DNA Adducts from N- Nitrosodiethanolamine and Related β-Oxidized Nitrosamines in Vivo: 32 P-Postlabeling Methods for Glyoxal- and O 6 -Hydroxyethyldeoxyguanosine Adducts

DNA Adducts from N-Nitrosodiethanolamine andRelated â-Oxidized Nitrosamines in Vivo:32P-Postlabeling Methods for Glyoxal- andO6-Hydroxyethyldeoxyguanosine Adducts

Richard N. Loeppky,* Qiuping Ye, Petra Goelzer, and Yuen Chen

Department of Chemistry, University of Missouri, Columbia, Missouri 65211

Received August 22, 2001

The mechanism by which environmentally prevalent N-nitrosodiethanolamine (NDELA) andrelated 2-hydroxyethyl- or other â-oxidized nitrosamines initiate the carcinogenic process hasremained obscure. 32P-Postlabeling assays for the pH sensitive glyoxal-deoxyguanosine (gdG)and the O6-2-hydroxyethyldeoxyguanosine (OHEdG) DNA adducts have been developed asprobes in this mechanistic investigation and used in both in vitro and in vivo experiments.The ready cleavage of the glyoxal fragment from gdG at pH 7 and greater has required methodsof optimization in order to achieve a detection limit of 0.05 µmol/mol of DNA. Nuclease P1treatment enhances the detection of gdG adducts but does not increase the detection limit forOHEdG. For OHEdG, best results were achieved using fraction collection from HPLC (0.3µmol/mol of DNA). Using radiochemical methods, both adducts could be detected either byHPLC or 2D TLC. NDELA, N-nitrosomorpholine (NMOR), N-nitrosomethyethanolamine(NMELA), and N-nitrosoethylethanolamine (NEELA) all produce both gdG and OHEdG adductsin rat liver DNA in vivo and are called bident carcinogens because fragments from both chainsof the nitrosamine are incorporated into DNA. N-Nitroso-2-hydroxymorpholine (NHMOR), ametabolite of NDELA and NMOR, generates gdG in DNA in vitro and in vivo. gdG DNA adductswere found in the range 1.1-6.5 µmol/mol of DNA. OHEdG DNA adducts were produced fromequimolar amounts of nitrosamines in rat liver in vivo over the range 4-25 µmol/mol of DNAand in the order NMELA > NEELA > NDELA > NMOR. Deuterated isotopomers of NDELAshowed a marked isotope effect on DNA OHEdG adduct formation. R-Deuteration markedlydecreased OHEdG adduct formation while â-deuteration had the opposite effect. These datasupport the hypothesis that NDELA and related nitrosamines are activated by both enzymemediated R-hydroxylation and â-oxidation. The formation of OHEdG adducts from NDELArequires R-hydroxylation of the 2-hydroxyethyl chain, and formation of gdG necessitates aâ-oxidation as well. The bident nature of these carcinogens may explain why they are relativelypotent carcinogens despite the fact that major proportions of doses are excreted unchanged.

Introduction

Despite significant advances in the treatment of vari-ous cancers, prevention of the onset of the carcinogenicprocess remains the best strategy for dealing with thesehorrific and costly diseases. Methods for the determina-tion and reduction of risk are powerful tools in the questfor enhanced cancer prevention. These methods rely ona combination of chemical, biochemical, and epidemio-logical approaches, which establish pathways and routesof carcinogen exposure, mechanisms of bio-activation, andthe consequences thereof in the human setting. Thetemporal separation of cause and effect in these diseasespresents the biggest challenge to successful prevention.An enormous research effort has focused attention on therole of mutations in the initiation, development, andprogression of neoplastic diseases. There is strong evi-dence linking DNA base modification, by carcinogenfragment adduction or other chemical transformations,to increased mutation frequency. Thus, for some years,

the identification of, and the determination of DNA orrelated protein adducts have been the focus of muchresearch and have served as valuable endpoints in boththe elucidation of mechanisms of carcinogenesis and thedevelopment of chemically based epidemiology to asseshuman risk. Here we report on the development of andthe utilization of two 32P-postlabeling assays for DNAadducts derived from two important environmental car-cinogens, N-nitrosodiethanolamine (NDELA)1 1 and N-

nitrosomorpholine (NMOR) 2, the glyoxal-deoxygua-nosine (gdG) adduct 6 and the O6-hydroxyethyldeoxy-guanosine (OHEdG) adduct 7.

* To whom correspondence should be addressed. Phone: (573) 882-4885. E-mail: [email protected].

470 Chem. Res. Toxicol. 2002, 15, 470-482

10.1021/tx0101393 CCC: $22.00 © 2002 American Chemical SocietyPublished on Web 03/20/2002

NDELA and NMOR have a common metabolite (1-3), 2-hydroxy-N-nitrosomorpholine (NHMOR) 3, which isa hemiacetal of an R-nitrosamino aldehyde. NHMOR andother R-nitrosamino aldehydes both deaminate bases,and form gdG adducts with nucleosides, nucleotides, andcalf thymus DNA in vitro (4-11), but the significance ofthese observations has been unclear. In the precedingpaper in this issue, we clearly show that NDELA ismetabolically activated by microsomes to give glyoxal andproducts arising from the 2-hydroxyethyldiazonium ion(12). The same is true for other 2-hydroxyethyl nitro-samines 4 and 5. These metabolites are also formedduring the microsomal incubation of NHMOR. Whilework on the mechanism of bioactivation of NDELA wasunderway, we sought methods for the sensitive detectionof DNA adducts from these metabolites, which we neededfor in vivo studies. Because Farrely et al. had reportedthat only low levels of 14C-labeled NDELA were incor-porated into rat liver DNA (13, 14), we looked for highlysensitive methodology and our attention was drawn tothe potential of 32P-postlabeling assays.

The development of 32P-postlabeling assays of DNAadducts was pioneered by R. C. Gupta, K. Randerath,and M. V. Reddy (15-17), and in the 20 years since theoriginal publication, the method has been utilized invarious forms by many research groups (18). The car-cinogen modified DNA is enzymatically hydrolyzed to 3′-mononucleotides and 5′-labeled with 32P, and resultingdiphosphates or enzymatically produced 5′-mononucle-otides are separated chromatographically and detectedby radiochemical methods, often employing autoradio-graphic analysis of 2D TLC plates. The method is mostpowerful when used with authentic nucleotide standardsand has the advantage of very high sensitivity. Theprincipal problems to be overcome in the development of32P-postlabeling methods for the gdG adduct revolvedaround its instability at neutral to basic pH (19-22), andin the case of the OHEdG adduct, the synthesis of thestandard (23). Here we report the successful developmentof assays for these adducts and their detection in thelivers of rats administered five different nitrosamines1-5.

Experimental Methods

Chemicals. Glyoxal was purchased from Aldrich as a 40%solution in water. The nitrosamines N-nitrosodiethanolamine(NDELA), N-nitrosoethylethanolamine (NEELA), N-nitrosom-ethyl-ethanolamine (NMELA), N-nitrosomorpholine (NMOR)and N-nitroso-2-hydroxymorpholine (NHMOR) are all known

compounds and were prepared by published methods. Theirpurity was determined by normal phase HPLC on Si-60 withhexane: ethyl acetate and UV detection (λ ) 254 nm) to be>99%. Caution: Most nitrosamines have been shown to beanimal carcinogens and have to be handled very carefully.Aqueous solutions are decontaminated with NaOH and alumi-num foil overnight. Concentrated nitrosamines in nonaqueoussolvents can be destroyed by HBr-glacial acetic acid followed bydilution. Deoxyguanosine (dG), guanine, glyoxal, 2′-deoxygua-nosine-3′-monophosphate sodium salt (dGMP), the correspond-ing 5′-monophosphate, spermidine, calf-thymus DNA (CT-DNA),isoamyl alcohol, sodium dodecyl sulfate (SDS), phosphodi-esterase I (venom exonuclease) (EC 3.1.4.1), spleen phosphodi-esterase (SPD, EC 3.1.16.1), proteinase K, ribonuclease A(RNase A), and RNase T1 were purchased from Sigma (St.Louis, MO). United States Biochemicals (Cleveland, OH) wasthe source for deoxyribonuclease I (DNase I) (EC 3.1.21.1),bacterial alkaline phosphatase from E. coli (EC 3.1.3.1), Exo-nuclease III (EC 3.1.11.2), DTT, micrococcal nuclease (MN),Nuclease P1 (NP1), and T4 polynucleotide kinase. γ32P-ATP(3000 Ci/mmol) was obtained from NEN. TLC plates: PEIcellulose CEL 300 were purchased from Macherey-Nagel (Du-eren, Germany), and CEL 300 plates from EM Science (Cincin-nati, OH). Medical X-ray film RX was purchased from Fuji(Tokyo, Japan).

Instrumentation. NMR spectra were taken on a BrukerAMX 500 instrument. HPLC separation was carried out on aWaters HPLC system with an autosampler WISP 710 B, twopumps model 510, a multiwavelength UV-detector 490, and aradiochemical detector â-flow A 250 from Radiomatic Instru-ments. The system was controlled with Millenium 2010 soft-ware. The radioactive spots on the TLC were quantified with aPhosphorimager 400A from Molecular Dynamics with Im-ageQuant software (3.15) or those from autoradiography byusing a system 200 imager scanner and an auto changer 1000from Bioscan, Inc. (Washington, DC).

Preparation of Standards. The glyoxal-guanine (gG), andguanosine adducts were synthesized as described by Shapiroet al. (20) The product (1 mg) was dissolved in 1 mg of D6DMSOand the structural assignment was confirmed by proton NMR.The procedure of Chung and Hecht (24) was used to preparethe deoxyguanosine, adduct gdG. The corresponding glyoxal-nucleotide adducts were prepared as follows: 3′-dGMP (0.05mmol) was stirred with 0.25 mmol of glyoxal in 0.1 M phosphatebuffer pH 7 for 6 h at 37 °C. The adduct was separated on HPLCusing a Alltech Synchropak C-18 column (10 × 250 mm) withwater as the eluent and a flow of 2 mL/min. The adduct fractionwas collected and lyophilized. To confirm the structure, 10 µgof the respective adduct was hydrolyzed with 0.1 M HCl at 90°C for 1 h and the neutralized sample was submitted to HPLC(same conditions as above) and compared with an authenticglyoxal guanine standard prepared according to Shapiro. Tofurther confirm the identity of the nucleotide standard it washydrolyzed enzymatically to the nucleoside under much milderconditions: The nucleotide adducts (50 µg) were incubated with200 units of alkaline phosphatase in 100 µL of a 50 mM Tris-buffer pH 8 for 1 h at 37 °C. The sample was co-chromato-graphed with a nucleoside standard prepared according toChung (24). HPLC conditions: Gradient of methanol and 0.1M ammonium acetate pH 4.2 from 7 to 30% in 20 min on anAlltech Synchropak column (10 × 250 mm), flow 2 mL/min, andUV detection (254 nm).

