Effect of chlorinated hydrocarbons on expression of cytochrome P450 1A1, 1A2 and 1B1 and 2- and 4-hydroxylation of 17ß-estradiol in female Sprague–Dawley rats

Alaa F. Badawi, Ercole L. Cavalieri and Eleanor G. Rogan1

Eppley Institute for Research in Cancer, 986805 University of Nebraska Medical Center, Omaha, NE 68198-6805, USA


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Chlorinated hydrocarbons (CHCs) are environmental contaminants that bioaccumulate and hence are detected in human tissues. Epidemiological evidence suggests that the increased incidence of a variety of human cancers, such as lymphoma, leukemia and liver and breast cancers, might be attributed to exposure to these agents. The ability of CHCs to disrupt estrogen homeostasis is hypothesized to be responsible for their biological effects. The present study examined the effect of CHCs on the expression of cytochrome P450 (CYP)1A1, CYP1A2 and CYP1B1 mRNAs and the consequent 2- and 4-hydroxylation of 17ß-estradiol (E2) in female Sprague–Dawley rats. Animals were administered a single dose of the LD50 of 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) (25 µg/kg), 2,4-dichlorophenoxyacetic acid (2,4-D) (375 mg/kg) and dieldrin (DED) (38 mg/kg) by gavage. Seventy-two hours after treatment, increased expression of CYP1A1, CYP1A2 and CYP1B1 was observed in the liver, kidney and mammary tissue. Since CYP1A and CYP1B1 are the major enzymes catalyzing 2- and 4-hydroxylation of E2, respectively, the effect of these CHCs on the metabolism of E2 was investigated in rat tissues. Formation of 2- and 4-catechol estrogens was increased in a tissue-specific manner in response to treatment. TCDD was the most potent inducer for CYP1 enzyme mRNA and for the 2- and 4-hydroxylation of E2. 2,4-D and DED induced similar responses, but less than that of TCDD. These results suggest that induction of CYP1 family enzymes and consequent increases in estrogen metabolism by CHCs in target tissues may be factors contributing to the biological effects associated with exposure to these agents.

Abbreviations: AhR, aryl hydrocarbon receptor; BSA, bovine serum albumin; CHCs, chlorinated hydrocarbons; CYP, cytochrome P450; 2,4-D, 2,4-dichlorophenoxyacetic acid; DED, dieldrin; E1, estrone; E2, 17ß-estradiol; IS, internal standard; rc, recombinant; TCDD; 2,3,7,8-tetrachlorodibenzo-p-dioxin.


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Chlorinated hydrocarbons (CHCs) are a heterogeneous group of man-made organic compounds that are widely present in the environment (1). Many CHCs have been used as insecticides, herbicides, flame retardants and in a variety of other industrial uses. The chemical stability and lipophilicity of CHCs and their resistance to degradation results in their persistence in the environment and concentration in food chains (2) as well as their bioaccumulation in human adipose tissue (3), blood (4) and breast milk (5).

Epidemiological evidence has linked occupational exposure to CHCs to a high incidence of a variety of human cancers (see below). However, the consensus of available information from studies on experimental animals is that many of these agents are not likely to be carcinogenic to humans per se (6,7). For example, the chlorophenoxy herbicide 2,4-dichlorophenoxyacetic acid (2,4-D) (Figure 1Go) was implicated in susceptibility to non-Hodgkin's lymphoma (810) and lung (11) and breast cancer in humans (10). 2,4-D, however, has led to no excess incidence of tumors when administered to male or female mice or rats under current experimental guidelines (6,12,13). Likewise, exposure to the insecticide dieldrin (hexachloroepoxyoctahydro-endo,exo-dimethanonaphthalene, DED) (Figure 1Go) has been implicated in the increased incidence of human liver (14) and breast (15) cancers. The marginal carcinogenic potency of DED in animal models, however, has led recently to the recognition that DED is not likely to be a human carcinogen (7).



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Fig. 1. Chemical structures of the chlorinated hydrocarbons studied.

