Quantification of Urinary Aflatoxin B1 Dialdehyde Metabolites Formed

752

Chem. Res. Toxicol. 2008, 21, 752–760

Quantification of Urinary Aflatoxin B1 Dialdehyde Metabolites Formed by Aflatoxin Aldehyde Reductase Using Isotope Dilution Tandem Mass Spectrometry Denise N. Johnson,† Patricia A. Egner,† Greg OBrian,‡ Norman Glassbrook,§ Bill D. Roebuck,| Thomas R. Sutter,⊥ Gary A. Payne,‡ Thomas W. Kensler,† and John D. Groopman*,† Department of EnVironmental Health Sciences, Bloomberg School of Public Health, Johns Hopkins UniVersity, Baltimore, Maryland 21205, Departments of Plant Pathology and EnVironmental and Molecular Toxicology, North Carolina State UniVersity, Raleigh, North Carolina 27695, Department of Pharmacology and Toxicology, Dartmouth Medical College, HanoVer, New Hampshire 03755, and W. Harry Feinstone Center for Genomic Research, UniVersity of Memphis, Memphis, Tennessee 38152 ReceiVed NoVember 7, 2007

The aflatoxin B1 aldehyde reductases (AFARs), inducible members of the aldo-keto reductase superfamily, convert aflatoxin B1 dialdehyde derived from the exo- and endo-8,9-epoxides into a number of reduced alcohol products that might be less capable of forming covalent adducts with proteins. An isotope dilution tandem mass spectrometry method for quantification of the metabolites, C-8 monoalcohol, dialcohol, and C-6a monoalcohol, was developed to ascertain their possible role as urinary biomarkers for application to chemoprevention investigations. This method uses a novel 13C17-aflatoxin B1 dialcohol internal standard, synthesized from 13C17-aflatoxin B1 biologically produced by Aspergillus flaVus. Chromatographic standards of the alcohols were generated through sodium borohydride reduction of the aflatoxin B1 dialdehyde. This method was then explored for sensitivity and specificity in urine samples of aflatoxin B1-dosed rats that were pretreated with 3H-1,2-dithiole-3-thione to induce the expression of AKR7A1, a rat isoform of AFAR. One of the two known monoalcohols and the dialcohol metabolite were detected in all urine samples. The concentrations were 203.5 ( 39.0 ng of monoalcohol C-6a/mg of urinary creatinine and 10.0 ( 1.0 ng of dialcohol/mg of creatinine (mean ( standard error). These levels represented about 8.0 and 0.4% of the administered aflatoxin B1 dose that was found in the urine at 24 h, respectively. Thus, this highly sensitive and specific isotope dilution method is applicable to in ViVo quantification of urinary alcohol products produced by AFAR. Heretofore, the metabolic fate of the 8,9-epoxides that are critical for aflatoxin toxicities has been measured by biomarkers of lysine-albumin adducts, hepatic and urinary DNA adducts, and urinary mercapturic acids. This urinary detection of the alcohol products directly contributes to the goal of mass balancing the fate of the bioreactive 8,9-epoxides of AFB1 in ViVo. Introduction 1

Aflatoxin B1 (AFB1), a secondary fungal product of the mold Aspergillus flaVus, is a potent experimental and human liver carcinogen (1–3). AFB1 requires metabolic activation to exert its toxic and carcinogenic effects, and the cytochrome P450 1A2 and 3A4 pathways forming the AFB1 8,9-exo- (e.g., 1A2 and 3A4) and 8,9-endo-epoxides (e.g., 1A2) are critical to this process (4). Only the exo isomer reacts with DNA, forming AFB1 N7-guanine and it derivative AFB1 formamidopyrimidine (FAPY) adducts (5). If not repaired, these DNA adducts or the * To whom correspondence should be addressed. E-mail: jgroopma@ jhsph.edu. † Johns Hopkins University. ‡ Department of Plant Pathology, North Carolina State University. § Department of Environmental and Molecular Toxicology, North Carolina State University. | Dartmouth Medical College. ⊥ University of Memphis. 1 Abbreviations: AFB1, aflatoxin B1; AFB1-N7-Gua, aflatoxin B1-N7guanine; AFM1, aflatoxin M1; AFP1, aflatoxin P1; HPLC, high-performance liquid chromatography; LC-MS/MS, liquid chromatography-tandem mass spectrometry; ESI, electrospray ionization; SPE, solid-phase extraction; D3T, 3H-1,2-dithiole-3-thione.

abasic sites that remain from the spontaneous depurination of the N7-guanine adduct can ultimately lead to mutational events. The endo-8,9-epoxide does not form DNA adducts, but it is capable of forming other macromolecular adducts with proteins (6). Both AFB1 epoxides are substrates for glutathione-Stransferases (GST) and are further metabolized to urinary mercapturic acids (7). A potentially deleterious fate of the exoand endo-epoxides are the formation of AFB1 protein adducts. The epoxides can nonenzymatically convert into a proteinbinding species, AFB1 dialdehyde (Figure 1). This metabolite can either react with lysine residues to form protein adducts, or AFB1 dialdehyde can be reduced into two partially reduced monoalcohols and one fully reduced dialcohol product by aflatoxin aldehyde reductase (AFAR) (8). While there are analytical tools currently available to determine the fate in ViVo of the AFB1 epoxides by measuring DNA adducts, serumalbumin-derived AFB1-lysine protein adducts, and the urinary mercapturic acids of AFB1 by mass spectrometry (9–12), there is no comparable mass spectrometry methodology for the AFAR-derived alcohols of AFB1. AFAR is a NAD(P)H-dependent enzyme that is a member of the aldo-keto reductase (AKR) superfamily. Members of the

10.1021/tx700397n CCC: $40.75  2008 American Chemical Society Published on Web 02/12/2008

Urinary AFB1 Dialdehyde Metabolites

Chem. Res. Toxicol., Vol. 21, No. 3, 2008 753

Figure 1. AFB1 metabolic scheme showing the formation of the AFAR substrate, AFB1 dialdehyde. AFAR reduces the dialdehyde into three alcohol products, the C-8 monoalcohol, C-6a monoalcohol, and the fully reduced dialcohol.

