Characterization and Functional Analysis of Two Common Human Cytochrome P450 1B1 Variants

Archives of Biochemistry and Biophysics Vol. 378, No. 1, June 1, pp. 175–181, 2000 doi:10.1006/abbi.2000.1808, available online at http://www.idealibrary.com on

Characterization and Functional Analysis of Two Common Human Cytochrome P450 1B1 Variants Roman A. McLellan,* Mikael Oscarson,* Mats Hidestrand,* Brith Leidvik,† Eva Jonsson,† Charlotta Otter,† and Magnus Ingelman-Sundberg* ,1 *Division of Molecular Toxicology, National Institute of Environmental Medicine, Karolinska Institutet, Box 210, SE-171 77 Stockholm, Sweden; and †Department of Molecular Biology, AstraZeneca R&D, SE-90736 Umeå, Sweden

Received December 16, 1999, and in revised form March 6, 2000

Cytochrome P450 1B1 (CYP1B1) is a human extrahepatic P450 that activates procarinogens, metabolizes 17␤-estradiol, and may well have a role in the pathogenesis of some forms of cancer. Besides rare deleterious mutations reported for the CYP1B1 gene, six single-nucleotide polymorphisms have been reported, of which four cause amino acid exchanges. We have expressed two of the common CYP1B1 alleles in yeast cells and mammalian COS-1 cells in order to functionally characterize the alleles with respect to kinetic properties and protein stability. The CYP1B1.2 variant contains the two linked amino acid substitutions R48G and A119S compared to CYP1B1.1. The kinetic parameters of two structurally unrelated CYP1B1 substrates for the two variants were examined. No kinetic differences were seen of 17␤-estradiol hydroxylation activities between the two CYP1B1 variants and an only minor increase in the apparent K m for ethoxyresorufin was observed for CYP1B1.2. It therefore appears that they have very similar catalytic properties and the substitutions do not appear to alter CYP1B1 catalytic function. The two CYP1B1 variants were similarly stable when expressed in mammalian COS-1 cells, indicating that the substitutions have no effect on protein folding or stability. The combined results indicate that these two CYP1B1 variants show very similar properties with respect to catalytic activities and protein stability and do not alter CYP1B1 function. © 2000 Academic Press Key Words: cytochrome P450; CYP1B1; 17␤-estradiol; genetic polymorphism; cancer pathogenesis.

1 To whom correspondence should be addressed. Fax: ⫹46-8-33 73 27. E-mail: [email protected].

0003-9861/00 $35.00 Copyright © 2000 by Academic Press All rights of reproduction in any form reserved.

The cytochrome P450s are a superfamily of enzymes that are mainly involved in the oxidative metabolism of many drugs, foreign chemicals, and carcinogens and endogenous substrates such as steroids, fatty acids, and vitamins. Cytochrome P450 1B1 (CYP1B1) 2,3 is an extrahepatic human P450 belonging to the CYP1 family and the amino acid sequence is ⬃40% identical to that of CYP1A1 and CYP1A2 (1). It has catalytic activities that overlap CYP1A1 and CYP1A2 with respect to some drugs such as caffeine and theophylline (2). Unlike the CYP1A1 and CYP1A2 genes, which are located on chromosome 15, the CYP1B1 gene has been localized to chromosome 2 in the region 2p21-22 (3, 4). The gene structure consists of three exons, with the open reading frame starting in exon two (4). CYP1B1 is the only known member of the CYP1B subfamily and Southern blot analysis has revealed that it is unlikely that other CYP1B genes exist in humans (3, 4). CYP1B1 can be induced by aryl hydrocarbon receptor ligands (3, 5– 8) and is expressed constitutively in several extrahepatic tissues, including the kidney, steroidogenic tissue (adrenal, ovary, and testis), and steroidresponsive tissue (breast, uterus, and prostate) (3, 9). It is also expressed at high levels in human breast tumors (10, 11) as well as a wide range of other malignant tumors (12). Similar to CYP1A1 and CYP1A2, CYP1B1 can activate many structurally diverse environmental and dietary procarcinogens such as polycyclic aromatic hy2

