Published online 5 March 2009
Nucleic Acids Research, 2009, Vol. 37, No. 8 2539–2548
doi:10.1093/nar/gkp100
Oxidative stress induces degradation of
mitochondrial DNA
Inna Shokolenko, Natalia Venediktova, Alexandra Bochkareva,
Glenn L. Wilson and Mikhail F. Alexeyev*
Department of Cell Biology and Neuroscience, University of South Alabama, Mobile, AL 36688, USA
Received September 4, 2008; Revised January 25, 2009; Accepted February 8, 2009
ABSTRACT
Mitochondrial DNA (mtDNA) is located in close proximity of the respiratory chains, which are the main
cellular source of reactive oxygen species (ROS).
ROS can induce oxidative base lesions in mtDNA
and are believed to be an important cause of the
mtDNA mutations, which accumulate with aging
and in diseased states. However, recent studies
indicate that cumulative levels of base substitutions
in mtDNA can be very low even in old individuals.
Considering the reduced complement of DNA
repair pathways available in mitochondria and
higher susceptibility of mtDNA to oxidative
damage than nDNA, it is presently unclear how
mitochondria manage to maintain the integrity of
their genetic information in the face of the permanent exposure to ROS. Here we show that oxidative
stress can lead to the degradation of mtDNA
and that strand breaks and abasic sites prevail
over mutagenic base lesions in ROS-damaged
mtDNA. Furthermore, we found that inhibition of
base excision repair enhanced mtDNA degradation
in response to both oxidative and alkylating
damage. These observations suggest a novel mechanism for the protection of mtDNA against oxidative
insults whereby a higher incidence of lesions to the
sugar–phosphate backbone induces degradation of
damaged mtDNA and prevents the accumulation
of mutagenic base lesions.
INTRODUCTION
Mutations in mitochondrial DNA (mtDNA) are an
underlying factor in many mitochondrial diseases (1)
and have been associated with cancer (2,3), neurodegenerative disorders (4,5), diabetes (6–8) and aging (9,10).
Therefore, mechanisms of mtDNA mutagenesis are of
considerable interest. Mutations in mtDNA can arise
following exposure to environmental mutagens, from
DNA polymerase errors during replication, from failure
of the mtDNA repair machinery, and from defects in
the mechanisms of degradation of damaged mtDNA.
However, perhaps the most recognized source of
mtDNA mutagenesis is reactive oxygen species (ROS),
which are produced by the mitochondrial electron transport chain (ETC). Rates of oxygen reduction to the principal ROS, the superoxide anion (O2.), have been
reported to be as high as 1–5% of total consumed
oxygen (11,12). The charged and relatively unstable O2.
is membrane-impermeable and, depending on the site of
production, can be released into either the mitochondrial
matrix or the intermembrane space. There, it is quickly
converted to hydrogen peroxide (H2O2), the principal cellular mediator of oxidative stress. This conversion occurs
either spontaneously or enzymatically, with the help of
superoxide dismutases. The relative stability and membrane permeability of H2O2 allows it to freely diffuse
throughout the cell, where it can generate, by means of
the Fenton reaction, the extremely reactive hydroxyl radical, which can efficiently damage DNA (13,14). It has been
reported that the main products of mtDNA base damage
are thymine glycol among pyrymidines (15) and 7,
8-dihydro-8-oxo-20 -deoxyguanosine (8-oxoG) among purines (16–18). The former has low mutagenicity, whereas
the latter upon replication can cause characteristic G!T
transversions (15).
Mitochondrial production of ROS and the ensuing
damage inflicted by these ROS on biological macromolecules, including DNA, constitutes the basis of the
free radical/mitochondrial theory of aging (19–21).
According to this theory, the production of ROS by mitochondria leads to mtDNA damage and mutations which
in turn lead to progressive respiratory chain dysfunction
and to a further increase in ROS production as a consequence of this dysfunction. The exponential escalation
of these processes is commonly referred to as a ‘vicious
cycle’, and the theory predicts that the rise in mtDNA
mutations and ROS eventually reach levels that are
incompatible with life.
In this study, we examine the effects of oxidative stress
on the integrity of mtDNA and conclude that oxidative
*To whom correspondence should be addressed. Tel: +1 251 460 6789; Fax: +1 251 460 6771; Email: malexeye@jaguar1.usouthal.edu
ß 2009 The Author(s)
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/
by-nc/2.0/uk/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.
2540 Nucleic Acids Research, 2009, Vol. 37, No. 8
stress can result in the loss of damaged mtDNA molecules.
This phenomenon is unique to the mitochondrial compartment, and is enabled by the high redundancy of
mtDNA.
