 s to 1 min; 6872°C for 1 min/kb; and a final cycle of 94°C for 1540 s, 5060°C for 30 s to 1 min, and 72°C for 510 min. Melting curve analyses were performed immediately following the final PCR cycle to differentiate between the desired amplicons and any primer-dimers or DNA contaminants.
For quantification of PCR results, Ct (the cycle at which the fluorescence signal is first significantly different from background) was determined for each reaction. Ct values for each gene of interest were normalized by division by the Ct for the endogenous control gene to produce Ct. Therefore, the difference between
Ct values for a control and a chemically exposed culture (designated
Ct) represent the degree of induction or inhibition of the gene of interest. Moreover, the degree of induction or inhibition can be calculated as a fold difference using the following relationship:
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Statistical analysis. Statistical analyses of gene expression profiles were conducted using SYSTAT 10 (SPSS Inc., Chicago, IL). Differences in gene expression were evaluated by ANOVA followed by Tukey's test. Differences with p 0.05 were considered significant. Statistical analysis of sequence homologies between amplicons and the GenBank database were conducted using the BLAST algorithm on the National Center for Biotechnology Information website (http://www.ncbi.nlm.nih.gov/).
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RESULTS |
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Due to the relatively short length of the amplified DNA, some bands could not be sequenced. While some differences were detected from the published sequences, these differences are not likely to be significant given that only a single sequence determination was conducted and the possibility that genetic variants different from the published sequences could occur.
The PCR methods were also optimized to ensure optimum efficiency (100%) over a range of tested RNA concentrations. Relative efficiencies were also determined to ensure that quantification of sequences of interest relative to housekeeping genes would remain constant even at a wide range of relative message concentrations (Fig. 4). The determination of all sequences of interest could be achieved quantitatively with 100% efficiency over a range of at least four orders of magnitude.
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To further elucidate the effects of forskolin and PMA on gene expression, cells were exposed to different concentrations of these compounds over time periods up to 48 h (Fig. 6). In general, treatment with PMA resulted in greater alteration in gene expression at 12 than at 24 h for both 10 and 40 nM PMA. As was observed in Exposure 1, PMA reduced the expression of CYP11A and CYP17; the inhibition of CYP17 was not apparent until 24 h, whereas the reduction of CYP11A was initially evident at 12 h and continued on to 24 h. In cells treated with 10 nM PMA, the expression of CYP11A was somewhat greater at 24 than at 12 h. Furthermore, when CYP11A levels at 24 h in the 10 nM-PMA group were compared to levels at 12 and 24 h in the 40-nM PMA treatment group, no significant differences were observed. These results suggest some recovery for this gene may have occurred, but the exact mechanism of this recovery is unknown at this time. The most significant effect of exposure to PMA was the large increase in CYP19 and 3ßHSD2 gene expression at 12 h for both the tested concentrations. The concentration of CYP19 mRNA was increased 240- and 274-fold by 10 and 40 nM PMA, respectively. Also, 13ßHSD2 gene expression was increased 43.2- and 23-fold by 10 and 40 µM PMA, respectively. The expression of these genes was approximately 10-fold less at 24 h than it was at 12 h, with the expression levels of both genes being less than 1.5-fold different between concentrations. This general pattern of greater gene expression at 12 compared to 24 h occurred for most of the genes analyzed. Furthermore, at 24 h there was little difference in gene expression between cells treated with 10 or 40 nM PMA for genes monitored in the exposure. The consistency of this result among genes and between concentrations as well as to the results of the previous exposures adds to the validity of the great levels of mRNA induction observed.
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As was observed in Exposure 1, 3ßHSD2, CYP11B2, and CYP19 were the genes for which expression was increased to the greatest extent over the three time periods. However, several specific time- and concentration-related differences in gene expression were noted among these three genes. For instance, a 40-fold induction in CYP19 gene expression was observed at 12 h in cells exposed to 10 or 50 µM forskolin. This was followed by a reduction in gene expression to approximately 20-fold induction at 24 and 48 h sampling time in both treatment groups. For CYP11B2 at 12 h, there was a 68- or 34-fold increase in gene expression in cell treated with 10 or 50 µM, respectively. Thereafter, there was a decrease in CYP11B2 expression that resulted in expression levels that were similar among dose groups for the 24 and 48 h time points. In contrast to CYP11B2, there was no concentration-related difference in expression of 3ßHSD2 with time but there was a general trend of reduced gene expression (11-fold reduction) over the experimental time period. Interestingly, while all previous exposures caused little effect on HMGR expression at 12 or 24 h of exposure to either 10 or 50 µM, an exposure to forskolin for 48 h resulted in a considerable and similar increase in the expression of this gene.