Abbreviations used: CYP or P450, cytochrome P450; PAH, polycyclic aromatic hydrocarbon; E 2, 17␤-estradiol; 2-OHE 2, 2-hydroxyestradiol; 4-OHE 2, 4-hydroxyestradiol; SNP, single-nucleotide polymorphism; SRS, substrate recognition site; EROD, ethoxyresorufin O-deethylation. 3 The present study uses the nomenclature system for CYP1B1 alleles recommended by the Human Cytochrome P450 Allele Nomenclature Committee; see http://www.imm.ki.se/CYPalleles/. 175

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drocarbons (PAHs) and their dihydrodiol derivatives, heterocyclic and aryl amines, and nitroaromatic hydrocarbons to reactive intermediates that can damage DNA in cells (9, 13). This was demonstrated in a recent study where mice were administered the potent carcinogen 7,12-dimethylbenz[␣]anthracene (DMBA) and Cyp1b1-null mice were protected against DMBA-induced tumors, whereas wild-type animals developed malignant lymphomas (14). This showed that extrahepatic CYP1B1 mediated the carcinogenicity of DMBA and demonstrated the importance of extrahepatic P450s such as CYP1B1 in determining susceptibility to chemical carcinogens. Human CYP1B1 is involved in the metabolism of the physiological steroid 17␤-estradiol (E 2). It is the most catalytically efficient E 2 hydroxylase characterized to date, largely due to its low apparent K m for E 2. CYP1B1 catalyzes the hydroxylation of E 2 primarily at the C-4 position and to a lesser extent at C-2 (15). This 4-OHE 2 activity has been shown to be elevated in human mammary and uterine tumors compared to surrounding tissue (16, 17) and 4-OHE 2 was found to be carcinogenic in some animal models (18). It is very likely that CYP1B1 plays a role in E 2 homeostasis in extrahepatic tissues such as the breast and may control local E 2 concentrations. Other steroid hormones, including estrone, testosterone, and progesterone, have been shown to be metabolized by CYP1B1 (19). Another physiological role that CYP1B1 appears to be involved in is ocular development and differentiation in humans. In support of this, rare mutations in the CYP1B1 gene have been identified as a molecular basis for primary congenital glaucoma, an autosomal recessive eye disease (20 –22). To date, 17 different truncating or missense mutations have been found. Besides these rare deleterious mutations reported for the CYP1B1 gene that have been found exclusively in patients with primary congenital glaucoma, six singlenucleotide polymorphisms (SNPs) have also been reported (21). Five of the SNPs are located in the coding region, of which four result in the amino acid substitutions R48G, A119S, L432V, and N453S. The sixth SNP is located in intron one, 13 base pairs upstream from the ATG start codon. Of the substitutions, R48G and A119S are always found together, indicating that the two SNPs causing these substitutions are linked, in contrast to the other two, which are not (21). (R. A. McLellan, unpublished observations). Some CYP1B1 variants have been recently functionally examined where slight differences were observed in catalytic properties, although not all of the combinations used actually exist in the human population (19). Since this enzyme activates procarcinogens, metabolizes E 2, and may well have a role in cancer pathogenesis, it is of great importance to examine the different alleles and determine if the amino acid substitutions