MATERIALS AND METHODS
Cell growth and treatment
HCT116 cells (a kind gift of Drs Vogelstein and Kinzler)
were grown in Dulbecco’s modified Eagle medium
(DMEM) containing 10% Fetal Bovine Serum and
50 mg/ml gentamycin in a humidified atmosphere containing 5% CO2 at 378C. For oxidative damage and repair
experiments cells were grown in 100 mm dishes until
50–80% confluence. Then cultures were washed with
warm HBSS and exposed to the indicated concentrations
of H2O2 in HBSS for the indicated periods of time at 378C
in a humidified atmosphere containing 5% CO2. Control
cultures were mock-treated with HBSS alone. After incubation, cells were either lysed immediately or allowed
to repair for the indicated periods of time in complete
growth medium. Cell treatment with hypoxanthine/xantine oxidase (XO) was performed for 1h using indicated
concentrations of enzyme and 0.5 mM hypoxanthine in
PBS. Methyl methane sulfonate (MMS) treatments were
performed in PBS for 30 min at 378C in a humidified
atmosphere containing 5% CO2. Where indicated, methoxyamine (30 mM) was included both during treatment
and recovery periods.
For long-term rotenone treatment, cells were seeded
in three T-75 flasks, allowed to attach for 20 h, and the
medium was replaced with fresh medium containing
200 nM rotenone. From this point on, the medium was
replaced every 48 h, and cells were split as necessary as
they approached confluency. At the end of the experiment,
cultures were allowed to undergo at least two cell divisions
to allow for the fixation of mtDNA mutations prior to
DNA extraction.
Mitochondrial ROS measurement
To measure mitochondrial ROS production in response
to rotenone, cells were collected by trysinization,
washed with PBS, and loaded with 5 mM of MitoSOX
(Invitrogen, Carlsbad, CA) in the presence of indicated
concentrations of rotenone for 30 min at 378C in a humidified atmosphere containing 5% CO2 followed by flow
cytometry.
Quantitative Southern blotting
Quantitative alkaline Southern blotting was performed
essentially as described earlier (22). Neutral Southern
blots were performed similarly, except that there was no
alkaline pretreatment, and NaOH was not included in the
loading dye, agarose gel or the electrophoresis buffer.
When blotting BamHI-digested total DNA, after transfer
the membrane was cut at the level of the 9-kb band of
lambda/HindIII marker. The upper portion was then
hybridized with the mtDNA probe (16.5-kb fragment),
and the lower portion was hybridized with the 18S
rDNA probe (5-kb fragment). After hybridization, membranes were exposed to an imaging screen to determine
band intensity. The number of pixels per band was determined by encompassing bands with identical rectangular
regions of interest and subtracting the background.
The break frequency was determined using the Poisson
expression (s = –ln P0, where s is the number of breaks
per fragment and P0 is the fraction of fragments free
of breaks).
PCR-cloning-sequencing
The isolation of total cellular DNA was described previously (22). The amplification of mtDNA was performed
in triplicate or quadruplicate 100 ml reactions per each
independent experimental condition. Each reaction contained 50 ml of 2 high fidelity PCR master mix (New
England Biolabs, Ipswich, MA), 500 ng of total DNA,
1 mM of each primer (forward AATGTCTGCACAGCC
ACTTTCCAC and reverse AGGGTTTGCTGAAGATG
GCGGTAT), and the balance of water. The PCR was
performed for 25 cycles (denaturation for 10 s at 988C,
annealing for 5 s at 558C and extension for 30 s at 558C).
The PCR product was directly purified from the reaction
mixture using PCR purification kit (Qiagen, Valencia,
CA) and ligated to the EcoRV digested plasmid
pBluescriptII SK+ for 4 h at 258C. The ligated mixture
was introduced into Escherichia coli cells, allowed to incubate for 1 h at 378C and then 25% of the total mixture was
diluted 1:4 and 100 ml aliquots of this dilution were plated
on LB agar plates containing 200 mg/ml ampicillin, 40 mg/
ml X-gal and 1 mM IPTG. Typically, no more than 16%
of electroporated cells were plated. White colonies were
picked, inoculated into TB medium (1.2% Tryptone, 2.4%
yeast extract, 0.4% glycerol, 72 mM K2HPO4 and 17 mM
KH2PO4), and grown overnight to OD600 = 5–8. At this
point, bacterial cells were pelleted by centrifugation, and
plasmid DNA was extracted using Qiaprep spin kit
(Qiagen). 0.5–1.0 mg of this DNA was used in 10 ml
cycle-sequencing reaction (BigDye v3.1., Applied
Biosystems) in 96-well PCR plates using T3 and
T7-sequencing primers. Sequencing products were
purified using 96-well DTR plates (Edge Biosystems,
Gaithersburg, MD), and submitted to high throughput
capillary sequencing runs at either the DNA-sequencing
facility at the Iowa State University, or Functional
Biosciences (Madison, WI). The resulting chromatograms
were aligned against the consensus sequence of amplified
fragments.
