Abstract

There is increasing evidence that human atherosclerosis is associated with damage to the DNA of both circulating cells, and cells of the vessel wall. Reactive oxygen species are the most likely agents inducing DNA damage in atherosclerosis. DNA damage produces a variety of responses, including cell senescence, apoptosis and DNA repair. This review summarises the evidence for DNA damage in atherosclerosis, the cellular responses to damage and the mechanisms of signalling DNA damage.

1. Introduction

Atherosclerotic plaques consist of an accumulation of vascular smooth muscle cells (VSMCs) (for full list of abbreviations, see Table 1) and inflammatory cells, together with lipid and extracellular matrix proteins–advanced plaques also show calcification. Although the plaque develops as a chronic inflammatory reaction, there is increasing evidence that DNA damage to cells within the lesion plays an important role in both atherogenesis and the behaviour of established lesions. DNA damage ranges from ‘macro’ damage, including deletions or additions of whole chromosomes or parts of chromosomes, to ‘micro’ damage, which includes DNA strand breaks, mutations of single bases, modified bases (including oxidation) or DNA adducts. This review outlines the evidence for DNA damage in atherosclerosis, the inducers of damage, the response to damage and repair pathways, the consequences of damage and potential therapies to prevent/reduce damage.

Table 1

List of abbreviations used

8-Oxo-G 8-Oxo-guanine, an oxidative modification of guanine in DNA 
9–1–1 Multiprotein complex consisting of Rad9–Rad1–Hus1 
A–T Ataxia–telangiectasia 
ATM Ataxia–telangiectasia mutated (protein) 
ATR ATM and Rad3-related (protein) 
ChK-1 Checkpoint kinase 1 
ChK-2 Checkpoint kinase 2 
DSBs Double-stranded (DNA) breaks 
γ-H2AX Phosphorylated form of the histone protein H2AX 
HMGCoA 3-Hydroxy-3-methylglutaryl coenzyme A 
MDM2 Murine double minute 2 gene 
MRN Multiprotein complex consisting of MRE11–RAD50–NBS1 
NBS1 Protein mutated in Nijmegen breakage syndrome 
PIKK Phosphatidylinositol 3-kinase 
ROS Reactive oxygen species 
SIPS Stress-induced premature senescence 
SSBs Single-stranded (DNA) breaks 
VSMC Vascular smooth muscle cell 
8-Oxo-G 8-Oxo-guanine, an oxidative modification of guanine in DNA 
9–1–1 Multiprotein complex consisting of Rad9–Rad1–Hus1 
A–T Ataxia–telangiectasia 
ATM Ataxia–telangiectasia mutated (protein) 
ATR ATM and Rad3-related (protein) 
ChK-1 Checkpoint kinase 1 
ChK-2 Checkpoint kinase 2 
DSBs Double-stranded (DNA) breaks 
γ-H2AX Phosphorylated form of the histone protein H2AX 
HMGCoA 3-Hydroxy-3-methylglutaryl coenzyme A 
MDM2 Murine double minute 2 gene 
MRN Multiprotein complex consisting of MRE11–RAD50–NBS1 
NBS1 Protein mutated in Nijmegen breakage syndrome 
PIKK Phosphatidylinositol 3-kinase 
ROS Reactive oxygen species 
SIPS Stress-induced premature senescence 
SSBs Single-stranded (DNA) breaks 
VSMC Vascular smooth muscle cell 

Evidence for DNA damage in atherosclerosis

DNA damage is present both in the circulation of patients with atherosclerosis and the plaques themselves. For example, patients with coronary artery disease have a higher micronucleus index (a marker of genetic instability) than healthy controls, which correlates with disease severity [1,2]. There is also a significantly higher incidence and extent of a common mitochondrial DNA deletion (mtDNA4977) [3]. Premature atherosclerosis is a feature of defects in DNA repair pathways, such as Werner syndrome, a disease characterised by predisposition to cancer and early onset of symptoms related to normal aging including osteoporosis, ocular cataracts, graying and loss of hair, diabetes mellitus, and atherosclerosis. Werner protein guards the genetic stability of cells, playing an integral role in base excision repair and at telomere ends [4].

There is also evidence of ‘macro’ DNA damage within atherosclerotic plaques. Cytogenetic analysis of primary cell cultures from human plaques identified loss of the Y chromosome and del13q14, XXY karyotype, and trisomy 7, 10 and 18 [5], whilst unstable carotid plaques demonstrated trisomy and tetrasomy of chromosome 7, and monosomy of chromosome 11 [6]. Although it was not clear from these studies which cells had chromosomal defects, VSMCs demonstrate microsatellite instability (mutations in microsatellite regions that may affect gene expression) [7,8] and loss of heterozygosity [8], suggesting that genomic destabilization/mutation may play a pivotal role in atherosclerosis. Indeed, monoclonality of VSMCs within lesions suggests that atherosclerosis may be characterised by a sub-population of VSMCs that undergo selective replication [9]. DNA damage can also occur in the mitochondria. Indeed, mitochondrial DNA damage correlates with the extent of atherosclerosis in human specimens and aortas from apolipoprotein E(− / −) mice [10].

