From the Department of Biochemistry and Molecular Biology, University College London, Gower Street, London WC1E 6BT, United Kingdom
Received for publication, October 30, 2002, and in revised form, December 11, 2002
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Metabolites of vinyl chloride react with cytosine
in DNA to form 3,N4-ethenocytosine.
Recent studies suggest that ethenocytosine is repaired by the base
excision repair pathway with the ethenobase being removed by
thymine-DNA glycosylase. Here single turnover kinetics have been used
to compare the excision of ethenocytosine by thymine-DNA glycosylase
with the excision of thymine. The effect of flanking DNA sequence on
the excision of ethenocytosine was also investigated. The 34-bp
duplexes studied here fall into three categories. Ethenocytosine
base-paired with guanine within a CpG site (i.e.
CpG· It has been known for nearly 30 years that exposure to vinyl
chloride can cause cancer in humans (1). Vinyl chloride is metabolized
by cytochrome P450 2E1 to form chloroethylene oxide (2) which can
rearrange spontaneously to give chloroacetaldehyde (3). Both these
metabolites react in vitro with DNA to form ethenoadducts of
adenine, guanine, and cytosine (Fig. 1).
Three of these four possible ethenobases have been detected in animals exposed to vinyl chloride (reviewed in Ref. 4). Ethenobases have also
been found in the DNA of rats and humans not exposed to vinyl chloride.
These are probably formed endogenously by the reaction of lipid
peroxidation products with DNA (5). The ethenobases cause mutations by
misincorporating during DNA replication, and there is evidence that
these mutations are responsible for the carcinogenicity of vinyl
chloride and related chemicals (6).
C-DNA) was by far the best substrate having
a specificity constant (k2/Kd) of 25.1 × 106 M
1 s
1. The next
best substrates were DNA duplexes containing
TpG·
C, GpG·
C, and
CpG·T. These had specificity constants 45-130 times smaller than
CpG·
C-DNA. The worst substrates were DNA
duplexes containing ApG·
C and TpG·T,
which had specificity constants, respectively, 1,600 and 7,400 times
lower than CpG·
C-DNA. DNA containing
ethenocytosine was bound much more tightly than DNA containing a
G·T mismatch. This is probably because thymine-DNA glycosylase
can flip out ethenocytosine from a G·
C base
pair more easily than it can flip out thymine from a G·T mismatch. Because thymine-DNA glycosylase has a larger specificity constant for the removal of ethenocytosine, it has been suggested its
primary purpose is to deal with ethenocytosine. However, these results
showing that thymine-DNA glycosylase has a strong sequence preference
for CpG sites in the excision of both thymine and ethenocytosine suggest that the main role of thymine-DNA glycosylase in
vivo is the removal of thymine produced by deamination of
5-methylcytosine at CpG sites.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (7K):
[in a new window]
Fig. 1.
Formation of
3,N4-ethenocytosine. The vinyl
chloride metabolites chloroethylene oxide (i) and
chloroacetaldehyde (ii) react with cytosine bases in DNA to
form 3,N4-ethenocytosine. Similar reactions
occur at the 1,N2- and
N2,3-positions of guanine and at the
1,N6-position of adenine (4).
Extracts from human cells remove all four ethenoadducts from DNA (7).
Because they are released from the DNA as the ethenobases, it is likely
that they are repaired by the base excision repair pathway. The base
excision repair pathway (reviewed in Refs. 8 and 9) involves initial
removal of the damaged base by a DNA glycosylase. In the short-patch
repair pathway the resultant abasic site is cut by an apurinic
endonuclease, probably human apurinic endonuclease 1 (APEX1; also known as HAP1,
APE1, or Ref-1). The single nucleotide gap is filled by DNA polymerase
which also removes the abasic sugar-phosphate. Finally, the
phosphate backbone is restored by a DNA ligase. Thymine-DNA glycosylase
(TDG) is the enzyme believed to repair G·T mismatches arising
from spontaneous deamination of 5-methylcytosine (10). Support for this
comes from the observation that TDG excises thymine from G·T
mismatches at sites of cytosine methylation (i.e. CpG) much
more efficiently than from other DNA sequences (11-14). Recently, two
groups (15, 16) independently found that TDG can also remove
ethenocytosine from DNA.
A common feature of many DNA glycosylases is their tight binding to the
abasic sites that they produce (14, 17-21). Failure to consider this
product inhibition led to the so-called single strand-selective monofunctional
uracil-DNA glycosylase (SMUG1) originally being
incorrectly designated as a single-stranded DNA glycosylase (18, 22).
