Identification of the major tamoxifen–DNA adducts in rat liver by mass spectroscopy

Heli Rajaniemi3, Ilpo Rasanen1, Pertti Koivisto2, Kimmo Peltonen2 and Kari Hemminki

Department of Biosciences at Novum, Karolinska Institute, Novum 141 57 Huddinge, Sweden,
1 University of Helsinki, Department of Forensic Medicine, FIN-00014 Helsinki and
2 Institute of Occupational Health, Topeliuksenkatu 41 aA, FIN-00250 Helsinki, Finland


    Abstract
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
We present here the first mass spectroscopic (MS) identification of the main tamoxifen-induced DNA adducts in rat liver. The two main adducts were isolated by high-performance liquid chromatography (HPLC) and identified by MS, MS–MS and ultraviolet spectroscopy. Adduct 1 was the N-desmethyltamoxifen–deoxyguanosine adduct in which the {alpha}-position of the metabolite N-desmethyltamoxifen is linked covalently to the amino group of deoxyguanosine. Adduct 2 was confirmed to be the trans isomer of {alpha}-(N2-deoxyguanosinyl)tamoxifen, as previously suggested by co-chromatography.

Abbreviations: dGMP, deoxyguanosine 5'-monophosphate; HPLC, high-performance liquid chromatography; MS, mass spectroscopy; NP1, nuclease P1.


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Tamoxifen [(Z)-1-{4-[2-(dimethylamino)ethoxy] phenyl}-1,2-diphenyl-1-butene], a non-steroidal anti-oestrogen, is widely used in the treatment of breast cancer. Endometrial cancer is a recognized side-effect of tamoxifen treatment in breast cancer patients (1). In rats tamoxifen is a potent liver carcinogen (1,2). It forms DNA adducts in several rat organs, the highest levels being found in liver (1,3,4). DNA damage in the form of tamoxifen-induced DNA adducts has also been shown in human endometrial and leukocyte samples using the 32P-post-labelling technique coupled to high-performance liquid chromatography (HPLC) and flow-through radioactivity detection (5,6).

Tamoxifen requires metabolic activation by liver microsomes before binding to DNA. It is extensively metabolized to N-demethylated and hydroxylated derivatives by hepatic cytochrome P450 enzymes in humans and various other mammalian species (7,8). One probable intermediate in the activation of tamoxifen to liver carcinogen is {alpha}-hydroxytamoxifen (911). It has been identified as a metabolite in rat hepatocytes (11), human liver (12) and plasma of patients treated with tamoxifen (12). However, the {alpha}-hydroxyl group requires either protonation or conjugation to provide a good leaving group, the loss of which would generate a highly reactive carbocation (9,13,14). This conjugation is suggested to be catalysed by liver phase II enzymes, such as sulphotransferases or UDP-glucuronosyltransferases (15). Formation of tamoxifen–DNA adducts has been shown to be inhibited by sulphotransferase inhibitors (16) and recently, {alpha}-hydroxytamoxifen was shown to be a substrate of hydroxysteroid (alcohol) sulphotransferase resulting in tamoxifen DNA adducts (17,18).

{alpha}-Acetoxytamoxifen, a model compound for the reactive metabolite of tamoxifen, has been shown in vitro to produce a DNA adduct that is identical to one of the two main adduct peaks in tamoxifen-treated rat liver samples (19). This product was identified as (E)-{alpha}-(N2-deoxyguanosinyl)tamoxifen in which the {alpha} position of tamoxifen was linked covalently to the exocyclic amino group of deoxyguanosine (13). The reaction of {alpha}-sulphate tamoxifen with DNA produced the same major product (20). The minor products of reaction of DNA with {alpha}-acetoxytamoxifen were identified as diastereoisomers of dG–N2–tamoxifen and as tamoxifen–deoxyadenosine adduct, attached through the amino group of adenine (21). The earlier identifications were based on co-chromatography of the 32P-post-labelled samples with the standards, whereas direct structural assignment of the DNA adducts in vivo has not been presented.