AKR superfamily are present in species ranging from bacteria and yeast to plants and mammals, highlighting the biological importance of detoxification of aldehydes and ketones in ViVo (13). Multiple rat and human isoforms of AFAR exist, with AKR7A1 being the first characterized rat isoform of AFAR (14). The expression of human AKR7A-related RNA has been detected in liver, kidney, stomach, pancreas, and other organ tissues (15). The rat AFAR protein, AKR7A1, is cytosolic and exhibits high expression in rat liver tissue (14). Hepatic AKR7A1 protein levels have been shown to be induced in ViVo by a number of agents including ethoxyquin, β-napthoflavone, and 3H-1,2-dithiole-3-thione (D3T) in rats (16–18). AKR7A1 has high catalytic efficiency toward the substrates 4-nitrobenzaldehyde and 9,10-phenanthrenquinone but was characterized and named for its activity toward the AFB1 dialdehyde (19). To date, endogenous substrates of AKR7A1 have yet to be elucidated. Previous studies examining AFAR activity toward AFB1 dialdehyde have monitored the resulting alcohol formation in in Vitro incubations with recombinant AFAR or in ex ViVo incubations with human liver cytosols (8). Limited studies, however, have been conducted to measure the in ViVo generation of AFAR alcohol products. With the goal of measuring AFAR alcohol products in urine of animals and humans, we have developed a quantitative highpressure liquid chromatography (HPLC), stable isotope dilution mass spectrometry method for measuring the in ViVo generation of AFB1-derived AFAR alcohols. This method is based on a novel, stable-isotope-labeled, 13C17AFB1-dialcohol internal standard derived from 13C17-AFB1 biosynthetically produced in an Aspergillus flaVus strain and uses solid-phase extraction (SPE)aflatoxin-specific immunoaffinity technology together with liquid chromatography-electrospray ionization tandem mass spec-

trometry (LC-ESI/MS/MS) analysis. This work provides an analytical tool needed to explore the biological significance of the AFAR pathway in aflatoxin toxicology.

Experimental Procedures Caution: Solid aflatoxins are hazardous, and some haVe been demonstrated to be human carcinogens. Extreme care needs to be exercised when handling aflatoxins including gloVes, respiratory masks, and well-Ventilated fume hoods. Aflatoxin residues can be destroyed using 3% bleach. Chemicals. AFB1, diazald, oxone, sodium borohydride, 2-(Ncyclohexylamino)ethanesulfonate (CHES), and N-(2-hydroxyethyl)piperazine-N-(2-ethanesulfonate) (HEPES) used in the preparation of standards or in kinetic incubations were purchased from Sigma Chemical Co. (St. Louis, MO). Dimethyldioxirane was prepared as previously described (20). 3H-1,2-Dithiole-3thione used in animal studies was obtained from DCPC Repository (Rockville, MD). All other chemicals and solvents were of analytical/reagent grade or higher. Urinary creatinine levels were determined by Eagle Diagnostics Direct Creatinine Reagent Set (Eagle Diagnostics, De Soto, TX). Recombinant Aflatoxin Aldehyde Reductase, AKR7A1. Escherichia coli recombinant oligo-His-tagged AKR7A1 clones were a generous gift of Dr. John D. Hayes (University of Dundee, Dundee, Scotland). AKR7A1 clones were prepared from BL21pLysS expression strain as previously described (16). The oligo-His tag was cleaved with a Biotinylated Thrombin Kit (Novagen, Madison, WI) according to directions of the manufacturers. Catalytic activity of cleaved AKR7A1 was confirmed monitoring NADPH oxidation at 340 nm with 4-nitrobenzaldehyde, 9,10-phenanthrenequinone, and AFB1 di-