Product binding is particularly strong for TDG and is so tight that
in vitro each glycosylase molecule can only process one
G·T mismatch (14). The next enzyme in the repair pathway,
APEX, can relieve this product inhibition and increase the turnover of
TDG by releasing the glycosylase from the abasic site (23). The
mechanism of this release is not yet known. The initial experiments
showing that TDG can remove ethenocytosine from DNA did not consider
product inhibition of the glycosylase (15, 16), and so we decided to
study TDG excision of ethenocytosine using single turnover experiments.
Because the base pair flanking the mismatched guanine has a remarkably
strong effect on the rate of thymine excision from G·T
mismatches, the effect of the flanking base pair on the excision of
ethenocytosine was also measured.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Synthesis and Purification of Oligodeoxynucleotides--
34-Base
deoxynucleotides of the general sequence AGC TTG GCT GCA GGX
GGA CGG ATC CCC GGG AAT T (where X is A, C, G,
or T) were annealed with the complementary strand that had either
thymine or ethenocytosine opposite the underlined G. The nomenclature used in the following text is XpG·T-DNA for the
guanine·thymine mismatch containing duplexes and
XpG·C-DNA for the guanine·ethenocytosine
containing duplexes. X is either A, C, G, or T and refers to
the base 5' to the mismatched guanine. Normal base-containing
oligodeoxynucleotides were synthesized on an Applied Biosystems 391 DNA
synthesizer and purified as described previously (24).
Deoxyethenocytidine was synthesized from deoxycytidine using the
protocol of Zhang et al. (25). The correct structure of the
synthesized deoxyethenocytidine was confirmed by UV and 1H
NMR spectroscopy, which gave data agreeing with that published previously (25). The 5'-dimethoxytrityl-protected deoxyethenocytidine phosphoramidite was prepared by standard procedures (26).
Oligodeoxynucleotides containing ethenocytosine opposite the underlined
G were synthesized using the standard DNA synthesis protocol except
that the coupling time for the ethenocytidine phosphoramidite was
increased to 2 min. The coupling yield, as judged by trityl cation
release, was the same for the ethenocytidine phosphoramidite as for the
unmodified phosphoramidites. Full-length oligodeoxynucleotides were
separated from failure sequences using Nensorb columns (DuPont),
further purified by ion exchange chromatography at pH 12 (27) using a Mono-Q column (Amersham Biosciences), and finally desalted.
Oligodeoxynucleotides prepared in this way were >95% pure as judged
by 260 nm absorbance of their ion exchange chromatography traces.
Samples of the oligodeoxynucleotides were digested with nuclease P1
plus alkaline phosphatase and analyzed by reverse-phase high pressure
liquid chromatography. A fifth peak eluted after the four natural
deoxynucleosides. This had the absorbance expected for a single
deoxyethenocytidine and co-eluted with standard deoxyethenocytidine,
thus confirming the presence of ethenocytosine in the oligodeoxynucleotides.
Enzymes-- Thymine-DNA glycosylase was expressed in Escherichia coli from the pT7-hTDG plasmid as described previously (28) and was purified in three chromatographic steps (14). The concentrations of the dilute TDG samples used for the kinetic experiments were determined accurately using a bandshift assay. For this, five different amounts of TDG were incubated for 30 min with 32P-labeled CpG·T-DNA in binding buffer (25 mM Hepes (pH 7.6), 50 mM KCl, 1 mM EDTA, 2 mM dithiothreitol, 0.5 mg/ml bovine serum albumin, and 4% Ficoll 400). Electrophoresis was performed as described (14). The amount of DNA bound was plotted against the volume of TDG added, and the concentration of the glycosylase was determined from linear regression analysis of this plot.
Human APEX was a gift from Dr. I. Hickson and Dr. D. Rothwell (Oxford University, UK).
Glycosylase Assays-- 34-Base pair DNA duplexes were 5'-labeled with 32P in the strand containing the mismatched thymine or ethenocytosine. They were reacted at room temperature with TDG in reaction buffer (25 mM Hepes (pH 7.6), 2.5 mM MgCl2, 2 mM dithiothreitol, 0.2 mM EDTA, 0.5 mg/ml bovine serum albumin) containing either 50 or 140 mM KCl. Because of the sensitivity of the TDG reaction to salt, consistent results can only be attained if extra care is taken to ensure that the concentrations of MgCl2 and KCl are kept constant (i.e. salt present in the protein and DNA stock solutions must be allowed for). For the inhibition experiments, labeled DNA and inhibitor were mixed first, and TDG was then added to start the reaction. Samples from the G·T reactions were removed at various times and quenched by addition of NaOH and EDTA to a concentration of 0.1 M and 10 mM, respectively. Abasic sites produced by the glycosylase were cleaved by heating at 90 °C for 30 min. Because of the greater lability of deoxyethenocytidine to alkali, milder conditions must be used for the ethenocytosine oligonucleotides. Ethenocytosine samples were heated at 90 °C for 30 min in 30 mM piperidine, 10 mM EDTA. This completely cleaved abasic sites, whereas cleavage of the parent DNA containing ethenocytosine was kept to less than 2%. The cleaved DNA was separated from full-length, unreacted DNA by perfusion chromatography using a 2.1 × 30 mm Q HyperD (Biosepra) anion exchange column as described (14). Radiolabeled DNA was detected by Cerenkov counting using a Berthold LB 506 C-1 monitor and was quantified by integration of the peaks.
