University of Minnesota Cancer Center, Minneapolis, MN 55455, USA
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Abstract |
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Abbreviations: 2-OH-THF, 2-hydroxytetrahydrofuran; 2-OH-THP, 2-hydroxy-tetrahydro-2H-pyran; GC-PICI-MS, gas chromatography-positive ion chemical ionization-mass spectrometry; HPLC-UV-radioflow, high performance liquid chromatography with in-line ultraviolet absorbance and radioflow detection; lactol, 2-hydroxy-5-(3-pyridyl)tetrahydrofuran; NAB, N'-nitrosoanabasine; NBzMA, N-nitrosobenzylmethylamine; NDMA, N-nitrosodimethylamine; NNN, N'-nitrosonornicotine; NPIP, N-nitrosopiperidine; NPYR, N-nitrosopyrrolidine; P450, cytochrome P450; [S], substrate concentration; Sf9, Spodoptera frugiptera
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Introduction |
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Nitrosamines require metabolic activation to exert their carcinogenic effects. The key activation pathway is cytochrome P450 (P450)-catalyzed hydroxylation of the carbon to the nitroso group. Extensive studies in laboratory animals have demonstrated that tissue-specific metabolic activation leads to tissue-specific DNA adduct formation and carcinogenesis by a number of nitrosamines (11,12). Kleihues and co-workers clearly demonstrated that the esophageal carcinogenicity of methylalkylnitrosamines in rats correlated with high levels of promutagenic methylated DNA adducts in target tissues (1315). Other studies showed that rat esophageal microsomes catalyzed the
-hydroxylation of esophageal carcinogens such as N'-nitrosonornicotine (NNN), N-nitrosobenzylmethylamine (NBzMA), N-nitrosodiethylamine, and N-nitrosomethyl-n-pentylamine with low KM values ranging from 5.1 to 64 µM (1619). Taken together, these results suggest that preferential metabolic activation of NPIP compared with NPYR in the rat esophagus might be one factor leading to its higher carcinogenicity in this tissue.
The metabolic activation of NPIP, leading to DNA adduct formation, is shown in Figure 1 (20,21). The unstable
-hydroxy-NPIP spontaneously decomposes to electrophilic intermediates that react with nucleophilic sites of DNA bases forming DNA adducts. Previous studies have identified several DNA adducts resulting from reactions of NPIP with DNA or deoxyguanosine (2225). The major water-trapped end product of NPIP
-hydroxylation is 5-hydroxypentanal, which cyclizes to 2-hydroxytetrahydro-2H-pyran (2-OH-THP) (20).
Metabolic -hydroxylation of NPYR, analogous to NPIP, generates the corresponding electrophilic intermediates shown in Figure 2
(2630). DNA adducts generated from
-hydroxylation of NPYR have been identified (22,23, 3135). The major water-trapped end product of NPYR
-hydroxylation is 4-hydroxybutanal, which exists predominantly as 2-hydroxytetrahydrofuran (2-OH-THF) (29).
In the present study, -hydroxylation rates (2-OH-THP versus 2-OH-THF formation) of NPIP and NPYR by rat esophageal or liver microsomes were compared to determine whether tissue-specific differences in metabolic activation exist for these two structurally related cyclic nitrosamines. Metabolism of these two nitrosamines in rat esophageal microsomes has not been studied previously. Highly sensitive methods were developed to address this question. Metabolism of NPYR by S9 fractions (containing microsomes) from rat esophagus and non-glandular stomach has been reported (36). However, it was not determined whether enzymes in the rat esophagus were involved in NPYR metabolism. Little is known about the kinetics of NPIP and NPYR
-hydroxylation. To our knowledge, there are only two reports in the literature concerning the kinetics of NPYR
-hydroxylation by liver microsomes; KM values were 14.1 and 16.5 mM (27,30). Further, the apparent kinetic parameters of NPIP
-hydroxylation catalyzed by liver microsomes have not been reported.