754

Chem. Res. Toxicol., Vol. 21, No. 3, 2008

aldehyde substrates on a Beckman Coulter DU 800 spectrophotometer (Beckman Coulter, Somerset, NJ). Preparation of 13C17-AFB1. A total of 1 × 106 conidia/mL A. flaVus strain NRRL 3357 (ATCC 200026; SRRC 167), a wild-type A. flaVus, was inoculated into 100 mL of modified Adye and Mateles (A&M) medium (21). Modified A&M medium per 100 mL contained 3 g of 13C-glucose, 0.3 g of ammonium sulfate, 1 g of potassium monophosphate, 0.2 g of magnesium sulfate, and 100 µL of metal mix (22). Cultures were grown at 28 °C with shaking at 200 rpm for 3 days and vacuum-filtered through miracloth before the medium was extracted to isolate aflatoxin. Batches of the fungal medium were extracted with chloroform, concentrated by rotary evaporation, and purified by reverse-phase HPLC. Chromatographic separation was performed on a Waters 515 HPLC system coupled to a Waters 996 photodiode diode array (PDA) detector (Waters Corp., Milford, MA). Separations were conducted at ambient temperature using a flow rate of 1 mL/min on a 4.6 mm × 250 mm × 5 µm Luna C18 HPLC column (Phenomenex, Torrance, CA). Mobile-phase compositions were as follows: (A) 0.1% formic acid in water and (B) 0.1% formic acid in acetonitrile. Under isocratic conditions of 70.0% A and 30.0% B, 13C17AFB1 eluted at a retention time of 26 min, the identical retention time of an authentic AFB1 standard. The chromatographic peak corresponding to authentic AFB1 standard was analyzed by mass spectrometry to confirm the production of 13C17-AFB1 and isotopic incorporation into the aflatoxin ring structure. Preparation of Monoalcohols, Dialcohol, and 13C17-AFB1 Dialcohol Standards. Monoalcohol and dialcohol chromatographic standards were prepared using a modification of a previously reported method by Guengerich et al. (8). Alcohol standards were made by synthesizing the AFB1 8,9-epoxide through oxidation of AFB1 with dimethyldioxirane (20). AFB1 8,9-epoxide was hydrolyzed to the dihydrodiol with water prior to incubation in 20 mM sodium 2-(N-cyclohexylamino)ethanesulfonate (CHES, pH 10.0) to form AFB1 dialdehyde, the immediate precursor in alcohol synthesis. Alcohol standards were synthesized by reacting AFB1 dialdehyde (0.14 mL, 5.20 mM) with a solution of sodium borohydride in methanol (0.10 mL, 0.13 M) dried over 4 Å, 1.6 mm molecular sieve pellets under a constant nitrogen stream. After 30 min, the reaction was quenched with the addition of 1 M formic acid and the alcohol products were isolated by reverse-phase HPLC on a Thermo-Finnigan Surveyor pumping and PDA detector system (ThermoElectron Corporation, San Jose, CA) using a 250 mm × 4.6 mm × 5 µm Luna C18 HPLC column (Phenomonex, Torrance, CA). UV was monitored at 362 nm. Mobile-phase compositions for alcohol method 1 were as follows: (A) 0.01% glacial acetic acid in water and (B) 100.0% reagent alcohol. Under isocratic conditions of 90.0% A and 10.0% B, C-8 monoalcohol, dialcohol, and C-6a monoalcohol eluted at retention times of 22.0, 31.0, and 37.5 min, respectively. HPLC peaks were collected individually, concentrated under high-purity nitrogen, and characterized by mass spectrometry analysis. Structures of monoalcohol products were also analyzed by mass spectrometry following sodium periodate cleavage of monoalcohol-derived deuterated dialcohol as previously described (8). 13 C17-AFB1 dialcohol internal standard was prepared and purified as described above for dialcohol with the exception that 13C17-AFB1 was substituted to yield the 13C17-AFB1 dialdehyde precursor. AKR7A1-AFB1 Dialdehyde Incubations. Incubation mixtures consisted of 10 µg of recombinant AKR7A1 protein, 50 mM potassium N-(2-hydroxyethyl)piperazine-N-(2-ethane-

Johnson et al.

sulfonate) (HEPES) buffer at pH 7.4, 5 mM MgCl2, 0.2 mM NADPH, and AFB1-dialdehyde in a total volume of 100 µL. Reaction mixtures were incubated at 37 °C and terminated by the addition of 10 µL of glacial acetic acid. Kinetic constants were derived from a computer-generated best fit to the Michaelis– Menten equation of plots experimentally obtained from linear velocities versus the dialdehyde substrate concentration, using the Solver feature of Excel (Microsoft). Solver precision and tolerance requirements were set to 0.001 and 5.0%, respectively. Least sum of squared differences between experimental and calculated data was used to determine the closeness of calculated data to experimental data. Animals, Diets, and Treatments. Male Sprague–Dawley rats (75–100 g; Harlan, Indianapolis, IN) were housed under controlled conditions of temperature, humidity, and lighting. Food and water were available ad libitum, and a purified diet of the AIN-76A formulation (Teklad, Madison, WI) (without the recommended addition of 0.02% ethoxyquin) was used. Rats were acclimated to the AIN-76A diet for 1 week prior to dosing. Four rats were dosed by gavage with three daily doses of 0.3 mmol/kg body weight 3H-1,2-dithole-3-thione, 48 h prior to administration of a single dose of 80 nmol (25 µg) of AFB1 by gavage. Animals were housed in glass metabolic cages for 24 h immediately following administration of AFB1 for collection of urine. Animal experiments were performed in accordance with protocols approved by the ACUC of the Johns Hopkins Medical Institutions. Isolation of AFAR Alcohols from Urine. Urine samples from animals were thawed, and a 5 mL aliquot was adjusted to pH 3.5 with acetic acid prior to centrifugation at 500g for 10 min. Samples were then spiked with 2 ng of 13C17-AFB1 dialcohol internal standard and extracted using 6 cm3 Waters MCX Oasis SPE columns (Waters Corp., Milford, MA) equilibrated with 1 column volume of methanol followed by 1 column volume of water prior to loading the urine sample. AFAR alcohols were eluted from the SPE columns with 10 mL of 100% methanol and reduced to a final volume of approximately 50 µL using high-purity nitrogen. This sample was then diluted to approximately 600 µL with water and loaded onto an aflatoxin-specific preparative monoclonal antibody immunoaffinity column, as previously described (10). The affinity column was washed with PBS and water to remove nonspecifically bound materials, and the AFAR alcohols were eluted from the immunoaffinity column with 6 mL of 70% dimethylsulfoxide/water (v/v) followed by another 2 volumes of water. The dimethylsulfoxide and water fractions were combined, diluted with water, and then applied to a 3 cm3 Varian Bond-Elut LRC C18 SPE column (Varian, Inc., Walnut Creek, CA). AFAR alcohols were eluted from the SPE with 5 mL of a 50/50 mixture of 1% acetic acid/methanol followed by 5 mL of 100% methanol and concentrated to approximately 40 µL under a nitrogen stream. A total of 1 µL of final urinary extracts was diluted with 24 µL of HPLC initial mobile phase prior to LC/MS/MS analysis. β-Glucuronidase and Sulfatase Treatment of Rat Urine. Thawed 5 mL urine aliquots were adjusted to pH 3.5 with acetic acid, centrifuged, and extracted with 6 cm3 Waters MCX Oasis SPE columns (Waters Corp., Milford, MA) as described above. Final SPE eluents were brought up to a volume of 500 µL with 70 mM sodium acetate at pH 5.0 buffer and digested with 2000 units of bovine liver β-glucuronidase and 300 units of sulfatase from Helix pomatia (Sigma Chemical, St. Louis, MO) for 12 h at 37 °C. Digested samples were brought to room temperature before applying to aflatoxin-affinity columns and processing as