For the determination of kinetic constants, reactions were carried out
using equimolar concentrations of TDG and DNA at seven different
concentrations as follows: 0.1, 0.2, 0.5, 1, 2, 5, and 10 times the
approximate value of Kd. The reaction was analyzed
using the reaction model given in Scheme
I that assumes no product is released by the glycosylase during the
time that the reaction is monitored. Data for all seven concentrations
were fitted simultaneously using the differential equation solving program Berkeley Madonna (version 8.0.1; www.berkeleymadonna.com). The program was used to obtain the best fit of the theoretical lines to
all of the experimental data by varying the values of k1, k1, and
k2.
|
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Single Turnover Excision of Ethenocytosine by TDG Is Slower Than
the Excision of Thymine--
The single turnover excision of
ethenocytosine from CpG·C-DNA was compared with
the excision of thymine from CpG·T-DNA (Fig. 2). Under the conditions used for this
experiment (10 nM DNA + 10 nM TDG in buffer
containing 2.5 mM MgCl2 and 50 mM
KCl), the initial rate of excision for ethenocytosine (0.72 nM min
1) was more than four times slower than
the initial rate of excision for thymine (3.2 nM
min
1). Decreasing the concentration of enzyme and DNA had
no effect upon the rate of excision of ethenocytosine showing that in
Fig. 2 all the enzyme is bound to the
CpG·
C-DNA. The value of Kd
must therefore be much less than 10 nM. Also, the initial
rate for CpG·
C-DNA in Fig. 2 must be
Vmax, giving a value of 0.0012 s
1
for k2. In contrast, reducing the concentration
of TDG and CpG·T-DNA significantly reduced the rate of thymine
excision (data not shown). Thus, under the conditions of Fig. 2, not
all the glycosylase is bound to the CpG·T-DNA. Therefore, the
initial rate for CpG·T-DNA in Fig. 2 is less than
Vmax, and so k2 for
CpG·T-DNA must be greater than 0.0053 s
1.
|
Determination of Kd and k2 for TDG Excision of
Ethenocytosine and Thymine--
An attempt to get accurate values for
Kd and k2 in the buffer
conditions used in Fig. 2, by measuring the rates of excision at
different concentrations of glycosylase and substrate, was unsuccessful
when CpG·C-DNA was used, because there was no
change in the rate of excision even at the lowest concentration
detectable by our assay (0.01 nM). Thus,
Kd values for the ethenocytosine oligonucleotide must be <0.01 nM under these buffer conditions. Increasing
the concentration of salt decreases the DNA binding affinity of most proteins, mainly because the protein displaces cations from the DNA
phosphates when it binds to the DNA (29, 30). By increasing KCl to 140 mM (which is isotonic to mammalian tissue),
Kd and k2 could be measured
for the action of TDG on both
CpG·T-DNA and CpG·
C-DNA (Fig.
3 and Table
I). Data for the base excision at seven different equimolar concentrations of DNA and TDG were fitted simultaneously to the reaction model in Scheme I. When fitting was
performed allowing all three rate constants to vary,
k1 reached the maximum value expected for a
diffusion-controlled reaction (109
M
1 s
1). However, equally good
fits to the experimental data were obtained with
k1 fixed at values up to 2 orders of magnitude
lower and allowing just k
1 and
k2 to vary. These fits gave essentially identical values for k2 and
k
1/k1 (i.e.
Kd). Although the absolute values of
k1 and k
1 were poorly
defined by the experimental data, the results do show that
k
1
k2, and so the
reaction is not limited by association of TDG with the substrates. In
close agreement with the results in 50 mM KCl, the maximum
rate of excision (k2) for ethenocytosine is six
times slower than the excision of thymine (Table I). Most strikingly Kd for the CpG·
C-DNA
substrate is nearly 800 times smaller than the Kd of
CpG·T-DNA.
|
|
TDG Flips Ethenocytosine Out of DNA More Easily Than It Flips
Thymine--
The binding of DNA, represented by Kd
in Scheme I, involves flipping the mismatched base out of the DNA helix
into a pocket of TDG (31). There are two possible explanations of why
CpG·C-DNA has a much smaller
Kd than CpG·T. 1) TDG binds CpG·
C-DNA more tightly by making more and/or
better contacts to the DNA substrate, or 2) less energy is required to
flip out the ethenocytosine from a G·
C base
pair than to flip out a thymine from a G·T mismatch. In terms
of potential contacts, the only difference between the two substrates
is the base to be excised. The first explanation therefore implies that
TDG binds ethenocytosine more tightly than thymine. This was tested by
investigating whether free ethenocytosine base inhibits the TDG
reaction more than free thymine base. Addition of either thymine or
ethenocytosine up to a concentration of 5 mM had no effect
upon the reaction of TDG (data not shown). Higher concentrations of
free base were not practical because of their poor solubility. TDG
therefore has little affinity for either thymine or ethenocytosine.