The reproducibility of the gene expression in the H295R bioassay was evaluated with forskolin- and PMA-treated cells from Exposures 1 and 3 (Table 5). In cells exposed to forskolin, the only significant differences in gene expression between the two experiments were for CYP21, CYP19, StAR, and CYP11B2. Of these genes, only CYP21 and CYP19 had interassay differences that were greater than 2-fold. In general, the expression of these genes was greater in Exposure 3 than in Exposure 1, and overall significances of these gene activities relative to solvent control values were consistent between assays. That is, if there was a significant alteration in gene expression in one assay it was also significant in the second assay. In the PMA exposures, only CYP17, CYP21, CYP19, 3ßHSD2, and CYP11B2 significantly differed between assays. As was observed with forskolin, the level of gene expression in Exposure 3 was generally greater than that measured in Exposure 1, with all fold differences being greater than 2.5. Again, while the magnitude of gene expression activity differed between the two assays, the significances of gene expression as a consequence of PMA exposure were similar, indicating that the gene expression profile remained the same between assays. Overall, this analysis indicates that while there is some interassay variability in absolute expression, the overall conclusion that can be drawn relative to gene expression profiles is consistent between assays.
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DISCUSSION |
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The production of steroids is a complex process with multiple sensitive control points. Given the complexity of the system and the number of enzymes and substrates involved, the potential for xenobiotic chemicals to interfere with this process is relatively great. Indeed the presence of genetic deficiencies in these steroidogenic enzymes leads to a condition known as congenital adrenal hyperplasia (CAH), which is often fatal (Richmond et al., 2001). While this condition is most frequently caused by a deficiency in CYP21B (Chiou et al., 1990
), deficiencies in StAR and other steroidogenic enzymes are also capable of causing CAH (Richmond et al., 2001
).
Mobilization of cholesterol to CYP11A, also known as CYP450SCC, and its conversion to pregnenolone are the first and rate-limiting steps in the conversion of cholesterol to steroid hormones and is a point of both acute and chronic control (Hu et al., 2001; 2002
). In our study, only 8BrcAMP and forskolin resulted in significant increases in CYP11A1 expression; PMA significantly decreased CYP11A expression, while lovastatin appeared to have little effect on this enzyme. Alterations in CYP11A are also noteworthy since some studies indicate that the expression of other steroidogenic enzymes is coordinated with CYP11A. For example, it has been demonstrated that CYP11A activity may be coordinated with CYP11B1 activity by the physical proximity of the two enzymes (Cauet et al., 2001
). Such physical interrelationships between enzymes may be of greater significance in vivo than in vitro due to the tissue-specific expression of some enzymes. In fact, some tissues, particularly the adrenal gland, exhibit differential enzyme expression within different regions of the tissue (Gazdar et al., 1990
; Sanderson et al., 2000
; Staels et al., 1993
).
Some of the chemicals tested resulted in some increase in CYP21 gene expression. The CYP21 gene product is required for the synthesis of both aldosterone and corticosteroids. Deficiency of this enzyme in CAH results in deficiencies in both cortisol and aldosterone that are also accompanied by overproduction of androgens (Chiou et al., 1990, Richmond et al., 2001
). The overproduction of androgens is due to a combination of the general adrenal hyperplasia and substrate accumulation related to inhibition of the gluco- and mineralocorticoid pathways. In our experiments, increases in CYP21 would be expected to lead to increased synthesis of cortisol and aldosterone and may result in decreased substrate availability for androgen and estrogen production.
CYP17 catalyzes the conversion of aldosterone to corticosteroid substrates and ultimately to sex steroid substrates. Therefore, it is possible that this enzyme could redirect steroid output from mineralocorticoids to glucocorticoids or weak androgens. Inhibition of CYP17 would have the opposite effect. Supporting this hypothesis is the observation that treatment with PMA, which results in an almost complete inhibition of CYP17 expression, results in the greatest increase in CYP19 expression (p < 0.01). CYP19 is responsible for the final conversion of androgens to estrogens.
