Carcinogenesis vol.19 no.5 pp.939–943, 1998
SHORT COMMUNICATION
2,3-Epoxy-4-hydroxynonanal, a potential lipid peroxidation
product for etheno adduct formation, is not a substrate of human
epoxide hydrolase
Hauh-Jyun Candy Chen1,3,5, Frank J.Gonzalez2,
Magang Shou2,4 and Fung-Lung Chung1,5
1Division
of Carcinogenesis and Molecular Epidemiology, American Health
Foundation, 1 Dana Road, Valhalla, NY 10595, 2Laboratory of Molecular
Carcinogenesis, National Cancer Institute, Bethesda, MD 20892, USA
3Present
address: Department of Chemistry, National Chung Cheng
University, Ming-Hsiung, Chia-Yi, Taiwan, ROC
4Present
address: Department of Drug Metabolism, Merck Research
Laboratories, West Point, PA, USA
5To
whom correspondence should be addressed (either of these authors)
Our previous studies have shown that 2,3-epoxy-4-hydroxynonanal, a reactive epoxy aldehyde capable of forming
etheno adducts with DNA bases, is mutagenic and tumorigenic (Carcinogenesis, 14, 2073). The epoxy aldehyde can be
generated from trans-4-hydroxy-2-nonenal, a lipid peroxidation product of ω-6 polyunsaturated fatty acids, by
autoxidation or by incubation with fatty acid hydroperoxides or hydrogen peroxides (Chem. Res. Toxicol., 9, 306).
These are plausible in vivo pathways for the formation
of 2,3-epoxy-4-hydroxynonanal. The possibility that 2,3epoxy-4-hydroxynonanal is a tumorigen of endogenous
origin is suggested by recent observations that etheno bases
are detected as background DNA lesions in untreated
rodents and humans. A metabolic pathway critical for
detoxification of 2,3-epoxy-4-hydroxynonanal involves the
ring-opening by epoxide hydrolase, which abolishes its
ability to form cyclic etheno DNA adducts. In this study,
we examined whether 2,3-epoxy-4-hydroxynonanal is a
substrate of cDNA expressed human epoxide hydrolase.
Human epoxide hydrolase was expressed in TK– 143 cells
(thymidine kinase-deficient human embryoblast) infected
with recombinant vaccinia virus encoding human epoxide
hydrolase cDNA. Controls consisted of the cells infected
with vaccinia virus in the absence of human epoxide
hydrolase cDNA. No hydrolysis occurred when [2,3-3H]2,3epoxy-4-hydroxynonanal was incubated at 37°C for 30 min
at pH 7.4 with cells expressing human epoxide hydrolase,
as indicated by the presence of a pair of radioactive
peaks in reversed-phase HPLC chromatography, which
comigrated with the UV standards of the two diastereomers
of the epoxy aldehyde. The identity of these compounds
as the intact epoxy aldehyde was further supported by
derivatization to the 2,4-dinitrophenylhydrazones followed
by reversed phase HPLC analysis. Similar results were
observed with the control cells or with the heat deactivated
human epoxide hydrolase. The epoxide hydrolase activity
in the expressed cells was demonstrated by their ability to
convert benzo[a]pyrene-4,5-dihydroepoxide to benzo[a]pyrene-trans-4,5-dihydrodiol under the same conditions.
*Abbreviations: B[a]P-4,5-diol, benzo[a]pyrene-trans-4,5-dihydrodiol;
B[a]P-4,5-epoxide, benzo[a]pyrene-4,5-dihydroepoxide; cDNA, complementary DNA; DNP, 2,4-dinitrophenylhydrazine; EH, 2,3-epoxy-4-hydroxynonanal; h-EHase, human epoxide hydrolase; HNE, trans-4-hydroxy-2-nonenal;
PUFAs, polyunsaturated fatty acids.
© Oxford University Press
These results clearly indicate that 2,3-epoxy-4-hydroxynonanal is not a substrate of human epoxide hydrolase, and,
thus strengthen its possible endogenous role in the formation of promutagenic exocyclic etheno adducts in vivo.