result in any alterations to enzyme function. To this end, we have expressed the CYP1B1*1 allele and the common CYP1B1*2 allele having R48G and A119S amino acid substitutions, the first one adjacent to the PPGP region of importance for protein folding (23) and the second in substrate recognition site (SRS) one (24), in yeast cells and mammalian COS-1 cells. These heterologous expression systems allowed for the functional characterization of the enzyme variants with respect to kinetic properties and protein stability. MATERIALS AND METHODS Cloning of CYP1B1 cDNA variants into the pYeDP60 expression vector. As the CYP1B1 gene only contains two coding exons, the 5⬘ and 3⬘ parts of CYP1B1*1 and CYP1B1*2 cDNAs were cloned from human genomic DNA (Clontech). The central part of the CYP1B1*1 cDNA was cloned from a human liver cDNA library using PCR primers and the Marathon cDNA amplification kit (Clontech) and a synthetic oligonucletide was used to introduce a stop codon and the correct cloning site at the 3⬘ end. The CYP1B1*2 5⬘ and 3⬘ parts were joined with a synthetic oligonucleotide and the 3⬘ end was constructed, as described for CYP1B1*1. The full-length variant cDNAs were cloned into the pYeDP60 (V60) yeast expression vector (25), which is under the control of a galactose-driven promoter, using the restriction enzymes BamHI and EcoRI. An AAA sequence was introduced in front of the translation initiation sites as it has previously been shown to increase expression levels of CYP2D6 (26). All cDNAs were sequenced using the ABI Prism Big Dye terminator cycle sequencing kit and analyzed with an ABI Prism 377 DNA sequencer to ensure correct constructs and to exclude any potential PCR artifacts. Expression of CYP1B1 cDNA variants in yeast cells. Yeast strain INVSc1-HR MAT ␣his3⌬1 leu2 trp1-289 ura3-52 (pFL-35 human reductase), a gift from the LINK project (a program of the University of Dundee/Biotechnology and Biology Research Council/Department of Trade and Industry/Pharmaceutical Industry), was transfected with the V60-CYP1B1 expression plasmids described above. Yeast cells were inoculated into selective medium (6.7 g/L yeast nitrogen base, 5 g/L glucose, 20 ␮g/ml histidine, 20 ␮g/ml leucine) and grown at 30°C overnight in a shaking flask. Fifty milliliters of preculture was inoculated in fermentation medium (20 g/L yeast extract, 20 g/L peptone, 2% glucose) and grown in a fermentor with stirring at 500 rpm, temperature at 28.5°C, air flow at 0.5 L/min, and pH controlled at 6.6. After 12 h of fermentation, 30 ml of 50% glucose and ethanol was added to a final concentration of 3%. Following an additional fermentation for 24 h, cells were induced by the addition of galactose to a final concentration of 2% and fermented for another 7 h. Cells were harvested by centrifugation, washed in distilled water, and resuspended in TEK buffer (50 mM Tris–HCl buffer, pH 7.4, 1 mM EDTA, 100 mM KCl). After 5 min of incubation at 20°C and recentrifugation, the pellet was resuspended in 10 mM Tris–HCl buffer, pH 7.5, containing 2 M sorbitol, 0.1 mM dithiothreitol, 0.1 mM EDTA, and 5 mg/ml yeast lytic enzyme. The suspension was gently shaken for 1 h at 30°C and then centrifuged. The pellet was resuspended in TES buffer (50 mM Tris–HCl buffer, pH 7.4, 1 mM EDTA, 0.6 M sorbitol) containing 10% glycerol and 4 mM Pefabloc SC (Roche). Cells were broken in a Microfluidizer (Microfluidics Corp.), where the yeast suspension was passed seven times at 20,000 psi. The suspension was centrifuged 5 min at 7000 rpm. The supernatant was centrifuged 10 min at 15,000 rpm and the pellet discarded, with this being repeated three times. The supernatant was diluted to 30 ml with TES buffer containing 0.4 mM Pefabloc SC. NaCl (5 M, 0.75 ml) was added, followed by 7.5 ml of 50% aqueous PEG 4000. The mixture was placed on ice for 15 min and then centrifuged 10 min at 12,000g. The surface of the pellet was washed with 2 ml of TEG