described above to produce final urinary extracts for LC/MS/ MS analysis. Tandem LC/MS Analyses of Sample Analytes and Chromatographic Standards. Analyses of sample analytes and chromatographic standards were carried out on a ThermoFinnigan Deca ion-trap mass spectrometer coupled to a ThermoFinnigan Surveyor Plus HPLC and autoinjector (ThermoElectron Corporation, San Jose, CA). Samples were maintained at 4 °C before injection. Chromatographic separation was carried out at ambient temperature using a flow rate of 25 µL/min on a 1 mm × 150 mm × 5 µm Luna C18 HPLC column (Phenomenex, Torrance, CA). Mobile-phase compositions for alcohol method 2 were as follows: (A) 1% aqueous acetic acid, (B) methanol, and (C) acetonitrile. The initial composition (75% A, 23% B, and 2% C) changed to (70% A, 28% B, and 2% C) over 5 min, to (65% A, 33% B, and 2% C) over 15 min, and to (60% A, 38% B, and 2% C) over 10 min and held for 15 min before returning to initial composition over 5 min. Column flow was diverted away from the ESI ion source during the initial 3.0 min. Under these conditions, C-8 monoalcohol, dialcohol, C-6a monoalcohol, and dihydrodiol (acidic form of AFB1dialdehyde) eluted at retention times of ∼26.0, ∼30.0, ∼34.0, and ∼37.0 min, respectively. The 13C17-AFB1 dialcohol internal standard eluted at ∼30.0 min. Positive ESI/MS/MS was conducted with the capillary temperature set at 225 °C, the sheath gas at 90 arbitrary units, and the spray voltage at 4.5 kV using He collision gas. Analyte and chromatographic standards were detected in the full-scan mode at 40% collision energy, monitoring the dialcohol transitions at m/z 351 f m/z 100–500, the monoalcohols transitions at m/z 349 f m/z 100–500, the dihydrodiol transitions at m/z 329 f m/z 100–500, and the 13C17-AFB1 dialcohol internal standard transitions at m/z 368 f m/z 100–500. Predominant daughter ions, representing 100% relative abundance in the m/z 100–500 range are m/z 333.0, m/z 331.0, m/z 329.0, m/z 350 for the dialcohol, monoalcohols, dihydrodiol, and the 13C17AFB1 dialcohol internal standard.

Results Synthesis of the 13C17-AFB1 Dialcohol Internal Standard. Initial experiments were performed to synthesize a 13C17-AFB1dialcohol internal standard, the final reduction product of the AFAR pathway (Figure 1), thereby providing a basis for developing a quantitative mass spectrometry method. The 13C17AFB1 used in the preparation of the internal standard was biosynthetically produced in A. flaVus using 13C-glucose as the sole carbon source. LC-MS and MS/MS analysis confirmed the production of 13C17-AFB1 as well as the completeness of 13 C-incorporation into the AFB1 molecule. Full mass spectrometric analysis of AFB1 and 13C17-AFB1 yielded parent molecular ions of [M + H]+ ) 313.0 and [M + H]+ ) 330.0, respectively (parts A and B of Figure 2). The incorporation of the 13C isotope determined by full-scan MS analysis was >99.0%. The 13C17-AFB1 dialcohol internal standard was synthesized by incubating purified 13C17-AFB1 with dimethyldioxirane to form the 13C17-AFB1 epoxides. Subsequent aqueous hydrolysis of these epoxides in water yielded the 13C17-AFB1 dihydrodiol product. The percent yield of 13C17-AFB1 dihydrodiol determined by HPLC was >90.0%. HPLC-purified 13C17-AFB1 dihydrodiol was then transformed to the immediate internal standard precursor, 13C17-AFB1 dialdehyde, by incubation in a basic buffer. To generate the 13C17-AFB1 dialcohol internal standard, the resulting dialdehyde precursor was reduced using