While only a limited number of chemicals were tested in this study, distinct gene expression profiles are apparent (Table 7). In particular, similar expression patterns were observed for 8BrcAMP and forskolin. These patterns included relatively great increases in the expression of 3ßHSD2 and CYP11B2 and moderate increases in expression of CYP11A, CYP17, CYP19, CYP21, HMGR, and StAR. In contrast, PMA resulted in decreases in CYP11A and CYP17, moderate increases in CYP11B2 and CYP21, and a greater increase in CYP19. This variety of responses demonstrates the utility of the H295R cell line for the detection of both induction and downregulation of gene expression for steroidogenic enzymes (Heneweer et al., 2004). Lovastatin resulted in only moderate increases of 3ßHSD2, CYP11B2, CYP21, and HMGR expression. We hypothesize that the expression profiles observed for forskolin and 8BrcAMP, which were similar, resulted from increased signaling through the cAMP pathway and that other chemicals causing a similar alteration would result in a similar expression profile, as reported previously (Sanderson et al., 2002
). It has been shown that forskolin is able to increase cellular cAMP concentrations in H295R cell line (Sanderson et al., 2002
). In contrast, the expression profiles observed for PMA and lovastatin appear to have been produced by a signaling pathway other than the cAMP pathway and were distinct from each other. PMA exerts effects on steroidogenesis primarily through the MAPKC pathway and so would be expected to have an expression profile distinct from the cAMP-dependent pathways. Lovastatin is known to specifically inhibit HMG-CoA reductase activity and, as expected, treatment with this chemical increased the expression of HMG-CoA reductase in H295R cells.
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The ability to assess all of the key enzymes involved in steroidogenesis in a single assay procedure will clearly be of great interest to those studying the effects of xenobiotics on steroidogenesis. While initial work has focused on specific enzymes such as aromatase, the assay we have presented allows for more general assessment of steroidogenesis by evaluating both enzymes that determine the overall rate of steroidogenesis as well as those specific enzymes that can influence the overall fate or balance of steroid production. The H295R cell line has been previously used in such a bioassay approach, but the end points in those studies were either mRNA species and one or two specific enzymes (Sanderson et al., 2001, 2002
) or were a variety of enzyme activities (Ohno et al., 2002
).
Our findings demonstrate that the genes within the steroidogenesis pathway are not expressed to the same extent and that different chemicals result in different relative changes in the expression of various genes. Chemical agents have the potential to alter gene expression profiles and, potentially, the steroids produced by this pathway. The changes in patterns of relative expression can be used to classify chemicals of unknown mechanisms of action on the steroidogenic pathways. In this way, chemicals can be grouped for further testing of a reduced set of model chemicals and for risk assessments.
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ACKNOWLEDGMENTS |
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NOTES |
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1 To whom correspondence should be addressed at Michigan State University, 224 National Food Safety and Toxicology Center, East Lansing, MI 48824-1311. Fax: (517) 432-2310. E-mail: jonespa7{at}msu.edu.
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REFERENCES |
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Cauet, G., Balbuena, D., Achstetter, T., and Dumas, B. (2001). CYP11A1 stimulates the hydroxylase activity of CYP11B1 in mitochondria of recombinant yeast in vivo and in vitro. Eur. J. Biochem. 268, 40544062.
Chiou, S. H., Hu, M. C., and Chung, B. C. (1990). A missense mutation at Ile172Asn or Arg356Trp causes steroid 21-hydroxylase deficiency. J. Biol. Chem. 265, 35493552.
Connor, K., Howell, J., Chen, I., Liu, H., Berhane, K., Sciarretta, C., Safe, S., and Zacherewski, T. (1996). Failure of chloro-s-triazinederived compounds to induce estrogen receptormediated responses in vivo and in vitro. Fundam. Appl. Toxicol. 30, 93101.[CrossRef][ISI][Medline]
EDSTAC Final Report (1998). Endocrine Disruptor Screening and Testing Advisory Committee Final Report. U.S. Environmental Protection Agency. www.epa.gov/opptintr/opptendo/finalrpt.htm.
Gazdar, A. F., Oie, H. K., Shackleton, C. H., Chen, T. R., Triche, T. J., Myers, C. E., Chrousos, G. P., Brennan, M. F., Stein, C. A., and La Rocca, R. V. (1990). Establishment and characterization of a human adrenocortical carcinoma cell line that expresses multiple pathways of steroid biosynthesis. Cancer Res. 50, 54885496.[Abstract]
Hayes, T. B. Collins, A., Lee, M., Mendoza, M., Noriega, N., Stuart, A. A., and Vonk, A. (2002). Hermaphroditic, demasculinized frogs after exposure to the herbicide atrazine at low ecologically relevant doses. Proc. Nat. Acad. Sci. USA 99, 54765480.