FUNCTIONAL CHARACTERIZATION OF HUMAN CYP1B1 VARIANTS buffer and the pellet homogenized in 1 ml of TEG buffer for each 500 ml of cultured media. Microsomes were stored at ⫺80°C until use. Microsomal protein concentration (27), reductase levels (28), and total cytochrome P450 content (29) were determined employing previously described methods. Variant CYP1B1 17␤-estradiol hydroxylation activities in yeast microsomes. Yeast microsomal E 2 hydroxylase activities were determined in an incubation mixture (final volume 0.5 ml) containing 20 pmol of P450, 100 mM potassium phosphate buffer, pH 7.4, 200 ␮g of NADPH, and varying amounts (0 –12.8 ␮M) of [4- 14C]E 2 (NEN Life Science Products) as substrate. All incubations contained equivalent amounts of P450 and total microsomal protein, which was done by adjusting protein levels with microsomes from yeast transfected with empty V60 plasmid. After 5 min of incubation at 37°C, reactions were terminated by the addition of 2 ml of dichloromethane. All measurements were performed with an incubation time and with an amount of P450 that would ensure linear reaction rates. Samples were vortexed and then centrifuged for 10 min at 3500 rpm, and the organic phase was subsequently removed and evaporated to dryness under a stream of nitrogen. The residue was dissolved in mobile phase and subjected to a reversed-phase HPLC system with a radioactive detection method. Separation of the compounds was done at room temperature on a C 18 reversed-phase column (250-4, 5 ␮M, Merck). The elution was performed with a mobile phase consisting of 38% acetonitrile (v/v) in 1% acetic acid (v/v) at a flow rate of 1 ml/min. The metabolites formed were identified by comparing their retention times with authentic standards and product amounts were calculated based on the specific activity of [4- 14C]E 2 substrate. Variant CYP1B1 ethoxyresorufin O-deethylation (EROD) activities in yeast microsomes. Yeast microsomal EROD activities were determined in an incubation mixture (final volume 0.5 ml) containing 4 pmol of P450, 100 mM potassium phosphate buffer, pH 7.4, 200 ␮g of NADPH, and varying amounts (0 –12.8 ␮M) of ethoxyresorufin (Sigma) as substrate. All incubations contained equivalent amounts of P450 and total microsomal protein, which was done by adjusting protein levels with microsomes from yeast transfected with empty V60 plasmid. After 2 min of incubation at 37°C, reactions were terminated by the addition of 2 ml of ether. All measurements were performed with an incubation time and with an amount of P450 that would ensure linear reaction rates. Samples were vortexed and then centrifuged for 2 min at 3500 rpm, and the organic phase was subsequently removed and evaporated to dryness under a stream of nitrogen. The residue was dissolved in mobile phase and subjected to a reversed-phase HPLC system with fluorescence detection at 530 (excitation) and 580 nm (emission). Analysis of the metabolite resorufin was done at room temperature on a C 18 reversed-phase column (125-4, 5 ␮M, Merck). The elution was performed with a mobile phase consisting of 25 mM potassium phosphate buffer, pH 7.4/methanol (60:40, v/v) at a flow rate of 1 ml/min. Resorufin was quantified by comparison with a standard curve using authentic resorufin (Sigma). Cloning of CYP1B1 alleles from human genomic DNA into the pCMV4 expression vector. The entire coding region of both CYP1B1*1 and CYP1B1*2 alleles was amplified from human genomic DNA with the GeneAmp XL PCR kit (Perkin Elmer) using the primers 5⬘-GACAGATCTAGCATGGGCACCAGCCTCAG-3⬘ and 5⬘-GACAAGCTTGCACACCTCACCTGATGGAC-3⬘ (restriction sites underlined). The PCR fragments were subsequently cloned into the mammalian cell expression vector pCMV4 (30) using BglII and HindIII restriction sites. The exons and exon–intron junctions were sequenced as described above to exclude potential PCR artifacts. Expression of CYP1B1 alleles in mammalian COS-1 cells. COS-1 cells were transiently transfected with the CYP1B1*1 and CYP1B1*2 plasmids in parallel as described previously (31). After a 40-h incubation, cells received fresh medium without serum containing 10 ␮g/ml cycloheximide. Cells were then harvested in 100 mM sodium phosphate buffer, pH 7.4, at times 0, 2, 4, 6, and 8 h following

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cycloheximide addition. Cellular disruption was achieved by sonicating for 20 ⫻ 1 s and the resulting homogenates were analyzed for CYP1B1 content by Western blot analysis. Quantification of CYP1B1 apoprotein levels in COS-1 cells. Cell homogenate corresponding to 15 ␮g of protein was subjected to sodium dodecyl sulfate gel electrophoresis using 8.5% polyacrylamide gels. The proteins were transferred to a nitrocellulose filter (Bio-Rad) and incubated with anti-CYP1B1 antibody (Gentest) and then with a secondary horseradish peroxidase linked goat anti-rabbit antibody (DAKO AS). To visualize the proteins, the enhanced chemiluminescence method (Amersham) was used. Protein quantification was done using a personal densitometer (Molecular Dynamics). Data analysis. The apparent kinetic parameters (K m and V max) for E 2 hydroxylase and EROD activities were estimated using the Enzfitter (Biosoft) computer program by fitting the data to the Michaelis–Menten equation. Statistical analysis was performed using Student’s t test. All data are expressed as the mean ⫾ SE.