 
This discrepancy between experimental and epidemiological evidence may be related to the ability of a variety of CHCs to disrupt endocrine-regulated homeostasis by up-regulating the transcription of an array of genes, including genes involved in cell growth and differentiation (16,17) and genes coding for estrogen metabolism (18). It is known that the estrogens, 17ß-estradiol (E2) and estrone (E1), are metabolized via two major pathways: formation of the 2- and 4-catechol estrogens [2- and 4-OH(E1)E2] and 16{alpha}-hydroxylation (19,20). The formation of 2-OH(E1)E2 is predominantly mediated by cytochrome (CYP)1A1 and CYP1A2, which also catalyze 4-hydroxylation to a lesser extent, i.e. <10% of 2-OH(E1)E2, while the production of 4-OH(E1)E2 is mainly catalyzed by CYP1B1 (21). Unless detoxified, catechol estrogens may be oxidized to electrophilic metabolites, catechol estrogen quinones, that can react with DNA to form depurinating and stable adducts. These adducts, particularly depurinating adducts, can lead to oncogenic mutations and may subsequently initiate many human cancers (2224). Thus, it can be hypothesized that induction of CYP enzymes, particularly CYP1B1, may lead to increased synthesis of 4-catechol estrogens, their oxygenated quinone metabolites and depurinating DNA adducts. In support of this hypothesis, 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) (Figure 1Go), a tetrachloro heterocyclic compound, induced the transcription of CYP1B1 in a variety of human and animal models (2531). This effect was suggested to be related to the etiology of cancer (25).

To investigate the influence of CHCs on the endogenous synthesis of catechol estrogens, the present study examines the effect of these agents on the expression of CYP1 enzyme genes that are involved in estrogen biotransformation and on the subsequent 2- and 4-hydroxylation of E2 in the liver, kidney and mammary gland of female Sprague–Dawley rats.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Animals and treatments
Pathogen-free female Sprague–Dawley rats (7-week-old, weight 211 ± 10 g) purchased from Harlan Industries (Indianapolis, IN) were housed at 24 ± 2°C and 50% humidity, with a 12 h light/dark cycle. Animals were acclimatized for 7 days with food and water provided ad libitum. Animals were then randomized into four groups (4 animals/group) and were orally administered the vehicle as a control group or the LD50 dose of TCDD (25 µg/kg) (Supelco, Bellefonte, PA), 2,4-D (375 mg/kg) (Aldrich Chemical Co., Milwaukee, WI) or DED (37 mg/kg) (Aldrich Chemical Co.). CHCs were dissolved in toluene and diluted into 9 vol of corn oil; the toluene was then evaporated under argon. Animals were killed 72 h following the treatment and the liver, kidney and mammary tissues were removed, frozen in liquid N2 and stored at –80°C. Tissues from individual animals were divided to provide ~1 g for the isolation of total RNA and the remaining portion for preparation of microsomes. If needed, the fourth animal in each group was used in the preparation of supplementary kidney and mammary microsomes.

Preparation of microsomes and isolation of RNA
Homogenates of thawed tissues were prepared, as previously described (32), in 10 mM Tris–HCl buffer (pH 8) containing 0.25 M sucrose, 0.1 mM EDTA, 0.1 mM DTT and 20 µM butylated hydroxytoluene using a Brinkmann Polytron homogenizer (two passes). Homogenates were centrifuged at 10 000 g at 4°C for 15 min. The supernatant was then centrifuged at 105 000 g at 4°C for 60 min and the resulting microsomal fraction was resuspended in 50 mM potassium phosphate buffer (pH 7.4). The microsomal protein content was quantified with Bio-Rad reagent (Bio-Rad Laboratories, Hercules, CA), measuring the absorbance at 595 nm using a Perkin Elmer spectrophotometer (Perkin Elmer, Norwalk, CT) with bovine serum albumin (BSA) as the standard (33). CYP content was measured from the sodium dithionite-reduced carbon monoxide difference spectrum with a molar extinction coefficient of 91/mM/cm (34) using a Varian-CARY 300 scan spectrophotometer (Varian Medical Systems, Palo Alto, CA).