The main in vivo tamoxifen DNA adduct has remained unidentified. As it was not present in DNA reacted with {alpha}-acetoxytamoxifen, it is likely to be produced by another metabolite of tamoxifen. We show here by mass spectroscopic analysis of rat liver DNA, that the major adduct is an N-desmethyltamoxifen–deoxyguanosine product in which the {alpha} position of the metabolite N-desmethyltamoxifen is linked covalently to the amino group of deoxyguanosine.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Animal treatment and DNA isolation
Tamoxifen citrate was synthesized and supplied by Orion Corporation Medipolar (Oulu, Finland). Adult female Sprague–Dawley rats were given tamoxifen citrate by gavage, daily 45 mg/kg for 2 weeks. The vehicle was a propylene glycol solution in water. DNA from rat liver samples was isolated as previously described (6). In brief, the rat liver tissue (0.6–1.2 g) was homogenized in 5 ml of 1 mM MgCl2, 10 mM Tris–HCl (pH 8.0), after which the suspension was treated with 0.5% Triton X-100. The nuclei were collected by centrifugation at 3000 r.p.m. for 10 min at 2°C. DNA was purified with RNase A and RNase T1 treatment followed by proteinase K digestion and extraction with phenol and chloroform–isoamyl alcohol. DNA was precipitated and washed with ethanol. This procedure ensured efficient removal of RNA, as evident in the HPLC analysis of the hydrolysed DNA.

Hydrolysis of DNA and labelling
The monophosphate modification of the post-labelling procedure was described by Randerath et al. (22) and was applied as described (19). DNA (5 µg) was hydrolysed with a mixture of nuclease P1 (0.2 µg/µg DNA) and prostatic acid phosphatase (20 mU/µg DNA) at pH 5 at 37°C for 45 min. The modified nucleotides were converted to 5'-32P-labelled dinucleotides in the labelling mixture (2 µl) containing 2.4 U T4 polynucleotide kinase and 2.3 pmol ATP (7 µCi [{gamma}-32P]ATP, 3000 Ci/mmol). The reaction was carried out at pH 9.6. Snake venom phosphodiesterase (0.8 mU/µg DNA) was added for 30 min at 37°C to yield 32P-labelled monophosphate adducts.

For preparation of material for mass spectroscopy (MS) DNA (100 or 50 µg) was hydrolysed to nucleosides by incubating with nuclease P1 (0.1 µg/µg DNA) at pH 5 for 1 h, with alkaline phosphatase (Sigma, Poole, UK; ammonium sulphate suspension, 60 mU/µg DNA) at pH 9 for 1 h and with snake venom phosphodiesterase I (Sigma type II; 0.8 mU/µg DNA, pH 9) for 45 min.

HPLC analysis of adducts
For HPLC analysis the labelled mixtures were diluted to 10 µl and nucleoside hydrolysates to 200 µl with water and injected into the Beckman HPLC system Gold. The volume of the sample loop was 20 or 250 µl, respectively. A pre-column filter and a Phenomenex Kromasil C18 (2x50 mm, particle size 5 µm) pre-column were installed in front of the analytical column. Normal nucleosides as well as the labelled normal nucleotides, inorganic phosphate and residual ATP in the labelled samples were separated from adducts in pre-columns and diverted to waste using a four-port switching valve (Valco Instruments, Houston, TX). Adducted nucleotides/nucleosides were separated in an analytical column, Phenomenex Prodigy ODS (2x250 mm, particle size 5 µm). Radioactivity was measured on-line with a Beckman 171 Radioisotope detector. The size of the Teflon sample loop in the flow cell was 50 µl and it was folded into a scintillation tube containing scintillation liquid (Ready Safe; Beckman, High Wycombe, UK). Absorbance at 254 nm was measured, and scanning at wavelengths from 200 to 350 nm was performed on-line using a Beckman 168 diode array detector.

Separations were carried out at ambient temperature using a binary gradient with ammonium formate buffer (0.5 M ammonium formate, 20 mM phosphoric acid, pH 4.6) and acetonitrile. Nucleosides and 32P-labelled adducted nucleoside monophosphates were analysed using a gradient that had initial conditions of 20% acetonitrile for 10 min. During the first 9.5 min the sample was eluted only through the pre-column. After 10 min the proportion of acetonitrile was increased linearly to 29% in 25 min, maintained isocratic for 15 min and after that further increased to 43% in 35 min and to 100% in 5 min. Flow rate was 0.2 ml/min. Each sample was analysed at least twice. The total recovery of the 32P-post-labelling method was ~70%.