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Materials and methods |
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Chemicals
[3,4-3H]NPIP (10 Ci/mmol), [3,4-3H]NPYR (2 Ci/mmol), and [5-3H]NNN (27 Ci/mmol) were obtained from Moravek Biochemicals (Brea, CA). Tritium-labeled NPIP and NPYR were purified by normal-phase high performance liquid chromatography (HPLC) using system 1. Using reverse-phase HPLC with radioflow detection of purified NPIP (system 4) and NPYR (system 2), the radiochemical purity of both nitrosamines was 99.8% with no radioactivity co-eluting with 2-OH-THP and 2-OH-THF, respectively. [5-3H]NNN was used without further purification. The radiochemical purity of [5-3H]NNN was 99.7% by reverse-phase HPLC with radioflow detection using system 6. The chemical purity of the radiolabeled substrates was >99%. Unlabeled NPIP and NPYR (99% purity) were purchased from Sigma Chemical Co. (St Louis, MO) and were used without further purification. 5-Hydroxypentanal (2,4-dinitrophenyl)hydrazone, 4-hydroxypentanal(2,4-dinitrophenyl)hydrazone, NNN, 4-hydroxy-1-(3-pyridyl)-1-butanone, and 2-hydroxy-5-(3-pyridyl)tetrahydrofuran (lactol) were prepared as described previously (3741). Pefabloc® SC was purchased from Roche Molecular Biochemicals (Indianapolis, IN). Monoflo 5TM scintillation fluid was obtained from National Diagnostics (Atlanta, GA). Isobutane and methane (research grade 99.9%) were purchased from Matheson (Irving, TX) and Air Products (Allentown, PA), respectively. All other chemicals were purchased from Sigma Chemical Co., Aldrich Chemical Co. (Milwaukee, WI), or Mallinckrodt Chemical Co. (Paris, KY).
Apparatus
HPLC analyses were carried out on a Waters Associates (Waters Division, Millipore, Milford, MA) system equipped with an in-line Gilson Model 116 UV detector (Gilson Medical Electronics, Middleton, WI) and a ß-Ram radioactivity HPLC flow-through detector (Model 2B, IN/US Systems, Tampa, FL). Gas chromatography-positive ion chemical ionization-mass spectrometry (GC-PICI-MS) was carried out with a HP5890 series II GC (Hewett-Packard) using a DB-17MS (50% dimethyl/50% diphenylpolysiloxane) capillary column (0.25 x 30 m, 0.15 mm film thickness; J & W Scientific, Folsom, CA) coupled to a Thermo Finnigan TSQ-7000 instrument (San Jose, CA). The emission current and voltage were 700 µA and 150 V, respectively. The source temperature was 150°C. Isobutane and methane were used as the reagent gases for mass analysis of NPIP and NPYR, respectively. 1H and 13C NMR data were obtained on a 300 MHz Varian spectrometer (San Jose, CA).
HPLC analysis
Tritium-labeled NPIP and NPYR were purified by a normal-phase system with a Waters Econosil C18 column (250 x 4.6 mm; Deerfield, IL). Compounds were eluted isocratically with methylene chloride. Impurities were then washed off the column with 95:5 methylene chloride/methanol (system 1). A reverse-phase system consisting of a Metasil© ODS column (250 x 4.6 mm, Metachem, Torrance, CA) was utilized for analysis of NNN, NPIP and NPYR metabolism and 2-OH-THP/2-OH-THF isolation. Analytes from NPYR metabolism catalyzed by esophageal and hepatic microsomes were eluted isocratically for 10 min with 1% methanol and 99% 20 mM sodium phosphate buffer, pH 6, followed by a linear gradient to 50% methanol over 30 min (system 2). NPYR metabolites from incubations with P450 2A3 were eluted isocratically for 10 min with 0% methanol and 100% 25 mM ammonium acetate buffer, pH 4.5, followed by a linear gradient to 20% methanol over 15 min. Analytes were then eluted isocratically with 20% methanol and 80% 25 mM ammonium acetate buffer, pH 4.5 (system 3). For analysis of NPIP metabolites, compounds were eluted isocratically for 20 min with 4% methanol and 96% 20 mM sodium phosphate buffer, pH 6, followed by a linear gradient to 50% methanol over 10 min (system 4). NPIP metabolites from incubations with P450 2A3 were eluted isocratically for 20 min with 1% methanol and 99% 25 mM ammonium acetate buffer, pH 4.5, followed by a linear gradient to 15% methanol over 15 min. Analytes were then eluted isocratically with 15% methanol and 85% ammonium acetate buffer, pH 4.5 (system 5). NNN and its metabolites were eluted isocratically for 5 min with 5% acetonitrile and 95% 25 mM ammonium acetate buffer, pH 4.5, followed by a linear gradient to 35% acetonitrile over 40 min (system 6). To confirm the identity of metabolites, 2,4-dinitrophenylhydrazone derivatives of 2-OH-THP and 2-OH-THF were analyzed using isocratic 30% acetonitrile and 70% water (system 7). For 2-OH-THP and 2-OH-THF recovery studies, metabolites were purified using isocratic 100% water. The column was pre-washed with 50% methanol and 50% water (system 8). The flow rate for all HPLC systems was 1 ml/min. Reference metabolites and internal standard were analyzed by UV absorbance at 254 nm. Radiolabeled metabolites of parent compounds were measured by radioflow detection (scintillation fluid-to-HPLC elutant ratio of 3).