RESULTS
Validation of PCR-cloning-sequencing assay
Several methods have been described for the mutational
analysis of mtDNA. Among those are a single-molecule
PCR (23), a random mutation capture (RMC) assay (24)
and PCR-cloning-sequencing (25). The first of these methods is difficult to adapt for the detection of low levels
of heteroplasmy, while the second samples mutation frequencies in very short palindromic sequences and may
underreport mutation frequency due to the relaxed
Nucleic Acids Research, 2009, Vol. 37, No. 8 2541
specificity of the restriction enzyme used to eliminate wild
type restriction sites. Therefore, we have adopted a PCRcloning-sequencing strategy for our studies. To experimentally determine the level of PCR-induced mutations
in our system, we amplified and cloned an 1111-bp
DNA fragment of the mtDNA control region in a kanamycin-resistant derivative of pBluescriptII SK+,
extracted this test plasmid DNA, and subjected it to
PCR-cloning-sequencing. To approximate the starting
template quantity, we assumed that mtDNA constitutes
1% of the total cellular DNA [or 4000 copies per cell (26)].
In PCR-cloning-sequencing we used 500 ng of total DNA
for the amplification, which corresponds to 5 ng of
mtDNA (1% of total DNA). The size of the template
plasmid (4.1 kb) is roughly 25% of that of the mtDNA
molecule (16.5 kb). Therefore, 1.25 ng of plasmid DNA
was used as the template. The template was subjected
to 25 cycles of PCR with Phusion DNA polymerase,
and the PCR product was ligated to ampicillin-resistant
pBluescriptII SK+ (this manipulation excludes contamination with kanamycin resistant template plasmid DNA).
The resulting clones were subjected to sequencing
as described in the ‘Materials and methods’ section.
The level of PCR-induced mutagenesis under these experimental conditions was 5.6 106 5.6 106 (n = 3), or
a little less than 1 mutation per 10 mitochondrial genomes
(Figure 1A). This is in very good agreement with the calculated upper limit of PCR-induced mutagenesis under
these conditions of 1 105 (see below).
There are three methodological concerns associated
with the PCR-cloning-sequencing approach:
(i) The presence of nuclear ‘pseudogenes’ for the mitochondrial genome. The amplification and sequencing of these pseudogenes results in overreporting
mutation frequencies (27). Therefore, the putative
PCR product was homology-searched for its
absence in nuclear DNA (nDNA).
(ii) Sequencing multiple copies of the same PCR fragment. This is only relevant for DNA ligations that
produce a low yield of bacterial colonies. Indeed,
during a 1-h incubation in the medium for the
expression of a selective marker following electroporation, E. coli cells can divide at least once.
Therefore, exhaustive sequencing of all clones
would result in sequencing of two to four copies
of each cloned PCR fragment. To control for this
problem, no more than 16% of electroporated bacteria were spread on selective plates and then only a
fraction of white colonies was sequenced.
(iii) PCR-induced
mutations.
DNA-polymerasemediated errors accumulate in PCR fragments as a
function of the cycle number and the fidelity of the
DNA polymerase. With each PCR cycle new mutations are introduced by the DNA polymerase. Also,
PCR errors introduced in earlier cycles are faithfully
copied in each subsequent cycle so that their
number grows exponentially. To model this process,
Lin et al. (25) have derived a formula for the
estimation of an average number of mutations
introduced into each strand of PCR product.
<k> = cfLZ/(1 + Z), where <k> is the average
number of errors per strand, c is the number of
PCR cycles, f is the fidelity of DNA polymerase
(4 107 for Phusion polymerase), L is the length
of the amplified fragment and Z is a fraction of
template strands amplified in each cycle. Making
substitutions for a 1111-bp fragment amplified
with Phusion polymerase, we obtain <k> =
25 4 107 1111 Z/(1 + Z) = 0.01111
Z/(1 + Z). Assuming that the fraction of template
strands amplified in each cycle Z is somewhere
between 0.3 and 1, we obtain <k> values 0.0026
and 0.0056, respectively, for each strand, or for a
double-stranded PCR fragment 0.005 and 0.01,
respectively. This means that PCR-introduced
errors will be present at the most in 1 fragment
per every 100 clones sequenced, or an error rate
of 105 per base pair.
Chronic superoxide exposure and aging do not result in
significant mtDNA mutagenesis
The mitochondrial ETC complex I is an important site
for ROS production. When inhibited with rotenone, this
complex releases superoxide to the matrix side of the inner
mitochondrial membrane, in close proximity to mtDNA
(28,29). Chronic systemic rotenone infusion has been utilized to induce symptoms in one of the most faithful
animal models of Parkinsonism, a disease associated
with mtDNA mutations and advanced age (30).