This ‘macro’ DNA damage is also associated with biomarkers of carcinogen exposure–such as DNA adducts or modifications to specific bases in atherosclerotic plaques. ‘Bulky’ aromatic DNA-adducts in VSMCs (most likely related to environmental exposure to genotoxic chemicals) are a predictor of atherosclerosis extent in humans even after adjustment for age, smoking, obesity, heart weight and genetic susceptibility markers [11]. DNA strand breaks, oxidised pyrimidines and altered purines are also significantly higher in patients with coronary artery disease than controls [12], and human plaques show markers of oxidative damage, including DNA strand breaks, expression of 8-oxo-G (an oxidative modification of guanine residues in DNA) and activation of DNA repair enzymes [13]. Strong nuclear and cytoplasmic immunoreactivity for 8-oxo-G is detected in plaque VSMCs, macrophages and endothelial cells, but not in VSMCs of adjacent normal media or normal arteries [13]. DNA damage is also a direct correlate of extent of atherosclerosis in experimental animals. For example, cholesterol feeding of rabbits induces oxidative damage in plaques, manifested by 8-oxo-G staining [14], DNA strand breaks, and apoptosis.

Causes of DNA damage–risk factors for atherosclerosis

DNA can be damaged in numerous ways. Spontaneous damage due to replication errors, deamination, or depurination has to be repaired in addition to damage derived from oxidation and environmental chemicals. Many of the risk factors associated with atherogenesis, such as smoking and diabetes mellitus, may directly induce DNA damage. For example, smoking can cause oxidative DNA damage, inhibit DNA repair, and induce the production of advanced glycation end products, which themselves cause DNA mutation (reviewed in [15]). Similarly, advanced glycation end products have been implicated in the oxidation of low-density lipoprotein [16] and elevated levels of 8-oxo-G. Although direct damage from specific risk factors or environmental agents may contribute to DNA damage in atherosclerosis, the most likely triggers of damage are reactive oxygen species.

Cause of DNA damage in atherosclerosis–reactive oxygen species

It has been estimated that approximately 2 × 104 DNA damaging events occur in every cell/day [17]; a major portion of these occur via reactive oxygen species (ROS) [18]. ROS include the superoxide anion (*O2), hydrogen peroxide, hydroxyl radical, peroxynitrite and lipid peroxides (Fig. 1), with their reactivity and half-life varying according to species. ROS are constantly produced within the cell, in particular via mitochondrial oxidative metabolism and pathological processes such as inflammation [18]. In vascular cells, the primary source of ROS may be xanthine oxidase and NAD(P)H oxidases (Noxs) [19,20]. Other sources of ROS include cytochrome P450 isoenzymes, lipoxygenase, cyclooxygenase, hemoxygenase, and glucose oxidase. Myeloperoxidase is also produced by invading macrophages (reviewed in [21]). ROS are also secondary messengers in specific signalling pathways, generated by specific plasma membrane oxidases in response to growth factors and cytokines.

Fig. 1

Schema of generation of free radicals and protective enzymes within the cell.

Fig. 1

Schema of generation of free radicals and protective enzymes within the cell.

DNA lesions caused by ROS include double- and single-stranded breaks (DSBs and SSBs), DNA–DNA and DNA–protein cross links and base modifications, including thymine glycol, 8-hydroxyguanine and 8-oxo-guanine. Certain types of damage can be traced back to specific insults. Whilst superoxide and hydrogen peroxide are normally not reactive to DNA, both can be converted to the extremely reactive hydroxyl radical via the Fenton reaction. The hydroxyl radical can induce a vast array of damage to both nuclear and mitochondrial DNA [22]. Hydroxyl radicals may induce base or nucleotide loss, adducts, and single- and double-strand breaks. In particular, hydroxyl radicals can interact with pyrimidine double bonds causing glycolytic damage (e.g. thymine glycol, uracil glycol) that can cause mis-pairing, transcriptional interrupts and, if left un-repaired, induce cell cycle arrest. Similarly, interaction of OH radicals with purines will generate formamidopyrimidines and other purine oxidative products that are usually recognised by the DNA repair mechanism. Thymidine oxidation, or the addition of methyl and alkyl groups, is frequently attributed to ROS of all types. Interestingly, 8-oxo-G, which frequently causes oxidative damage to deoxyguanines, is not recognised by the DNA repair system and is thus highly deleterious to cells. Un-repaired 8-oxo-G will mis-pair with dA, leading to an increase in G to T transition mutations.

The cell has a number of mechanisms of protection against oxidative damage (reviewed in [23]), including direct interaction with antioxidants such as α-tocopherol, ascorbic acid and glutathione (scavengers). Glutathione exists in both its oxidised form (GSSG) and reduced (GSH) forms, and can function alone or in association with enzyme activities. In addition, antioxidant enzymes are present such as superoxide dismutases (which convert the superoxide radical to hydrogen peroxide and water), and glutathione S transferases that conjugate reduced GSH to hydrophobic organic compounds such as lipid peroxides. In general, the balance between ROS levels and the activity of these defence mechanisms determines the degree of oxidative stress encountered by the cell.