RESULTS

Expression of CYP1B1 variants in yeast. To achieve amounts of protein necessary for kinetic analysis, the two full-length CYP1B1 cDNA variants were constructed and cloned into the V60 expression vector, which is under the control of a galactose-driven promoter. The CYP1B1 plasmids were transfected into a yeast strain which was modified by genome insertion of the human reductase gene. Microsomal fractions containing either of the CYP1B1 variants had similar levels of reductase, 145 ⫾ 28 and 117 ⫾ 3 pmol/mg of protein for CYP1B1.1 and CYP1B1.2 fractions, respectively. Levels of CYP1B1 holoprotein were determined spectrophotometrically from the reduced CO-bound form. The P450 levels were 7 and 24 pmol/mg of protein for CYP1B1.1 fractions and 7 and 14 pmol/mg of protein for CYP1B1.2 microsomal fractions. These low levels of CYP1B1 protein have been previously reported when unmodified full-length CYP1B1 cDNA is used for in vitro expression studies (15). Despite this, it was desired to analyze unmodified CYP1B1 variants containing the entire amino acid sequence. Kinetic analysis of CYP1B1 variants. Kinetic studies were done using E 2 in order to characterize the CYP1B1 variants with respect to their catalytic properties for this physiological substrate. As shown in Table I, there were no significant differences between the apparent K m and V max for the CYP1B1.1 and CYP1B1.2 variants. E 2 4-hydroxylase activity was much higher than 2-hydroxylase activity for both of the variants. The V max ratios of E 2 4-hydroxylation versus 2-hydroxylation were found to be 4.6 ⫾ 0.5 and 4.8 ⫾ 0.6 for CYP1B1.1 and CYP1B1.2, respectively. These results are in good agreement with the range of K m and V max values and the E 2 4-hydroxylation 2-hydroxylation ratio determined in a previous study using yeast-expressed CYP1B1 (15). The V max/K m ratios for E 2 4-hydroxylation and 2-hydroxylation were 0.27 ⫾ 0.01 and 0.02 ⫾ 0 for CYP1B1.1 and 0.24 ⫾ 0.01 and 0.02 ⫾ 0.01

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Kinetic Properties for E 2 4- and 2-Hydroxylation by Human CYP1B1 Variants K ma (␮M)

a V max (pmol/min/pmol of P450)

CYP1B1 variant

4-Hydroxylation

2-Hydroxylation

4-Hydroxylation

2-Hydroxylation

V max ratio 4-OH/2-OH

CYP1B1.1 CYP1B1.2

3.5 ⫾ 0.6 3.3 ⫾ 0.3

10.6 ⫾ 2.3 10.2 ⫾ 2.7

0.94 ⫾ 0.13 0.80 ⫾ 0.05

0.22 ⫾ 0.06 0.17 ⫾ 0.02

4.6 ⫾ 0.5 4.8 ⫾ 0.6

a

Mean ⫾ SE, determined from three experiments using two independent preparations of microsomes for each variant.

for CYP1B1.2, respectively, demonstrating that they were similar between the variants. To further investigate the catalytic properties of the two variants, ethoxyresorufin O-deethylation activity was measured using a sensitive HPLC method. The velocities of CYP1B1.1 and CYP1B1.2 were determined at seven different concentrations of ethoxyresorufin (Fig. 1). The resulting apparent kinetic parameters of K m and V max are shown in Table II. While the V max values of EROD activities were nearly identical, a small but significant increase was observed for the K m values, 0.57 ⫾ 0.06 and 0.79 ⫾ 0.03 ␮M for CYP1B1.1 and CYP1B1.2, respectively. The minor difference observed between K m values is also reflected in the V max/K m ratios, which were somewhat altered, with CYP1B1.1 having a value of 20.9 ⫾ 2.3 and CYP1B1.2 a value of 14.8 ⫾ 2.4. Stability of CYP1B1 variants in mammalian COS-1 cells. The coding region of the two alleles, CYP1B1*1 and CYP1B1*2, were amplified from human genomic DNA using a long PCR protocol, cloned into the pCMV4 expression vector, and expressed in COS-1 cells. In order to study protein stability, the degradation of the two CYP1B1 variants was followed after the addition