Preparation of Glyoxal Modified CT-DNA Samples. CT-DNA, 1 mg, was dissolved in 1 mL of 0.1 M phosphate buffer,pH 7. The concentration of the DNA solution was determinedby UV. The DNA solution (100 µL) was shaken at 37 °C with 1µL (0.5 mg, 8.6 × 10-6 mol) of 40% aqueous glyoxal solution for6 h. The glyoxal-modified DNA was precipitated by addition of0.1 vol (10 µL) of 3 M sodium acetate, pH 5, and 2.5 vol (250µL) of 95% cold ethanol to the aqueous layer and kept overnightat -20 °C. It was washed twice with 80% cold ethanol and driedby vacuum centrifuge at room temperature. Control DNA, and

1 Abbreviations: 3′-dGMP, 3′-glyoxaldeoxyguanosine monophos-phate; CT-DNA, calf-thymus DNA; dG, deoxyguanosine; DTT, dithio-threitol; gdG, glyoxal-deoxyguanosine; gdGMP, 5′-glyoxaldeoxygua-nosine monophosphate; gDNA, glyoxal modified DNA; gG, glyoxal-guanine; MN, microccoal nuclease; NDELA, N-nitrosodiethanolamine;NEELA, N-nitrosoethylethanolamine; NHEG, N-nitroso-N-2-hydroxy-ethylglycine; NHMOR, N-nitroso-2-hydroxymorpholine; NMELA, N-nitrosomethylethanolamine; NMOR, N-nitrosomorpholine; NP1, nu-clease P1; OHEdG, O6-(2-hydroxyethyl)-deoxyguanosine; SPD, spleenphosphodiesterase; SSB, single strand breaks; TBAP, tetrabutylam-monium phosphate.

DNA Adducts from N-Nitrosodiethanolamine Chem. Res. Toxicol., Vol. 15, No. 4, 2002 471

DNA used for the spiking experiments with the glyoxal-deoxyguanosine-3′-monophosphate (3′-gdGMP) standard wereobtained using the same procedure except no glyoxal was added.

Optimization of Steps for the 32P-Postlabeling of thegdG Adduct. Because the gdG adduct stability is a function ofpH, all procedures, beginning with DNA isolation, and goingon to enzymatic hydrolysis, nuclease P1 enrichment, postlabel-ing, and chromatography, had to be carefully optimized withrespect to pH, concentration, time, and other variables. Theseoptimization procedures are given in the Supporting Informationfor the interested reader. NP1 has been employed for theremoval of unadducted nucleotides by Reddy and Randerath toincrease the sensitivity in 32P-postlabeling (25). We found thatthe gdG 3′-nucleotide was sufficiently resistant to nuclease P1treatment to permit significant enrichment. The 32P-postlabelingmethodology for the gdG adduct was developed in two stages.At the end of Stage 1, the 32P-labeled 3′,5′-diphosphonucleotideswere chromatographed, detected, and quantitated by the opti-mized method reported below. Later, we found that we couldshorten the DNA hydrolysis time, increase the labeling efficiencyby an increase of the pH, and detect the adduct as the5′-mononucleotide. The optimized procedure with these revisionsis reported below as the Stage 2 method. The tabulated data inthe Results clearly indicate which analytical method wasutilized.

32P-Postlabeling Method for the gdG DNA Adduct(Stage 1). The glyoxal modified DNA sample (100 µg), obtainedfrom either in vitro or in vivo experiments, was hydrolyzed with100 units of MN at 37 °C for 10 h after it was dissolved in 50µL of 50 mM Tris-HCl buffer, pH 7, and 50 µL of 20 mM CaCl2.The hydrolysis to the nucleotide 3′-monophosphates was com-pleted by addition of 0.4 units of SPD at 37 °C for 10 h. Theresulting solution, 10 µL, was added to a mixture of 6 µL ofH2O, 2 µL of 2 mM ZnCl2, and 2 µL of a buffered solution ofNP1 (1 µg/µL), which was prepared by dissolving 1 mg in 1 mLof 0.3 M sodium acetate, pH 7, to give a final volume was 20µL. This mixture was incubated at 37 °C for 12 h and dried byvacuum centrifuge at room temperature. The pellet was dis-solved in 4 µL of H2O, then 1 µL of ATP (20 ng/µL), 1 µL of 10µCi/µL [γ-32P]ATP, 2 µL of bicine mixture (100 mM bicine, pH7, 100 mM MgCl2, 10 mM spermidine and 100 mM DTT), and2 µL of T4 polynucleotide kinase (3 units/µL) were added to thesolution. The reaction mixture (10 µL) was incubated at 37 °Cfor 2 h. The labeled nucleotides were separated and quantitatedby two methods, RP ion-paired HPLC and 2D TLC. For HPLC,the sample was injected onto a 4.6 mm × 25 cm ODS ZorbaxLC-18 column, using 50 mM phosphate buffer containing 2 mMtetrabutylammonium phosphate (TBAP), pH 7 (A), and aceto-nitrile (B) as eluents, and a program, 0-10% B in 25 min, then10-30% B in 10 min at a flow rate 1.0 mL/min. Elution wasmonitored by UV absorbance at 254 nm and through the use ofa âradioflow detector. For TLC, a PEI-cellulose TLC plate wasemployed, and the labeled reaction mixture, 1 µL, was applied2.5 cm from the bottom and left-hand edges of a clean PEI-cellulose TLC plate. The spot was dried by cool air. Theprocedure was repeated until 5 µL of the reaction mixture hadbeen applied at the origin. The TLC plate was developed in D1with 6 M urea containing 1.25 M ammonium formate, pH 7, to15 cm from the origin spot at room temperature, then washedwith water and dried completely by cool air. The TLC plate wasthen developed in D2 with 4 M lithium formate, pH 7, to 15 cmfrom the origin spot at room temperature. For autoradiography,the TLC plate was washed with water, dried and Kodak XLS-5X-ray film exposed using intensifying screens at room temper-ature or -20 °C for 1 h or up to several days depending uponthe activity level. The spot of radioactive adduct was locatedfrom the autoradiogram and cut off the plate, then cut into smallpieces and transferred to a counting vial. CytoScint liquidscintillation cocktail, 2 mL, was added to the vial. The spot wasthen counted for 1 min by using a liquid scintillation counter(efficiency ) 99%). In each case, HPLC and TLC, the amount

of the gdG adduct was determined using standard addition(spiking) of the authentic labeled nucleotide.

Optimization of the 32P-Labeling as a Function of pHfor the gdG Adduct (Stage 2). Standards for this part of thework were prepared as described above. For the developmentof HPLC and TLC conditions, a 5′-adduct standard was preparedsimilarly, from dGMP. The phosphorylation reaction was carriedout in total volume of 10.0 µL by adding to a plastic tube either2.0 µL of 3′-gdGMP (0.5 ng/µL) or 3′-dGMP (0.5 ng/µL) standardsand 8.0 µL of a mixture containing 10.0 µCi [γ-32P]ATP, 0.2 µL(30 units/µL) cloned phosphatase-free T4 polynucleotide kinase,distilled water and 2.0 µL of a buffer solution at three differentpHs (7.0, 8.0, and 9.5). The buffer solutions were made by mixing1.45 mg of spermidine, 15.4 mg of DTT, 500 µL of MgCl2 (0.2M), and 500 µL of bicine buffer (0.2 M), pH 7.0, 8.0, or 9.5. Themixtures were incubated for 30 min at 37 °C. After labeling, 10µL of NP1 (1 µg/µL) and 1 µL of NaOAc/HOAc buffer (1 M), pH4.5, were added to the reaction solution to bring the pH to 5.This mixture was incubated for 30 min at 37 °C to ensuremaximal conversion of the 3′,5′-diphosphates to the 5′-phos-phates. The solutions were diluted to 50 µL with distilled water,and 45 µL of aliquots were injected for analysis by HPLC usinga â radioflow detector. HPLC conditions: 4.6 mm × 25 cm ODSZorbax LC-18 column, and mobile phase programs of 50 mMphosphate buffer containing 2 mM TBAP, pH 7, 0-10% aceto-nitrile in 25 min, then 10-30% acetonitrile in 10 min at a flowrate 1.0 mL/min. The maximum yields of labeled gdG adductwere obtained at pH 8 and the method described here wasutilized at this pH.

Optimized Method for 32P-Postlabeling of gdG DNAAdduct (Stage 2). DNA (100 µg) was dissolved in 50 µL of Trisbuffer (50 mM, pH 7) and 50 µL of CaCl2 (20 mM). MN (100units) and 0.4 units of SPD were added and the samples wereincubated for 4 h at 37 °C. Both enzymes, MN and SPD weredialyzed against water before use. The digested DNA (6 µL ) 6µg), 2 µL of ZnCl2 (2 mM), and 6 µL of NP1 solution (1 mg/mLin water) were incubated for 2 h at 37 °C. Then 3 µL of a labelingmix containing 8 µCi 32P-ATP, 6 units of T4-polynucleotidekinase in bicine buffer (100 mM bicine, 100 mM MgCl2, 10 mMspermidine, 100 mM DTT, pH 8) was added, and the sampleswere incubated for 30 min at 37 °C. After this treatment, 1 µLof sodium acetate buffer (1 M, pH 4.5) was added, and themixture (pH 5) incubated for another 30 min. The samples werediluted to 50 µL and injected in the HPLC. If no NP1 was usedfor the enrichment (e.g., the enrichment step was omitted fordirect labeling of standard nucleotides), 2 µL of NP1 was addedafter the labeling together with the acetate buffer. If highlymodified digested CT-DNA was labeled, 10 ng of nonradioactiveATP was added before the labeling to secure an excess of ATP.The labeled samples were separated by HPLC on a Zorbax C-18column (4.6 × 250 mm), flow 1 mL/min, with a gradient of eluentA (50 mM potassium phosphate buffer pH 5 with 2 mM TABP)and B (acetonitrile): 0-25 min, 0 to 10% B; 25-35 min, 10 to30% B; 35-45 min, 30% B; 45-50 min, 30 to 0% B. For CT-DNA samples eluent A consisted of 50 mM sodium phosphatebuffer (pH 7). For TLC, the polyethyleneimine cellulose plateswere washed with deionized water. A 4 µL sample was appliedand the plate was developed in D1 with 1 M lithium formatepH 3.5. The plate was washed in water, air-dried, turned 90°and developed in D2 with 1 M ammonium formate, 1 M urea,(pH 3.5). On cellulose plates, 4 × 1 µL samples were appliedand developed 2 times in isobutyric acid/NH3/H2O (99/1.2/30).The plate was air-dried overnight, turned 90° and developed in(NH4)2SO4 saturated/2-propanol/sodium acetate 1 M (100/2.5/20). Plates were developed to 15 cm in each direction. Afterdrying the plates were autoradiographed with an intensifyingscreen up to 24 h. To quantify the adduct spots the origin ofthe plates holding a lot of activity was cut off. The remainingplate with the adduct spot was exposed for 3 h to a phosphorusimager screen and scanned afterward.