Preparation of 2-OH-THP and 2-OH-THF
2-OH-THF was prepared as previously described (42). Briefly, 2,3-dihydrofuran (132 mmol) was reacted with 50 ml of 0.2 N HCl for 15 min at 4°C followed by 30 min at room temperature. The mixture was extracted with chloroform (6 x 50 ml). The organic extracts were washed with 5% NaHCO3, dried (MgSO4), and concentrated. The resulting oil was purified by distillation and flash chromatography (silica gel, 70230 mesh, 60 Å) using chloroform and methanol. The identity and purity of the product were confirmed by HPLC using system 2, 1H NMR, 13C NMR, and GC-PICI-MS. 1H NMR (CDCl3): 5.52 (m, 1H, OCH-OH), 4.04 (m, 1H, H
of OCH2), 3.85 (s, 1H, -OH), 3.82 (m, 1H, Hß of OCH2), 1.82.1 (m, 4H, CH2CH2). 13C NMR (CDCl3): 98.4 (C-2), 67.4 (C-4), 34.1, and 23.5. MS [m/z (relative intensity)]: 89([M+1]+, 98), 71 (100), 43 (8), 39 (7).
2-OH-THP was prepared in similar fashion using 3,4-2H-dihydropyran (109.6 mmol) as the starting material (43). The identity and purity of the product were confirmed by HPLC using system 4, 1H NMR, 13C NMR, and GC-PICI-MS. 1H NMR (CDCl3): 4.88 (m, 1H, OCH-OH), 4.00 (m, 1H, H of OCH2), 3.52 (m, 1H, Hß of OCH2), 3.34 (s, 1H, -OH), 1.82 (m, 2H, Hs on C-3), 1.50 (m, 4H, Hs on C-4 and C-5). 13C NMR (CDCl3):
94.6 (C-2), 64.0 (C-5), 32.0, 25.3, and 20.3. MS [m/z (relative intensity)]: 103 ([M+1]+, 3), 101 (10), 85 (100).
Isolation of esophageal microsomes
Esophageal microsomes were prepared by the method of Murphy and Spina with a few exceptions (16). Briefly, esophageal mucosa from male F344 rats (age 1517 weeks) were obtained from Harlan Bioproducts (Indianapolis, IN). Dry-ice frozen tissues (eight per preparation) were crushed in a liquid nitrogen-cooled Bessman tissue pulverizer (Fisher Scientific, Springfield, NJ). All subsequent steps in the preparation of microsomes were performed at 4°C. The tissues were homogenized in 50 mM Tris buffer, pH 7.4, containing 1 mM EDTA and 0.4 mM Pefabloc® SC using a glass Tenbroeck tissue homogenizer. The homogenate was centrifuged at 9000 g for 30 min. The supernatant was centrifuged at 105 000 g for 90 min. The microsomal pellet was resuspended in 50 mM Tris buffer, pH 7.4, containing 1 mM EDTA. The microsomes were assayed immediately for NNN -hydroxylation as described previously (16). The remaining microsomal suspension was stored at 80°C.
The assay for NNN -hydroxylation was carried out as follows: Tritium-labeled NNN (0.1 µM; 0.54 µCi), 10 mM MgCl2, an NADPH-generating system (0.4 mM NADP+, 10 mM glucose-6-phosphate, 0.4 units/ml glucose-6-phosphate dehydrogenase), microsomes (5070 µg microsomal protein) in 100 mM sodium phosphate, pH 7.4, were incubated for 30 min at 37°C. The total volume was 0.2 ml. Glucose-6-phosphate dehydrogenase was purchased from Sigma (Type XII from Torula yeast, St Louis, MO). The reaction was terminated with 20 µl of 0.3 N Ba(OH)2 and 20 µl of 0.3 N ZnSO4. 4-Hydroxy-1-(3-pyridyl)-1-butanone metabolite standard was added prior to centrifugation of the mixture at 8000 g for 10 min. The supernatant was analyzed by HPLC with in-line ultraviolet absorbance and radioflow detection (HPLC-UV-radioflow) using system 6. Microsomal preparations that
-hydroxylated 0.5% or more of NNN under the above conditions were considered active.