This is in contrast to uracil-DNA glycosylase where 2 mM
uracil inhibited the reaction by 58% (32).
Inhibition of the TDG reaction by single-stranded oligonucleotides
containing either ethenocytosine or thymine was also investigated. TDG
does not excise thymine (23) or ethenocytosine (16) from single-stranded DNA, and so these oligonucleotides would act as reversible inhibitors. The results in Fig.
4 show that the rate of excision of
thymine from 25 nM CpG·T-DNA can be reduced to ~50% by inhibition with 250 nM single-stranded thymine
containing oligonucleotide. A similar but slightly lower level of
inhibition was given by 60 nM single-stranded
ethenocytosine oligonucleotide. The results in Fig. 4 suggest that TDG
binds single-stranded DNA containing ethenocytosine approximately three
times more tightly than single-stranded DNA containing thymine.
|
Excision of Ethenocytosine Is Very Dependent Upon the Flanking DNA
Sequence--
The rate of excision of thymine from G·T
mismatches is dependent upon the base pair 5' to the mismatched guanine
(11-14). To see whether the excision of ethenocytosine exhibited a
similar dependence upon flanking sequence, Kd and
k2 values for the excision of ethenocytosine
from TpG·C-DNA,
GpG·
C-DNA, and
ApG·
C-DNA were determined. For comparison,
Kd and k2 were also measured
for the excision of thymine from TpG·T-DNA under identical
buffer conditions (the excision of thymine from other sequence contexts
was too slow to allow accurate determination of Kd).
The results in Table I show that excision of ethenocytosine also is
very dependent upon the 5' base pair. In terms of the specificity
constant, k2/Kd, an
ethenocytosine in a CpG·
C site is 45 times more
efficiently removed than the next best ethenocytosine substrate,
TpG·
C-DNA. This is very similar to the
difference in specificity constant between CpG·T-DNA and
TpG·T-DNA. However, although with the G·T substrates
this drop in specificity is entirely because of a reduced k2, the drop in specificity for ethenocytosine
is the result of TpG·
C-DNA having both reduced
k2 and decreased binding to TDG
(larger Kd).
The other ethenocytosine oligonucleotides are even worse substrates;
GpG·C-DNA and ApG·
C-DNA
had 77- and 1630-fold, respectively, lower specificity constants than
CpG·
C-DNA. Again these reduced specificities
are because of both decreased k2 and increased
Kd. An earlier study using slightly different buffer
conditions found that k2 for the excision of thymine from G·T mismatches depended upon the base 5' to the
mismatched guanine in the order C
T > G > A (14).
Here, excision of ethenocytosine from G·
C base
pairs follows a similar trend except that k2 is
faster with a 5'-guanine than with a 5'-thymine (i.e. C
G > T > A).
Effect of APEX on Product Release by TDG--
In vitro,
the reaction of TDG with G·T mismatches is limited by
extremely slow release of the abasic DNA product (14, 33, 34). This
dissociation is so slow that each TDG molecule removes only one
thymine. The apurinic endonuclease, APEX, increases the turnover number
of TDG by displacing the glycosylase from the abasic site. The effect
of APEX on the turnover of TDG with CpG·T-DNA was compared
with its effect on the turnover of TDG with
CpG·C-DNA (Fig.