Evidence for oxidant stress in atherosclerosis

Increased levels of ROS are found in all layers of the atherosclerotic arterial wall, and particularly in the plaque itself [24]. Although ROS are associated with upregulation of several vascular NAD(PH) oxidase subunits and increased enzyme activity [25], confirmation of the role of ROS in atherosclerosis comes from studies with genetic disruption of the p47phox subunit of NAD(P)H oxidase, which causes dramatic inhibition of atherogenesis in ApoE− / − mice [26]. Increased levels of xanthine oxidase and reduced ecSOD are found in coronary arteries and plasma of patients with coronary artery disease [27]. Similarly, low levels of glutathione peroxidase activity are an independent risk factor for cardiovascular events in patients with coronary artery disease [28].

Oxidative damage to the mitochondrial genome with resultant mitochondrial dysfunction is also an important consequence of increased intracellular ROS. Hydrogen peroxide and peroxynitrite induce mitochondrial DNA damage and dysfunction in endothelial cells and VSMCs, with endothelial cells being particularly sensitive [29]. As mentioned above, mitochondrial DNA damage correlates with the extent of atherosclerosis in human specimens and aortas from ApoE(− / −) mice, whilst disruption of manganese superoxide dismutase, a mitochondrial antioxidant enzyme, show increased mitochondrial DNA damage and accelerated atherogenesis at arterial branch points [10].

The DNA damage response pathway

The plethora of DNA damage lesions in cells is rapidly detected by a complex web of signalling pathways called the DNA damage response pathway. This pathway involves three groups of evolutionarily conserved proteins, which act in concert to translate the DNA damage signal into a specific cellular response (Figs. 2 and 3). These are:

  1. 1.

    Sensors: recognise the lesions themselves or chromatin alterations that follow DNA damage. Examples include the MRN and 9–1–1 complexes.

    Fig. 3

    Signalling pathways for DNA repair after DNA damage. Abbreviations are as described in the text.

    Fig. 3

    Signalling pathways for DNA repair after DNA damage. Abbreviations are as described in the text.

    Fig. 2

    Outline of the DNA damage response pathway in mammalian cells.

    Fig. 2

    Outline of the DNA damage response pathway in mammalian cells.

  2. 2.

    Transducers: initiate a signal transduction cascade that propagate and amplify the signal. Examples include ATM and ATR.

  3. 3.

    Effectors: execute the specific cellular response. Examples include ChK1 and ChK2.

Sensors

MRE11–RAD50–NBS1 (MRN)

Double-stranded DNA breaks (DSBs) may cause chromosomal aberrations and disrupt genetic integrity, leading to carcinogenesis. DNA replication errors are the major cause of DSBs in dividing cells, although they may also arise from ionising radiation and genotoxic agents. The two mechanisms for DSB repair are homologous recombination between sister chromatids and the rapid, error prone non-homologous end joining [30]. The MRN complex plays a key role in the cellular response to DSBs [31] as well as other aspects of DNA metabolism such as telomere maintenance [32]. The MRE11 protein has an amino terminal phosphoesterase domain and two DNA binding sites. The phosphoesterase domain not only contains the NBS1 binding site, but also functions as both a single and double stranded DNA endonuclease as well as a 3′-5′ double strand exonuclease [33]. RAD50 contains a bipartite ATP binding cassette joined by two coiled-coil domains, with a Cys-X–X-Cys motif located in the middle of the coiled-coil domain, which serves as a dimerization domain [34]. NBS1 is essential for the nuclear transportation of the MRN complex and binding of phospho-H2AX, and is responsible for MRN recruitment to sites of DSBs [35].

The key role of the MRN complex in the DSB response has been emphasized by genetic studies. Hypomorphic mutations in MRE11 lead to ataxia–telangiectasia-like disease, which like ataxia–telangiectasia (A–T–see below), includes progressive cerebellar degeneration, hypersensitivity to ionising radiation and radioresistant DNA synthesis [36]. Hypomorphic mutations in NBS1 lead to the Nijmegen breakage syndrome (NBS), which is characterised by microcephaly, mental deficiency, immunodeficiency, hypersensitivity to irradiation, chromosomal instability and cancer predisposition. A variant of this disease is caused by RAD50 mutations.

The similarities noted amongst ataxia–telangiectasia, ataxia–telangiectasia-like disease, and NBS suggest a functional link between MRN and ATM. This link is complex and is explained by a two-step model in which NBS1 is firstly recruited to sites of DSB by binding to MRE11 in an ATM-dependent manner. ATM activation and phosphorylation of its downstream targets such as γ-H2AX are thought to assist in the recruitment and retention of further MRN molecules by NBS1–γ-H2AX interaction [37].

Rad9–Rad1–Hus1 (9–1–1)

The 9–1–1 complex is present in the nucleus of both damaged and undamaged cells. The complex resembles the proliferating nuclear antigen, a doughnut shaped homotrimeric complex, which forms a sliding clamp at sites of DNA replication and serves as a sliding platform for the accumulation of the replicative machinery [38]. The 9–1–1 complex is loaded around the DNA by the Rad17-replication factor C in response to genotoxic stresses such as ionising radiation, UV light and replication inhibitors [39]. The 9–1–1 complex is involved in the ATR/ChK1 signalling axis [40] and is loaded onto DNA following single strand DNA damage. The Rad9 tail, which contains a consensus phosphatidylinositol 3-kinase (PIKK) site, is then phosphorylated by ATR and in turn facilitates the phosphorylation and activation of ChK1 [41].