Isolation of hepatic microsomes
Liver tissues were excised from male F344 rats purchased from Charles River Laboratories (Wilmington, MA). The rats were housed in the specific pathogen-free animal quarters of the University of Minnesota Cancer Center, fed a standard diet, and given water ad libitum. The tissues were frozen in liquid nitrogen and stored until the time of preparation. Three separate preparations of microsomes were isolated as described previously using one liver per preparation (26); preparations were designated as RLM101, RLM102, and RLM103. Microsomes were suspended in 10 mM Tris buffer, pH 7.4, containing 1 mM EDTA and 20% glycerol and stored at 80°C. Protein concentrations were determined using the Microprotein 610-A kit (Sigma, St Louis, MO) and a UV-Vis spectrophotometer (Beckman DU-7400, Fullerton, CA).
In vitro metabolism by esophageal microsomes
Metabolic studies were carried out using tritium-labeled NPIP or NPYR. Substrate solutions were prepared using the following general procedure as described previously for N-nitrosomethyl-n-pentylamine (17). Tritium-labeled NPIP or NPYR in methylene chloride and water were added to a 4 ml glass vial. Transfer of tritium-labeled nitrosamine to the aqueous layer was achieved by application of a stream of nitrogen via a Pasteur pipette to the surface of the mixture until 10 min after the methylene chloride layer disappeared.
Based on the requisite amount of microsomes needed for each metabolic study, four to eight preparations (eight esophagi per preparation) of esophageal microsomes were thawed and pooled prior to use. Each pool of microsomes was used for only one metabolic study. In all, five pools were prepared. The pools were designated as REM A, REM B, REM C, REM D, and REM E. Protein concentrations of each pool were determined as described above for liver microsomes.
NPIP and NPYR metabolism, catalyzed by esophageal microsomes at varying substrate concentrations, was carried out as follows: [3,4-3H]NPIP or [3,4-3H]NPYR (1.54 µCi) and the corresponding unlabeled nitrosamine (in amounts to attain final substrate concentrations of 4, 7, 10, 20, 40, 70, and 100 µM), 10 mM MgCl2, an NADPH-generating system (0.4 mM NADP+, 10 mM glucose-6-phosphate, 0.4 units/ml glucose-6-phosphate dehydrogenase), and microsomes (95 µg microsomal protein; REM B) in 100 mM sodium phosphate, pH 7.4, were incubated from 15 to 30 min at 37°C. The reaction was initiated by addition of microsomes. The specific activities of [3,4-3H]NPIP and [3,4-3H]NPYR varied from 0.0110 and 0.012 Ci/mmol, respectively. The total volume was 200 µl. The reaction was terminated with 20 µl of 0.3 N Ba(OH)2 and 20 µl of 0.3 N ZnSO4. For studies with NPIP, 2-OH-THP (100 µg) metabolite standard and lactol (3 µg, internal standard) were added prior to centrifugation of the mixture at 8000 g for 10 min. The supernatant was analyzed by HPLC-UV-radioflow using system 4. For NPYR experiments, the procedure was repeated with the following exceptions: 2-OH-THF standard (10 mg) was added instead of 2-OH-THP and system 2 was used for HPLC analysis.
NPIP and NPYR metabolism was studied at varying microsomal protein concentrations. Samples were prepared as described above with the following modifications: incubations contained 15, 30, 60, 90, or 120 µg microsomal protein (REM A), 1.6 µCi of the appropriate radiolabeled substrate, and the corresponding amount of unlabeled nitrosamine to attain the final substrate concentration of 4 µM NPIP or NPYR. Saturation kinetics study of NPIP -hydroxylation was conducted as described above using final substrate concentrations of 0.5, 1, 2, 4, 7, 10, 20, 30, 40, 55, 70, 100, 150, 250, 500, 1000, and 2000 µM and microsomes from REM C. The specific activities of [3,4-3H]NPIP varied from 0.0110 and 0.012 Ci/mmol, respectively.
The following control studies in esophageal microsomes were performed. To determine if a component in the aqueous [3,4-3H]NPYR solution inhibited microsomal metabolic activity, the rates of NPIP -hydroxylation in the presence or absence of NPYR were compared. [3,4-3H]NPIP (0.42 Ci/mmol) with or without [3,4-3H]NPYR (2 Ci/mmol) was incubated with microsomes (REM D) and co-factors at 37°C for 15 min and then analyzed as above. To determine whether metabolites reacted with esophageal microsomal protein during the incubation, the rates of 2-OH-THP and 2-OH-THF metabolism were determined. [3H]2-OH-THP (10 Ci/mmol) or [3H]2-OH-THF (2 Ci/mmol) were isolated from incubations of rat liver microsomes and tritium-labeled NPIP or NPYR, respectively, by reverse-phase HPLC using system 8, incubated with esophageal microsomes (REM E) and co-factors at 37°C for 30 min, and then analyzed as described above for esophageal microsomal metabolism. All aforementioned incubations with NPIP and NPYR were performed in triplicate (n = 3), i.e. three individual samples incubated under the same reaction conditions and each analyzed once.