5). With both substrates in the absence
of APEX, the reaction stops after a stoichiometric amount of base has
been removed. In the presence of APEX, the turnover of both substrates is increased. This increase in turnover is dependent upon the concentration of APEX and is essentially the same for
CpG·T-DNA and for CpG·
C-DNA.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In this paper the effect of flanking DNA sequence on the excision
of ethenocytosine by TDG has been examined. To avoid the problem of
product inhibition often encountered with DNA glycosylases and which is
particularly strong for TDG, single turnover kinetics were used to
determine the kinetic constants k2 and
Kd (Scheme I). Saparbaev et al. (16)
previously calculated kcat and
Km values for the reaction of TDG with
oligonucleotides containing GpG·T or
GpG·C mismatches assuming that the reaction
followed Michaelis-Menten kinetics. Their conclusion that
G·
C is a much better substrate for TDG than
G·T broadly agrees with our results for the mismatches in this
sequence context, but, although the conclusion is correct, since
product release by TDG is so slow a Michaelis-Menten analysis is not
appropriate. It was originally believed that the physiological purpose
of TDG was to remove thymine from the G·T mismatches produced
by deamination of 5-methylcytosine (10), but the observation that TDG
removes ethenocytosine faster than thymine led to the suggestion that the real in vivo role of TDG is to remove ethenocytosine and
that the removal of thymine from G·T mismatches was a
fortuitous accident (15, 16). However, from Table I it is clear that
TDG has evolved to have a strong sequence preference for base excision at CpG sites. Because G·T mismatches produced by the
deamination of 5-methylcytosine occur exclusively in the sequence
context CpG·T, the main role of TDG in cells must therefore be
to remove deaminated 5-methylcytosine from CpG sites. In E. coli the mismatch-specific uracil-DNA glycosylase, which is a
homologue of TDG, appears to be the only glycosylase that removes
ethenocytosine efficiently (35). In humans, besides TDG there are two
other enzymes known to remove ethenocytosine. These are SMUG1
(36),2 and the methyl-CpG
binding domain protein 4 (MBD4, also known as MED1) which has a weak
ethenocytosine glycosylase activity (20). It is not yet known which (if
any) of these enzymes is responsible for removing ethenocytosine
in vivo.
The specificity constant for CpG·T-DNA mismatches is 56 times
higher than for the next best G·T substrate,
TpG·T-DNA (Table I). This difference in specificity is almost
entirely because of a drop in k2 with virtually
no change in Kd. In an earlier study we deduced that
when it binds to a G·T mismatch in the sequence
CpG·T, the glycosylase makes cooperative contacts to the
mismatched guanine and to the C·G base pair on the 5' side of the
mismatched guanine (14). The fact that Kd does not
change when the 5'-flanking base pair is changed shows that the binding
energy of the contacts to the mismatched guanine and to the 5'-C·G
base pair is used to stabilize the transition state (to lower
k2) and not to stabilize the enzyme-substrate
complex. There is a similar drop in specificity (45-fold) on going from CpG·C-DNA to TpG·
C-DNA
suggesting that TDG makes the same contacts to the mismatched guanine
and the 5'-C·G base pair of the CpG·
C-DNA
substrate. However, in this case the drop in specificity is due both to
a lower k2 and to an increase in
Kd, and so the binding energy of these contacts is
used to stabilize both the transition state and the enzyme-substrate complex.
TDG excised ethenocytosine from three of the four DNA sequences
containing G·C tested with specificity
constants (k2/Kd) that are at
least as good as the physiologically relevant G·T
oligonucleotide, CpG·T-DNA. However, in all of the
G·
C substrates ethenocytosine was excised more
slowly than thymine, and the increased specificity constants result
from much lower Kd values. TDG binds
CpG·
C-DNA nearly 800 times more tightly than
CpG·T-DNA, corresponding to a difference in binding energy of
~4 kcal/mol. In the enzyme-substrate complex the base to be excised
would be flipped out of the DNA helix into a binding pocket of the
glycosylase (31, 33). In this complex the enzyme makes contacts to the
DNA backbone, to the mismatched guanine, and to the flipped out base
itself. Because the contacts to the mismatched guanine and to the DNA
backbone would be the same for CpG·T-DNA and for
CpG·
C-DNA, the 800-fold lower
Kd of CpG·
C-DNA may result
from TDG binding the flipped out ethenocytosine more tightly than the
flipped out thymine. If ethenocytosine was bound very strongly one
might expect free ethenocytosine to inhibit the TDG reaction, but up to
5 mM ethenocytosine had no effect. Sub-micromolar
concentrations of single-stranded DNA containing either an
ethenocytosine or a thymine did inhibit the TDG reaction (Fig. 4), but
the single-stranded DNA containing an ethenocytosine was only three
times more effective than the single-stranded DNA containing a thymine.
This suggests that the majority of the 800-fold tighter binding of
CpG·
C-DNA compared with CpG·T-DNA is
not because of TDG binding the flipped out ethenocytosine more tightly
than thymine. An alternative explanation for the lower
Kd of CpG·
C-DNA is that less
energy is needed to flip out the ethenocytosine from a
G·
C base pair than is needed to flip out
thymine from a G·T mismatch. Melting temperature studies show
that a G·
C base pair is less stable than a
G·T base pair, although the magnitude of this difference in
stability varies considerably between different authors. One melting
temperature study found that on average a G·T mismatch
contributes 3.5 kcal/mol less energy than a G·C base pair to
the stability of a DNA duplex (37). In another study, 13 bp DNA
duplexes containing a G·
C base pair were
13.4-15.3 kcal/mol less stable than the parental G·C-containing duplexes (38). A further study directly
comparing the melting temperature of 15-bp DNA duplexes containing
either a G·
C base pair or a G·T
mismatch found that the G·
C containing duplexes
were 0.43-1.63 kcal/mol less stable than the corresponding
G·T-containing duplex (39). Interestingly, the difference in
stability between G·
C and G·T was
greatest when the mismatch was in a CpG site. Structural studies also
indicate that a G·
C base pair is considerably
less stable than a G·T mismatch. Both crystal (40) and NMR
(41) structures show that a G·T mismatch forms a stable
"wobble" base pair that involves two good hydrogen bonds (Fig.