Reaction of NHMOR with DNA in Vitro. A series of 100µg of CT-DNA samples were dissolved in 100 µL of 0.1 M

472 Chem. Res. Toxicol., Vol. 15, No. 4, 2002 Loeppky et al.

phosphate buffer solution, pH 7.4, (The concentration of theDNA solution was determined by UV). NHMOR (1 × 10-5 mol)in a small amount of DMSO was added to each sample and theresulting mixtures were allowed to stand in the dark at 37 °Cfor 6, 12, 24 and 48 h, respectively. The NHMOR modified DNAwas precipitated with 0.1 volume (10 µL) of 3 M sodium acetate,pH 5, and 2.5 volume (250 µL) of 95% cold ethanol. Theprecipitation was kept at -20 °C overnight, then washed twicewith 80% cold ethanol and dried by vacuum centrifuge at roomtemperature. The modified DNA was hydrolyzed and labeledwith [32P]ATP. The 32P-labeling reaction mixture, 5 µL, wasseparated and quantitated on 2-D PEI-cellulose TLC as de-scribed above. In a second set of experiments, a batch of 100 µgof CT-DNA solutions (1 µg/µL) as above stood in the dark with1 µL of 1 × 10-6 mol and 1 × 10-9 mol of NHMOR (dissolved inDMSO) at 37 °C for 6, 12, 24, and 48 h. The NHMOR modifiedDNA was purified, hydrolyzed, labeled with 32P and quantitatedon HPLC as described above. All samples were prepared induplicate. Control DNA was incubated with 1 µL of DMSOunder the same conditions as each of the samples. In a thirdset of experiments, 100 µg of CT-DNA solutions (1 µg/µL) weremixed with 1 µL each of 1 × 10-9, 1 × 10-10, 1 × 10-11, 1 ×10-12, or 1 × 10-13 mol NHMOR (dissolved in DMSO) at 37 °Cin the dark, respectively. After 24 h, the NHMOR-modified DNAwas purified, hydrolyzed, labeled with 32P and separated andquantitated on TLC as described in the section above. Allreactions and analyses were run in duplicate.

32P-Postlabeling Procedure for Detection and Deter-mination of the OHEdG DNA Adduct. The DNA wasprocessed and hydrolyzed as described above (Stage 2). Aliquots(25 µg as determined by UV) of the digested DNA were injectedonto a Waters C-I8 delta-pak HPLC column (3.9 × 150 mm)and chromatographed: flow, 1 mL/min; UV, 254 nm; eluent A,20 mM ammonium formate pH 5.4; eluent B, acetonitrile,gradient 0 to 7% B in 30 min. Under these chromatographicconditions, the retention times of the 3′-OHEdG and 5′-OHEdGmononucleotides were similar; however, use of the 3′-nucleotidestandard and UV detection to find the retention time of thedesired fraction led to column contamination. As a result, the5′-32P-labeled OHEdG standard was prepared and used for thispurpose (radiochemical detection). Ten fractions of 1 mL eachwere collected, at the retention time of the standard. Fractions1-3, 4-7, and 8-10 were combined to make three differentfractions, the middle one of which was expected to contain the3′-OHEdG nucleotide. Each of these fractions was lyophilizedand redissolved in water (concentration, 1 µg DNA/µL). Eachfraction (5 µL) was then labeled with 8 µCi of [γ32P]ATP and 6units of T4-polynucleotide kinase in 3 µL of bicine-mixture, pH8.5 (100 mM bicine, 100 mM MgCl2, 10 mM spermidine, 100mM DTT), for 30 min at 37 °C. Then, 1 µL sodium acetate buffer(1 M, pH 4.5) and 2 µL of NP1 (2 µg) were added and the mixturewas incubated for another 30 min. The samples were diluted to50 µL with water and injected onto the HPLC column (C-18Zorbax 4.5 × 250 mm), and separated (flow, 1 mL/min; A, 50mM phosphate buffer with 2 mM tetrabutylammonium phos-phate pH 7), B, acetonitrile, 0-30 min 0 to 30% B, 30-45 min30% B, 45-55 min, 30% B, 55-70 min 0% B. Spiking a knownamount of adduct into the digested DNA before HPLC collectionresulted in a recovery of 15% compared to the direct labeling ofthe same quantity. Most of this loss is due to the HPLCcollection step. For TLC, the OHEdG adduct was separated onPEI plates. They were washed overnight with water, then 4 µLof the sample was applied and they were developed in D1 with1.25 M formic acid, then washed with water, dried and devel-oped in D2 with 2-propanol/water/NH3 (14/6/1).

Animal Experiments. Experiment Set 1: three 9 week old(200 g) male Wister rats were used for the study. One was thecontrol, one was given NDELA, and the other was givenNHMOR at a single dose of 0.8 mmol/kg body weight. Water(control) or aqueous solutions (500 µL) were given the rats bygavage. After 4 h, the animals were killed by cervical dislocation,and the liver was excised. The liver was washed with digestion

buffer (10 mM Tris-HCl, pH 7, 10 mM EDTA, pH 7, 100 mMNaCl) and then used immediately or stored in liquid nitrogenuntil needed. Experiment Set 2 and Set 3: the experiments weredone in the same way except that the rats were 7 weeks old.Dosing was the same but the nitrosamines used were N-nitrosodiethanolamine, N-nitrosomorpholine, N-nitrosoethyl-ethanolamine, and N-nitrosomethylethanolamine. ExperimentSet 4: this experiment used stored, frozen, liver tissue anddeuterated nitrosamines as we have described previously (26).

Isolation of DNA from Rat Liver. Rat liver (5 g) waswashed with buffer (10 mM Tris-HCl, 10 mM EDTA, 100 mMNaCl, pH 7), minced, and homogenized in a hand potter. To 10mL of the suspension, 1 mL of SDS 10% and 11 mg of proteinaseK were added and the sample was incubated for 2-3 h at 37°C. NaCl (0.1 volume, 5 M) was added and the protein wasextracted with an equal volume of phenol (saturated with 2 MTris buffer, pH 7.4)/chloroform/isoamyl alcohol (25/24/1). Thesample was centrifuged at 1700g for 10 min, the aqueous layerwas transferred to a fresh tube. The extraction was repeatedwith an equal volume of chloroform until no protein was visible.Ethanol (100%, -20 °C), in an amount equal to 2.5× the originalvolume, was carefully added to form a top layer. Then a glassrod was used by slowly moving it up and down between the twolayers to precipitate the DNA. The DNA was washed with 70%ethanol, dried, and redissolved in 1 mL of buffer (10 mM Tris,0.1 mM EDTA at pH 7). RNA was removed by using 20 µL ofRNAse A (10 mg/mL in 10 mM Tris-buffer at pH 7 heated to 70°C for 10 min, cooled) and 35 µL of RNAse T1 (1 unit/µL), andthe RNA was digested for 1 h at 37 °C. Then, the protein wasextracted as described before, 0.1 vol of 3 M sodium acetate wasadded, the DNA was precipitated with ethanol, washed, dried,and redissolved in 50 mM Tris-buffer, pH 7. The concentrationof the DNA was determined by UV at 260 nm.

Results

gdG 32P-Postlabeling Assay. The glyoxal-guanine(gG) adduct, (1,4,6,7-tetrahydro-6,7-dihydroxy-9H-imi-dazo[1,2-a]purin-9-one), as well as the correspondingderivative of guanosine, was first studied extensively byShapiro and his colleagues (19-21). It, or various deriva-tives, such as gdG or mononucleotides thereof, is easilyprepared by the reaction of the guanine compound withglyoxal by either Shapiro’s method (20) or a modificationof the procedure used by Chung and Hecht to preparegdG (24). Shapiro demonstrated that the gG adductstability decreases as the pH increases, but that it isreasonably stable between pH 4 and 6.5 (21). In prelimi-nary work, we found that more than 35% of the gGadduct is hydrolyzed to guanine under the conditions ofa typical DNA acid hydrolysis procedure (0.1 M HCl at65-70 °C, 1 h). Additional studies with gdG demon-strated that cleavage of the nucleosidic linkage alsooccurs under these same conditions. [After the develop-ment of the 32P-postlabeling assay reported here, we havemade a more careful study of the equilibrium and rateconstants for dissociation of the adduct as a function ofpH (22).]

To facilitate our work, we developed HPLC-UV meth-ods for the detection of all nucleosides and nucleotidesprepared. In preliminary experiments (data not given),these analytical methods were used to assess the time-dependent stability of various glyoxal-guanine adductsas a function of pH, temperature, and enzymatic protocol.Our results showed that we could recover the gdG adductas the nucleoside from either acid or enzymatic hydroly-ses of glyoxal modified DNA. The recoveries, however,were low, and the gdG nucleoside undergoes competitiveacid-catalyzed cleavage of either glyoxal from the adduct


or cleavage of the nucleosidic linkage. We attempted todevelop a number of derivatization procedures as we havedelineated elsewhere (22), but were not successful. Also,extraction into alcohols, the use of a boronate column,and the employment of microporous filters only decreasedour recovery. As a result we sought to carefully optimizeindividual steps and work toward the development of asuitable 32P-postlabeling procedure.