Trans-4-hydroxy-2-nonenal (HNE*) is an α,β-unsaturated
aldehyde produced by peroxidation of ω-6 polyunsaturated
fatty acids (PUFAs) (1–4). It is believed that HNE contributes
to the cytotoxicity associated with peroxidation of liver microsomal lipids (1). HNE is not only cytotoxic, it also possesses
growth inhibiting, genotoxic and chemotactic activity, and
may actually play a role in atherogenesis (5). Since lipid
peroxidation has been implicated in tumorigenesis, it is postulated that HNE may also be involved in this process. Our
previous studies showed that HNE is readily converted to
its epoxide, 2,3-epoxy-4-hydroxynonanal (EH), by tert-butyl
hydroperoxide or by biological oxidants such as hydrogen
peroxide and fatty acid hydroperoxides (6,7). EH can also be
formed by autoxidation of HNE (8). EH is reactive toward
nucleic acid bases yielding etheno adducts (Scheme 1) (6,9).
Consistent with its reactivity, EH, unlike HNE, was shown to
be mutagenic in Salmonella tester strains TA100 and TA104
and tumorigenic in CD-1 mice (10). These results suggest that
EH could be an activated species of HNE. Recent studies
showed that a number of exocyclic DNA adducts, including
etheno adducts, are present as background lesions in tissue
DNA of humans and untreated rodents (11–14). These observations raised the possibility that EH is generated by lipid
peroxidation which subsequently forms cyclic etheno adducts
with tissue DNA (15).
Scheme 1. Formation and reactions of EH.
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H.-J.C.Chen et al.
Fig. 1. HPLC chromatograms obtained from analysis of the incubation mixture of [2,3-3H]EH with (a) h-EHase; (b) boiled h-EHase; (c) wild type h-EHase at
37°C for 30 min. Two peaks eluted at 29 and 32 min represent the diastereomers of EH. The small radioactive peak eluted at 3 min, which accounted for a
very small percentage of radioactivity, was not identified.
Fig. 2. Kinetics of hydrolysis of [2,3-3H]EH and B[a]P-4,5-epoxide in the presence of h-EHase expressed in human TK–143 cells. A 50 µl reaction mixture
containing potassium phosphate buffer, h-EHase and B[a]P-4,5-epoxide in 1.0 µl ethanol or [3H]EH with final concentration 0.38 mM was incubated at 37°C
for 10, 30, 60, or 120 min. The reaction was quenched by the addition of cold ethanol (50 µl) and centrifuged at 13 000 g for 10 min. The supernatant was
analyzed by reversed phase HPLC consisting of a Supelcosil LC-18-DB 5 µ column (Bellefonte, PA) which was eluted with H2O and CH3CN gradient (0–5
min, 5% CH3CN; 5–50% CH3CN at a flow rate of 1.0 ml/min) and a UV detector set at 190 nm. For each time point, triplicates were performed.
While the potential role of EH in the formation of endogenous etheno DNA lesions is recognized, little is known about
its metabolism. Possible pathways by which EH can be
metabolized are the conjugation with glutathione and oxidation
and reduction of aldehyde (16,17). Another pathway involves
the ring-opening of the epoxide by epoxide hydrolase. Since
the epoxide is the critical functional group for the covalent
binding to DNA bases, ring-opening by epoxide hydrolase is
believed to be an important detoxification step (18,19). As
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examples, the addition of human epoxide hydrolase to cell
cultures reduced the binding of reactive metabolites of
benzo[a]pyrene to DNA (20) and the purified epoxide hydrolase
from rat liver hydrolyzed 2-chlorooxirane, the epoxide of vinyl
chloride and, consequently, inhibited the formation of N2,3ethenoguanine (21). To better assess the role of EH in endogenous etheno adduct formation, it is important to know
whether EH serves as a substrate for epoxide hydrolase. In
this study, we examined the metabolic fate of EH upon
2,3-Epoxy-4-hydroxynonanal and human epoxide hydrolase
Fig. 3. HPLC chromatograms showing comigration of radioactive peak with the UV standard of EH-DNP derivative after DNP derivatization of the
incubation mixture with (a) h-EHase; (b) boiled h-EHase; (c) wild type h-EHase. Portions of the supernatant from incubation of [3H]EH at 37°C for 30 min
was derivatized with 2% DNP in methanol and 1.0 N HCl at 37°C for 1 h. The reaction mixture was then analyzed by a radioflow HPLC system consisting
of a supekosil LC-18-DB 5µ column which was eluted with an H2O and CH3CN gradient (0–5 min, 5% CH3CN; 5–60 min, 5–100% CH3CN) at a flow rate
of 1.0 ml/min and a UV detector set at 370 nm.