The identification of 2-OH-THP as a metabolite of NPIP with esophageal microsomes and co-factors was confirmed by co-elution with a standard using system 4. For further confirmation, tritium-labeled co-elutant corresponding to the 2-OH-THP standard (system 4) was collected and derivatized with an acidic solution of 2,4-dinitrophenylhydrazine. A radiolabeled peak co-eluting with the 5-hydroxypentanal(2,4-dinitrophenyl)hydrazone standard was detected by reverse-phase HPLC-UV-radioflow using system 7.
In vitro metabolism by liver microsomes
NPIP and NPYR substrate solutions were prepared as described above for metabolic studies with esophageal microsomes. To determine the KM and Vmax of NPIP -hydroxylation for three different preparations of liver microsomes, NPIP was incubated with RLM101, RLM102, and RLM103. Kinetic parameters for NPYR
-hydroxylation were determined using one microsomal preparation, RLM102. The final substrate concentrations were 1, 5, 10, 20, 50, 100, 200, 350, 500, and 750 µM. The specific activities of [3,4-3H]NPIP and [3,4-3H]NPYR ranged from 0.01310 and 0.0132 Ci/mmol, respectively. NPIP and NPYR incubations contained 2570 µg and 25120 µg microsomal protein, respectively. The work-up procedure and HPLC analysis were repeated as described above for esophageal microsomes. All incubations were conducted in triplicate (n = 3).
The identification of 2-OH-THP as a metabolite of NPIP with liver microsomes and co-factors was confirmed as described above following metabolism by esophageal microsomes. The identity of 2-OH-THF from liver microsomal incubations with NPYR was confirmed by similar methods. System 2 was used to isolate the metabolite and analyze with 2-OH-THF standard.
In vitro metabolism by P450 2A3
Microsomes from Spodoptera frugiperda (Sf9) cells transfected with cDNA for P450 2A3 (2.4 pmol/µl in 50 mM Tris buffer, pH 7.4 containing 20% glycerol) were a generous gift from Dr Xinxin Ding (Wadsworth Center, Albany, NY). Microsomes from Sf9 cells transfected with cDNA for NADPH-P450 oxidoreductase were purchased from PanVera (Madison, WI). Incubations were performed essentially as described previously (18) with some exceptions. Briefly, [3,4-3H]NPIP or [3,4-3H]NPYR (0.44 µCi) and the corresponding unlabeled nitrosamine (in amounts to attain final substrate concentrations of 1, 3, 10, 20, 50, 100, 250, 500, 1000, and 1500 µM), MgCl2 (2 µmol), P450 2A3 (24 pmol), NADPH-P450 oxidoreductase (25-molar excess relative to P450 2A3), and Tris buffer, pH 7.4 (10 µmol), in a volume of 180 µl were incubated for 5 min at 37°C and then placed on ice. This first incubation served to facilitate the association of the proteins prior to initiation of reaction. The specific activities of [3,4-3H]NPIP and [3,4-3H]NPYR ranged from 0.01310 and 0.0132 Ci/mmol, respectively. Reactions were initiated by the addition of an NADPH-generating system (0.08 µmol of NADP+, 2 µmol of glucose-6-phosphate, 0.08 units of glucose-6-phosphate dehydrogenase in a volume of 20 µl) and were carried out for 30 min at 37°C. The total reaction volume was 200 µl. Reactions were terminated with 20 µl of 0.3 N Ba(OH)2 and 20 µl of 0.3 N ZnSO4. The samples were prepared and analyzed as described above for in vitro metabolism by esophageal microsomes except that systems 3 and 5 were used for analysis of NPYR and NPIP metabolism, respectively.
Data analysis
Velocities were quantitated by accounting for the percent recovery of lactol (internal standard) for substrate concentrations below 500 µM. For concentrations at or above 500 µM, velocities were calculated using the percent product formed relative to the total radioactivity. For saturation kinetics studies, the mean and standard deviation of product velocities from incubations conducted in triplicate for each substrate concentration were calculated and represented graphically. Except for the saturation kinetics study of NPIP metabolism to 2-OH-THP catalyzed by esophageal microsomes, KM, Vmax, and their standard deviations of goodness of fit were calculated by SigmaPlot® 2000 (SPSS, Chicago, IL) computer application program. The software program employs nonlinear least-squares regression analysis to fit a MichaelisMenten curve to the data plotted on a velocity (v) versus substrate concentration ([S]) graph. For the saturation kinetics study of esophageal microsomal metabolism of NPIP, kinetic parameters and their standard deviations were calculated by linear least-squares regression analysis (SigmaPlot® 2000) of the data plotted on a HanesWoolf plot ([S]/v versus [S]). The limits of detection for NPIP and NPYR metabolites were 0.02 and 0.1 pmol, respectively (signal-to-noise ratio after background subtraction = 3; sp. act. = 10 Ci/mmol for NPIP and 2 Ci/mmol for NPYR).