of cycloheximide, a protein synthesis inhibitor (32). Cells remained viable upon visual inspection at the latest time point used. The levels of CYP1B1 apoprotein were measured by Western blot analysis. Following transfection and a 40-h incubation period, both CYP1B1.1 and CYP1B1.2 yielded immunodetectable apoprotein immediately prior to cycloheximide treatment. A representative Western blot is shown in Fig. 2, which demonstrates the amounts of CYP1B1 variant apoprotein levels when isolated at different times after the addition of cycloheximide. Densitometric analysis of the Western blots revealed that the relative levels of the two CYP1B1 variants were very similar over the entire time course when expressed as a percentage of pretreatment levels at time 0 h (Fig. 3). DISCUSSION

In this study, we have examined and compared the functional characteristics of two common human CYP1B1 variants encoded by CYP1B1*1 and CYP1B1*2. The CYP1B1.2 variant contains the two linked amino acid substitutions R48G and A119S compared to CYP1B1.1, and the SNPs causing these substitutions are present at a frequency of 29% in the Caucasian population (21). We observed no major differences in either the catalytic activities or protein stability of the variants, which suggests that these substitutions do not significantly alter CYP1B1 function. SNPs leading to amino acid substitutions in the P450 enzymes have been demonstrated to alter the in

TABLE II

Kinetic Properties for Ethoxyresorufin O-Deethylation by Human CYP1B1 Variants CYP1B1 variant

FIG. 1. Dependence of CYP1B1.1 (Œ) and CYP1B1.2 (■) EROD activities on ethoxyresorufin concentration. Yeast microsomal incubation mixtures contained 4 pmol of either CYP1B1 variant and 0.0, 0.2, 0.4, 0.8, 1.6, 3.2, 6.4, or 12.8 ␮M ethoxyresorufin as substrate. Each point represents the mean ⫾ SE of three determinations.

CYP1B1.1 CYP1B1.2

K ma (␮M)

a V max (pmol/min/pmol of P450)

0.57 ⫾ 0.06 0.79 ⫾ 0.03 b

11.7 ⫾ 0.2 11.6 ⫾ 1.5

a Mean ⫾ SE, determined from three experiments using two independent preparations of microsomes for each variant. b Significantly different from CYP1B1.1, P ⬍ 0.05.

FUNCTIONAL CHARACTERIZATION OF HUMAN CYP1B1 VARIANTS

FIG. 2. Representative Western blots demonstrating the apoprotein levels of CYP1B1.1 and CYP1B1.2 that were present in mammalian COS-1 cells when isolated at different times after addition of cycloheximide, a protein synthesis inhibitor. Each lane contained cell homogenate corresponding to 15 ␮g of protein, and a polyclonal anti-CYP1B1 antibody (Gentest) was used to detect CYP1B1.

vitro catalytic properties of the enzyme, which has translated into measurable in vivo effects in humans. One example is the CYP2C9*3 allele, which contains an SNP that results in an I359L substitution. The corresponding CYP2C9.3 enzyme has higher K m values for the substrates tolbutamide (33) and S-warfarin (34) while V max values are similar or lower compared to the wild-type enzyme. The presence of CYP2C9.3 has also been shown to decrease the clearance of CYP2C9 substrates such as S-warfarin in vivo (35). Another example of altered enzyme function involves CYP2D6.17, which contains three amino acid exchanges compared to the wild-type enzyme. The T107I substitution alone caused an increase in the K m for codeine but the R296C substitution was also needed to increase the K m for the substrate bufuralol (36). Decreased in vivo CYP2D6 activity in humans was caused by the altered catalytic properties of CYP2D6.17. When CYP2 amino acid sequences were aligned with those of bacterial P450 cam (CYP101), six putative SRSs were identified for the CYP2 family (24). Most amino acid substitutions shown to affect substrate binding are indeed located in these SRSs (37, 38). As illustrated in Fig. 4, the A119S substitution found in the CYP1B1.2 enzyme is located in SRS-1, suggesting a potential role for this amino acid in substrate binding. To examine this possibility, we examined the kinetic parameters of two structurally unrelated CYP1B1 substrates for the CYP1B1.1 and CYP1B1.2 variants. As there were no kinetic differences seen for E 2 hydroxylation activities between the two CYP1B1 variants and a only minor difference in the apparent K m for ethoxyresorufin, it appears that they have very similar catalytic properties and the amino acid substitutions do not appear to alter CYP1B1 catalytic function. In agreement, Shimada et al. (19) observed no significant differences when they compared E 2 hydroxylase activities between CYP1B1.1 and an A119S variant. There still remains the possibility, however, that other substrates such as the PAHs may display altered kinetic parameters when these CYP1B1 variants are com-