Fig. 4. Percentage of B[a]P-4,5-epoxide remaining in the reaction mixture
after reaction with h-EHase, boiled h-EHase, wild type h-EHase and
h-EHase 1 2.0 mM cyclohexene oxide at 37°C for 30 min.
incubation with microsomes obtained from human embryoblastoid cells containing cDNA-expressed human epoxide
hydrolase (h-EHase). The cDNA is for one of human microsomal epoxide hydrolases which acts on a wide range of
alkene and arene oxide (22).
The cDNA encoding h-EHase was removed from pUC9 by
partial digestion with Eco R1. The cDNA was made bluntended by treatment with DNA polymerase Klenow fragment
and inserted into the Sma I site of pSC11 vector (23).
Recombinant vaccinia viruses were constructed as described
previously (24). Human TK–143 (thymidine kinase-deficient
human embryoblast) cells were grown to .90% confluence
on 175-cm2 plastic flasks and infected with either wild type
or the recombinant vaccinia viruses (25). Microsomal fractions
of TK–143 cells were prepared by sonication of cells followed
by two different centrifugations as previously reported (26).
Protein concentrations of microsomes for both h-EHase and
control cells were 11.7 and 7.9 mg/ml as determined by the
method of Lowry et al. (27). A 50 µl reaction mixture
containing potassium phosphate buffer (100 mM, pH 7.4),
h-EHase (1.0 mg/ml protein), and [2,3-3H]EH (sp. act.
0.83mCi/mmol; 0.0165 µCi; 0.38 mM) was incubated at 37°C
for 10, 30, 60, or 120 min. [2,3-3H]EH was obtained by adding
100 mM H2O2 in 100 mM potassium phosphate buffer (pH 7.4)
to [2,3-3H]HNE and allowing it to stand at room temperature
for 23 h. The parent aldehyde [2,3-3H]HNE (sp. act. 375 mCi/
mmol) was obtained by acid hydrolysis of [2,3-3H]HNE
diethyl acetal (customer synthesized by Chemsyn Science
Laboratories, Lenexa, KS). The reaction was quenched by the
addition of cold ethanol followed by centrifugation. The
supernatant was spiked with EH as a UV marker and analyzed
by a radioflow-HPLC system. For the control experiments,
incubations were carried out under the same conditions, except
that cells infected with the wild type virus or boiled for 10 min
were used.
Figure 1 shows typical HPLC chromatograms obtained from
analysis of the reaction mixture incubated for 30 min. Two
radioactive peaks with retention time at 29 and 32 min, which
comigrated with the UV markers of EH isomers (1:1 ratio),
were observed in either the h-EHase expressing cells or the
controls. The amount of radioactivity of these EH isomers did
not decrease significantly in the presence of h-EHase, indicating
no significant extent of hydrolysis of EH had occurred during
incubation. Similar results were obtained with incubation
extended to 2 h, as shown in Figure 2. An ~10% decrease in
EH was observed after 2 h incubation. This decrease is likely
941
H.-J.C.Chen et al.
due to spontaneous hydrolysis of EH, since a similar loss of
EH was seen in the control experiments.