The gdG adduct is hydrolyzed to glyoxal and thecorresponding guanine fragment with increasing rate asthe pH is increased from 6.5 to 9 or greater. To securethe best conditions for the 32P-postlabeling of this adduct,we optimized each step in the procedure with respect tothe nature of the enzymes, their units, pH, time, thecomposition of the buffer, and several other variables asappropriate. Each step, DNA isolation, DNA hydrolysis,nuclease P1 enrichment of the adduct, kinase mediated5′-32P-labeling, and chromatography, was optimized. Thedetails of these optimization experiments are availablein the Supporting Information. As this methodology wasutilized in our laboratory, a refinement and improvementof the method became necessary as a result of differingnuclease P1 activity. The two optimized methods reportedhere for 32P-postlabeling of the gdG adduct are designatedas Stage 1, where we detected the adduct as the 3′,5′-dinucleotide, and Stage 2, in which case the adduct wasseparated and determined as its 5′-phosphonucleotide.The differences between these methods are summarizedin Table 1.

We determined relatively early in the investigationthat nuclease P1 hydrolyzed the unadducted 3′-nucle-otides to nucleosides much more rapidly than it catalyzedthe same transformation of the gdG adducted nucleotideat pH 7. Under optimized conditions the inclusion of thisstep led to a 5-fold increase in the gdG adduct comparedto normal nucleotide background. The most critical stepin the efficiency of the assay is the T4 kinase mediated32P-5′-labeling. The rate of this process is greatest at pH9.5, but the gdG adduct hydrolyzes rapidly at this pH.In Stage 1, the labeling was done at pH 7 for 2 h, butthe yields were low. This is the principal limitation ofthe 32P-postlabeling method for the gdG adduct.

As we used the methodology discussed above, it becameapparent that transformations were taking place assamples stood in the HPLC autosampler waiting foranalysis. We found that residual NP1 in the sample wascatalyzing the hydrolysis of the 3′,32P-5′-diphosphonucle-

tides to 32P-5′-monophosphates. At Stage 2, refinementof the methodology was made in three places. The DNAhydrolysis time was significantly shortened by combiningthe MN and SPD enzymes, and by allowing the incuba-tion to proceed for only 4 h. The kinase mediated labelingstep was improved by carrying out the process at pH 8for 30 min rather than the 2 h and lower pH used inStage 1. We also found during the course of theseexperiments that we achieved greater labeling efficiencyat lower concentrations of 3′-gdGP (<2.5 ng). Mostimportantly, a pH shift to 5, following the 32P-postlabel-ing, increases the NP1 activity and it efficiently catalyzesthe hydrolysis of the 3′-phosphate bond to cleanly givethe 5′-mononucleotides, which can be separated ef-ficiently by HPLC and TLC. HPLC conditions werechanged to maximize the separation of the 5′-phospho-nucletides and are described in the Experimental Meth-ods. These improvements were carried out using well-characterized standards. Spiking experiments permitteda determination of our sensitivity at 5 adducts in 107 innucleotides by HPLC with a recovery in the range of 50%.Samples were stable in the autosampler up to 10 h andthe residual enzymes were removed through the use ofa precolumn. The separation of 5′-phosphonucleotides,rather than the diphosphates also required changes inthe TLC methodology. Details are given in the Experi-mental Methods. Quantitation utilized a phosphorimageanalyzer and allowed a detection limit of 5 adducts in108 nucleotides.

Testing of gdG 32P-Postlabeling Assay Using theReaction of NHMOR and CT-DNA. As discussed inthe Introduction, Chung and Hecht (5, 24) and we haveindependently shown that NHMOR reacts with dG togive gdG adducts. We have demonstrated that thistransformation also occurs when other R-nitrosaminoal-dehydes are reacted with dG (5). To test whether ourassay could detect the formation of gdG adducts in DNAwhen it was reacted with NHMOR, we incubated amountsof NHMOR ranging from 10-6 to 10-13 mol/100 µg of CT-DNA at 37 °C for up to 48 h. The assays at developmentStage 1 or 2 were both able to detect the formation ofthe gdG adduct in the DNA. In Figure 1 the formationof the adduct is seen to increase with time, although therate of adduct formation is rather low. It increases withthe initial concentration of NHMOR, as anticipated butdoes not exceed 0.5 nmol of adduct. Figure 2 shows theeffect of [NHMOR] on the yield of gdG adduct in DNA.

Table 1. Experimental Parameters for the 32P-postlabeling Assays of the gdG and OHEdG DNA Adducts

parametergdG adduct

diphosphate (stage 1)gdG adduct

5′-monophosphate (stage 2)OHEdG adduct

5-monophosphate

DNA digestion (100 µg)a

MN 100 units, 12 h 100 units of MN and 0.4 unitsof SPD combined, 4 h

100 units of MN and 0.4 unitsof SPD combined, 4 h(SPD) 0.4 units, 10 h

enrichmentNP1 2-4 µg/10 µg of DNA, pH 7 6 µg, pH 7 N/A HPLC enrichmentNP1 time 2-12 h 2 h N/A

postlabelingT4 kinase 6 units/10 µg of DNA, pH 7 1 unit/1 µg of DNA, pH 8 1 unit/µg of DNA, pH 8.5T4 kinase time 2 h 30 min 30 min

NP1 conversion to 5′-monophosphatepH N/A 5 5

2 µg of NP1/ µg of DNAincubation time N/A 30 min 30 min

a Digestion time determined by inspection of HPLC of mixture (presence of polynucleotides). Longer times, up to 20 h were requiredfor highly modified samples.


At low concentrations (0.1-1 ng), the reaction is completeand the adduct level increases linearly with [NHMOR].At higher concentrations, saturation is reached becausethe limit of G sites is being reached. In these in vitroexperiments, using Stage 1 methodology, we could detectthe gdG adduct from 1 pmol of NHMOR/100 µg of DNA.Similar results were obtained using Stage 2 methodology;for example, at low concentrations of NHMOR (10-100pmol/100 µg of CT-DNA), we observed nearly quantitativeconversion of NHMOR to the gdG adduct.

32P-Postlabeling Assay for the O6-2-Hydroxyeth-yldeoxyguanosine Adduct. 2-Hydroxyethyl base DNAadducts arise from a number of different sources, andhave been characterized by several different methods(27-34), but no 32P-postlabeling method exists for the O6-2-hydroxyethyldeoxyguanosine adduct (OHEG). In con-trast to the gdG adduct, where the stability of the adductwas the major problem in the development of a goodassay, the principal difficulty related to OHEdG detectionwas the availability of an unambiguous standard of O6-2-hydroxyethyldeoxyguanosine-3′-phosphate (3′-OHEdG-MP) from which other standards could be made andutilized. We have solved this problem and the synthesis

is published elsewhere (23). A 1 ng sample of 3′-OHEdGMP was 32P-labeled using 8 µCi of [γ32P]ATP, atpH 8.5 in bicine buffer using T4 PNK for 30 min at 37°C. We determined, by comparing the hydrolysis of both3′-OHEdGMP and the diphosphate, with that obtainedfrom 3′-dGMP, that we could not observe any “enrich-ment” of the adduct using NP1. The adducted nucleotideunderwent 3′-dephosphorylation at rates similar to thoseof normal mononucleotides. The NP1 treatment of thelabeled diphosphate, at pH 5 (achieved by the additon ofacetate buffer to the mixture) led to its clean conversioninto 32P-5′-OHEdGMP, which was then used to developHPLC and TLC separation methods.

CT-DNA was treated with a mildly acidic solution ofethylene oxide to produce OHEdG adducted DNA. TheDNA was subjected to hydrolyses using the same condi-tions developed for the gdG adduct (Stage 2), but becauseenrichment of the adduct of interest could not be achievedthrough the use of NP1, we utilized HPLC to separatethe fractions containing 3′-OHEdGMP prior to labeling.Fractions located near the retention time of the 3′-OHEdG nucleotide were combined, lyophilized, redis-solved, 32P-labeled, and treated with NP1 as describedabove. After dilution, the samples were either separatedby RP-ion pair HPLC on a C-18 column, or applied toPEI-cellulose TLC plates and separated. The recovery,as determined by spiking with a known amount ofstandard prior to the DNA digestion, was 15%, with mostof the loss occurring during the HPLC fractionation.Although we did not “push” the method to its ultimatesensitivity, the lowest sample level detected was 0.3 µmol/mol of DNA. The data obtained (not shown) demonstratedthat the methodology was sensitive and, reasonablyefficient, as is demonstrated in the animal experimentsdescribed and discussed below.

Animal Experiments. As stated in the Introduction,the principal reason for the development of these 32P-postlabeling assays was to enable us to test varioushypotheses in whole animals regarding the mechanismof carcinogenic activation of NDELA and of structurallyrelated nitrosamines. Here we report on the use of thegdG and OHEdG 32P-postlabeling assays in four differentsets of animal experiments. Our first experiments weresolely directed at determining whether gdG adductformed in rat liver DNA after the administration ofNDELA or its metabolite, NHMOR. At the outset, we didnot know whether the assay was either sensitive enoughfor this purpose or if the adducts would survive the DNAisolation procedure. Because we had used male Wistarrats in DNA SSB work (26), we continued to use theseanimals and to use the same dosing regimen. MaleWistar rats (9 weeks, ∼200 g) were given 0.8 mmol ofNDELA or 0.34 mmol of NHMOR, in water by gavage orwater (control). The animals were sacrificed after 4 h,their livers excised, and DNA isolated from the livers asis described in the Experimental Methods. The DNAhydrolysis and postlableing was done as described abovefor Stage 1 development. Both HPLC and TLC wereutilized in the analysis, although adducts from bothNDELA and NHMOR were easily detected by HPLC(radiochemical detection). Typical data are shown inFigure 3. The spiking was done with a separate DNAsample from the same rat. Adduct levels were greaterfrom NDELA than they were from NHMOR (see Table2).

Figure 1. Time dependence of the gG adduct formation in calfthymus DNA (100 µg) as a function of [NHMOR] is shown.

Figure 2. Yields of gdG adduct produced from the reaction ofafter 24 h (37 °C) of reaction of CT-DNA (100 µg) with variousconcentrations of NHMOR are shown. (A) At low [NHMOR]adduct formation is linear, but at higher concentrations (B, seeinset) saturation is seen.


In the next two sets of experiments, we compared theadduct levels produced in rat liver DNA from theadministration of four different nitrosamines to maleWistar rats. The gdG adducts were detected as mono-nucleotides using the Stage 2 assay protocol. In thesecond experiment set, we detected only the gdG adductbut compared the values determined from HPLC withthose obtained from TLC. The latter two methods did notyield statistically different results and are presented inTable 2 for each of the nitrosamines.