sodium borohydride treatment. 13C17-AFB1 dialcohol internal standard was isolated by HPLC and then subjected to MS and MS/MS characterization. The percent yield of the internal standard from the dialdehyde starting material was determined by HPLC to be >75.0%. The purified 13C17-AFB1 dialcohol standard was stored under acidic conditions (∼pH 3.5) at 4 °C, and under these conditions, the alcohols shown in Figure 1 are stable for at least 1 year. Full-mass spectrometric analysis of dialcohol and 13C17-AFB1 dialcohol standards yielded parent molecular ions of [M + H]+ ) 351.0 and [M + H]+ ) 368.0, respectively (parts C and D of Figure 2). Isotope Dilution Tandem Mass Spectrometry Method. HPLC-gradient conditions provided baseline separation of the dialcohol, monoalcohols, dihydrodiol (acidic form of AFB1 dialdehyde), and 13C17-AFB1 dialcohol internal standard (parts A-D of Figure 3). Under these conditions, the dialcohol standard and 13C17-AFB1 dialcohol internal standard have virtually identical retention times, differing by less than 0.1 min (left side of parts A and D of Figure 3). The corresponding MS/MS spectra for the dialcohol, monoalcohols, dihydrodiol, and internal standard (right side of parts A-D of Figure 3) depict fragments that differ from the parent molecular ions by -18 mass units. This loss of 18 mass units represents the loss of a single water molecule: a signature MS/MS fragment of alcohol-containing molecules (see standard structures in Figure 1). MS/MS analyses across a mass range of m/z 100–500 yielded secondary daughter ions for dialcohol, monoalcohols, dihydrodiol, and internal standard of m/z 333 f 315.0, m/z 331 f 313.0, m/z 347 f 301.0, and m/z 368 f 332.0, respectively (data not shown). The relative percent abundance of these predominant secondary daughter ions for the dialcohol, monoalcohols, dihydrodiol, and internal standard were 21.2, 11.1, 24.9, and 22.6%, respectively. MS/MS spectra shown in Figure 3 detail major daughter ions with percent relative abundances of 100.0%. A collision energy of 40.0% was used to induce MS/ MS fragmentation. 13 C17-AFB1 dialcohol served as a surrogate internal standard for monoalcohol and dihydrodiol analytes. The dialcohol is the ultimate reduction product of the AFAR pathway (Figure 1) and the least capable of all analytes to form protein adducts; therefore, use of a dialcohol-based internal standard to account for sample analyte recoveries eliminates any spurious reduction and/or protein binding that may occur with monoalcohol- or dihydrodiol-based internal standards. Linear 10-point isotopic dilution standard curves were generated for dialcohol (r2 ) 0.998), C-8 monoalcohol (r2 ) 0.994), C-6a monoalcohol (r2 ) 0.999), and dihydrodiol (r2 ) 0.999). The limits of detection for dialcohol, C-8 monoalcohol, C-6a monoalcohol, and dihydrodiol were determined to be 10.0, 16.0, 4.0, and 6.0 pg injected on the column, respectively. Stopped-flow quantitation and LC-MS characterization of AFAR alcohol products formed from AFB1 dialdehyde incubations with AKR7A1 have been previously reported (8). To test the applicability of the isotope dilution LC-MS/MS method to quantify AFAR alcohol products, AFB1-dialdehyde incubations were conducted with AKR7A1 and the resulting AFAR alcohol products were measured. For method validation purposes, kinetic monoalcohol parameters obtained from this method were compared to kinetic monoalcohol parameters obtained from the previous report (8). We monitored and quantified monoalcohol formation resulting from the addition of 0–100 µM AFB1 dialdehyde substrate to 0.25 µM AKR7A1, as described in the Experimental Procedures. Analyses of the obtained linear velocities versus the AFB1 dialdehyde substrate

756


Figure 2. Full-scan MS spectra for (A) AFB1 [M + H]+ ) 313.0, (B) and (D) 13C17-AFB1 dialcohol internal standard [M + H]+) 368.0.

concentration were best-fit with the Michaelis–Menten equation to calculate kinetic parameters for the monoalcohols. The kcat (min-1) and Km (µM) kinetic parameters for the C-8 monoalcohol are in direct agreement with the previous literature report. Our results are as follows for kcat (min-1) 12 ( 2 and for Km (µM) 112 ( 2 [mean ( standard deviation (SD) from three independent experiments]. Detection of Urinary AFB1 Mono- and Dialcohols in Rats. Urine samples from male Sprague–Dawley rats pretreated with a oral dose of D3T, an inducer of AKR7A1 protein expression, prior to oral administration of AFB1 were used to

Johnson et al.

C17-AFB1 [M + H]+) 330.0, (C) AFB1 dialcohol [M + H]+ ) 351.0,

13

identify and quantify detectable levels of AFAR alcohols excreted in urine, 24 h following a single AFB1 dose. An aflatoxin-specific preparative monoclonal antibody immunoaffinity column was used in sample processing to capture AFAR alcohols from applied urine samples, as described in the Experimental Procedures. Immunoaffinity methodology has been used for the urinary extraction of AFB1-N7-guanine, AFB1mercapturic acid, aflatoxin M1, aflatoxin P1, and other aflatoxin metabolites from human and animal samples (9, 10, 23, 24). The cross-reactivity of the AFAR alcohols with the immunoaffinity columns was tested with synthesized alcohol standards



Figure 3. LC chromatograms (left) and MS/MS spectra (right) of predominant daughter ions of synthesized method standards. (A) Dialcohol (m/z 351 f 333); ion intensity, 1.35 × 106. (B) C-8 and C-6a monoalcohols (m/z 349 f 331); ion intensity, 1.81 × 106. (C) Dihydrodiol (m/z 347 f 329); ion intensity, 2.53 × 106. (D) 13C17-AFB1 dialcohol internal standard (m/z 368 f 350); ion intensity, 1.50 × 106.

prior to applying this methodology to rat urine samples and yielded immunoaffinity percent recoveries >90.0%. For quantitation and percent recovery determinations, 2 ng of 13C17-AFB1 dialcohol internal standard was added into all urine samples prior to processing. Representative chromatograms and MS/MS spectra obtained from rat urine are shown in Figure 4. Chromatographic peaks corresponding in both retention time and MS fragmentation patterns of external chromatographic dialcohol and C-6a monoalcohol standards (shown in Figure 3) were observed (parts A-C of Figure 4). To further confirm the identity of the urinary peaks, dialcohol and C-6a monoalcohol standards were spiked into final urine extracts prior to LC-MS/MS analysis to monitor for coelution of spiked standards and increased signal with detected urinary peaks. Standard and sample peak co-elution was observed; thus, the detected peaks in parts A and B of Figure 4 correspond to urinary dialcohol and C-6a monoalcohol metabolites. These peaks correspond to 200 pg of dialcohol and 4 ng of C-6a monoalcohol on the column. We did not detect the C-8 monoalochol in any of the final urinary extracts. To test whether glucuronide conjugation could account for the lack of detection of this alcohol metabolite, all urinary samples were