Therefore, to approximate exposure of mtDNA to ROS
over a lifetime, HCT116 cells and immortalized mouse
embryonic fibroblasts (MEFs) were treated with 200 nM
and 400 nM rotenone, respectively, for 30 days. 200 nM
and 400 nM have been determined to be the maximum
concentrations tolerated by HCT116 and MEF cells,
respectively, without induction of apoptosis. These treatments did not result in a statistically significant increase in
the rate of mtDNA mutagenesis over untreated controls
(Figure 1A and C). If, during the treatment period, a selection of cells unable to produce O2. in response to rotenone inhibition had occurred, this artifact could have
manifested itself as a lack of mtDNA mutagenesis in
response to rotenone treatment. To explore this possibility, the production of O2. in response to rotenone treatment was measured in both HCT116 and MEF cells
before and after the completion of experiments. Both
types of cells preserved their ability to respond to rotenone
treatment with superoxide production (Figure 1B and D).
It is often postulated that a significant accumulation of
ROS-induced mtDNA mutations should be observed at
the end of the lifespan. Accordingly, mtDNA from the
lymphocytes of two female centenarians and one male
non-agenarian (98 years old), available as a panel of samples from the NIA repository at the Coriell Institute (samples NG11426, NG11467 and NG11716), was subjected
to PCR-cloning-sequencing. The frequency of mtDNA
mutations in these samples was not statistically different
from that observed in the control experiment with amplified plasmid or in untreated HCT116 cells (Figure 1A).
2542 Nucleic Acids Research, 2009, Vol. 37, No. 8
B
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E
Figure 1. Levels of mtDNA mutations in response to ROS. (A) mtDNA cloned in a plasmid vector (Test plasmid) as well mtDNA from very old
individuals (98–101 years old) and mtDNA from HCT116 cells, which were either left untreated or were treated for 30 days with the ROS generator
rotenone, were subjected to PCR-cloning-sequencing analysis. No significant increase was detected under any of the conditions tested (n = 3, one-way
ANOVA with Tukey post-test). (B and D) Prolonged treatment of HCT116 and MEF cells with rotenone does not result in the selection of cells
unable to generate ROS. Control and rotenone-treated (30 days, 200 nM, see ‘Materials and Methods’ section) HCT116 cells were loaded with 5 mM
of the mitochondrial superoxide radical indicator MitoSOX, treated with 200 nM rotenone for 30 min, and subjected to FACS analysis; (C) chronic
treatment of MEFs with rotenone (400 nM, 30 days) does not cause a significant increase in the mutations frequency (n = 3, two-tailed Student’s
t-test assuming unequal variances). (E) A representative quantitative Southern blot illustrating both mtDNA damage immediately after treatment
(0 h), and repair over 6-h period (6 h). HCT116 cells were treated with 400 mM H2O2 for 60 min, and total BamHI-digested genomic DNA was
subjected to both quantitative Southern blotting under the alkaline conditions and PCR-cloning-sequencing. (F) Relative frequency of combined
SSBs and abasic sites versus mutagenic base lesions in mtDNA of control (untreated) cells, in mtDNA of cells from (E) (H2O2—0 h), and in mtDNA
of cells treated with H2O2 four times with 24 h intervals between treatments (4 H2O2). The frequency is expressed on a per mtDNA molecule basis
(n = 3).
Nucleic Acids Research, 2009, Vol. 37, No. 8 2543
H2O2 has low mutagenicity in HCT116 cells
Several factors can underlie the apparent lack of a
significant accumulation of mtDNA mutations in aging
cells, including the low mutagenicity of ROS in mitochondria. This possibility is especially interesting in the light
of recent reports that susceptibility of mtDNA to the accumulation of oxidative base lesions may have been overestimated (31,32).
Hydrogen peroxide is a universal intracellular mediator
of oxidative stress. Therefore, we studied the effect of
H2O2 on mtDNA mutagenesis in HCT116 cells.
HCT116 cells were treated with 400 mM of H2O2 for 1 h,
after which they were either lysed or allowed to repair for
6 h, and total DNA was extracted and subjected to a
quantitative alkaline Southern blotting and PCR-cloning-sequencing analysis.
H2O2 treatment induced strand breaks and abasic sites
in mtDNA (Figure 1E). The prevalence of abasic sites is
impossible to evaluate because they are readily converted
into single-strand breaks (SSBs) by mitochondrial apurinic/apyrimidinic endonuclease (APE/Ref-1) and therefore
manifest themselves as SSB in quantitative Southern blotting under denaturing conditions. Importantly, mtDNA
analyzed immediately after H2O2 treatment contained
ten times more combined strand breaks and abasic sites
than mutagenic base lesions (Figure 1F). This ratio does
not take into account the background of PCR-induced
mutagenesis (0.09 mutations per mitochondrial genome),
and therefore represents a conservative estimate. Indeed,
subtraction of the PCR-induced mutagenesis rate
from experimentally observed values would result in a
ratio of 14.