In experiment set 3, we also detected the OHEdGadduct for each of the four nitrosamines. The sameanimals were used as in set 2. Typical HPLC data areshown in Figure 4 for the OHEdG adducts produced inrat liver from NDELA. We compared quantitation de-rived from addition of the standard after isolation fromthe liver, but before hydrolysis, with the use of anexternal standard, and obtained comparable data. Thedata given in Table 2 are derived from quantitation usingthe internal standard. All of the nitrosamines give boththe gdG and OHEdG adduct, but the relative amountsdepend on the nitrosamine structure.

In the fourth animal experiment set, we analyzed onlyfor the OHEdG adduct in rat liver samples which werederived from a previous study (26). These liver tissue

samples had been kept for 3-3.5 years in a freezer priorto being used in this work. Our aim in this study was toemploy deuterated isotopomers of NDELA and NHMORin conjunction with a DNA SSB assay to discover whichC-H bonds were being broken in the bioactivation ofthese compounds. In this case we sought to determinewhether the position of the deuterium substitution inNDELA affects the levels of OHEdG adducts arising fromthese nitrosamines. The gdG adduct was too unstable inthese samples for this determination. We analyzed theDNA from rats which had been given either H2O,NDELA, 1,1,1′1′-tetradeutero-N-nitrosodiethanolamine(R-D4-NDELA, the OH groups are in the 2, and 2′positions), or 2,2,2′2′-tetradeutero-N-nitrosodiethanola-mine (â-D4-NDELA). In each case, the DNA from morethan one rat was used for each isotopomer to improvethe certainty of our observations. The data are presentedin Table 3. Despite the fact that the adduct levels aredifferent in the two sets of animals, the data clearly showthe same pattern with respect to deuterium substitution.Compared to NDELA, the levels of OHEdG adduct aresignificantly diminished by R-deuteration and signifi-cantly enhanced by â-deuteration. Isotope effects arereported for each data set. A value greater than 1 meansthat deuteration at that position retards adduct forma-tion, while the converse is true for values less than 1.

Discussion32P-Postlabeling Methodology. We have made ex-

tensive use of the literature and the experience of othersin the development of the 32P-postlabeling assays for thegdG and OHEdG DNA adducts (18). There is nothing newin the DNA hydrolysis methodology. Similarly, NP1 hasbeen used for adduct enrichment and to cleave diphos-phates to monophosphates to permit better chromato-graphic separation and detection (25, 35). However, theuse of HPLC in both methods development and detectionfor 32P-postlabeling is less common (36, 37). HPLC wasvery valuable in our hands because of the many optimi-zation steps that were necessary for the sensitive detec-tion of the gdG adduct. Moreover, with the standards inhand, the use of the radiochemical detector in combina-tion with HPLC was sufficient for both the detection andfacile quantitation of adduct levels in all of our animal

Figure 3. Partial HPLC chromatograms, using radiochemical detection, showing the formation of the gdG adduct (as thediphosphonucleotide) in rat liver DNA following the administration of 0.8 mmol of NDELA. The left-most panel shows results fromthe control animal, and those at the center and right are data from the same dosed animal. The right-most panel shows the increaseof gdG adduct on spiking with the standard before DNA hydrolysis.

Table 2. gdG and OHEdG Rat-Liver DNA Adducts fromthe in Vivo Administration of â-Oxidized Nitrosaminesd

DNA adducts per 106 nucleotides

nitrosamine set gdG (HPLC) gdG (TLC) OHEdG

NDELA 1 5 NDa

2b 4.8 ( 0.2 6.5 ( 0.2 ND3c 4 ( 0.3 5.1 ( 0.1 10 ( 2

NHMOR 1 3 NDNMOR 2 2.6 ( 0.3 2.1 ( 0.3

3 1.6 ( 0.4 3.4 ( 0.1 4 ( 2NMELA 2 3.2 ( 0.3 4.8 ( 0.2 ND

3 3.7 ( 0.2 3.7 ( 0.2 25 ( 14NEELA 2 1.7 ( 0.1 1.4 ( 0.1 ND

3 0.9 ( 0.07 1.1 ( 0.05 11 ( 2a ND ) not determined. b In sets 2, and 3 the error represents

standard deviation of two to four replicate analyses of same DNAhydrolysate. c In set 3 the same animals were used as set 2, butthe adduct analyses were from a completely different workup ofthe liver tissue to give the DNA hydrolysates. d Control animalsshowed no gdG or OHEdG adducts.


experiments. The use of TLC in combination with HPLCallows the use of a second chromatographic method toconfirm adduct formation. Modern phosphorimage analy-sis, which is in common use in molecular biology labo-ratories, permits sensitive and reasonably accurate de-tection of adduct levels by TLC in combination with theuse of standards.

We have taken some pains to provide a description ofthe numerous optimization experiments that were usedin the development of the gdG 32P-postlabeling methodsin the Supporting Information. The OHEdG 32P-postla-beling assay is much easier to carry out once the standardnucleotide (23) is in hand. The low recovery of the gdGadduct is principally due to its low labeling efficiency.Significant hydrolytic cleavage of the glyoxal bishemiac-etal fragment from the guanine occurs at the optimallabeling pH of 8 for this adduct. This pH choice is acompromise between the increased labeling rate and therelatively rapid base-catalyzed rate of conversion of theadduct to dG. At pH 7, as was used in Stage 1 methodol-ogy, the labeling reaction is quite slow and residual NP1becomes more active in the hydrolysis of the required 3′-nucleotide to the nucleoside. NP1 enrichment, whichdepends on the more rapid dephosphorylation of normal3′nucleotides, compared to the adducted nucleotide, waspossible for gdG but not OHEdG. After the 32P-labeling

of either adduct, NP1-mediated hydrolysis of the 3′-phosphate from the 3′,5′-diphosphonucleotide could beaffected by shifting the pH to 5 by means of an acetatebuffer. Then the labeled adducts could be separated moreeasily and detected as 5′-32P-nucleotides.

In general, even in the hands of experienced andskillful workers, the gdG 32P-postlabeling assay is prob-lematic, laborious, and very time-consuming. It was notuncommon for us to encounter inconsistencies withcommercial biochemical preparations, which requiredeither reoptimization or did not lead to fruitful results.The extreme pH sensitivity of the adduct was the majorcause. Despite the fact that we were able to profitablyuse the gdG 32P-postlabeling assay in our mechanisticwork, we believe that it is not suitable for work incombination with epidemiology. Because of this, and thedesire to pursue additional mechanistic experiments, wehave been active in pursuing other sensitive methods forthe detection of the gdG adduct and will soon report onthe development and use of a sensitive LC/MSMS assayfor gdG, which is easier and much more reliable. Theseconcerns and limitations, however, do not apply to the32P-postlabeling assay for the OHEdG adduct. It isreliable and reasonably easy to use, although, like all 32P-postlabeling assays, requires the use of highly radioactivematerials and is somewhat laborious. We also had someproblems with HPLC column contamination and memoryeffects, resulting from the use of the same HPLC columnin the cleanup phase and the analytical phase of thedetermination. These problems could be circumvented.

In Vivo Adduct Formation from â-Oxidized Nit-rosamines. The most significant findings to emerge fromthis work are that all 2-hydroxyethynitrosamines, ex-amined so far, produce both gdG and OHEdG adductsin the DNA of rat liver (Table 2), and, in the case ofNDELA, OHEdG adduct formation is subject to deute-rium isotope effects. As is discussed below, these isotopeeffects parallel those observed for this compound in otherbioassays (12, 26). To our knowledge, this is the firstreport of the formation of gdG adducts in the DNA ofmammals following the in vivo administration of any

Figure 4. HPLC radiochromatogram of 32P-postlabeled DNA derived from rat liver following administration of 0.8 mmol of NDELA/kg is shown. The inset shows the control (rat given water), and the OHEdG adduct peaks from the DNA of the rat given NDELA,without the standard spike, and with the standard spike.

Table 3. Isotope Effects on the Production of OHEdGRat-Liver DNA Adducts from the in Vivo Administration

of Deuterated Isotopomers of NDELA

OHEdG adducts per106 nucleotidesa,b isotope effectc

nitrosamine set 4/7 set 11/7 set 4/7 set 11/7

NDELA 2.3 24 ( 15R-D4NDELA 0.62 2.2 ( 0.4 3.7 11â-D4NDELA 9 ( 3 58 ( 22 0.26 0.42

a Adduct analyses are from two different sets of animalsreceiving two different doses of NDELA and its deuteratedisotopomers. b Errors are standard deviations of means determinedfrom two to five replicate analyses of the same DNA sample. Incases where there is no error reported only a single sample gaveusable data. c Calculated by dividing the adduct level by thatproduced by NDELA in the given set.


carcinogen. While glyoxal has long been known to be amutagen, it is not carcinogenic. It is probable that glyoxalis rapidly cleared from the mammalian blood stream byoxidative metabolism in the liver. Here, however, we haveconclusively demonstrated the formation of glyoxal-DNAadducts in the form of gdG, and while we do not yet knowwhether the feeding of glyoxal results in DNA adducts,we must consider how gdG formation can occur in vivo.Our general hypothesis is that glyoxal arises by theoxidative processes shown in Scheme 1 at sites withinthe cell that facilitate its reaction with DNA. While it islikely that it is competitively detoxified, it is obvious thatits site of origin and its concentration are sufficient forreaction with DNA. In the previous paper in this issue(12), we show that R-hydroxylation of the 2-hydroxyethylside chain of NDELA or NEELA results in the formationof glycolaldehyde and a reactive diazonium ion (seeScheme 1, 8 f 9 f 14 + 15). We have demonstrated thatmicrosomes (probably P4502E1) catalyze the rapid oxida-tion of glycolaldehyde to glyoxal. We also have shown thatNDELA and NEELA are competitively oxidized to theircorresponding aldehydes (8 f 10) which are furtherR-hydroxylated in the same P450 system to give glyoxaland the diazonium ion 15. Because two reactive DNAbinding molecules arise from this process, we call thesecompounds bident (two-toothed) carcinogens (12).