6). Also, the guanine and thymine are
stacked well with the adjacent bases in the DNA helix. Although the NMR
structure of a G·
C base pair reveals a similar
wobble geometry, this base pair is much more distorted, and there is
only one hydrogen bond between the ethenocytosine and the guanine (42).
In addition to this weaker hydrogen bonding, the ethenocytosine base is
very poorly stacked with the adjacent bases, suggesting that the
G·
C base pair is considerably less stable than
a G·T mismatch.
|
An investigation of the reaction of chloroacetaldehyde with DNA found
that, although the distribution of ethenoadducts was not random, there
was no obvious DNA sequence preference for the formation of
ethenoadducts (43). Ethenocytosine is therefore expected to occur in
all sequence contexts. If TDG were the sole enzyme responsible for
repairing ethenocytosine, the results in Table I suggest that
ethenocytosine formed at ApG·C sites would be
very poorly repaired. Thus one might expect a predominance of mutations
of G·C base pairs at ApG sites in vinyl chloride-treated
animals. Analysis of liver tumors from rats exposed to vinyl chloride
found that 3 of 25 angiosarcomas had a mutation of a G·C base
pair in their p53 gene (44). In another study, analysis of
hepatacellular carcinomas taken from workers exposed to vinyl chloride
found that 5 of 18 had a mutated G·C base pair in their p53
gene (45). Although the number of mutations studied so far is very
small, it is perhaps significant that none of these G·C mutations occur at ApG sequences. This suggests that poor repair of ethenocytosine at ApG·
C sites by TDG
is not an important factor in vinyl chloride-induced carcinogenesis.
Acting alone, TDG removes a stoichiometric amount of mismatched base
because the glycosylase remains bound to the abasic site product, but
the next enzyme in the base excision repair pathway, the apurinic
endonuclease APEX, releases the TDG from the abasic site (Fig. 5 and
Refs. 23 and 46). Tight binding of abasic DNA and the stimulatory
effect of apurinic endonucleases has now been found for several other
DNA glycosylases (21, 47-49). The mechanism of the release of the
glycosylase by APEX is unknown and the subject of some controversy. One
theory that has been proposed is that the apurinic endonuclease acts
passively by simply "mopping up" free abasic DNA to
prevent re-binding of the glycosylase to the abasic DNA, thus allowing
it to react with more substrate DNA (21, 50). This is probably true for
some glycosylases but not for TDG. The fact that APEX increases the
turnover of TDG to more than 40 times the observed rate of TDG
dissociation from abasic DNA suggests that APEX actively
displaces the glycosylase from the abasic site (23, 46), either by
interacting directly with the bound TDG to displace the glycosylase or
by binding to the DNA and distorting the DNA structure in such a way
that disrupts the TDG-DNA complex (51, 52). The recent discovery
that mouse TDG interacts, albeit weakly, with mouse apurinic
endonuclease 1 also supports an active displacement mechanism (53). As
shown in Fig. 5, APEX increases the turnover of TDG with
CpG·C-DNA to the same extent as with
CpG·T-DNA. Because the reaction of TDG with
CpG·
C-DNA gives the same glycosylase-abasic DNA
complex as the reaction of TDG with CpG·T-DNA, this is
consistent with a model where APEX actively displaces the TDG from the
abasic site.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Professor Peter Swann (University College London, UK) for helpful discussions and for critical reading of the manuscript.
![]() |
FOOTNOTES |
---|
* This work was supported by the Wellcome Trust, UK.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed. Tel.: 44-20-7679 2323;
Fax: 44-20-7679 7193; E-mail: t.waters@biochem.ucl.ac.uk.