treated with β-glucuronidase as described in the Experimental Procedures. β-Glucuronidase treatment did not alter the alcohol metabolic profile seen in Figure 4. The internal standard was readily detected in final urine extracts (Figure 4C). The apparent recovery of the method, which takes into account losses during cleanup and matrix suppression in the ESI source, was estimated to be 85–100%. β-Glucuronidase or sulfatase treatment of urinary samples did not change the alcohol metabolic profile. The concentrations of rat urinary alcohols were 203.5 ( 39.0 ng of monoalcohol C-6a/mg of urinary creatinine and 10.0 ( 1.0 ng of dialcohol/mg of creatinine (mean ( standard error, four animals). These levels at 24 h post-dosing represent 0.81 and 0.04% of the aflatoxin B1 dose administered, respectively. Previous experiments in rats have shown that about 10% of the administered dose of aflatoxin B1 is excreted into urine by 24 h; therefore, the monoalcohol C-6a and the dialcohol represents about 8.0 and 0.4% of the aflatoxin metabolites in urine, respectively (9). The limit of detection for these alcohol metabolites was calculated to be 0.75 ng/mg creatinine. Thus, successful detection of these alcohol products in urine with the isotope dilution MS methodology allows for a more complete mass balance of the fate of the 8,9-epoxides in ViVo and permits

758


Johnson et al.

Figure 4. Representative LC chromatograms (left) and MS/MS spectra (right) from a processed D3T-AFB1 rat urine sample. (A) Detected urinary dialcohol peak; ion intensity, 6.08 × 105. (B) Detected urinary C-6a monoalcohol peak; ion intensity, 7.07 × 106. (C) Detected 13C17-AFB1 dialcohol; ion intensity, 9.11 × 104. Detected urinary peaks correspond to retention times and MS/MS fragmentation patterns of standards shown in Figure 3.

the exploration of these aflatoxin alcohols as biomarkers of pharmacodynamic effects of modulators of the AFAR pathway.

Discussion The determination of the mechanistic role of enzymes in bioactivating and detoxifying pathways of AFB1 has long been an important strategy for devising chemoprevention interventions (4, 25). The role of the AFAR system in AFB1 metabolism has been explored in Vitro and ex ViVo by monitoring AFB1 dialdehyde substrate-based AFAR kinetics of rat and/or human isozymes and using human liver cytosols (8). To date, however, only limited data on in ViVo AFAR activity have been reported. The metabolic pathway described in Figure 1 highlights that AFAR has the potential to modulate the formation of covalent protein adducts by AFB1, through altering the disposition of the aflatoxin dialdehyde. It is therefore important to conduct in ViVo analyses of the products across the AFB1 protein adductforming pathway; the protein adducts themselves as well as the AFAR alcohol products begin to fully characterize the biological significance and fate of AFB1 dialdehyde and the role of AFAR in its disposition. Quantitative isotope dilution MS methodologies, as well as an array of other analytical techniques, are currently available to quantify levels of in ViVo AFB1 protein adducts by monitoring serum-albumin-derived AFB1-lysine adducts (26-30). This paper reports an isotope dilution MS methodology for quantification of in ViVo AFAR alcohol products and is an important addition in our goal to establish specific, sensitive, and quantitative mass spectrometry methods for the detection and measurement of in ViVo AFB1 biomarkers formed by the critical AFB1 epoxides.

A 13C17-AFB1 dialcohol internal standard provides the basis for quantification of AFAR alcohol products in this described method. To generate the 13C17-AFB1 precursor needed for internal standard synthesis, uniformly labeled 13C-glucose was provided to A. flaVus cultures as their sole carbon source for biosynthesis. This approach yielded >99.9% incorporation of the 13C-isotope into the AFB1 ring structure. Use of a fully reduced 13C17-AFB1 dialcohol internal standard eliminated the possibility of a secondary reaction with proteins and its in Vitro reductive metabolism when method validation was conducted with kinetic incubations of AFB1 dialdehyde with AKR7A1. Indeed, although the application of this method to the in ViVo analyses of AFAR alcohol products is examined in this report, this method can also be used for LC-MS/MS quantification of AFB1 dialdehyde substrate-based kinetics of AFARs. The kinetics-based approach to method validation resulted in Km and kcat parameters for C-8 monoalcohol that are in direct agreement with previously reported kinetic parameters for this AFAR alcohol (8). These data assured the accuracy of the method in quantifying AFAR alcohol products and provided confidence in applying this LC-MS/MS method to urinary analyses of AFAR alcohols from rats experimentally treated with AFB1. Application of the isotope dilution MS method to in ViVo analyses of AFAR alcohol products revealed the detection and quantification of the dialcohol and C-6a monoalcohol products in the urine of dosed male Sprague–Dawley rats. Prior studies to analyze AFAR alcohol products in urine samples noted that low constitutive protein levels of AKR7A1 in rat livers may result in very low levels of AFAR alcohols produced in rat urine (8). To promote the in ViVo formation of these alcohols, rats were pretreated with D3T to enhance AKR7A1 protein expres-