While in vivo production of glyoxal adducts from thenitrosamines used in this work has not been reportedpreviously, all of them except NMOR, have previouslybeen reported to produce OHEdG adducts (34, 38, 39).The specific pathways leading to DNA adducts arediscussed below for each nitrosamine. From a generalperspective, microsome (presumably P450)-mediated R-hy-droxylation of the nitrosamine 8 leads to either 9 or 11,which decompose to give aldehydes and diazonium ions,15 or 17, respectively. In the case of 9, the aldehyde isglycolaldehyde 14, which is further oxidized to glyoxal13. We presume that gdG DNA adduct formation (Scheme2) occurs from glyoxal generated by either this pathwayor the one that involves â-oxidation to give 10, which isthen R-hydroxylated on the same chain to give 12. Theintermediate decomposes to glyoxal and the diazoniumion 15. In the case of NDELA, the side chains are thesame, and this ensures the production of glycolaldehydeand the 2-hydroxyethyldiazonium ion (17) by any of the

pathways. Diazonium ions are well-known to be potentalkylating agents, and 17 is the likely precursor ofOHEdG (Scheme 2).

The data of Table 2, experiment set 3, give a compari-son of gdG and OHEdG adduct levels as a function ofthe nitrosamine. In all cases, the levels of OHEdGadducts are greater than gdG adducts. We must becautious, however, in making conclusions regarding theseobservations, because of the difficulties associated withthe gdG assay and its relatively low recoveries. A bettercomparison, at this time, concerns the relative amountof OHEdG adduct as nitrosamine structure changes,which is NMELA . NEELA = NDELA > NMOR.OHEdG DNA adducts have previously been detected forNDELA, NMELA, and NEELA. Scherer et al. examinedthe ability of some of the same nitrosamines to produceOHEdG, which was detected using rabbit antibodies tothe deoxynucleoside and employed an immuno-slot-blotassay (34). Because of possible cross-reactivity, the lattermethod could not be considered definitive, but our datasignificantly corroborate its validity. The adduct levels(µmol/mol DNA) measured by Scherer et al. were NME-LA, 72.1; NEELA, 48.7; NDELA, 6.7; NHMOR, 2.0; andthe control, 1.7. We have not yet determined OHEdGlevels from NHMOR, but have data for NMOR. Both theability of the nitrosamines to produce OHEdG and theadduct levels generated are consistent with the previousfindings.

In general, nitrosamines without oxygen substituentsin their side chains are better substrates for P450s thantheir hydroxylated counterparts. It is reasonable toexpect, therefore, that NMELA and NEELA, with N-bound methyl, and ethyl substituents, respectively, will

Scheme 1a

a This scheme shows pathways for the microsome-mediated oxidation of various nitrosamines. Unstable compounds are shown in brackets.

Scheme 2


be preferentially hydroxylated at these groups ratherthan the oxidation of the 2-hydroxyethyl side chain ateither the R- or â-position. In other words, oxidation, asdepicted in the conversion 8 f 11, is preferred for thesecompounds and results in greater yields of the 2-hy-droxyethyldiazonium ion 17 and thereby greater amountsof OHEdG. NEELN is also known to be â-hydroxylatedat the ethyl group to give NDELA. This competitivetransformation will diminish the extent of OHEdG fromthis nitrosamine.

NDELA. R-Hydroxylation of NDELA at either sidechain will produce 17, which leads to OHEdG (Scheme2). As discussed in the previous paper in this issue andin related literature, NDELA and some other â-hydroxy-nitrosamines were not thought to be activated to proxi-mal carcinogens by R-hydroxylation (12). Our first indica-tions that there were problems with this suppositionarose from our comparison of in vivo DNA SSB producedby NDELA and its deuterated isotopomers, R-D4NDELA,and â-D4NDELA (26). R-Deuteration significantly inhib-ited DNA SSB, but â-deuteration did not. NDELA andâ-D4NDELA were cytotoxic to cells transfected withCYP2E1, but R-D4NDELA was not (26). We have dem-onstrated that R-deuteration significantly inhibits gly-colaldehyde 14 formation from NDELA by microsomesand increases the amount of NHMOR which is formed(12). â-Deuteration has the opposite effect. Here we haveshown (Table 3) that R-deuteration significantly de-creases the formation of OHEdG adducts in vivo, aprocess which requires R-hydroxylation, but â-deutera-tion significantly increases OHEdG production fromNDELA in vivo. These observations are all internallyconsistent and strongly support the hypothesis thatOHEdG adducts arise from NDELA by the P450-medi-ated R-hydroxylation.

Deuterium isotope effects result from the double massof deuterium compared to a proton. In classical chemicalor biochemical mechanism experiments, rates of CH vsCD bond breakage are measured and the ratio of rateconstants is expressed as the kinetic isotope effect. Thein vivo isotope effect experiments we have done, whichhave involved DNA SSB and OHEdG adduct formation,do not permit rate determinations because the experi-ment requires the sacrifice of the animal. The observationof an isotope effect in such an experiment requires some“luck” in choosing the sacrifice time of the animal. If thetransformation that gives rise to the DNA SSB andmodification is completely over, for all isotopomers, atthe sacrifice time, then no isotope effect can be observed.This could be true in the DNA SSB experiments for â-D4-NDELA, where we saw only a slight increase of DNA SSBfor this isotopomer (26). However, the sacrifice time (4h) we chose did allow us to see an isotope effect for R-D4-NDELA. Not only that, we assayed the same DNA forSSB and OHEdG formation, providing a strong connec-tion between the two. The fact that â-D4NDELA resultsin significantly more OHEdG than NDELA, undoubtedlyresults from enzymatic switching with respect to theposition of oxidation. The same phenomena were ob-served in our microsomal oxidation studies of NDELA(12). Deuteration at the â-position inhibits the conversionof NDELA to NHMOR and increases the conversion ofNDELA to glycolaldehyde and the 2-hydroxyethyl dia-zonium ion, the progenitor of OHEdG. Diazonium ions,because of their high reactivity, are rather indiscriminatewith respect to nucleophilic DNA sites that are alkylated.

In the case of guanine, we can also expect significantalkylation at N-7, although we have not yet made anyattempt to detect it. The charged purine generated bysuch a process leads to apurinic sites by depurination.These lesions are repaired by excision repair and thisprocess is likely a strong contributor to DNA SSB. BaseO-alkyl adducts are repaired principally by O-dealkyl-ases, a process which does not involve cleavage of theDNA phosphodiester backbone. There are no data on therepair of gdG adducts.

Several other mechanisms have been proposed for theactivation of NDELA and related 2-hydroxyethylnitro-samines. Most prominent among these is the sulfationhypothesis, which is shown in Scheme 3 (38-46). Thishypothesis foresees the formation of a sulfate ester 19 ofthe alcohol group in the nitrosamine 18. It has beenproposed that this cyclizes to generate the oxadiazoliniumion 20, which then alkylates DNA to incorporate the2-hydroxyethyl group 21 or the R group 22, as shown.We have shown in the case where R ) CH3 that the majorreaction product of 20 with dG or DNA oligomers resultsfrom nucleophilic attack at the starred carbon andincorporation of the entire nitrosamine fragment at N-7of dG (23). No hydroxyethylation is observed. We did seea very small amount of methylation. Furthermore wehave shown that oxadiazolinium ions are very reactivetoward thiols and give a completely different set ofreaction products including disulfides from oxidation ofthe SH (47). The process shown in Scheme 3 will not showa deuterium isotope effect for incorporation of the 2-hy-droxyethyl group into DNA because no C-H (C-D) bondsare broken. Our collective data cast large shadow of doubton the validity of the sulfation hypothesis as a mecha-nism for the activation of NDELA and other relatednitrosamines.

NHMOR, the cyclic hemiacetal form of the R-nitro-saminoaldehyde â-oxidation product and metabolite ofNDELA, reacts in vitro with DNA to deaminate the basesand form gdG adducts. This is also true of other R-nit-rosaminoaldehydes, which are highly reactive substances(11). We have proposed that these compounds are im-portant intermediates in the carcinogenic activation of2-hydroxyethylnitrosamines. Since, formation of the gly-oxal from these nitrosamines requires both â-, andR-oxidation, this hypothesis has validity. It is discussedmore directly below.

NMOR and NHMOR. Like NDELA, NMOR has beenshown to undergo both microsome mediated R- andâ-oxidation (2, 48-50). The â-oxidation product of NMORis NHMOR, which generates the gdG DNA adduct invitro. While in vivo experiments employing [14C]NMORdemonstrated the formation of at least seven DNAadducts, the only one identified was 7-(2-hydroxyethyl)-guanine (48). Prior to this work, this is the only known

Scheme 3


DNA adduct of NMOR. Here we show that NMOR givesrise to both gdG and OHEdG DNA adducts in vivo. Theformation of either OHEdG or 7-(2-hydroxyethyl)-guaninemust be preceded by â-hydroxylation to give NHMOR.The latter is a known biotransformation of NMOR. Theopen chain isomer of this hemiacetal is also a 2-hydroxy-ethylnitrosamine as well as being an R-nitrosaminoal-dehyde. P450-mediated oxidation of NHMOR by the path10 f 12 gives both glyoxal and the 2-hydroxyethydiazo-nium ion (here 15 ) 17). Nevertheless, the requirementof a prior oxidative transformation of NMOR to NHMOR,which competes with the R-hydroxylation of NMOR, andadditional oxidative pathways open to NHMOR, explainwhy the OHEdG level is relatively low in this case.Several oxidative pathways, other than one discussedabove, can lead from NMOR to gdG adducts, but at somepoint there must be an oxidation at one of the carbonsadjacent to what was the ring O of NMOR.

We have only a single observation for DNA adductformation by NHMOR, and clearly it must be investi-gated more extensively. The level of gdG produced fromNHMOR in vivo is less than that obtained from NDELA.We have known for some time that R-nitrosamino alde-hydes decompose in H2O or more rapidly in the presenceof amines to give glyoxal or its equivalents. In vitro,NHMOR forms gdG adducts with guanosine and itsnucleotides, and here we demonstrate small concentra-tions of NHMOR form gdG adducts in DNA, althoughthe process is rather slow. Because of the relatively slowrates of the reactions giving glyoxal equivalents fromNHMOR and other R-nitrosamino aldehydes, we havequestioned the importance of this transformation incarcinogenesis.