Heneweer, M., Van den Berg, M., and Sanderson, J. (2004). A comparison of human H295R and rat R2C cell lines as in vitro screening tools for effects on aromatase. Toxicol. Lett. 146, 183194.[CrossRef][ISI][Medline]
Hu, M. C., Chiang, E. F.-L., Tong, S. K., Lai, W., Hsu, N. C., Wang, L. C.-K., and Chung, B. C. (2001). Regulation of steroidogenesis in transgenic mice and zebrafish. Mol. Cell. Endocrinol. 171, 914.[CrossRef][ISI][Medline]
Hu, M. C., Hsu, N. C., El Hadj, N. B., Pai, C. I., Chi, H. P. C, Wang, K. L., and Chung, B. C. (2002). Steroid deficiency syndromes in mice with targeted disruption of CYP11A1. Mol. Endocrinol. 16, 19431950.
Kavlock, R. T., Daston, G. P., De Rosa, C., Fenner-Crisp, P., Gray, L. E., Kaattari, S., Lucier, G., Luster, M., Mac, M. J., Maczka, C., et al. (1996). Research needs for the risk assessment of health and environmental effects of endocrine disruptors: A report of the U.S. EPA sponsored workshop. Environ. Health Perspect. 104, 715740.[ISI][Medline]
Ohno, S., Shinoda, S., Toyoshima, S., Nakazawa, H., Makino, T., and Nakajin, S. (2002). Effects of flavonoid phytochemicals on cortisol production and on activities of steroidogenic enzymes in human adrenocortical H295R cells. J. Steroid Biochem. Mol. Biol. 80, 355363.[CrossRef][ISI][Medline]
Rainey, W. E., Bird, I. M., Sawetawan, C., Hanley, N. A., McCarthy, J. L., McGee, E. A., Wester, R., and Mason, J. I. (1993). Regulation of human adrenal carcinoma cell (NCI-H295) production of C19 steroids. J. Clin. Endocrinol. Metab. 77, 731737.[Abstract]
Richmond, E. J., Flickinger, C. J., McDonald, J. A., Lovell, M. A., and Rogol, A. D. (2001) Lipoid congenital adrenal hyperplasia (CAH): Patient report and a mini-review. Clin. Pediatr. (Phila.) 40, 403407.[ISI][Medline]
Sanderson, J. T., Boerma, J., Lansbergen, G. W., and Van den Berg, M. (2002). Induction and inhibition of aromatase (CYP19) activity by various classes of pesticides in H295R human adrenocortical carcinoma cells. Toxicol. Appl. Pharmacol. 182, 4454.[CrossRef][ISI][Medline]
Sanderson, J. T., Heneweer, M., Seinen, W., Giesy, J. P., and Van den Berg, M. (1999). Chloro-s-triazine herbicides and certain metabolites induce aromatase (CYP19) activity in H295R human adrenocortical carcinoma cells. Organohalogen Compounds 42, 58.
Sanderson, J. T., Seinen, W., Giesy, J. P., and Van den Berg, M. (2000). 2-Chloro-S-triazine herbicides induce aromatase (CYP19) activity in H295R human adrenocortical carcinoma cells: A novel mechanism for estrogenicity. Toxicol. Sci. 54, 121127.
Sanderson, J., Thomas, R. J., Letcher, M., Heneweer, Giesy, J. P., and Van den Berg, M. (2001). Effects of chloro-S-triazine herbicides and metabolites on aromatase (CYP19) activity in various human cell lines and on vitellogenin production in male carp hepatocytes. Environ. Health Perspect. 109, 10271031.[ISI][Medline]
Staels, B., Hum, D. W., and Miller, W. L. (1993). Regulation of steroidogenesis in NCI-H295R cells: A cellular model of the human fetal adrenal. Mol. Endocrinol. 7, 423433.[Abstract]
Villeneuve, D. L., Blankenship, A. L., and Giesy, J. P. (1998). Estrogen receptors environmental xenobiotics. In Toxicant-Receptor Interactions and Modulation of Gene Expression. (M. S. Denison and W. G. Helferich, Eds.), pp. 6999. Lippincott-Raven Publishers, Philadelphia.
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