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Results |
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Metabolism of NPIP and NPYR by rat liver microsomes
The formation of 2-OH-THP and 2-OH-THF increased linearly over a 60 min period and over a protein concentration up to 0.5 mg/ml in incubations with tritium-labeled NPIP and NPYR, respectively. These metabolites were not present in control incubations, carried out with heat-treated microsomes or without co-factors. Concentration-dependent formation of 2-OH-THP from NPIP was determined following incubation with three different preparations of hepatic microsomes (RLM101, RLM102, and RLM103; microsomes isolated from one liver per preparation). A representative velocity versus substrate concentration plot is shown in Figure 4A for substrate concentrations between 1 and 750 µM with RLM102 microsomes. The KM and Vmax values for 2-OH-THP formation from NPIP metabolism by RLM101, RLM102, and RLM103 are summarized in Table I
. The KM values among the three microsomal preparations were comparable (Table I
). Saturation kinetics of 2-OH-THF formation from tritium-labeled NPYR were carried out with one preparation of hepatic microsomes (RLM102). A velocity versus substrate concentration plot of 2-OH-THF is illustrated in Figure 4B
for the same concentration range with RLM102 microsomes. The KM and Vmax values for 2-OH-THF formation from NPYR are presented in Table I
. Catalytic efficiencies (V/K) were similar for the two nitrosamines (Table I
).
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Depending on the amount of microsomes required for each metabolic study, active preparations (between 48) were pooled and immediately used. Each pool of microsomes was used for only one study. To ensure an accurate comparison of NPIP versus NPYR metabolism, tritium-labeled NPIP and NPYR were incubated with the same pool of microsomes. In all, five pools were prepared for studies with esophageal microsomes.
The formation of 2-OH-THP and 2-OH-THF from NPIP and NPYR, respectively, were determined over a protein concentration range of 00.6 mg/ml as shown in Figure 5. The initial concentrations of NPIP and NPYR were 4 µM. With increasing microsomal protein concentration, 2-OH-THP formed linearly whereas little 2-OH-THF was produced. The amounts of 2-OH-THF detected for all samples over this microsomal protein concentration range were no more than twice the limit of detection (0.1 pmol). The slope of 2-OH-THP formation from NPIP versus protein concentration was 40-fold higher than that of 2-OH-THF production from NPYR. Neither metabolite was present in control incubations, carried out with heat-treated microsomes or without co-factors. Also, 2-OH-THP formation increased over a 30 min period (data not shown).
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The kinetics of 2-OH-THP formation over the concentration range of 0.5 to 2000 µM were determined as shown in Figure 7. Standard MichaelisMenten kinetics predicts a linear relationship of the data in the HanesWoolf plot; however the data fit along two different slopes with an inflection point at 180 µM. The KM and Vmax values calculated from the fitted line for the data at higher substrate concentrations (1502000 µM) were 1600 ± 312 µM and 824 ± 119 pmol/min/mg, respectively. The KM and Vmax values calculated from the fitted line for the data for concentrations from 0.5 to 150 µM were 312 ± 50 µM and 226 ± 35 pmol/min/mg, respectively. HanesWoolf analysis suggests the presence of biphasic kinetics. Further, analysis using an EadieHofstee (v vs. v/[S]) or velocity versus substrate plot gave similar values for the two sets of kinetic parameters.
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Discussion |
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Rat esophageal microsomes -hydroxylate NPIP at a considerably higher rate than NPYR. Microsomes converted NPIP (4 µM) to 2-OH-THP at a 40-fold higher rate than NPYR (4 µM) to 2-OH-THF at five different microsomal concentrations (Figure 5
). Further, microsomes activated NPIP with at least a seven-fold higher rate compared with NPYR over the substrate concentration range of 4100 µM (Figure 6
). In contrast, the
-hydroxylation rates of these nitrosamines by rat liver microsomes were similar (Figure 4
, Table I
).