To further support that the comigrating radioactive peaks
were indeed the intact EH isomers, portions of the incubation
supernatant were derivatized with 2% 2,4-dinitrophenylhydrazine (DNP) in 100 µl methanol and 40 µl 1.0 N HCl (37°C
for 1 h). Under these conditions, EH is converted to the
previously characterized 3,4-dihydroxy-2-methoxynonanal
(29,4’-dinitrophenyl)hydrazone (7). The hydrazone derivative
of EH isomers, which collapsed into a single peak under the
HPLC conditions used, was again comigrating with the UV
standard of the DNP derivative (Figure 3). These results
showed that EH remained mostly unchanged in the presence
of h-Ehase. The activity of h-EHase in the expressed TK–143
cells was demonstrated by the conversion of benzo[a]pyrene4,5-dihydroepoxide (B[a]P-4,5-epoxide) to benzo[a]pyrenetrans-4,5-dihydrodiol (B[a]P-4,5-diol). Only 9.8% of B[a]P4,5-epoxide remained when it was incubated with h-EHase at
37°C for 30 min at pH 7.4., whereas 90.9% remained with
the wild type and 74.4% with the boiled cells. Furthermore,
2.0 mM of cyclohexene oxide, a known epoxide hydrolase
inhibitor, inhibited the hydrolysis of B[a]P-4,5-epoxide
(Figure 4).
The distinct kinetics of h-EHase toward the hydrolysis of
EH and B[a]P-4,5-epoxide illustrates the specific structural
requirements of this enzyme for its substrates. The substrate
specificity and species and tissue differences in epoxide
hydrolase activity have been described (28). The presence of
multiple forms of microsomal epoxide hydrolase in liver
microsomes of rats and humans has been suggested and each
appears to have a significant substrate specificity (29). It is
plausible that other forms of epoxide hydrolase in microsomes
or cytosols may be able to hydrolyze EH. Species differences
were noted in the hydrolysis of 2-cyanoethylene oxide, the
epoxide metabolite of acrylnitrile, as the epoxide hydrolase in
human liver microsomes was clearly active, whereas, that in
mouse and rat was not (30,31). Furthermore, the genetic
variations in quantities of epoxide hydrolase could play an
important role in the susceptibility to chemical carcinogenesis
(32). It was reported that a variant epoxide hydrolase allele is
significantly over-represented in patients with hepatocellular
carcinoma and that higher levels of aflatoxin B1-albumin
adducts were found (33). However, the recent biochemical
study showed that h-EHase did not hydrolyze aflatoxin B1 8,9epoxide, the ultimate metabolite of the genotoxic human liver
carcinogen aflatoxin B1 (36). Like exposure to exogenous
carcinogens, the metabolism by h-EHase of potential endogenous epoxides such as EH could have important implications
in carcinogenesis.
The present study showed that h-EHase did not hydrolyze
EH. These results provide a new line of support for the
potential of EH in the formation of endogenous etheno DNA
adducts in humans. Similarly, other homologous epoxides of
enal generated by lipid peroxidation may also contribute to
their total formation (15). Although the reaction of EH or
other epoxy aldehydes with DNA and RNA bases yielded both
unsubstituted and alkyl substituted etheno adducts (6,9,35,36),
so far, only the unsubstituted etheno adducts have been
detected in rodents and humans (12). The detection of the 1,2dihydroxyheptyl substituted etheno adducts in tissue DNA
would provide the strongest evidence for the involvement of
EH in the formation of etheno adducts in vivo. Since these
adducts can only be formed from HNE, if detected in vivo,
942
they could be useful specific markers of lipid peroxidation of
ω-6 PUFAs.
Acknowledgement
This work was supported by grant CA 43159 from National Institutes of
Health. Part of this study was presented at the 87th AACR meeting,
Washington, D.C., April, 1996.
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Received on November 26, 1997; revised on January 20, 1998; accepted on
January 23, 1998
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