Published, JBC Papers in Press, December 18, 2002, DOI 10.1074/jbc.M211084200
2 J. E. A. Wibley, T. R. Waters, K. Haushalter, G. L. Verdine, and L. H. Pearl, manuscript in preparation.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: APEX, apurinic endonuclease 1; TDG, thymine-DNA glycosylase; SMUG1, single strand-selective monofunctional uracil-DNA glycosylase.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Creech, J. L., Jr., and Johnson, M. N. (1974) J. Occup. Med. 16, 150-151[Medline] [Order article via Infotrieve] |
2. | Barbin, A., Bresil, H., Croisy, A., Jacquignon, P., Malaveille, C., Montesano, R., and Bartsch, H. (1975) Biochem. Biophys. Res. Commun. 67, 596-603[Medline] [Order article via Infotrieve] |
3. | O'Neill, I., Barbin, A., Friesen, M., and Bartsch, H. (1986) IARC Sci. Publ. 70, 57-73[Medline] [Order article via Infotrieve] |
4. | Barbin, A. (2000) Mutat. Res. 462, 55-69[Medline] [Order article via Infotrieve] |
5. | Nair, J. (1999) IARC Sci. Publ. 150, 55-61[Medline] [Order article via Infotrieve] |
6. | Marion, M. J., and Boivin-Angele, S. (1999) IARC Sci. Publ. 150, 315-324[Medline] [Order article via Infotrieve] |
7. | Dosanjh, M. K., Chenna, A., Kim, E., Fraenkel-Conrat, H., Samson, L., and Singer, B. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 1024-1028[Abstract] |
8. | Krokan, H. E., Standal, R., and Slupphaug, G. (1997) Biochem. J. 325, 1-16[Medline] [Order article via Infotrieve] |
9. | Schärer, O. D., and Jiricny, J. (2001) Bioessays 23, 270-281[CrossRef][Medline] [Order article via Infotrieve] |
10. |
Neddermann, P.,
and Jiricny, J.
(1993)
J. Biol. Chem.
268,
21218-21224 |
11. | Griffin, S., Branch, P., Xu, Y.-Z., and Karran, P. (1994) Biochemistry 33, 4787-4793[Medline] [Order article via Infotrieve] |
12. | Sibghat-Ullah, and Day, R. S., III (1995) Biochemistry 34, 6869-6875[Medline] [Order article via Infotrieve] |
13. | Sibghat-Ullah, Gallinari, P., Xu, Y.-Z., Goodman, M. F., Bloom, L. B., Jiricny, J., and Day, R. S., III (1996) Biochemistry 35, 12926-12932[CrossRef][Medline] [Order article via Infotrieve] |
14. |
Waters, T. R.,
and Swann, P. F.
(1998)
J. Biol. Chem.
273,
20007-20014 |
15. |
Hang, B.,
Medina, M.,
Fraenkel-Conrat, H.,
and Singer, B.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
13561-13566 |
16. |
Saparbaev, M.,
and Laval, J.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
8508-8513 |
17. |
Miao, F.,
Bouziane, M.,
and O'Connor, T. R.
(1998)
Nucleic Acids Res.
26,
4034-4041 |
18. |
Nilsen, H.,
Haushalter, K. A.,
Robins, P.,
Barnes, D. E.,
Verdine, G. L.,
and Lindahl, T.
(2001)
EMBO J.
20,
4278-4286 |
19. | Noll, D. M., Gogos, A., Granek, J. A., and Clarke, N. D. (1999) Biochemistry 38, 6374-6379[CrossRef][Medline] [Order article via Infotrieve] |
20. | Petronzelli, F., Riccio, A., Markham, G. D., Seeholzer, S. H., Genuardi, M., Karbowski, M., Yeung, A. T., Matsumoto, Y., and Bellacosa, A. (2000) J. Cell. Physiol. 185, 473-480[CrossRef][Medline] [Order article via Infotrieve] |
21. |
Vidal, A. E.,
Hickson, I. D.,
Boiteux, S.,
and Radicella, J. P.
(2001)
Nucleic Acids Res.
29,
1285-1292 |
22. | Haushalter, K. A., Stukenberg, P. T., Kirschner, M. W., and Verdine, G. L. (1999) Curr. Biol. 9, 174-185[CrossRef][Medline] [Order article via Infotrieve] |
23. |
Waters, T. R.,
Gallinari, P.,
Jiricny, J.,
and Swann, P. F.
(1999)
J. Biol. Chem.
274,
67-74 |
24. | Waters, T. R., and Swann, P. F. (1997) Biochemistry 36, 2501-2506[CrossRef][Medline] [Order article via Infotrieve] |
25. | Zhang, W., Rieger, R., Iden, C., and Johnson, F. (1995) Chem. Res. Toxicol. 8, 148-156[Medline] [Order article via Infotrieve] |
26. | Gait, M. (ed) (1984) Oligonucleotide Synthesis: A Practical Approach , IRL Press at Oxford University Press, Oxford |
27. | Xu, Y.-Z., and Swann, P. F. (1992) Anal. Biochem. 204, 185-189[CrossRef][Medline] [Order article via Infotrieve] |
28. |
Neddermann, P.,
Gallinari, P.,
Lettieri, T.,
Schmid, D.,
Truong, O.,
Hsuan, J. J.,
Wiebauer, K.,
and Jiricny, J.