Total RNA was isolated using the TRIzol reagent (Life Technologies, Grand Island, NY). RNA was resuspended in 100 µl of RNase-free water, quantified by measuring the absorbance at 260 nm, diluted to 50 ng/µl with RNase-free water and stored at –80°C for RT–PCR analysis.

Competitive RT–PCR analysis
Quantitative RT–PCR was carried out using a modified competitive titration assay, which uses heterologous internal standards (IS) of recombinant (rc)RNA (35,36). For quantification of CYP1A1, CYP1A2, CYP1B1 and ß-actin mRNA transcripts, PCR primers (Table IGo) were used that have been described for the IS and target genes (26). Primers were synthesized and purified by the Eppley Institute Molecular Biology Core Laboratory.


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Table I. Primer pairs for target and internal standards used for competitive RT–PCR
 
Preparation of rcRNA internal standards.
Specific IS were generated by PCR amplification of the rcRNA forward and reverse primers, followed by RNA transcription of the amplified products. PCR reactions were carried out essentially as described previously (26,35). PCR products were purified using the Wizard PCR DNA system (Promega, Madison, WI) and transcribed into RNA by the T7 RNA polymerase promotor using the RiboMax large-scale RNA production system-T7 (Promega). The rcRNA was subsequently treated with RNase-free DNase to remove the DNA template and then extracted with water-saturated phenol:chloroform:isoamyl alcohol (24:25:1). The rcRNA for IS was washed with 70% ethanol, suspended in RNase-free water, quantified by measuring absorbance at 260 nm and stored at –80°C until analysis. The IS amplicons were smaller than the target mRNAs and were, therefore, distinct when separated by gel electrophoresis.

RT–PCR reaction.
Each tissue RNA sample (100 ng) was reverse transcribed in six separate equal aliquots in the presence of one of six increasing concentrations of IS rcRNA (Figure 2Go). The reaction was carried out, as described previously (26), in 20 µl of 1x reaction buffer (80 µg/ml BSA, 16.6 mM ammonium sulfate, 67 mM Tris–HCl, pH 8.8, 6.8 µM EDTA, pH 8.0, 50 mM ß-mercaptoethanol) containing 5 mM MgCl2, 1 mM each dNTP, 0.75 U/µl RNase inhibitor, 5 µg/ml oligo(dT)18 and 4 U/µl MMLV reverse transcriptase at 37°C for 15 min. After terminating the reactions by heating at 99°C for 5 min and cooling at 4°C, 20 µl of PCR mix were added (2.5 mM MgCl2 for CYP1B1 and ß-actin, 4 mM for CYP1A1 and 5 mM for CYP1A2; 0.5 mM each forward and reverse primers and 0.5 U Taq polymerase). Reactions were incubated at 95°C for 5 min and then amplified using a melting temperature of 94°C for 30 s, annealing temperature of 55 (CYP1A1) or 60°C (CYP1A2, CYP1B1, ß-actin) for 30 s and extension temperature of 75°C for 30 s, followed by a final extension step at 75°C for 5 min in a GeneAmp PCR system 2400 thermal cycler (Perkin Elmer). PCR products were separated by 2% agarose gel electrophoresis in 1x TEA buffer (40 mM Tris–acetate, 1 mM EDTA) and visualized by ethidium bromide staining (1 µg/ml). DNA product sizes were estimated relative to a 100 bp DNA ladder (Promega). All RNA samples were quantitated at least twice in two different dilution ranges (35,36). Control reactions were run to rule out contamination of RNA with genomic DNA, in which reverse transcriptase was omitted from the reaction mixtures that contained both IS rcRNA and the sample RNA. To rule out other sources of contamination, control PCR reactions were carried out in reaction mixtures containing no cDNA.



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Fig. 2. Quantitative RT–PCR analysis of CYP1A1, CYP1A2, CYP1B1 and ß-actin in rat liver. The blot is representative of observations in the liver, kidney and mammary gland of at least three animals. Varying amounts of the rcRNA were spiked into aliquots of total tissue RNA prior to reverse transcription. DNA product sizes were estimated against a 100 bp DNA ladder (M). The log ratio of the intensities of target:IS and log number of copies of IS were fitted by linear regression and the level of the target mRNA was quantified as described in Materials and methods.