Mass spectroscopy
Mass spectra were recorded with a Perkin–Elmer Sciex (Norwalk, CT) API 365 bench-top Triple-Quad mass spectrometer with the Turbo Ion Spray ionization probe. Experiments were performed on a positive ion mode with the orifice voltage set at 30 V. The resolution was set on both quadrupoles at 0.2 mass units in all experiments. The mass spectrum of the products was recorded and the mass spectrometer was then programmed to transmit the protonated molecular ions [M+H] (m/z 637 and m/z 623, respectively) through the first quadrupole and following collision-induced fragmentation in the Q2 collision cell operating with nitrogen as the collision gas and a collision energy of 30 eV. Full scan product ion spectra were recorded in Q3. Mass spectra were collected in a continuous flow mode by connecting a Harward infusion pump directly to the Turbo Ion Spray probe. The adducts were infused in methanol:water (1:1 v/v) at a flow rate of 8 µl/min.


    Results
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 Materials and methods
 Results
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Exposure of female rats to daily doses of tamoxifen for 2 weeks resulted in the formation of two main adducts and several minor adducts in liver DNA (Figure 1Go); the mean adduct level was 20/106 nucleotides. A sample of {alpha}-acetoxytamoxifen-reacted DNA, prepared by Osborne et al. (13) was also digested, labelled and analysed using the same method, and the main peak was shown to co-chromatograph with peak 2 (Figure 1Go) in the current and many other chromatographic systems. A large amount (1.2 mg) of rat liver DNA was digested to adducted nucleosides and analysed by a similar HPLC method using absorbance detection at 254 nm. The HPLC profiles of pre-purified DNA hydrolysates from tamoxifen-treated and control rat liver samples are shown in Figure 2Go. The two main adducts, 1 and 2 in rat liver sample, had a similar UV spectrum that was indistinguishable from the spectrum of (E)-{alpha}-(N2-deoxyguanosinyl)tamoxifen, with absorbance maxima at 250 nm (pH 5), which suggests that both were N2-deoxyguanosine adducts.



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Fig. 1. HPLC-radioactivity detector analysis of tamoxifen-treated rat liver DNA (one-third of the sample of 5 µg was injected). The major adduct peaks are numbered. Peak number 2 is identical to dGMP derivative of (E)-{alpha}-(N2-deoxyguanosinyl)tamoxifen.

 


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Fig. 2. HPLC-absorbance detection (254 nm) analysis of rat liver DNA (1.2 mg) from a tamoxifen-treated animal (A) and rat liver DNA (1.0 mg) from control animal (B). Peaks 1 and 2 correspond to the monophosphate derivatives shown in Figure 1Go. Peak 2 is identical to (E)-{alpha}-(N2-deoxyguanosinyl)tamoxifen.

 
Adduct peaks were collected in water/acetonitrile mixture for mass spectroscopic analysis. Tamoxifen standard (E)-{alpha}-(N2-deoxyguanosinyl)tamoxifen gave a clear protonated molecular ion at m/z 637 (Figure 3CGo). The collision-induced fragmentation of the protonated molecular ion resulted in ions at m/z 521, 370, 344, 178 and 72 and 71. The fragment at m/z 521 is a result of a loss of the sugar moiety from the tamoxifen–deoxyguanosine adduct (Figure 4Go). The fragment at m/z 370 corresponds to tamoxifen moiety. The fragment at m/z 344 is formed as a result of the bond breakage between the {alpha}-carbon and the unsaturated carbon in tamoxifen. The fragment at m/z 178 gives structural information regarding the binding site of tamoxifen in deoxyguanosine. The fragment may be formed by a bond breakage between the {alpha}-carbon and the unsaturated carbon in tamoxifen moiety followed by intramolecular hydrogen rearrangement and a subsequent ring closure to N1 position of guanine. This fragment indicates that tamoxifen is bound to the N2 position of guanine as described by Osborne et al. (13). The signal at m/z 72 is the fragment of the aliphatic ether chain of tamoxifen, i.e. –O–C{alpha}'–Cß'–N(CH3)2. Collision-induced fragmentation of adduct 2 resulted in all the same fragments as in the standard and they are present at almost equal intensity (Figure 3BGo), thus this adduct was assigned as (E)-{alpha}-(N2-deoxyguanosinyl)tamoxifen.