sion prior to administering AFB1. The level of D3T administered to animals has been previously estimated to elevate hepatic AKR7A1 protein levels approximately 18-fold relative to control animals (15). Using this enzyme induction protocol, the LC-MS/MS data from rat urinary samples clearly showed the detection of urinary AFB1 dialcohol and AFB1 C6-a monoalcohol metabolites. The C6-a monoalcohol was the predominant urinary alcohol metabolite, with detected levels comparable to the oxidative metabolite, AFM1, in urine of AFB1-dosed Sprague–Dawley rats (unpublished results). Studies have demonstrated that, 24 h following a single dose of AFB1, albumin adducts and total urinary excretion products account for 1–2 and 10% of the administered dose, respectively, in rats (31). AFM1 has been reported as the major urinary metabolite in AFB1-treated rats and has been detected at levels that represent 1–2% of an administered single dose of AFB1; however, minor urinary metabolites, such as aflatoxin N7-guanine, have been detected in rat urinary samples at levels that represent 0.1% of the total AFB1 dose (32). Thus, the levels of AFAR alcohols, monoalcohol C-6a (0.81%) and the dialcohol (0.04%), detected in this investigation straddle the previously reported concentration range of AFB1 urinary metabolites. The remaining dose is either macromolecularly bound or excreted by the biliary/fecal pathway. The C-8 monoalcohol was not detected in the studied samples. To confirm that our urinary cleanup method retained this monoalcohol for its detection and quantification in final urinary extracts, C-8 monoalcohol was spiked into control rat urine and yielded a 30% recovery. Because hepatic cytosols from D3Tdosed male Sprague–Dawley rats form the same levels of C-8 monoalcohol and C-6a monoalcohol when AFB1 dialdehyde is administered as a substrate (unpublished results), there is a discontinuity between the in Vitro and in ViVo observation. Given the high urinary levels of the C-6a monoalcohol seen in this report, 30% recovery of the C-8 monoalcohol does not account for the total lack of detection of this alcohol product. Preliminary studies, treating urinary samples with either β-glucuronidase or sulfatase, revealed no changes in the urinary alcohol profile; thus, it does not appear that the C-8 monoalcohol forms a glucuronide or a sulfate conjugate. The method described herein results in robust LC-MS/MS detection and quantitation of AFAR alcohols produced in ViVo, thereby adding to the repertoire of technologies available to mass balance the fate of the AFB1 epoxides. Using this method, exploration of the AFAR pathway, monitoring for both urinary AFAR alcohols and modulation of the AFB1 protein adducts (as measured by serum-albumin-derived AFB1 lysine adducts), when AFAR is either induced by treatment with an inducer or overexpression in transgenic animals, can be conducted. The development of biomarkers reflecting the fate of the aflatoxin epoxide-derived metabolites requires quantitative technologies that can be applied to chemoprevention interventions in highrisk populations. The tools developed in these current studies have applicability for assessing the role of AFAR enzymes in human AFB1 metabolism and in evaluating chemoprevention paradigms.


(2) (3) (4) (5)

(6) (7) (8)

(9)

(10)

(11)

(12)

(13) (14)

(15)

(16)

(17)

(18) (19) (20) (21)

Acknowledgment. This work was supported by NIH Grants CA39416 and ES06052, NIEHS Training Grant T32ES07141, NIEHS Center Grant ES03819, and NAS Ford Foundation Predoctoral Fellowship.

(22) (23)

References (1) Groopman, J. D., and Cain, L. G. (1990) Interactions of fungal and plant toxins with DNA: Aflatoxins, sterigmatocystin, safrole, cycasin,

(24)

and pyrrolizidine alkaloids. In Handbook of Experimental Pharmacology, Chapter 10, Vol. 94/I, Springer-Verlag, Berlin Heidelberg, Germany. Busby, W. F. J., and Wogan, G. N. (1984) Aflatoxins. In Chemical Carcinogens, 2nd ed. (Searle, C. B., Ed.) pp 945–1136, American Chemical Society, Washington, D.C. Council for Agricultural Science and Technology. (2003) Mycotoxins: Risks in plant, animal, and human systems. In Task Force Report, number 139. Guengerich, F. P. (2003) Cytochrome P450 oxidations in the generation of reactive electrophiles: Epoxidation and related reactions. Arch. Biochem. Biophys. 409, 59–71. Johnson, W. W., and Guengerich, F. P. (1997) Reaction of aflatoxin B1 exo-epoxide with DNA: Kinetic analysis of covalent binding and DNA-induced hydrolysis. Proc. Natl. Acad. Sci. U.S.A. 94, 6121– 6125. Sabbioni, G., Skipper, P. L., Buchi, G., and Tannenbaum, S. R. (1987) Isolation and characterization of the major serum albumin adduct formed by aflatoxin B1 in vivo in rats. Carcinogenesis 8, 819–824. Scholl, P. F., Musser, S. M., and Groopman, J. D. (1997) Synthesis and characterization of aflatoxin B1 mercapturic acids and their identification in rat urine. Chem. Res. Toxicol. 10, 1144–1151. Guengerich, F. P., Cai, H., McMahon, M., Hayes, J. D., Sutter, T. R., Groopman, J. D., Deng, Z., and Harris, T. M. (2001) Reduction of aflatoxin B1 dialdehyde by rat and human aldo-keto reductases. Chem. Res. Toxicol. 14, 727–737. Walton, M., Egner, P., Scholl, P. F., Walker, J., Kensler, T. W., and Groopman, J. D. (2001) Liquid chromatography electrospray-mass spectrometry of urinary aflatoxin biomarkers: Characterization and application to dosimetry and chemoprevention in rats. Chem. Res. Toxicol. 14, 919–926. Egner, P. A., Groopman, J. D., Wang, J., Kensler, T. W., and Friesen, M. D. (2006) Quantification of aflatoxin-B1-N7-guanine in human urine by high-performance liquid chromatography and isotope dilution tandem mass spectrometry. Chem. Res. Toxicol. 19, 1191–1195. Scholl, P. F., Turner, P. C., Sutcliffe, A. E., Sylla, A., Diallo, M. S., Friesen, M. D., Groopman, J. D., and Wild, C. P. (2006) Quantitative comparison of aflatoxin B1 serum albumin adducts in humans by isotope dilution mass spectrometry and ELISA. Cancer Epidemiol., Biomarkers PreV. 15 (4), 823–826. Egner, P. A., Yu, X., Johnson, J. K., Nathasingh, C. K., Groopman, J. D., Kensler, T. W., and Roebuck, B. D. (2003) Identification of aflatoxin M1-N7-guanine in liver and urine of tree shrews and rats following administration of aflatoxin B1. Chem. Res. Toxicol. 16, 1174–1180. Jez, J. M., Flynn, T. G., and Pennings, T. M. (1997) A new nomenclature for the aldo-keto reductase superfamily. Biochem. Pharmacol. 54, 639–647. Hayes, J. D., Judah, D. J., and Neal, G. E. (1993) Resistance to aflatoxin B1 is associated with the expression of a novel aldo-keto reductase which has catalytic activity towards a cytotoxic aldehydecontaining metabolite of the toxin. Cancer Res. 53, 3887–3894. Knight, L. P., Primiano, T., Groopman, J. D., Kensler, T. W., and Sutter, T. R. (1999) cDNA cloning, expression and activity of a second human aflatoxin B1-metabolizing member of the aldo-keto reductase superfamily, AKR7A3. Carcinogenesis 20 (7), 1215–1223. Ellis, E. M., Judah, D. J., Neal, G. E., and Hayes, J. D. (1993) An ethoxyquin-inducible aldehyde reductase from rat liver that metabolizes aflatoxin B1 defines a subfamily of aldo-keto reductases. Proc. Natl. Acad. Sci. U.S.A. 90, 10350–10354. Kwak, M. K., Egner, P. E., Dolan, P. M., Ramos-Gomez, M., Groopman, J. D., Itoh, K., Yamamoto, M., and Kensler, T. W. (2001) Role of phase 2 enzyme induction in chemoprotection by dithiolethiones. Mutat. Res. 480–481, 305–315. Primiano, T., Gastel, J. A., Kensler, T. W., and Sutter, T. R. (1996) Isolation of cDNAs representing dithiolethione-responsive genes. Carcinogenesis 17, 2297–2303. Ellis, E. M., and Hayes, J. D. (1995) Substrate specificity of an aflatoxin-metabolizing aldehyde reductase. Biochem. J. 312, 535–541. Baertschi, S. W., Raney, K. D., Stone, M. P., and Harris, T. M. (1988) Preparation of the 8,9-epoxide of the mycotoxin aflatoxin B1: The ultimate carcinogenic species. J. Am. Chem. Soc. 110, 7929–7931. Adye, J., and Mateles, R. I. (1964) Incorporation of labeled compounds into aflatoxins. Biochim. Biophys. Acta 88, 418–420. Buchanan, R. L., Hoover, D. G., and Jones, S. B. (1983) Caffeine inhibition of aflatoxin production: Mode of action. Appl. EnViron. Microbiol. 46, 1193–2000. Groopmam, J. D., Trudel, L. J., Donahue, P. R., Marshak-Rothstein, A., and Wogan, G. N. (1984) High affinity monoclonal antibodies for aflatoxins and their application to solid-phase immunoassays. Proc. Natl. Acad. Sci. U.S.A. 81, 7728–7731. Groopman, J. D., Donahue, P. R., Zhu, J., Chen, J., and Wogan, G. N. (1985) Aflatoxin metabolism in humans: Detection of metabolites and