Transducers

The two main transducers of DNA damage in mammals are ATM (ataxia–telangiectasia gene product) and ATR (ATM and Rad3-related) [42]. Both are members of the phosphatidylinositol 3-kinase-like family of serine/threonine protein kinases (PIKK) [42]. ATM functions particularly after DSBs, ATR after SSBs and separate N-glycosylases function in base excision repair.

ATM

ATM deficiency leads to ataxia–telangiectasia (A–T), an autosomal recessive syndrome with early childhood onset of progressive cerebellar ataxia, occulocutaneous telangiectasia, primary immunodeficiency (decreased IgA, IgE, and IgG2), retarded somatic growth, premature ageing, and predisposition to lymphoreticular malignancy [43]. In addition, these patients exhibit chromosomal aberrations, cell cycle checkpoint defects and increased rates of telomere shortening [44,45]. Patients with two mutant alleles develop new cases of cancer at approximately 100 times the age-specific population rate and have a median age of death of approximately 20years.

Over 400 different ATM mutations have so far been identified of which more than 85% are null mutations resulting in no detectable ATM protein [46,47]. A–T heterozygotes comprise approximately 1.4–2% of the general population [48]. ATM carriers have an increased risk of cancer, but also of ischaemic heart disease [49,50], dying on average 7–8years earlier. Importantly, the effect of ATM carrier status on cardiovascular mortality is similar to that of smoking or male gender. Although this suggests that ATM carriage may account for a significant proportion of the genetic inheritance of coronary artery disease, the mechanism by which ATM heterozygosity promotes coronary artery disease is unknown.

ATM is located within the nucleus of most cells as dimers or higher order multimers with the kinase domain of each molecule being blocked by the other [51]. In response to DSBs, each ATM molecule phosphorylates the other on ser1981 leading to monomerisation and activation of ATM [51]. The active ATM molecules are then recruited to DSB sites, an event which is facilitated by the MRN complex, serving as a platform for further enzymatic reactions [52] as well as initiating phosphorylation events leading to regulation of specific cell cycle checkpoints [53]. The main target for ATM in this pathway is the tumour suppressor gene p53, which is phosphorylated on ser15[54–56], enhancing its transcriptional activity [57]. ATM also phosphorylates and activates ChK2, which in turn phosphorylates p53 on ser20[58], and interferes with the p53–MDM2 interaction. Finally, ATM directly phosphorylates MDM2 on ser395 thereby inhibiting the nuclear export of the p53–MDM2 complex and the subsequent degradation of p53 [59]. The ATM mediated modulation of p53 is an illustrative example of the numerous mechanisms by which ATM can regulate a single protein within the same pathway.

In addition to its critical role in cell cycle checkpoints, and maintenance of telomere length and integrity [60], A–T cells show abnormal responses to oxidant stress and increased oxidative stress is evident in A–T. For example, A–T fibroblasts are more sensitive to oxidant stress-induced DNA damage than normal cells [61], and have constitutive activation of DNA damage repair pathways that are reversed in part by antioxidants [62]. Oxidant stress can induce ATM kinase activity, which in normal cells is associated with cell cycle arrest in G1 and G2/M; indeed, ATM is required for growth arrest after oxidative stress [63]. In addition, tissues from ATM knockout mice show increased oxidative stress, including elevated levels of 3-nitrotyrosine [64], lipid peroxidation [64], and accumulation of superoxide and hydrogen peroxide [65]. Whilst the most striking effects are seen in the brain, patients with A–T show reduced total plasma antioxidant capacity, suggesting that they may sustain constant systemic oxidative stress [66].

ATR

ATM-mediated pathways are not completely abolished in A–T cells, but are rather dampened and turned on slowly. This indicates the presence of parallel kinases, which act in concert with ATM. One such system is ATR, which can phosphorylate ATM substrates albeit with much slower kinetics [67]. ATR responds to UV-light, stalled replication forks and hypoxia [53]. The recruitment of ATR to DNA lesions is facilitated by the ATR interacting protein. This is accompanied by phosphorylation of some ATM substrates such as p53 [68] as well as ATR-specific targets such as ChK1 and RAD17 [69]. Mice lacking ATR die in the early stages of embryogenesis and ATR deficiency is not compatible with cellular viability in culture [70]. This is because the ATR–ChK1 axis is fundamental in S phase, preventing the progression of damaged DNA and hence preventing mitotic catastrophe. Mutations causing partial loss of ATR activity in humans have been associated with the autosomal recessive disorder Seckel syndrome, which shares features in common with A–T [71].

Effectors

ChK1

The ChK1 protein consists of a kinase domain and an SQ domain, containing multiple phosphorylation sites. ChK1 is present in an active form during the S and G2 phases of the cell cycle even in the absence of DNA damage [72], but is activated in response to DNA damage and stalled replication forks by ATR, which phosphorylates ChK1 at ser317[72]. Mice lacking ChK1 die early in embryogenesis [73] and embryonic stem cells lacking ChK1 are not viable in culture [74]. In human cells, ChK1 is essential in Cdc25A metabolism as well as the checkpoint response to ionising radiation and DNA damaging agents [75].