 
RT–PCR quantification.
Digitized images of the stained cDNA products were captured as 8 bit digital TIFF files using a DC120 Digital Camera (Eastman Kodak, Rochester, NY). The intensity of each band was measured using Kodak Digital Science 1D Image Analysis software (Eastman Kodak). Quantition of the amount of target mRNA present was determined as described (37). Briefly, log10 of the IS rcRNA copies added to each reaction was plotted against log10 of the ratio of intensity of target mRNA to IS rcRNA using linear regression. The plots were derived from at least six independent reactions. Where the ratio of target to IS RNA was equal to 0, the antilog of the corresponding concentration of the IS rcRNA represents the amount of target mRNA in the sample RNA.

Estradiol metabolism
The assay of metabolism of E2 was modified from previously described procedures (38,39). Fifty micrograms of microsomal protein were incubated, in a total volume of 500 µl, with 10 µM E2, 0.1 M sodium phosphate buffer (pH 7.4), 1.4 mM NADPH and 5 mM MgCl2. Additionally, 2 mM ascorbic acid was added to the reaction mixture to protect E2 metabolites from oxidative degradation without affecting the enzyme activity (40). After 15 min incubation at 37°C, reactions were terminated by extracting the metabolites twice in 1 vol ethyl acetate (31). Extracts were combined, evaporated under argon, reconstituted in 100 µl of methanol:50 mM ammonium acetate, pH 4.4 (4:1) and frozen at –80°C until analysis.

Metabolites were subjected to HPLC analysis (R.Todorovic, P.Devanesan, J.Zhao, M.Gross, E.G.Rogan and E.L.Cavalieri, Metabolic response to treatment of Syrian golden hamsters with 4-hydroxyestradiol: novel analysis of estrogenic compounds in urine, manuscript in preparation) on a reverse phase Luna-2 C18 column (250x4.6 mm, 5 µm; Phenomenex) in a system equipped with dual ESA Model 580 solvent delivery modules, an ESA Model 540 autosampler and a 12 channel CoulArray detector (ESA, Chelmsford, MA) (Figure 3Go). Metabolites were separated using two mobile phases, I (0.1 M ammonium acetate, pH 4.4, acetonitrile and methanol, ratio 80:15:5) and II (ratio 30:50:20), in a linear gradient starting with 100% mobile phase I and ending with 90% mobile phase II over 50 min at a flow rate of 0.1 ml/min. The serial array of 12 coulometric electrodes was set at potentials between –10 and 590 mV (Figure 3Go). The system was controlled and the data were acquired and processed using the CoulArray software package (ESA). Peaks were identified by both retention time and peak height ratios between the dominant peak and the peaks in the two adjacent channels. Metabolites were quantified by comparison of peak response ratios with known amounts of standards.



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Fig. 3. Separation of E2 metabolites by HPLC with a 12 channel ESA CoulArray electrochemical detector. A linear gradient starting at 100% mobile phase I (acetonitrile, methanol and ammonium acetate, pH 4.4, ratio 15:5:80) to 90% mobile phase 2 (acetonitrile, methanol and ammonium acetate, pH 4.4, ratio 50:20:30) over 50 min was employed at a flow rate of 0.1 ml/min. The oxidation potentials of the detector channels were set between –10 and 590 mV.

 
Statistical analysis
Logarithmic transformations and linear regression analysis were carried out using SigmaStat software (Jandel, Corte Madera, CA) to generate standard curves for the mRNA transcripts. Correlations between the levels of E2 metabolites and CYP1 mRNA transcripts were calculated using Pearson correlation analysis. For comparisons between groups versus controls, Student's t-test was used to determine whether the differences were statistically significant.