While the direct reaction of NHMOR with DNA to givegdG adducts requires more attention, we have shown inthe previous paper in this issue how relatively rapidmetabolic transformations of NDELA, NHMOR, andother 2-hydroxyethylnitrosamines can give glyoxal equiva-lents by means of both R- and â-oxidation of the 2-hy-droxyethyl side chain (Scheme 1). Scherer et al. failed toobserve OHEdG formation from NHMOR using theimuno-slot-blot assay (34). At relatively low doses, theonly ones tested, NHMOR is not carcinogenic to eitherA/J mice or F344 rats (51). However, NDELA and NMORare both carcinogenic. In the discussion above we haveascribed an important role to NHMOR in adduct forma-tion from both of these compounds. How can theseapparently contradictory observations be reconciled?N-Nitroso-2-hydroxyethylglycine (NHEG), the immediateoxidation product of NHMOR, is a major urinary me-tabolite of both NDELA and NMOR and is not carcino-genic. NHEG is also the in vivo oxidation product ofNHMOR (52). Like glyoxal, it is probable that NHMORis significantly detoxified by oxidation of the aldehydeto the acid in the liver and a significant quantity doesnot reach sites within the cell that would lead to itsR-hydroxylation. The microsomal oxidation of NDELA orNMOR to NHMOR place the latter compound at a sitewhere R-hydroxylation can occur and lead to DNA adductformation.

NMELA. Several mechanisms have been proposed forthe carcinogenic activation of NMELA. In particular,much attention has been given to the sulfation hypothesisdepicted in Scheme 3 and discussed above. NMELA givesrise to O6-methyl-, 7-methyl-, 7-(2-hydroxyethyl)-, and O6-hydroxyethylguanine adducts in DNA (39). Additionally,

we have found that gdG is a significant adduct from thisnitrosamine. Of the nitrosamines we have studied,NMELA produces the greatest levels of OHEdG, andthese adducts cannot arise from this nitrosamine by theroute postulated in Scheme 3. Although other mecha-nisms of activation may be operative, we believe that thisnitrosamine is principally activated by the pathwaysshown in Scheme 1. All of the adducts observed can arisethrough one of these routes. The pathways shown on theright side of Scheme 1 give rise to the methyldiazoniumion, from which methyl adducts arise, and glyoxal theprecursor to OHEdG. Thus, NMELA is also a bidentcarcinogen (12).

NEELA. Despite the fact that NEELA has beenextensively investigated as a model renal carcinogen inrodents (53), only two DNA modifications have previouslybeen reported for this compound, in vivo. Here we havefound both gdG and OHEdG DNA adducts from NEELA.OHEdG adducts were also detected using the immuno-slot-blot assay (34), and recently NEELA has been foundto give rise to 8-oxodeoxyguanosine (54). It has beenproposed to be activated by sulfation as shown in Scheme3 (34, 46). On the other hand, in the previous paper inthis issue we show the microsomal oxidation of NEELAoccurs by R-hydroxylation of both the ethyl, and the2-hydroxyethyl groups and also by oxidation at theâ-carbon of the 2-hydroxyethyl group to give the corre-sponding aldehyde (12). Glycolaldehyde, glyoxal, andacetaldehyde are products of these transformations.NDELA is also known to be a metabolite of NEELA. Thusthe activation of NEELA can be explained by the trans-formations shown in Scheme 1 and it can also beclassified as a bident carcinogen. The major urinarymetabolite of NEELA is N-ethyl-N-nitrosoglycine (55),showing that significant amount of â-oxidation is occur-ring. In vitro studies have demonstrated that N-ethyl-N-nitroso-2-aminoethanal, the aldehyde oxidation prod-uct, is an intermediate in this metabolic transformation(55). Like NDELA, much of NEELA is excreted in theurine unchanged. Thus, the bident nature of the activa-tion of these carcinogens may be very important in thecarcinogenic potency of these water soluble and easilyexcreted nitrosamines.

Conclusions and Summary

We have developed 32P-postlabeling assays for both thegdG and the OHEdG DNA adducts. The gdG adductstability is very pH sensitive. This property and adduct’sbis-hemiacetal structure present problems with the as-say’s use. Nevertheless we have fruitfully used the gdG,and the more rugged OHEdG, 32P-postlabeling assays inour mechanistic work to demonstrate the formation ofthese adducts in male Wistar rats administered equimo-lar doses of nitrosamines containing the 2-hydroxyethylchain (NDELA, NMELA, NEELA), as well as the cyclicâ-oxidized nitrosamine NMOR. This constitutes the firstreport of the formation of gdG DNA adducts in vivo. Thedemonstration that these nitrosamines also form OHEdGDNA adducts supports our contention that they are allbident carcinogens, which may provide an explanationfor their relatively potent carcinogenicity despite the factthat a very high percentage of the doses are known to beexcreted unchanged. We have demonstrated that theformation of OHEdG DNA adducts from NDELA issubject to deuterium isotope effects. R-Deuteration sig-


nificantly reduces OHEdG adduct formation while â-deu-teration results in a marked increase in the levels of thisadduct. These results are in accord with similar isotopeeffects on the production of DNA SSB, the microsomalmetabolism of NDELA and its isotopomers. All of thesedata strongly support our hypothesis that NDELA, andother 2-hydroxyethylnitrosamines are activated by R-hy-droxylation on the 2-hydroxyethyl chain. While the mostreactive electrophile generated is a diazonium ion, thisprocess also produces an aldehyde, either glycolaldehydeor glyoxal, which leads to the formation of gdG DNAadducts. These results significantly advance our under-standing of the way in which these environmentallyprevalent carcinogens are activated and provide pathsfor molecular epidemiological investigations of exposedpopulations.

Acknowledgment. We gratefully acknowledge thesupport of this research by the National Institute ofEnvironmental Health Sciences (NIH, DHHS) underGrant ES03953. We also express our appreciation to Dr.Anne Fuchs and to Maria Lorenz for their assistance withthe animal experiments with the deuterated nitro-samines, as well as the personnel of the University ofMissouri Laboratory of Animal Medicine for their as-sistance.

Supporting Information Available: Description of theoptimization procedures undertaken in the development of a32P-postlabeling method for the gdG DNA adduct. This materialis available free of charge via the Internet at http://pubs.acs.org.

References

(1) Airoldi, L., Bonfanti, M., Fanelli, R., Bove, B., Benfenati, E., andGariboldi, P. (1984) Identification of a nitrosamino aldehyde anda nitrosamino acid resulting from â-oxidation of N-nitrosodietha-nolamine. Chem.-Biol. Interact. 51, 103-113.

(2) Jarman, M., and Manson, D. (1986) The metabolism of N-nitrosomorpholine by rat liver microsomes and its oxidation bythe Fenton system. Carcinogenesis 7, 559-565.

(3) Hecht, S. S. (1984) N-Nitroso-2-hydroxymorpholine, a mutagenicmetabolite of N-nitrosodiethanolamine. Carcinogenesis (London)5, 1745-1747.

(4) Loeppky, R. N., Tomasik, W., Kovacs, D. A., Outram, J. R., andByington, K. H. (1984) Alternative bioactivation routes for â-hy-droxynitrosamines. Biochemical and chemical model studies. InN-Nitroso Compd: Occurrence, Biol. Eff. Relevance Hum. Cancer(Bartsch, H., O’Neill, I., and VanBorstal, Eds.) pp 429-436, IARC,Lyon.

(5) Loeppky, R. N., Tomasik, W., Denkel, E., and Eisenbrand, G.(1987) R-Nitrosaminoaldehydes: Highly reactive metabolites.IARC Sci. Publ. 84, 94-99.

(6) Loeppky, R. N., Tomasik, W., and Kerrick, B., E. (1987) Nitrosotransfer from alpha-nitrosamino aldehydes: Implications forcarcinogenesis. Carcinogenesis (London) 8, 941-946.

(7) Loeppky, R. N., Srinivasan, A., and Erb, E. (1992) Putativeproximate carcinogens derived from ethanolnitrosamines: Chemi-cal properties in Nitroso Compounds: Biological Mechanisms,Exposures and Cancer Etiology (Bartsch, H., and O’Neill, I., Eds.)pp 42a-42b, IARC, Lyon.

(8) Loeppky, R. N., Erb, E., and Srinivasan, A. (1993) The chemistryof putative intermediates in the bioactivation of â-oxidizednitrosamines. In The Chemistry and Biochemistry of Nitrosaminesand Other N-nitroso Compounds (Loeppky, R. N., and Michejda,C. J., Eds.) pp 334-336, American Chemical Society.

(9) Loeppky, R. N., Lee, M. P., and Mueller, S. (1993) DNA modifica-tion by R-nitrosamino aldehydes. In The Chemistry of DNAAdducts (Hemminki, K., and Bartsch, H., Eds.) pp 429-432,International Agency for Research on Cancer.

(10) Loeppky, R. N. (1999) The mechanism of bioactivation of N-nitrosodiethanolamine. Drug Metab. Rev. 31, 175-193.

(11) Park, M., and Loeppky, R. N. (2000) In Vitro DNA Deaminationby R-nitrosaminoaldehydes determined by GC/MS-SIM quanti-tation. Chem. Res. Toxicol. 13, 72-81.

(12) Loeppky, R. N., and Goelzer, P. (2001) The microsome mediatedoxidation of N-nitrosodiethanolamine (NDELA), a bident carcino-gen. Chem. Res. Toxicol. 15, 457-469.

(13) Farrelly, J. G., Thomas, B. J., and Lijinsky, W. (1987) Metabolismand cellular interactions of N-nitrosodiethanolamine. IARC Sci.Publ. 84, 87-90.

(14) Farrelly, J. G., Stewart, M. L., and Lijinsky, W. (1984) Themetabolism of nitrosodi-n-propylamine, nitrosodiallylamine andnitrosodiethanolamine. Carcinogenesis (London) 5, 1015-1019.

(15) Gupta, R. C., Randerath, E., and Randerath, K. (1976) Animproved separation procedure for nucleoside monophosphateson polyethyleneimine-(PEI-)cellulose thin layers. Nucleic AcidsRes. 3, 2915-2921.

(16) Reddy, M. V., Gupta, R. C., and Randerath, K. (1981) Phosphorus-32 base analysis of DNA. Anal. Biochem. 117, 271-279.

(17) Randerath, K., Reddy, M. V., and Gupta, R. C. (1981) Phosphorus-32 labeling test for DNA damage. Proc. Natl. Acad. Sci. U.S.A.78, 6126-6129.

(18) Phillips, D. H. (1997) Detection of DNA modifications by the 32P-postlabelling assay. Mutat. Res. 378, 1-12.

(19) Shapiro, R., Sodum, R. S., Everett, D. W., and Kundu, S. K. (1986)Reactions of nucleosides with glyoxal and acrolein. IARC Sci.Publ. 70, 165-173.

(20) Shapiro, R., and Hachmann, J. (1966) The reaction of guaninederivatives with 1,2-dicarbonyl compounds. Biochemistry 5, 2799-2807.