Oxidative stress induces in DNA premutagenic base
lesions rather than mutations. A fraction of these lesions
(mutagenic lesions) is fixed as mutations only upon replication by DNA polymerase. Since different DNA polymerases have different propensities for misincorporation
of wrong nucleotides across from the damaged base
during replication, there exists a formal possibility that
low mutation rates observed in mtDNA immediately
after H2O2 treatment are due to a higher fidelity of
Phusion polymerase as compared to mtDNA polymerase
g (Pol g). This possibility was explored by subjecting cells
to four consecutive treatments with H2O2 with 24 h intervals between treatments. At the end of experiment, cultures were allowed to undergo at least five cell divisions
prior to DNA extraction to allow for the fixation of mutations with Pol g. This mode of treatment did not result in
the increase in the frequency of mtDNA mutations over
untreated control or over mtDNA isolated immediately
after treatment (n = 3, Figure 1F), suggesting that differences in the propensities of Phusion and Pol g to induce
mutations at the site of oxidative DNA damage do not
confound our results.
mtDNA repair and degradation pathways compete for
the H2O2-damaged mtDNA
If accumulation of abasic sites and strand breaks, rather
than oxidative base lesions in mtDNA is the predominant
consequence of exposure to H2O2, the main cellular
mediator of oxidative stress, then what is the fate of
damaged mtDNA? This question was addressed by following the fate of H2O2-damaged mtDNA by Southern
blotting under denaturing (alkaline) and non-denaturing
(neutral) conditions. H2O2 induced lesions in both nDNA
and mtDNA (Figure 2A). However, specific patterns
observed in mtDNA under the denaturing and non-denaturing conditions were markedly different in that under
the denaturing conditions, the amount of intact DNA
was the lowest immediately after the treatment, and
grew over the 6 h allowed for repair (Figure 2A). In contrast, under the non-denaturing conditions, the greatest
amount of intact DNA was observed immediately after
treatment, and then it decreased over the repair period
(Figure 2B). This trend was preserved between different
concentrations of H2O2 (Figure 2A and B), in different
experiments, and in HeLa and A549 cell lines
(Supplementary Figures 1 and 2). The differences observed
in the same samples of mtDNA separated under the denaturing and non-denaturing conditions are attributable to
the fact that sample preparation and electrophoresis under
the alkaline conditions results in the separation of the two
DNA strands and in the conversion of abasic sites into
SSBs. Therefore, mtDNA containing oxidative damage in
the form of abasic sites or strand breaks will be fragmented resulting in the decrease in intensity of the band corresponding to intact mtDNA. In contrast, neither SSBs,
nor abasic sites will affect the quantity of intact mtDNA
under non-denaturing conditions, as under these conditions the complementary strand will bridge the nick thus
preventing fragmentation. Under the non-denaturing conditions, only double-strand breaks (DSBs) will result in
mtDNA fragmentation and in decrease in the quantity
of full-length mtDNA. The repair of DSBs in mtDNA
has not been described, and introduction of DSBs in
mtDNA of mammalian cells by restriction endonucleases
results in mtDNA elimination through degradation
(33,34). Therefore, trends in the quantity of the full
length mtDNA observed under the denaturing and nondenaturing conditions following oxidative stress can be
interpreted as follows. If a cell is unable to repair all the
damage inflicted by H2O2 in the mitochondrial genome, a
fraction of mtDNA molecules (presumably, those which
received greater damage) receives DSBs and is degraded
(hence the reduction in the amount of total mtDNA seen
on a neutral gel, Figure 2B), while moderately damaged
genomes are repaired, which is reflected in the increase in
mtDNA quantity between 0 h and 6 h seen in alkaline gels
(Figure 2A).
If mtDNA is indeed degraded in response to oxidative
stress, then one of the initial steps in this process is likely
to be the conversion of the circular mtDNA molecule into
a linear intermediate. The half-life (and therefore, the
steady-state level) of this intermediate is likely to be determined by a balance between formation and degradation.
In the absence of knowledge of these processes, it is difficult to predict or interpret specific steady-state levels of
linear mtDNA intermediates. However, the mere presence
of such intermediates would support the notion of
mtDNA degradation in response to oxidative stress. We
therefore sought to determine whether oxidative stress is
2544 Nucleic Acids Research, 2009, Vol. 37, No. 8
accompanied by an accumulation of linear mtDNA degradation intermediates. Indeed, H2O2 treatment resulted
in an increase in the steady-state content of linear mtDNA
(Figure 2C). This increase was transient, and linear intermediates disappear (degrade) within 1 h after the removal
of a stressor (Figure 2D).
mtDNA degradation can be induced by enzymatically
generated ROS
To see if degradation of oxidatively damaged mtDNA
can potentially be induced by a physiologically relevant
enzyme system, HCT116 cells were treated with 5, 10 or
20 mU of XO in the presence of hypoxanthine (0.5 mM).