ChK2

The ChK2 protein contains multiple ATM recognition sites and both dimerization and kinase domains. ChK2 is expressed throughout the cell cycle and is inactive in the absence of DNA damage [76]. Following DNA damage and ATM activation, ChK2 is phosphorylated at thr68 and forms a homodimer [77]. Mice deficient in ChK2 are viable, fertile and exhibit increased resistance to ionising radiation [74]. In humans, mutations in ChK2 are associated with a spectrum of malignancies including the Li Fraumeni syndrome [78], breast and colon cancer [79].

The ATM/ChK2 and ATR/ChK1 pathways are no longer considered as parallel branches of the DNA damage response pathway, but rather a system where there is a high degree of cross talk and connectivity. For instance, ATM can activate ChK1 in response to ionising radiation, and ChK2 can be activated independently of ATM [80].

Effects of DNA damage in atherosclerosis

DNA damage to cells of the vessel wall may have a number of effects. However, the most likely consequences are cell senescence, cell death and DNA repair.

Cell senescence

Extensive characterisation of lesion development has shown that cell proliferation is low in early atherosclerosis (Fatty streak, Stary Type I lesion), peaks in the intermediate lesions (Stary II–IV), and declines in advanced fibroproliferative, complicated plaques (Stary type V lesions) [81,82]. In vitro studies have confirmed that plaque VSMCs show reduced cell proliferation rates, increased population doubling times, and earlier failure to proliferate (senescence) than medial VSMCs [83].

Cell senescence may be triggered by two broadly different mechanisms. Replicative senescence may be induced by reduction of telomere length, changes in structure such as telomeric fusion or dicentrics, or loss of telomere-bound factors. Telomeres may trigger growth arrest via DNA damage responses at critical telomere lengths or structure. Cells subjected to sub-lethal stress due to DNA damage (UV and γ-irradiation, oxidative stress and treatment with histone deacetylase inhibitors) also undergo ‘stress-induced premature senescence’ (SIPS). SIPS resembles replicative senescence, cells demonstrating a similar morphology and pattern of cell cycle regulators.

ROS can induce both telomere-based senescence and SIPS. ROS induce DNA strand breaks, and base and nucleotide modifications, particularly in sequences with high guanosine content [84], such as telomeres. Indeed, telomeres are one of the DNA structures most sensitive to oxidative damage [85] and ROS can accelerate telomere loss in vitro [86]. Increased telomere loss/division can also occur in individual cells due to a telomere-specific deficiency in base excision repair, leading to preferential accumulation of ROS-induced single-stranded DNA breaks [85] preventing replication of distal telomeres when cells divide. Thus, it is possible that oxidative DNA damage to telomeres induces a DNA repair response that induces senescence. Not only do ROS induce senescence, but also senescent cells produce high levels of ROS, and contain higher levels of oxidatively damaged DNA [87]. In vitro, VSMCs from aged mice have decreased proliferation, yet generated higher levels of ROS in comparison with cells from younger mice, associated with decreased endogenous antioxidant activity, increased lipid peroxidation, and mitochondrial DNA damage [88].

Cell death

In cells where the DNA damage is too much to repair, or in cells that are driven to proliferate, DNA damage induces apoptosis. Apoptosis is evident in endothelial cells, macrophages and VSMCs in atherosclerotic plaques. Endothelial cell death is implicated in both atherogenesis and plaque erosion [89], whereas VSMC death may promote thinning of the fibrous cap and plaque rupture. Indeed, increased levels of VSMC apoptosis are seen in mature plaques when compared to control vessels [90], and in unstable versus stable plaques or patients [91,92]. Cell death can be induced by ROS and oxidised lipids in vascular cells, whereas p53 expression in an experimental neointima can induce apoptosis and promote plaque rupture [93].

DNA repair

Although there is evidence of activation of DNA repair pathways in human atherosclerosis, cells displaying DNA damage are much more frequent than cells displaying markers of apoptosis, implying that repair occurs to prevent death [13]. In human plaques markers of DNA damage are associated with activation of base excision repair or nonspecific repair pathways [13]. In cholesterol-fed rabbits plaques also manifest DNA damage associated with upregulation of DNA repair enzymes [14]. In these studies, DNA strand breaks normalized after 4weeks of dietary lipid lowering, but a significant reduction of 8-oxo-G immunoreactivity was only observed after a prolonged period of lipid lowering, emphasizing the longevity of DNA damage in atherosclerosis. In addition, repair pathways started to decline progressively when cholesterol-fed animals were placed on a normal diet [14]. These studies demonstrate that DNA damage and activation of repair pathways occur in atherosclerosis, and are also reversible, at least in the early stages of atherosclerosis.

Prevention/treatment of DNA damage in atherosclerosis

The presence and biological consequences of DNA damage in atherosclerosis mean that both prevention and reversal of damage are therapeutic aims. In vitro, antioxidants can ameliorate ROS-induced DNA damage, although antioxidant trials in humans have been disappointing. In contrast, cholesterol lowering by diet is associated with a reduction in DNA damage and markers of DNA repair, at least in animal models. Drugs that have been proven to alter plaque progression and patient events have also been shown to alter vascular oxidative stress. In particular, HMGCoA reductase inhibitors (‘Statins’) reduce NAD(P)H oxidase activation [94,95] and superoxide production in vitro, in part by inhibiting the membrane translocation (and thus activity) of the small GTP-binding protein Rac-1 [96,97], a regulatory component of vascular NAD(P)H oxidase. Statins can also reduce superoxide production, and mRNA expression of specific nox subunits in vivo [96,98]. This effect may underlie the observation that atorvastatin reduces the degree of DNA damage of peripheral lymphocytes as well levels of oxidant stress in hyperlipidaemic patients [99].