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pared. It could also be the case that a combination of particular substitutions may be necessary to alter the function of the enzyme, as has been shown for CYP2D6.17 (36). The R48G substitution in CYP1B1.2 is located in the N-terminal spanning domain of the protein only two amino acids upstream of the very well conserved PPGP region (Fig. 4). This motif is found in almost all P450s belonging to families 1 and 2 and is thought to be important for proper protein folding and stability (23). An example illustrating the importance of this prolinerich region is the CYP2D6*10 allele, which encodes a P34S substitution and leads to the expression of a more unstable enzyme (31). The frequent distribution of this allele in Asians explains the decreased capacity for CYP2D6-dependent metabolism in this population. Other reports have further established that the proline residues in this region are crucial for the formation of functional enzyme (39 – 41). It was therefore reasonable to expect that the R48G substitution could have an effect on protein stability due to the drastic nature of the amino acid exchange. In particular, glycine residues are often involved in protein bending and have important structural functions. Despite this, the two CYP1B1 variants were similarly stable when expressed in mammalian COS-1 cells, indicating that the substitutions in the CYP1B1.2 variant have no effect on protein folding or stability. There still remains the need to examine if the other two substitutions found in the CYP1B1 enzyme, namely, L432V and N453S, result in functional alterations either by themselves or in combination with the linked R48G and A119S substitutions characterized here. This is being currently evaluated in our laboratory. It must also not be overlooked that the various CYP1B1 alleles may differ in either constitutive ex-

FIG. 3. Relative amounts of CYP1B1 variant apoprotein levels in COS-1 cells when isolated at various times after cycloheximide treatment as determined by densitometric analysis. CYP1B1.1 levels are shown in black bars and CYP1B1.2 in gray bars. Amounts are expressed as a percentage of pretreatment levels at time 0 h and represent the mean ⫾ SE of three experiments.

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FIG. 4. Sequence alignment of the N-terminal region of human (h) CYP1B1 with mouse (m) Cyp1b1 and rat (r) CYP1B1 and other human P450s from families one and two. The well-conserved PPGP region and SRS-1 are indicated by the black boxes. The R48G and A119S amino acid substitutions found in CYP1B1.2 are indicated in bold.

pression or inducibility. This could be due to other genetic polymorphisms in regulatory regions which might be linked to the known SNPs. The functional characterization of the CYP1B1 variants is important because this enzyme is involved in procarcinogen activation and metabolism of E 2. The levels of E 2 and E 2-hydroxylated metabolites are of great significance as a woman’s lifetime exposure to E 2 is a risk factor for developing breast cancer and 4-OHE 2 has been shown to be elevated in human tumor tissue (16, 17) and genotoxic in some animal models (18). In conclusion, we have expressed and examined two common human CYP1B1 variants, CYP1B1.1 and CYP1B1.2 for potential functional differences with respect to kinetic parameters and protein stability. The results of the present study indicate that these two CYP1B1 variants share very similar properties with respect to catalytic activities and protein stability. Since CYP1B1 could possibly play a role in human cancer pathogenesis because of its catalytic functions, any potential alterations in CYP1B1 function must therefore be fully characterized. ACKNOWLEDGMENTS The authors are grateful to Dr. Ian Cotgreave for allowing generous access to his HPLC equipment. This study was supported by grants from Swedish Match, the Swedish Medical Research Council, and the Swedish Society for Medical Research.

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