XO is released into a bloodstream upon liver damage,
and XO/hypoxanthine reaction produces both H2O2 and
superoxide (35). This treatment has faithfully recapitulated trends observed upon H2O2 treatment in that
mtDNA repair was observed under the denaturing conditions (Figure 3A), which occurred simultaneously with
mtDNA degradation observed under the non-denaturing
conditions (Figure 3B), and that mtDNA degradation was
accompanied by the accumulation of linear mtDNA intermediates (Figure 3C). Importantly, all three phenomena
appear to be dose-dependent suggesting their specificity
(Figure 3).
Inhibition of base excision repair (BER) of oxidative
DNA damage enhances mtDNA degradation
Figure 2. Oxidative damage results in the degradation of mtDNA.
(A and B) HCT116 cells were either left untreated (C, control) or
treated with the indicated concentrations of H2O2 for 1 h, after which
total genomic DNA was either extracted immediately (0 h), or allowed
to repair for 6 h (6 h) prior to extraction. The DNA was digested with
BamHI, which has a single recognition site in human mtDNA, quantitated and separated in 0.6% agarose gel under either alkaline (A) or
neutral (B) conditions. After Southern blotting with mitochondrial
(Mito) and nuclear (18S) DNA probes, the intensity of bands corresponding to intact mtDNA was expressed as a percent of control
(untreated) values. (C) Oxidative DNA damage is accompanied by
accumulation of linear mtDNA intermediates. HCT116 cells were treated with 200 mM H2O2, total DNA was extracted, digested with Bgl II,
and subjected to Southern blotting under the neutral conditions. Bgl II
has no recognition sites in mtDNA, and therefore the banding pattern
is reflective of the native state of mtDNA. The positions of markers
corresponding to covalently closed circular (CCC), linear and relaxed
mtDNA molecules are indicated by arrows. (D) A time course of degradation of linear mtDNA intermediates. HCT116 cells were either
left untreated (C) or treated with 400 mM H2O2 for 30 min, and then
either lysed immediately, or allowed to recover for 30, 60, or 90 min.
The percentage of linear mtDNA as compared to untreated control
is indicated.
To study the effect of BER inhibition on mtDNA degradation, HeLa cells were subjected to oxidative stress in the
presence or absence of the BER inhibitor methoxyamine
(MA). This compound covalently modifies abasic sites
generated by lesion-specific DNA glycosylases and prevents their incision by APE/Ref-1 (36). Since APE/Ref-1
is a bi-functional enzyme (AP endonuclease and a redox
factor, which maintains the activity of numerous transcription factors), methoxyamine treatment is preferred
over knockdown because it does not affect the redox
function of APE/Ref-1. Methoxyamine treatment abrogated BER of oxidative damage in both mtDNA and
nDNA (Figure 4A) and enhanced mtDNA degradation
(Figure 4B).
Inhibition of BER of alkylating DNA damage induces
mtDNA degradation
To test whether mtDNA degradation can only be induced
by oxidative DNA damage, or this is a more general phenomenon, we studied the repair of alkylation mtDNA
lesions in the presence or absence of MA. At low MMS
doses, no mtDNA degradation was observed in the
absence of MA. However, inhibition of BER with MA
both impaired the repair (Figure 5A) and dramatically
induced mtDNA degradation (Figure 5B). In the presence
of MA, nDNA repair was also impaired (Figure 5A).
However, induction of DSBs in nDNA was much less
pronounced compared to mtDNA under these conditions
(Figure 5B).
Nucleic Acids Research, 2009, Vol. 37, No. 8 2545
A
Mito
5mU
10mU
0.3%
2%
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9%
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Time
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20mU
6h 0h
100%
12%
1.5%
14%
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0h
HeLa XO MA
7%
6h
HeLa XO
3%
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C 0h
Alkaline
17%
32%
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5.3%
19%
%C
100%
Alkaline
100%
A
5mU
10mU
0h 6h C 0h 6h
0h 6h
Mito
90%
14%
64%
25%
81%
40%
78%
100%
HeLa XO
HeLa XO MA
4%
43%
18%
84%
61%
10mU
5mU
0h 6h C 0h 6h
16%
5mU
C 0h 6h
38%
Enz
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65%
46%
%C
0h 6h 0h 6h
18S
100%
10mU 20mU
Mito
10mU
0h 6h
10mU 20mU
0h 6h 0h 6h
Figure 3. The effect of enzymatically induced oxidative stress on
mtDNA. HCT116 cells were either left untreated (C), or treated with
5, 10, or 20 mU of xanthine oxidase (XO) in the presence of hypoxanthine (0.5 mM). Cells were then either lysed immediately after treatment (0 h) or allowed for repair (6 h). Total DNA was extracted and
digested with BamHI, and (A), separated under denaturing (alkaline)
conditions, and subjected to Southern blotting with nuclear (18S) and
mtDNA probe. %C indicates percent intact mtDNA as compared to
untreated control. (B) The same samples as in A were run under nondenaturing (TBE buffer) conditions. Annotation is the same as in (A).