    Results
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
The effects of CHCs on the expression of CYP1A1, CYP1A2 and CYP1B1 mRNA and on the metabolism of E2 were studied in the liver, kidney and mammary gland of Sprague–Dawley rats administered TCDD, 2,4-D or DED at the LD50 level for 72 h. TCDD was included to serve as a prototype for the CHCs examined in the present study.

Expression of CYP1A1, CYP1A2 and CYP1B1 were increased significantly (P < 0.05) in the three organs in response to treatment with the CHCs (Figure 4Go). In control animals CYP1A1 was expressed at levels of ~4, 1.2 and 0.3x106 copies/µg total RNA, whereas CYP1B1 mRNA levels were ~7, 3 and 1x103 copies/µg RNA in the liver, kidney and mammary gland, respectively. However, CYP1A2 was expressed at levels of ~11 and 1.3x107 copies/µg RNA in the liver and mammary gland, respectively, but was not detected in the kidney in control animals. The level of CYP1A2 mRNA was 30-fold higher than CYP1A1 and 15 000-fold higher than CYP1B1.



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Fig. 4. Expression of CYP1A1, CYP1A2 and CYP1B1 in response to treatment with CHCs. The alterations in response to treatment were all significantly different from the corresponding control values (P < 0.05).

 
TCDD induced expression of the three CYP1 enzymes in the liver, kidney and mammary gland by 35- to 50-, 15- to 20- and 6- to 10-fold, respectively, over the corresponding control values. This induction was the maximal and most potent compared with the effects of the other CHCs tested. Induction of CYP1 enzyme mRNAs by TCDD was followed, to a similar extent, by the effects of 2,4-D and DED (Figure 4Go). Expression of ß-actin did not change significantly in relation to treatment with CHCs and varied only from ~6 to 8x108 copies/µg RNA between the control and treatment groups.

Metabolites of E2 were detected after incubation with microsomes from the liver, kidney and mammary gland of all control animals. As expected, liver microsomes had the highest capacity to metabolize E2, followed by kidney and then mammary gland microsomes. In the liver, TCDD, 2,4-D and DED significantly increased the 2-, 4- and 16{alpha}-hydroxylation of E2 (Table IIGo). However, in the kidney and mammary gland only 2- and 4-hydroxylation, but not 16{alpha}-hydroxylation, were significantly elevated in response to treatment. The average ratio 2-OH(E1)E2:4-OH(E1)E2 was ~14, 4 and 3, respectively, with liver, kidney and mammary gland microsomes (Table IIGo). This ratio was not changed appreciably in response to CHC treatment in any of the examined tissues. Interestingly, CHCs significantly reduced the ratio 2-OH(E1)E2:16{alpha}-OH(E1)E2 in the liver, but not in kidney or mammary tissue (Table IIGo).


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Table II. Effect of chlorinated hydrocarbons on the metabolism of E2 in tissues of female Sprague–Dawley rats
 
The results of correlation analyses across tissues showed that induction of CYP1A1 and CYP1A2 by the three CHCs was associated with the formation of 2-catechol estrogens (r = 0.66 and 0.72, P < 0.05, respectively) and to a lesser extent with the formation of 4-catechol estrogens (r = 0.5 and 0.53, P < 0.05, respectively). CYP1B1 expression, however, correlated well with the formation of 4-catechol estrogens (r = 0.74, P < 0.02), but not with 2-catechol estrogens (data not shown). In the three tissues, expression of CYP1A1 and CYP1A2 was significantly higher (P < 0.05) than that of CYP1B1 and 2-hydroxylation was higher than 4- or 16{alpha}-hydroxylation (P < 0.05) in the experimental and control groups.


    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
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 References
 
Induction of CYP enzymes by xenobiotics produces an imbalance between activation and detoxification pathways, leading to adverse effects associated with exposure to these agents (41). Analysis of CYP induction can, therefore, provide insight into the mechanisms by which a variety of environmental factors induce toxic and carcinogenic effects.