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Fig. 3. The collision-induced spectrum of adduct 1 (A), adduct 2 (B) and the standard (E)-{alpha}-(N2-deoxyguanosinyl)tamoxifen (C). Adducts 1 and 2 were isolated from tamoxifen-treated rat liver DNA.

 


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Fig. 4. The chemical structure of the tamoxifen–deoxyguanosine adducts in rat liver DNA. The collision-induced fragmentation is also included.

 
The collision-induced spectrum of adduct 1 is also similar except that all the fragments appear at 14 mass units lower than in the standard and adduct 2 (Figure 3AGo). This is a clear demonstration of desmethylation in one of the two alternative positions present in the tamoxifen molecule, one attached to the {alpha}-carbon of tamoxifen and the other attached to the nitrogen atom (Figure 4Go). Adduct 1 displays a fragment at m/z 58 instead of 72, which demonstrates that one of the two methyl groups attached to the nitrogen on the aliphatic ether chain was removed. An additional proof of this interpretation is that ion at m/z 178 is present in the spectrum of adduct 1. The formation of this fragment is not possible without the methyl group at the {alpha}-carbon of tamoxifen molecule. Thus adduct 1 was assigned as N2-deoxyguanosinyl-N-desmethyltamoxifen.


    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
We showed here by direct analysis of rat liver samples that the main tamoxifen adduct in this organ was the N-desmethyltamoxifen–deoxyguanosine adduct in which the {alpha} position of the metabolite N-desmethyltamoxifen is linked covalently to the amino group of deoxyguanosine. We also confirmed that the other main fraction was {alpha}-(N2-deoxyguanosinyl)tamoxifen, which was not shown before by direct mass-spectroscopic analysis of biological samples. As additional, independent evidence for this assignment we have shown elsewhere that adducts 1 and 2 only differ in the number of methyl groups present (19). When methylated with dimethyl sulphate the two adducts form an identical compound because of the methylation at the amino side chain of tamoxifen. Additionally, the pK determinations of the two adducts in a two-phase system support this assignment (data not shown).

Tamoxifen is extensively metabolized in humans and rodents to N-desmethyltamoxifen (7,8). Also, the metabolite {alpha}-hydroxy-N-desmethyltamoxifen has been identified in microsomal incubates with tamoxifen (23) as well as in plasma of human breast cancer patients (12). In rats, after long-term treatment with tamoxifen (6 or 15 months of continuous administration of 10 mg/kg orally), the ratio of tamoxifen:N-desmethyltamoxifen has been reported to be 1:0.5–1 in serum and 1:1.3–2.3 in liver (24), which is consistent with the present results showing somewhat higher levels of the N-demethylated adduct. In human breast cancer patients the administration of 40 mg/day tamoxifen for 15–940 days resulted in mean plasma concentrations of 300 ng/ml tamoxifen and 462 ng/ml N-desmethyltamoxifen (25), indicating that activation of N-desmethyltamoxifen to DNA-reactive species can also play an important role in humans.

Since tamoxifen requires metabolic activation, the identification of these reactive metabolites is essential in risk estimation. As the extent of DNA adduct formation in target tissues among structural analogues of tamoxifen may be related to their carcinogenic potency, understanding their DNA binding properties is important in the search for safe drugs. However, the identity of human tamoxifen adducts remains to be established.


    Acknowledgments
 
We thank Dr Eero Mäntylä for tamoxifen-treated rat liver samples and Dr Martin Osborne and Dr David Phillips for providing the tamoxifen DNA standard. This study was supported by the Orion-Farmos Company.


    Notes
 
3 To whom correspondence should be addressed Email: heli.rajaniemi{at}cnt.ki.se Back


    References
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received June 30, 1998; revised September 22, 1998; accepted October 9, 1998.