This blot shows intact and SSBs containing mtDNA and nDNA. (C)
Total DNA samples from the same experiment as in (A) and (B) were
digested with BglII and run under the non-denaturing conditions to
detect linear mtDNA intermediates. Annotation is the same as in (A)
and (B). The arrow indicates the position of the linear mtDNA species.
DISCUSSION
While considerable progress has been achieved towards
unraveling the process of nDNA repair, our current
understanding of mtDNA repair and mutagenesis is
much less detailed.
Here, we present evidence that the frequency of oxidative stress-induced base substitutions in mtDNA can be
61%
88%
83%
91%
100%
91%
18S
92%
89%
32%
97%
105%
96%
87%
548%
5mU
C 0h 6h
53%
Enz
Time
492%
98%
109%
298%
59%
100%
100%
Mito
%C
C
64%
Neutral
100%
C 0h 6h
19%
71%
24%
5mU
B
100%
Enz
Time
100%
%C
67%
Neutral
B
44%
100%
18S
68%
75%
62%
90%
74%
91%
80%
100%
18S
Figure 4. Effect of APE1 inhibition on mtDNA degradation in
response to oxidative stress. HeLa cells were treated with XO in the
presence of hypoxanthine and in the presence or absence of methoxyamine (MA). Cells were then either lysed immediately after treatment
(0 h) or allowed for repair (6 h) in the presence or absence of MA. Total
DNA was extracted and digested with BamHI, and (A), separated
under denaturing (alkaline) conditions, and subjected to Southern blotting with nuclear (18S) and mtDNA (Mito) probes. %C indicates percentage intact mtDNA as compared to untreated control. (B) The same
samples as in (A) were run under non-denaturing (TBE buffer) conditions. Annotation is the same as in (A). This blot shows both intact and
SSB-containing nDNA and mtDNA.
very low. This holds true for both human adenocarcinoma
and mouse embryonic fibroblast cell lines. This is also true
for oxidative stress induced by superoxide production
from mitochondrial complex I and by H2O2 treatment.
The finding that the combined frequency of H2O2induced strand breaks and abasic sites exceeds that of
base substitutions by at least a factor of 10 is rather unexpected and indicates that handling of these types of
mtDNA damage constitute an important aspect of the
maintenance of overall mtDNA integrity. There are two
reasons to believe that this observation is not likely to be
confounded by an inability to amplify mtDNA fragments
containing SSBs and abasic sites by PCR. First, in our
experiments H2O2 induced an average of 2.71 0.44
strand breaks/abasic sites per mtDNA molecule as measured immediately after the treatment (Figure 1E and F).
This means an average of one strand break/abasic site
per 6135 bp. Therefore, one can not expect a skewed
amplification to substantially affect quantitation of mutagenic base lesions as there is only 16% probability that the
2546 Nucleic Acids Research, 2009, Vol. 37, No. 8
A
Alkaline
MMS
Time
Mito
1mM
C 0h 6h
2mM
1mM
0h 6h C 0h 6h
0.2%
2%
0.7%
14%
100%
23%
ROS
HeLa MMS MA
50%
49%
86%
%C
100%
HeLa MMS
2mM
0h 6h
mtDNA
AP site
APE
B
14%
15%
21%
36%
100%
42%
64%
69%
86%
100%
18S
Neutral
MMS
Time
Mito
1mM
C 0h 6h
1mM
5%
94%
8%
96%
100%
2mM
0h 6h C 0h 6h
2mM
0h 6h
SSB
Degradation
Figure 6. Proposed interaction between mtDNA repair and degradation pathways. Glycosylase I and Glycosylase II, mono- and bi-functional DNA glycosylases. A bi-functional DNA glycosylase also
posseses an AP-lyase activity (makes an incision at an abasic site).
APE, apurinic/apyrimidinic endonuclease APE/Ref1. See ‘Discussion’
section for details.
78%
92%
75%
99%
100%
93%
101%
102%
100%
Glycosylase II
or
Glycosylase I + APE
DSB
18S
94%
Base damage
HeLa MMS MA
94%
93%
95%
96%
%C
100%
HeLa MMS
R
e
p
a
i
r
Figure 5. Effect of APE1 inhibition on mtDNA degradation in
response to alkylating damage. HeLa cells were treated with indicated
concentrations of MMS in the presence or absence of methoxyamine
(MA). Cells were then either lysed immediately after treatment (0 h) or
allowed for repair (6 h) in the presence or absence of MA. Total DNA
was extracted and digested with BamHI, and (A), separated under
denaturing (alkaline) conditions, and subjected to Southern blotting
with nuclear (18S) and mtDNA (Mito) probes. %C indicates percentage intact mtDNA as compared to untreated control. (B) The same
samples as in A were run under non-denaturing (TBE buffer) conditions. Annotation is the same as in (A). This blot shows both intact and
SSB-containing nDNA and mtDNA.