In the present study we have investigated the effect of CHCs on expression of CYP1 enzyme genes that are involved in estrogen biotransformation (i.e. CYP1A1, CYP1A2 and CYP1B1) in the liver, kidney and mammary gland of female Sprague–Dawley rats. TCDD serves as a prototype for chemicals that act via the aryl hydrocarbon receptor (AhR) to produce adverse health effects. However, a variety of other synthetic or naturally occurring lipophilic compounds of diverse structures can bind to the AhR and induce gene expression to varying extents (42).

TCDD, 2,4-D and DED elicited significant effects on the individual CYP1 mRNA transcripts, with TCDD, the high affinity agonist for the AhR, the most potent inducer of CYP1 mRNA transcripts, followed, to a similar extent, by the effects of 2,4-D and DED (Figure 4Go). Moreover, CYP1A1, CYP1A2 and CYP1B1 mRNAs were induced differently in response to the same CHC (Figure 4Go). This varying response was observed previously for CYP1 enzymes in the rat liver, kidney and lung following exposure to small doses of TCDD (43) or administration of agents other than CHCs, such as nicotine (44). The different effects of each CHC on the CYP1 enzymes and the distinctive pattern of responses of the individual CYP1 enzyme genes to CHCs could be due to the extent of AhR involvement in the process of transcriptional activation. CYP1 genes are controlled by the same AhR, although their transcription and mRNA stabilization and concentration respond differently to the same inducer in a tissue-specific manner (25). TCDD and related CHCs are also known to differ in their biological effects, although they invoke them via a common AhR-mediated process (45). The mechanism of induction of gene transcription by TCDD and related CHCs involves ligand recognition and binding by the AhR, nuclear translocation and dimerization with the AhR nuclear translocator protein (18). The nuclear heterodimer then recognizes and interacts with a specific DNA sequence, designated xenobiotic-responsive elements, located in the promoter/enhancer regions of multiple AhR-responsive genes (4648). This interaction occurs within the major groove of the DNA helix (46) and, therefore, leads to changes in chromatin structure and/or alterations in the basal transcriptional machinery, which results in the induction of gene expression (45).

Induction of CYP1A1, CYP1A2 and CYP1B1 mRNA transcripts in response to CHC treatment was accompanied by increases in the 2- and 4-hydroxylation of E2 in the liver, kidney and mammary gland (Table IIGo and Figure 4Go). Correlation analysis across tissues revealed that CHC-induced increases in CYP1A1 and CYP1A2 transcripts were accompanied by increased formation of 2-catechol estrogen and, to a lesser extent, 4-catechol estrogen, while CYP1B1 expression correlated well with the formation of 4-catechol estrogen, but not with 2-catechol estrogen (data not shown). The formation of 2-OH(E1)E2 is known to be mediated, to a large extent, by CYP1A1 and CYP1A2, which also have some 4- and 16{alpha}-hydroxylation activity (20,21,49). On the other hand, CYP1B1 is the major enzyme that catalyzes the synthesis of 4-catechol estrogen (20,21), with a minor 2-hydroxylation capacity (A.F.Badawi et al., unpublished data). Formation of 2- and 4-OH(E1)E2 usually responds coordinately to the same inhibitor and inducer (50). In support of this observation, the average ratio 2-OH(E1)E2:4-OH(E1)E2 was ~14, 4 and 3 in the liver, kidney and mammary gland, respectively, and was not altered significantly by CHC treatment (Table IIGo). This ratio varies from 4 (38) to 27 (38,39) in the livers of untreated rats. In contrast to the unchanged ratio of 2-catechol estrogens:4-catechol estrogens, the ratio 2-OH(E1)E2:16{alpha}-OH(E1)E2 was decreased in the liver following exposure to CHCs (Table IIGo) and was not affected in the kidney and mammary gland. Moreover, this ratio has been shown to be lower in extrahepatic tissues compared with values in the liver (Table IIGo). This tissue-specific pattern of 2-OH(E1)E2:16{alpha}-OH(E1)E2 may be linked to the abundant expression of hepatic CYP3A4/5 (51), the most active enzyme catalyzing synthesis of 16{alpha}-OH(E1)E2 (49) and the lower inducibility of CYP1A1 and CYP1A2 in the mammary gland and kidney compared with their hepatic levels (Figure 4Go and Table IIGo). CHCs increased the rate of 16{alpha}-hydroxylation of E2 in breast cancer cells (52), an effect thought to be involved in their potential to induce mammary carcinogenesis (52). However, several lines of evidence suggest that 4-OH(E1)E2, but not 16{alpha}-OH(E1)E2, is associated with increased risk of breast cancer (20,22). In support of this idea, the results reported here show that formation of 4-OH(E1)E2 was more pronounced than that of 16{alpha}-OH(E1)E2 in the mammary gland in response to CHCs.