Conclusions

DNA damage is increasingly recognised as being present in all cells within the atherosclerotic plaque. DNA damage may promote atherogenesis, and in advanced lesions induce phenotypic changes such as cell senescence and cell death that promote unstable plaques. Knowledge of both the triggers of DNA damage and the pathways underlying the DNA damage responses should lead to both prevention and treatment of DNA damage in atherosclerosis.

Acknowledgements

MM, JM and MB are supported by British Heart Foundation Grants FS/05/008 and RG/04/001.

References

[1]
Botto
N.
Rizza
A.
Colombo
M.
Mazzone
A.
Manfredi
S.
Masetti
S.
et al
Evidence for DNA damage in patients with coronary artery disease
Mutat Res
 
2001
493
23
30
[2]
Andreassi
M.G.
Botto
N.
Cocci
F.
Battaglia
D.
Antonioli
E.
Masetti
S.
et al
Methylenetetrahydrofolate reductase gene C677T polymorphism, homocysteine, vitamin B12, and DNA damage in coronary artery disease
Hum Genet
 
2003
112
171
177
[3]
Botto
N.
Berti
S.
Manfredi
S.
Al-Jabri
A.
Federici
C.
Clerico
A.
et al
Detection of mtDNA with 4977bp deletion in blood cells and atherosclerotic lesions of patients with coronary artery disease
Mutat Res
 
2005
570
81
88
[4]
Lee
J.W.
Harrigan
J.
Opresko
P.L.
Bohr
V.A.
Pathways and functions of the Werner syndrome protein
Mech Ageing Dev
 
2005
126
79
86
[5]
Casalone
R.
Granata
P.
Minelli
E.
Portentoso
P.
Giudici
A.
Righi
R.
et al
Cytogenetic analysis reveals clonal proliferation of smooth muscle cells in atherosclerotic plaques
Hum Genet
 
1991
87
139
143
[6]
Matturri
L.
Cazzullo
A.
Turconi
P.
Lavezzi
A.
Vandone
P.
Gabrielli
L.
et al
Chromosomal alterations in atherosclerotic plaques
Atherosclerosis
 
2001
15
755
761
[7]
McCaffrey
T.A.
Du
B.H.
Consigli
S.
Szabo
P.
Bray
P.J.
Hartner
L.
et al
Genomic instability in the type II TGF-beta 1 receptor gene in atherosclerotic and restenotic vascular cells
J Clin Invest
 
1997
100
2182
2188
[8]
Hatzistamou
J.
Kiaris
H.
Ergazaki
M.
Spandidos
D.A.
Loss of heterozygosity and microsatellite instability in human atherosclerotic plaques
Biochem Biophys Res Commun
 
1996
225
186
190
[9]
Chung
I.M.
Schwartz
S.M.
Murry
C.E.
Clonal architecture of normal and atherosclerotic aorta–implications for atherogenesis and vascular development
Am J Pathol
 
1998
152
913
923
[10]
Ballinger
S.W.
Patterson
C.
Knight-Lozano
C.A.
Burow
D.L.
Conklin
C.A.
Hu
Z.
et al
Mitochondrial integrity and function in atherogenesis
Circulation
 
2002
106
544
549
[11]
Binkova
B.
Smerhovsky
Z.
Strejc
P.
Boubelik
O.
Stavkova
Z.
Chvatalova
I.
et al
DNA-adducts and atherosclerosis: a study of accidental and sudden death males in the Czech Republic
Mutat Res
 
2002
501
115
128
[12]
Botto
N.
Masetti
S.
Petrozzi
L.
Vassalle
C.
Manfredi
S.
Biagini
A.
et al
Elevated levels of oxidative DNA damage in patients with coronary artery disease
Coron Artery Dis
 
2002
13
269
274
[13]
Martinet
W.
Knaapen
M.W.
De Meyer
G.R.
Herman
A.G.
Kockx
M.M.
Elevated levels of oxidative DNA damage and DNA repair enzymes in human atherosclerotic plaques
Circulation
 
2002
106
927
932
[14]
Martinet
W.
Knaapen
M.
De Meyer
G.
Herman
A.
Kockx
M.
Oxidative DNA damage and repair in experimental atherosclerosis are reversed by dietary lipid lowering
Circ Res
 
2001
88
733
739
[15]
Basta
G.
Schmidt
A.M.
De Caterina
R.
Advanced glycation end products and vascular inflammation: implications for accelerated atherosclerosis in diabetes
Cardiovasc Res
 
2004
63
582
592
[16]
Bucala
R.
Makita
Z.
Koschinsky
T.
Cerami
A.
Vlassara
H.
Lipid advanced glycosylation: pathway for lipid oxidation in vivo
Proc Natl Acad Sci U S A
 