(1996)
J. Biol. Chem.
271,
12767-12774 |
29. | Härd, T., and Lundbäck, T. (1996) Biophys. Chem. 62, 121-139[CrossRef] |
30. | Lohman, T. M. (1986) CRC Crit. Rev. Biochem. 19, 191-245[Medline] [Order article via Infotrieve] |
31. |
Barrett, T. E.,
Schärer, O. D.,
Savva, R.,
Brown, T.,
Jiricny, J.,
Verdine, G. L.,
and Pearl, L. H.
(1999)
EMBO J.
18,
6599-6609 |
32. | Slupphaug, G., Eftedal, I., Kavli, B., Bharati, S., Helle, N. M., Haug, T., Levine, D. W., and Krokan, H. E. (1995) Biochemistry 34, 128-138[Medline] [Order article via Infotrieve] |
33. | Barrett, T. E., Savva, R., Panayotou, G., Barlow, T., Brown, T., Jiricny, J., and Pearl, L. H. (1998) Cell 92, 117-129[Medline] [Order article via Infotrieve] |
34. |
Schärer, O. D.,
Nash, H. M.,
Jiricny, J.,
Laval, J.,
and Verdine, G. L.
(1998)
J. Biol. Chem.
273,
8592-8597 |
35. |
Lutsenko, E.,
and Bhagwat, A. S.
(1999)
J. Biol. Chem.
274,
31034-31038 |
36. |
Kavli, B.,
Sundheim, O.,
Akbari, M.,
Otterlei, M.,
Nilsen, H.,
Skorpen, F.,
Aas, P. A.,
Hagen, L.,
Krokan, H. E.,
and Slupphaug, G.
(2002)
J. Biol. Chem.
277,
39926-39936 |
37. | Allawi, H. T., and SantaLucia, J., Jr. (1997) Biochemistry 36, 10581-10594[CrossRef][Medline] [Order article via Infotrieve] |
38. | Gelfand, C. A., Plum, G. E., Grollman, A. P., Johnson, F., and Breslauer, K. J. (1998) Biochemistry 37, 12507-12512[CrossRef][Medline] [Order article via Infotrieve] |
39. | Sági, J., Perry, A., Hang, B., and Singer, B. (2000) Chem. Res. Toxicol. 13, 839-845[CrossRef][Medline] [Order article via Infotrieve] |
40. |
Hunter, W. N.,
Brown, T.,
Kneale, G.,
Anand, N. N.,
Rabinovich, D.,
and Kennard, O.
(1987)
J. Biol. Chem.
262,
9962-9970 |
41. |
Allawi, H. T.,
and SantaLucia, J., Jr.
(1998)
Nucleic Acids Res.
26,
4925-4934 |
42. | Cullinan, D., Johnson, F., Grollman, A. P., Eisenberg, M., and de los Santos, C. (1997) Biochemistry 36, 11933-11943[CrossRef][Medline] [Order article via Infotrieve] |
43. | Tudek, B., Kowalczyk, P., and Ciesla, J. M. (1999) IARC Sci. Publ. 150, 279-293[Medline] [Order article via Infotrieve] |
44. | Barbin, A., Froment, O., Boivin, S., Marion, M. J., Belpoggi, F., Maltoni, C., and Montesano, R. (1997) Cancer Res. 57, 1695-1698[Abstract] |
45. | Weihrauch, M., Lehnert, G., Köckerling, F., Wittekind, C., and Tannapfel, A. (2000) Cancer (Phila.) 88, 1030-1036 |
46. | Privezentzev, C. V., Saparbaev, M., and Laval, J. (2001) Mutat. Res. 480, 277-284 |
47. |
Hill, J. W.,
Hazra, T. K.,
Izumi, T.,
and Mitra, S.
(2001)
Nucleic Acids Res.
29,
430-438 |
48. | Sung, J. S., and Mosbaugh, D. W. (2000) Biochemistry 39, 10224-10235[CrossRef][Medline] [Order article via Infotrieve] |
49. |
Yang, H.,
Clendenin, W. M.,
Wong, D.,
Demple, B.,
Slupska, M. M.,
Chiang, J. H.,
and Miller, J. H.
(2001)
Nucleic Acids Res.
29,
743-752 |
50. | Hardeland, U., Bentele, M., Lettieri, T., Steinacher, R., Jiricny, J., and Schar, P. (2001) Prog. Nucleic Acids Res. Mol. Biol. 68, 235-253[Medline] [Order article via Infotrieve] |
51. |
Parikh, S. S.,
Mol, C. D.,
Slupphaug, G.,
Bharati, S.,
Krokan, H. E.,
and Tainer, J. A.
(1998)
EMBO J.
17,
5214-5226 |
52. | Waters, T. R., and Swann, P. F. (2000) Mutat. Res. 462, 137-147[Medline] [Order article via Infotrieve] |
53. | Tini, M., Benecke, A., Um, S. J., Torchia, J., Evans, R. M., and Chambon, P. (2002) Mol. Cell 9, 265-277[Medline] [Order article via Infotrieve] |