We developed an HPLC-radioflow method to directly quantitate 2-OH-THP or 2-OH-THF following reaction of tritium-labeled NPIP or NPYR, respectively, with microsomes or P450 2A3. Aldehydic products of NPIP and NPYR -hydroxylation previously have been characterized and quantitated via their 2,4-dinitrophenylhydrazone derivatives using HPLC with UV and radioflow detection (20,27,28,36,4448). Using nonlabeled NPYR, Chen et al. achieved a limit of detection of 0.2 nmol for 2-OH-THF, as measured by its derivative, for studies with rat liver microsomes in vitro (21). Several in vivo and in vitro studies employed 14C-labeled NPYR and NPIP to increase assay sensitivity (20,28,36,4448). The highest specific activity used in these studies for either NPIP and NPYR was 0.0188 Ci/mmol (44). In the present study, the specific activities of tritium-labeled NPIP and NPYR were 10 and 2 Ci/mmol, respectively, markedly higher than in previous studies with the 14C-labeled compounds. Because of the high specific activities, we were able to examine the metabolism of these nitrosamines at concentrations as low as 0.5 µM. The limits of detection for NPIP and NPYR metabolites were 0.02 and 0.1 pmol, respectively, substantially lower than in previous studies. Further, the recoveries were excellent: >90%.
From previous studies, the KM values of liver microsomal metabolism of NPYR were 14.1 and 16.5 mM for substrate concentrations between 1 and 90 mM (27,30). We report a KM for NPYR -hydroxylation in rat liver microsomes that is >100-fold lower over a substrate concentration range from 1 to 750 µM. These results suggest that there are at least two enzymes involved in NPYR
-hydroxylation in rat liver. P450 2E1, a human and rat liver microsomal enzyme, and P450 2A6, a human ortholog to P450 2A1/2, exhibited high NPYR
-hydroxylase activity (49). KM values of cDNA-expressed P450 2E1 and 2A6 were 34 µM and 1.26 mM (49). P450 2E1 and rat P450 2A1/2 may be involved in
-hydroxylation of NPYR by rat liver microsomes. The presence of more than one key liver NPYR
-hydroxylase would explain why the velocities of 2-OH-THF formation are higher than the calculated MichaelisMenten curve at the higher substrate concentrations (Figure 4
).
In contrast to NPYR, NPIP -hydroxylation kinetics by rat liver microsomes and individual P450s have not been studied. Our findings represent the first report on the apparent kinetic parameters of NPIP
-hydroxylation by rat liver microsomes (Table I
). It seems likely that in rat liver, the same P450 enzymes mediate NPIP and NPYR
-hydroxylation. Fujita and co-workers reported significant mutagenicity of NPIP in S.typhimurium strains expressing P450 2A6 and 2E1 (50). If P450 2E1 and a rat enzyme similar to P450 2A6 are primarily responsible for NPIP and NPYR activation in the liver, then the similar KM values observed for the microsomal metabolism of these cyclic nitrosamines would suggest that these enzymes do not preferentially activate either NPIP or NPYR.
In this study, there was a clear difference in NPIP and NPYR -hydroxylation catalyzed by rat esophageal microsomes (Figures 5 and 6
). Several lines of evidence suggest that P450 2A3 is an enzyme that may contribute to the preferential activation of NPIP in the esophagus. P450 2A3 has been detected in the esophagus, nasal mucosa, and lung, but not in the liver (5154). The KM values for NPIP and NPYR
-hydroxylation catalyzed by Sf9-expressed P450 2A3 were 61.6 ± 20.5 and 1198 ± 308 µM, respectively (19.4-fold difference; Table I
). P450 2A3 is an efficient
-hydroxylase of other esophageal carcinogens such as NNN, NBzMA, and N-nitrosomethyl-n-pentylamine (1618,55). Also, elevated levels of P450 2A3 mRNA corresponded with increased NBzMA activation in cultured esophagus from animals treated with N-(4-hydroxyphenyl)retinamide, an esophageal tumor enhancer (56).
However, levels of P450 2A3 mRNA in rat esophagus are minute compared with nasal mucosa (1600-fold less) (53). Rat esophageal microsomes do not catalyze coumarin 7-hydroxylation, an efficient P450 2A3-mediated pathway (18). Further, previous studies with NBzMA and NNN provided strong evidence that the esophageal enzyme responsible for their activation is not P450 2A3, but an enzyme not yet isolated (18,54). This yet-to-be-isolated enzyme may be the one responsible for the differential activation of NPIP and NPYR in the esophagus.
Recently, Swann and co-workers isolated P450 2B21 cDNA from the rat esophagus (57). P450 2B21 may be involved in nitrosamine activation, although preliminary data indicate that it does not catalyze the metabolism of NPIP. Also, P450 1A1 was detected in the rat esophagus (19,53). Purified rabbit P450 1A1 catalyzed NPYR -hydroxylation (58), but P450 1A1-mediated NPIP
-hydroxylation has not been studied. Therefore, it is unclear what role P450 1A1 plays in the esophageal metabolism of NPIP and NPYR. Further work is necessary to identify the rat esophageal enzyme(s) responsible for activation of NPIP.