1993
90
6434
6438
[17]
Ames
B.
Shigenaga
K.
Oxidative stress is a major contribution to aging
Ann NY Acad Sci
 
1992
663
85
96
[18]
Williams
G.M.
Jeffrey
A.M.
Oxidative DNA damage: endogenous and chemically induced
Regul Toxicol Pharmacol
 
2000
32
283
292
[19]
Mohazzab
K.M.
Kaminski
P.M.
Wolin
M.S.
NADH oxidoreductase is a major source of superoxide anion in bovine coronary artery endothelium
Am J Physiol
 
1994
266
H2568
H2572
[20]
Griendling
K.K.
Minieri
C.A.
Ollerenshaw
J.D.
Alexander
R.W.
Angiotensin II stimulates NADH and NADPH oxidase activity in cultured vascular smooth muscle cells
Circ Res
 
1994
74
1141
1148
[21]
Wassmann
S.
Wassmann
K.
Nickenig
G.
Modulation of oxidant and antioxidant enzyme expression and function in vascular cells
Hypertension
 
2004
44
381
386
[22]
Michalik
V.
Spotheim Maurizot
M.
Charlier
M.
Calculation of hydroxyl radical attack on different forms of DNA
J Biomol Struct Dyn
 
1995
13
565
575
[23]
Offord
E.
van Poppel
G.
Tyrrell
R.
Markers of oxidative damage and antioxidant protection: current status and relevance to disease
Free Radic Res
 
2000
33
S5
S19
Suppl
[24]
Warnholtz
A.
Nickenig
G.
Schulz
E.
Macharzina
R.
Brasen
J.H.
Skatchkov
M.
et al
Increased NADH-oxidase-mediated superoxide production in the early stages of atherosclerosis: evidence for involvement of the renin–angiotensin system
Circulation
 
1999
99
2027
2033
[25]
Sorescu
D.
Weiss
D.
Lassegue
B.
Clempus
R.E.
Szocs
K.
Sorescu
G.P.
et al
Superoxide production and expression of nox family proteins in human atherosclerosis
Circulation
 
2002
105
1429
1435
[26]
Barry-Lane
P.A.
Patterson
C.
van der Merwe
M.
Hu
Z.
Holland
S.M.
Yeh
E.T.
et al
p47phox is required for atherosclerotic lesion progression in ApoE(− / −) mice
J Clin Invest
 
2001
108
1513
1522
[27]
Landmesser
U.
Merten
R.
Spiekermann
S.
Buttner
K.
Drexler
H.
Hornig
B.
Vascular extracellular superoxide dismutase activity in patients with coronary artery disease: relation to endothelium-dependent vasodilation
Circulation
 
2000
101
2264
2270
[28]
Blankenberg
S.
Rupprecht
H.J.
Bickel
C.
Torzewski
M.
Hafner
G.
Tiret
L.
et al
Glutathione peroxidase 1 activity and cardiovascular events in patients with coronary artery disease
N Engl J Med
 
2003
349
1605
1613
[29]
Ballinger
S.
Patterson
C.
Yan
C.
Doan
R.
Burow
D.
Young
C.
et al
Hydrogen peroxide and peroxynitrite induced mitochondrial DNA damage and dysfunction in vascular endothelial and smooth muscle cells
Circ Res
 
2000
86
960
966
[30]
Barnes
D.
Non-homologous end joining as a mechanism of DNA repair
Curr Biol
 
2001
11
455
457
[31]
Mirzoeva
O.
Petrini
J.
DNA replication-dependent nuclear dynamics of the Mre11 complex
Mol Cancer Res
 
2003
1
207
218
[32]
Lundblad
V.
Telomere maintenance without telomerase
Oncogene
 
2002
21
522
531
[33]
D'Amours
D.
Jackson
S.
The Mre11 complex: at the crossroads of DNA repair and checkpoint signalling
Nat Rev Mol Cell Biol
 
2002
3
317
327
[34]
de Jager
M.
van Noort
J.
van Gent
D.
Dekker
C.
Kanaar
R.
Wyman
C.
Human Rad50/Mre11 is a flexible complex that can tether DNA ends
Mol Cell
 
2001
8
1129
1135
[35]
Kobayashi
J.
Tauchi
H.
Sakamoto
S.
Nakamura
A.
Morishima
K.
Matsuura
S.
et al
NBS1 localizes to gamma-H2AX foci through interaction with the FHA/BRCT domain
Curr Biol
 
2002
12
1846
1851
[36]
Stewart
G.S.
Maser
R.S.
Stankovic
T.
Bressan
D.A.
Kaplan
M.I.
Jaspers
N.G.
et al
The DNA double-strand break repair gene hMRE11 is mutated in individuals with an ataxia–telangiectasia-like disorder
Cell
 
1999
99
577
587
[37]
Kobayashi
J.
Antoccia
A.
Tauchi
H.
Matsuura
S.
Komatsu
K.
NBS1 and its functional role in the DNA damage response
DNA Repair (Amst)
 
2004
3
855
861
[38]
Venclovas
C.
Thelen
M.P.
Structure-based predictions of Rad1, Rad9, Hus1 and Rad17 participation in sliding clamp and clamp-loading complexes
Nucleic Acids Res
 