amplified 1000-bp fragment would contain a strand break
or abasic site. Second, if ROS-induced strand breaks were
to interfere with detection of point mutations by PCR
using DNA isolated immediately after treatment with
ROS, then one would expect an increase in the frequency
of base substitutions in samples which were allowed for
repair of strand breaks and abasic sites (thus eliminating
the amplification bias) as in the experiment where
HCT116 cells were treated repeatedly with H2O2
(Figure 1F). However, this increase was not observed
(Figure 1F). Importantly, in cells treated repeatedly with
H2O2 the initial very substantial oxidative stress may be
expected to be further amplified through a ‘vicious cycle’
thus enhancing mtDNA mutagenesis.
Our data provide evidence that both DNA repair and
degradation processes operate on oxidatively damaged
mtDNA (Figure 2A and B). The elimination of damaged
mtDNA is preceded by the accumulation of linear
mtDNA molecules, which may represent degradation
intermediates (Figure 2C). These intermediates, unlike
undamaged circular mtDNA molecules, are susceptible
to exonucleolitic degradation thus ensuring the specificity
of the process. mtDNA degradation can be induced by
a physiologically relevant enzyme system, and in this
case the amount of linear intermediates is proportional
to enzyme dose (Figure 3C). Inhibition of abasic site processing by APE/Ref-1 stimulated mtDNA degradation
following oxidative stress (Figure 4) suggesting that inability to repair mtDNA damage may be the signal for its
degradation. This notion was further supported by induction of mtDNA degradation by inhibition of BER at levels
of alkylation damage, which do not normally induce degradation (Figure 5). Importantly, inhibition of BER
enhanced degradation of mtDNA containing both oxidative and alkylation lesions suggesting that degradation
of mtDNA molecules, which failed to repair is a general
mechanism. The higher propensity of ROS to induce
mtDNA strand breaks and abasic sites, as compared to
mutagenic base lesions, provides a new mechanism
whereby mitochondria maintain the integrity of their
genetic information (Figure 6). According to this mechanism, under conditions of oxidative stress, the generation
of a single steady-state premutagenic lesion in mtDNA
is accompanied by generation of as many as 10 strand
breaks and abasic sites, which lead to elimination of
mtDNA molecules containing such premutagenic
lesions. Mitochondrial BER pathway components such
as lesion-specific DNA glycosylases may aid in generation
of abasic sites, which if left unrepaired, stimulate mtDNA
degradation (Figures 4 and 5). The formation of DSBs is
a commitment step, which may be mediated by stalled
DNA or RNA polymerase, or by another mechanism.
This model provides a mechanistic explanation for the
observations made by Suter and Richter (37) who have
found that 8-oxoG content of circular mtDNA is low
and does not increase in response to oxidative insult.
Nucleic Acids Research, 2009, Vol. 37, No. 8 2547
However, fragmented mtDNA had very high 8-oxoG content, which further increased after oxidative stress. This
model is consistent with observations of Yakes and van
Houten (38) who found higher incidence of polymeraseblocking lesions (i.e. SSBs and abasic sites) in mtDNA as
compared to nDNA. Observations of Ikeda and Ozaki
(39) who found that mitochondrial endonuclease G is
more active in vitro on oxidatively modified DNA compared to undamaged DNA provide a candidate, which
along with other yet unknown enzymes may be involved
in the degradation of oxidatively damaged mtDNA.
Finally, the notion of mtDNA’s higher susceptibility to
strand breaks in response to oxidative stress as a mechanism for the protection of the integrity of encoded genetic
information concurs with evolutionary theory. It suggests
that (in combination with the high redundancy of
mtDNA) this unique mechanism may have evolved in
response to mtDNA’s close proximity to the main cellular
source of ROS. Recent studies indicate that excessive
accumulation of mtDNA mutations in genetically altered
mice results in premature aging (40,41). However, levels of
mutations which accumulate in mtDNA naturally do not
appear to limit mammalian lifespan (42). These observations, together with mtDNA’s proximity to a major site
for the generation of potentially mutagenic ROS, suggest
the existence of very efficient mechanisms which prevent
excessive accumulation of ROS-induced mtDNA mutations. The degradation of oxidatively damaged mtDNA
described in this study may represent a component of
this machinery.
SUPPLEMENTARY DATA
Supplementary Data are available at NAR Online.
ACKNOWLEDGEMENTS
The authors are thankful to Bert Vogelstein and Kenneth
Kinzler for HCT116 cell line.
FUNDING
National Institutes of Health (grant numbers PO1
HL06629907, R21RR02396101 to M.A. and RO1
ES03456 to G.L.W.). Funding for open access charge:
R21RR02396101.
Conflict of interest statement. None declared.
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