(21) Shapiro, R., Cohen, B. I., Shiuey, S., and Maurer, H. (1969) Onthe reactions of guanine with glyoxal, pyruvaldehyde, and kethox-al, and the structure of the acylguanines. A new synthesis of N2-alkylguanines. Biochemistry 8, 238-245.

(22) Loeppky, R. N., Cui, W., Goelzer, P., Park, M., and Ye, Q. (1999)Glyoxal-guanine DNA adducts: Detection, stability and formationfrom nitrosamines. In Cyclic DNA Adducts II (Singer, B., andBartsch, H., Eds.) pp 155-168, IARC, Lyon.

(23) Loeppky, R. N., Yu, L., Gu, F., and Ye, Q. (1996) DNA GuanineAdducts from 3-methyl-1,2,3-oxadiazolinium ions. J. Am. Chem.Soc. 118, 10995-11005.

(24) Chung, F. L., and Hecht, S. S. (1985) Formation of the cyclic 1,N2-glyoxal-deoxyguanosine adduct upon reaction of N-nitroso-2-hydroxymorpholine with deoxyguanosine. Carcinogenesis (Lon-don) 6, 1671-1673.

(25) Reddy, M. V., and Randerath, K. (1986) Nuclease P1-mediatedenhancement of sensitivity of phosphorus-32-postlabeling test forstructurally diverse DNA adducts. Carcinogenesis 7, 1543-1551.

(26) Loeppky, R. N., Fuchs, A., Janzowski, C., Humberd, C., Goelzer,P., Schneider, H., and Eisenbrand, G. (1998) Probing the mech-anism of the carcinogenic activation of N-nitrosodiethanolaminewith deuterium isotope effects: In vivo induction of DNA singlestrand breaks and related in vitro assays. Chem. Res. Toxicol.11, 1556-1566.

(27) Liao, P.-C., Li, C.-M., Hung, C.-W., Chen, S.-H., Zhao, C., Kumar,R., and Hemminki, K. (2001) Quantitative detection of N7-(2-hydroxyethyl)guanine adducts in DNA using high-performanceliquid chromatography/electrospray ionization tandem mass spec-trometry: Measurement of 7-methyl- and 7-(2-hydroxyethyl)-guanine DNA adducts in white blood cells of smokers and non-smokers. J. Mass Spectrom. 36, 336-343.

(28) Ranasinghe, A., Scheller, N., Wu, K. Y., Upton, P. B., andSwenberg, J. A. (1998) Application of gas chromatography/electroncapture negative chemical ionization high-resolution mass spec-trometry for analysis of DNA and protein adducts. Chem. Res.Toxicol. 11, 520-526.

(29) Leclercq, L., Laurent, C., and Pauw, E. D. (1997) High-performance liquid chromatography/electrospray mass spectrom-etry for the analysis of modified bases in DNA: 7-(2-hydroxyethyl)-guanine, the major ethylene oxide-DNA adduct. Anal. Chem. 69,1952-1955.

(30) Kumar, R., Hemminki, K., and Zhao, C. (1996) Separation of7-methyl- and 7-(2-hydroxyethyl)-guanine adducts in human DNAsamples using a combination of TLC and HPLC: Measurementof 7-methyl- and 7-(2-hydroxyethyl)guanine DNA adducts in whiteblood cells of smokers and non-smokers. Carcinogenesis 17, 485-492.

(31) Allam, K., Saha, M., and Giese, R. W. (1990) Preparation ofelectrophoric derivatives of N7-(2-hydroxyethyl)guanine, an eth-ylene oxide-DNA adduct. J. Chromatogr. 499, 571-578.

(32) Ludeke, B., Schubert, M., Yamada, Y., Lijinsky, W., and Kleihues,P. (1991) DNA hydroxyethylation by hydroxyethylnitrosoureas inrelation to their organ specific carcinogenicity in rats. Chem.-Biol.Interact. 79, 207-216.

(33) Ludeke, B. I., and Kleihues, P. (1988) Formation and persistenceof O6-(2-hydroxyethyl)-2′-deoxyguanosine in DNA of various rat


tissues following a single dose of N-nitroso-N-(2-hydroxylethyl)-urea. An immuno-slot-blot study. Carcinogenesis (London) 9, 147-151.

(34) Scherer, G., Ludeke, B., Kleihues, P., Loeppky, R. N., andEisenbrand, G. (1991) Mutagenicity, DNA damage and DNAadduct formation by N-nitroso-2-hydroxyalkylamine and corre-sponding aldehydes. IARC Sci. Publ. 105, 339-342.

(35) Gorelick, N. J., and Wogan, G. N. (1989) Fluoranthene-DNAadducts: identification and quantification by an HPLC-32P-postlabeling method. Carcinogenesis 10, 1567-1577.

(36) Dunn, B. P., and San, R. H. C. (1988) HPLC enrichment ofhydrophobic DNA-carcinogen adducts for enhanced sensitivity ofphosphorus-32-postlabeling analysis. Carcinogenesis 9, 1055-1060.

(37) Gorelick, N. J. (1993) Application of HPLC in the phosphorus-32-postlabeling assay. Mutat. Res. 288, 5-18.

(38) Kroeger-Koepke, M. B., Koepke, S. R., Hernandez, L., andMichejda, C. J. (1992) Activation of a â-hydroxyalkylnitrosamineto alkylating agents: Evidence for the involvement of a sul-fotransferase. Cancer Res. 52, 3300-3305.

(39) Koepke, S. T. R., Kroeger-Koepke, M. B., Born, W., Thomas, B.J., Alvord, W. G., and Michejda, C. J. (1988) Alkylation of DNAin rats by N-nitrosomethyl-(2-hydroxyethyl)amine: Dose resposeand persistence of the alkylated lesions in vivo. Cancer Res. 48,1537-1542.

(40) Koepke, S. R., Kupper, R., and Michejda, C. J. (1979) Unusuallyfacile solvolysis of primary tosylates. A case for participation bythe N-nitroso group. J. Org. Chem. 44, 2718-2722.

(41) Michejda, C. J., and Koepke, S. R. (1978) Powerful anchimericeffect of the N-nitroso group. J. Am. Chem. Soc. 100, 1959-1960.

(42) Koepke, S. R., Cresia, D. R., Knutsen, G. L., and Michejda, C. J.(1988) Carcinogenicity of hydroxyalkylnitrosamines in F344rats: Contrasting behavior of beta- and gamma-hydroxyalkylnitrosamines. Cancer Res. 48, 1533-1536.

(43) Michejda, C. J., and Koepke, M. B. K. (1994) Carcinogen activationby sulfate conjugate formation. Adv. Pharmacol. (San Diego) 27,331-363.

(44) Michejda, C. J., Koepke, S. B., Kroeger, M. B., and Hernandez,L. (1994) Activation of â-hydroxyalkylnitrosamines. Evidence forinvolvement of a sulfotransferase. In Nitrosamines and relatedN-nitroso compounds: Chemistry and Biochemistry (Loeppky, R.N., and Michejda, C. J., Eds.) pp 195-210, Am. Chem. Soc.,Washington, DC.

(45) Denkel, E., Sterzel, G., and Eisenbrand, G. (1987) In-vivo activa-tion of N-nitrosodiethanolamine and other N-nitroso-2-hydroxy-

alkylamines by alcohol dehydrogenase and sulfotransferase. IARCSci. Publ. 84, 83-86.

(46) Sterzel, W., and Eisenbrand, G. (1986) N-Nitrosodiethanolamineis activated in the rat to an ultimate genotoxic metabolite bysulfotransferase. J. Cancer Res. Clin. Oncol. 111, 20-24.

(47) Loeppky, R. N., and Srinivasan, A. (1995) Thiol oxidation by 1,2,3-oxadiazolinium ions, presumed carcinogens. Chem. Res. Toxicol.8, 817-820.

(48) Manson, D., Cox, P. J., and Jarman, M. (1978) Metabolism ofN-nitrosomorpholine by the rat in vivo and by rat liver mi-crosomes and its oxidation by the Fenton system. Chem.-Biol.Interact. 20, 341-354.

(49) Hecht, S. S., and Young, R. (1981) Metabolic R-hydroxylation ofN-nitrosomorpholine and 3,3,5,5-tetradeutero-N-nitrosomorpho-line in the F344 rat. Cancer Res. 41, 5039-5043.

(50) Hecht, S. S., Young, R., Rivenson, A., and Hoffmann, D. (1982)On the metabolic activation of N-nitrosomorpholine and N′-nitrosonornicotine: effects of deuterium substitution. IARC Sci.Publ. 41, 499-507.

(51) Hecht, S. S., Lijinsky, W., Kovatch, R. M., Chung, F. L., andSaavedra, J. E. (1989) Comparative tumorigenicity of N-nitroso-2-hydroxymorpholine, N-nitrosodiethanolamine, and N-nitroso-morpholine in A/J mice and F344 rats. Carcinogenesis (London)10, 1475-1477.

(52) Airoldi, L., Bonfanti, M., Benfenati, E., Tavecchia, P., and Fanelli,R. (1983) Identification of an acidic metabolite of N-nitrosodi-ethanolamine isolated from rat urine. Biomed. Mass Spectrom.10, 334-337.

(53) Konishi, N., Hayashi, I., Nakaoka, S., Kitahori, Y., Tsuzuki, T.,Matsuda, H., Kitamura, M., Hiasa, Y., and Ohshima, M. (1994)Rodent species and strain difference in kidney and lung pathologyinduced by N-ethyl-N-hydroxyethylnitrosamine. J. Toxicol. Pathol.7, 67-71.

(54) Sato, M., Kitahori, Y., Nakagawa, Y., Konishi, N., Cho, M., andHiasa, Y. (1998) Formation of 8-hydroxydeoxyguanosine in ratkidney DNA after administration of N-ethyl-N-hydroxyethylni-trosamine. Cancer Lett. 124, 111-118.

(55) Hiasa, Y., Kitahori, Y., Kitamura, M., Nishioka, H., Yane, K.,Fukumoto, M., Ohshima, M., Nakaoka, S., and Nishii, S. (1991)Relationships between serum thyroid stimulating hormone levelsand development of thyroid tumors in rats treated with N-bis-(2-hydroxypropyl)nitrosamine. Carcinogenesis (London) 12, 873-877.

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DNA Adducts from N- Nitrosodiethanolamine and Related β-Oxidized Nitrosamines in Vivo: 32 P-Postlabeling Methods for Glyoxal- and O 6 -Hydroxyethyldeoxyguanosine Adducts