760

(25)

(26)

(27)

(28)

Chem. Res. Toxicol., Vol. 21, No. 3, 2008 nucleic acid adducts in urine by affinity chromatography. Proc. Natl. Acad. Sci. U.S.A. 82, 6492–6496. Guengerich, F. P., Shimada, T., Raney, K. D., Yun, C., Meyer, D. J., Ketterer, B., Harris, T. M., Groopman, J. D., and Kadlubar, F. F. (1992) Elucidation of catalytic specificities of human cytochrome P450 and glutathione S-transferases enzymes and relevance to molecular epidemiology. EnViron. Health Perspect. 98, 75–80. Scholl, P. F., Turner, P. C., Sutcliffe, A. E., Sylla, A., Diallo, M. S., Friesen, M. D., Groopman, J. D., and Wild, C. P. (2006) Quantitative comparison of aflatoxin B1 serum albumin adducts in humans by isotope dilution mass spectrometry and ELISA. Cancer Epidemiol., Biomarkers PreV. 15 (4), 823–826. Chapot, B., and Wild, C. P. (1991) ELISA for quantification of aflatoxin-albumin adducts and their application to human exposure assessment. In Techniques in Diagonostic Pathology (Van Warhol, M., Velzen, D., and Bullock, G. R., Eds.) Vol. 2, pp 139–155, Academic Press, Inc., New York. Gan, L.-S., Skipper, P. L., Peng, X., Groopman, J. D., Chen, J.-S., Wogan, G. N., and Tannenbaum, S. R. (1988) Serum albumin adducts in the molecular epidemiology of aflatoxin carcinogenesis: correlation

Johnson et al.

(29)

(30)

(31)

(32)

with aflatoxin B1 intake and urinary excretion of aflatoxin M1. Carcinogenesis 9, 1323–1325. Sabbioni, G., Ambs, S., Wogan, G. N., and Groopman, J. D. (1990) The aflatoxin-lysine adduct quantified by high-performance liquid chromatography from human serum albumin samples. Carcinogenesis 11, 2063–2066. Wild, C. P., Jiang, Y.-Z., Sabbioni, G., Chapot, B., and Montesano, R. (1990) Evaluation of methods for quantitation of aflatoxin-albumin adducts and their application to human to human exposure assessment. Cancer Res. 50, 245–251. Hsieh, D. P. H., and Wong, J. J. (1994) Pharmacokinetics and excretion of aflatoxins. In The Toxicology of Aflatoxins: Human Health, Veterinary, and Agricultural Significance (Eaton, D. L., and Groopman, J. D., Eds.) pp 73–88, Academic Press, San Diego, CA. Groopman, J. D., DeMatos, P., Egner, P. A., Love-Hunt, A., and Kensler, T. W. (1992) Molecular dosimetry of urinary aflatoxin N7guanine and serum aflatoxin albumin adducts predict chemoprotection by 1,2-dithiole-3-thione in rats. Carcinogenesis 13, 101–106.

TX700397N

Quantification of Urinary Aflatoxin B1 Dialdehyde Metabolites Formed

Recommend Documents