Induction of CYP1 mRNA usually occurs rapidly and in the absence of similar rates of protein synthesis (18). This primary, or adaptive, response may influence the mRNA and protein to respond differently to a given CYP inducer (43). Indeed, in the present study induction of CYP1 mRNAs was not paralleled by a similar increase in the activity of the enzymes, as was evident from E2 metabolism (Figure 4Go and Table IIGo). In agreement, Kimura et al. (43) reported that TCDD treatment caused a 27-fold increase in liver CYP1A1 mRNA, which was associated with only an 8-fold rise in the level of protein. Similarly, in the kidney a 12-fold increase in CYP1A1 mRNA was accompanied by a 2-fold rise in the protein content (43). Furthermore, CYP1B1 protein was only detected at doses of 35.7 ng TCDD/kg/day for 30 weeks, while its mRNA was significantly induced and detected at one-tenth of this dose (26). This varying response between CYP1 enzyme and the corresponding mRNA transcripts was not limited to TCDD or the related CHCs, but was also apparent in the lung, kidney and liver of male and female rats in response to inducers such as pyridine (53).

Occupational exposure to CHCs has been linked epidemiologically to a high incidence of a variety of human cancers. For example, 2,4-D has been suggested as a risk factor for non-Hodgkin's lymphoma (810) and lung (11) and breast cancer in humans (10), whereas DED was associated with an increased incidence of human liver (14) and breast (15) cancers. However, experimental evidence in animals stands in contrast to the epidemiology and indicates that these agents are not likely to be carcinogenic in humans (6,7,12,13). The results of the present study suggest that CHCs, although not carcinogenic in experimental animals, may still play an important role in neoplastic transformation in human populations exposed to these agents. CHCs can induce transcription of a variety of genes coding for estrogen metabolism, particularly CYP1B1, and can, therefore, increase the endogenous synthesis of 4-catechol estrogens. We have demonstrated previously that 4-catechol estrogens may be oxidized to quinones that react with DNA to form depurinating and stable adducts (2224), which may lead to a variety of oncogenic mutations and initiation of cancer. In addition, various studies have concluded that 4-catechol estrogens are associated with an increased risk of human breast cancer (54,55) and were found to be carcinogenic in hamster kidney (56,57). In support of this association, CYP1B1 was overexpressed in 92% of human breast cancer specimens, compared with normal tissue (58). The role of CYP1B1 in 4-catechol estrogen formation and in estrogen-induced carcinogenesis should be interpreted with caution, given the low levels of the enzyme in mammary tissue even after induction.

In conclusion, the present study demonstrates that the biological effects of CHCs may be associated with sustained changes in the expression of genes regulating estrogen biotransformation. These primary persistent alterations may elicit secondary compensatory changes, such as a steady-state up-regulation in estrogen metabolism and subsequent synthesis of catechol estrogens. Further oxidation of these metabolites could lead to the formation of catechol estrogen quinones and DNA adducts that can produce mutations leading to neoplastic transformation.


    Notes
 
1 To whom correspondence should be addressed Email: egrogan{at}unmc.edu Back


    Acknowledgments
 
We thank Dr R.Todorovic for her advice on the HPLC analysis of E2 metabolites. This work was supported by US Public Health Service grant no. P01CA 49210 from the National Cancer Institute. Core support to the Eppley Institute was provided by grant CA 36727 from the National Cancer Institute.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received February 8, 2000; revised April 18, 2000; accepted May 1, 2000.