Kinetic data for NPIP -hydroxylation, catalyzed by esophageal microsomes, adhered to two distinct lines in the HanesWoolf plot with an inflection point at 180 µM, which suggests biphasic kinetics (Figure 7
). We report KM values of 312 and 1600 µM for 2-OH-THP formation from NPIP. One explanation for the observed kinetics is the presence of more than one
-hydroxylase involved in NPIP metabolism. von Weymarn and co-workers observed the presence of at least two P450 enzymes involved in the methylene hydroxylation of NBzMA by rat esophageal microsomes (18). The KM and Vmax values for the high and low-affinity enzymes were 5.1 µM and 54 pmol/min/mg and 446 µM and 500 pmol/min/mg, respectively. Another possibility is the presence of two substrate-binding sites of one esophageal microsomal enzyme involved in NPIP metabolism. Biphasic kinetics involving two substrate-binding sites have been reported for P450 1A2, 2B6, 2C8, 2C9, 3A4, and 3A5 (5961). However, a fit of our data to the Hill equation (n < 1) indicated the presence of one binding site.
Rat esophageal microsomes efficiently catalyze the activation of several esophageal carcinogens. The KM values for the metabolic activation of NNN, N-nitrosodiethylamine, and N-nitroso-n-pentylamine were 49, 13, and 64 µM, respectively (16,17,19). The KM values of NBzMA and NPIP metabolic activation are given above. Taken together, rat esophageal microsomes do not activate NPIP as well as other esophageal carcinogens. Further work is necessary to understand these differences in activation. The observed low KM for NPIP was higher than expected, which suggests that there are contributing factors other than metabolic activation involved in its organospecificity. Also, we cannot exclude the possibility of a high-affinity NPIP -hydroxylase in the esophagus. A more sensitive quantitation method measuring
-hydroxylation rates for substrate concentrations <0.5 µM may reveal an enzyme with a KM value lower than reported in this study. However, it is clear that esophageal microsomes activated NPIP, but not NPYR. This substantial difference in activation is remarkable considering that they differ by only one carbon.
The results of the present study are consistent with those of previous investigations comparing metabolism of structurally related nitrosamines in the rat esophagus. In rats, methylalkylnitrosamines with alkyl chains of 36 carbons are potent esophageal carcinogens (95100% tumor incidence) whereas those nitrosamines having alkyl chains of 1, 2, 7, 8, 9, or 10 carbons are weakly carcinogenic or non-carcinogenic to the esophagus (62). Methylalkylnitrosamines are believed to elicit their carcinogenic potential via metabolic formation of a DNA methylating species (63,64). As discussed before, Kleihues and co-workers clearly demonstrated that esophageal carcinogenicity of these nitrosamines correlated with high levels of promutagenic methylated DNA adducts in target tissues (1315). Differences in tissue-specific metabolism leading to differences in tissue-specific carcinogenicities were also observed with NBzMA and N-nitrosodimethylamine (NDMA). In rats, NBzMA induces esophageal tumors while NDMA causes mainly liver tumors, but no esophageal tumors (62). Rat esophageal microsomes catalyzed NBzMA methylene hydroxylation to benzaldehyde and a methylating agent (64). In contrast, NDMA demethylation, which generates formaldehyde and a methylating agent, was not detected (64).
Metabolic activation of structurally homologous NNN and N'-nitrosoanabasine (NAB) was also compared. NNN is a potent rat esophageal carcinogen whereas NAB is weakly carcinogenic in the esophagus (65). Hydroxylation at the 2' carbon leading to the generation of a pyridyloxobutylating agent is considered to be the key metabolic activation pathway of NNN (66). Cultured rat esophagus primarily metabolized NNN via 2'-hydroxylation and NAB via 6'-hydroxylation (67). Significantly lower levels of 2'-hydroxylation of NAB were detected compared with that of NNN (67). These findings and those of the present study strongly support the hypothesis that local metabolic activation in the esophagus leads to tissue-specific tumor induction.
In summary, this study clearly shows that rat esophageal microsomes activate NPIP, but not NPYR, while rat liver microsomes activate both NPIP and NPYR. This is likely a contributing factor to the organospecific carcinogenicities of NPIP and NPYR in rats. We expect that these differences in esophageal metabolism will translate to differences in esophageal DNA adduct levels for rats treated with NPIP or NPYR; these studies are ongoing in our laboratory.
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