2000
28
2481
2493
[39]
Bermudez
V.P.
Lindsey-Boltz
L.A.
Cesare
A.J.
Maniwa
Y.
Griffith
J.D.
Hurwitz
J.
et al
Loading of the human 9–1–1 checkpoint complex onto DNA by the checkpoint clamp loader hRad17-replication factor C complex in vitro
Proc Natl Acad Sci U S A
 
2003
100
1633
1638
[40]
Roos-Mattjus
P.
Hopkins
K.M.
Oestreich
A.J.
Vroman
B.T.
Johnson
K.L.
Naylor
S.
et al
Phosphorylation of human Rad9 is required for genotoxin-activated checkpoint signaling
J Biol Chem
 
2003
278
24428
24437
[41]
Roos-Mattjus
P.
Vroman
B.T.
Burtelow
M.A.
Rauen
M.
Eapen
A.K.
Karnitz
L.M.
Genotoxin-induced Rad9–Hus1–Rad1 (9–1–1) chromatin association is an early checkpoint signaling event
J Biol Chem
 
2002
277
43809
43812
[42]
Nyberg
K.A.
Michelson
R.J.
Putnam
C.W.
Weinert
T.A.
Toward maintaining the genome: DNA damage and replication checkpoints
Annu Rev Genet
 
2002
36
617
656
[43]
Swift
M.
Heim
R.A.
Lench
N.J.
Genetic aspects of ataxia–telangiectasia
Adv Neurol
 
1993
61
115
125
[44]
Metcalfe
J.A.
Parkhill
J.
Campbell
L.
Stacey
M.
Biggs
P.
Byrd
P.J.
et al
Accelerated telomere shortening in ataxia–telangiectasia
Nat Genet
 
1996
13
350
353
[45]
Young
B.
Painter
R.
Radioresistant DNA synthesis and human genetic diseases
Hum Genet
 
1989
82
113
117
[46]
Gilad
S.
Chessa
L.
Khosravi
R.
Russell
P.
Galanty
Y.
Piane
M.
et al
Genotype–phenotype relationships in ataxia–telangiectasia and variants
Am J Hum Genet
 
1998
62
551
561
[47]
Becker-Cantania
S.
Chen
G.
Hwang
M.
Wang
Z.
Sun
X.
Sanal
O.
et al
Ataxia–telangiectasia: phenotype/genotype studies of ATM protein expression, mutations, and radiosensitivity
Mol Genet Metab
 
2000
70
122
133
[48]
Swift
M.
Morrell
D.
Cromartie
E.
Chamberlin
A.
Skolnick
M.
Bishop
D.
The incidence and gene frequency of ataxia–telangiectasia in the United States
Am J Hum Genet
 
1986
39
573
583
[49]
Swift
M.
Chase
C.
Cancer and cardiac deaths in obligatory ataxia–telangiectasia heterozygotes
Lancet
 
1983
1
1049
1050
[50]
Su
Y.
Swift
M.
Mortality rates among carriers of ataxia–telangiectasia mutant alleles
Ann Intern Med
 
2000
133
770
778
[51]
Bakkenist
C.
Kastan
M.
DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation
Nature
 
2003
421
499
506
[52]
Andegeko
Y.
Moyal
L.
Mittelman
L.
Tsarfaty
I.
Shiloh
Y.
Rotman
G.
Nuclear retention of ATM at sites of DNA double strand breaks
J Biol Chem
 
2001
276
38224
38230
[53]
Abraham
R.
Cell cycle checkpoint signaling through the ATM and ATR kinases
Genes Dev
 
2001
15
2177
2196
[54]
Banin
S.
Moyal
L.
Shieh
S.
Taya
Y.
Anderson
C.
Chessa
L.
et al
Enhanced phosphorylation of p53 by ATM in response to DNA damage
Science
 
1998
281
1674
1677
[55]
Canman
C.
Lim
D.
Cimprich
K.
Taya
Y.
Tamai
K.
Sakaguchi
K.
et al
Activation of the ATM kinase by ionising radiation and phosphorylation of p53
Science
 
1998
281
1677
1679
[56]
Khanna
K.
Keating
K.
Kozlov
S.
Scott
S.
Gatei
M.
Hobson
K.
et al
ATM associates with and phosphorylates p53: mapping the region of interaction
Nat Genet
 
1998
20
398
400
[57]
Dumaz
N.
MeeK
D.
Serine 15 phosphorylation stimulates p53 transactivation but does not directly influence interaction with HDM2
EMBO J
 
1999
18
7002
7010
[58]
Bartek
J.
Falck
J.
Lukas
J.
ChK2 kinase: a busy messenger
Nat Rev Mol Biol
 
2001
2
877
886
[59]
Maya
R.
Balass
M.
Kim
S.
Shkedy
D.
Leal
J.
Shifman
O.
et al
ATM-dependent phosphorylation of Mdm2 on serine 395: role in p53 activation by DNA damage
Genes Dev
 
2001
15
1067
1077
[60]
Pandita
T.
The role of ATM in telomere structure and function
Radiat Res
 
2001
156
642
647
[61]
Rosin
M.
Anderson
C.
Response of fibroblast cultures from ataxia–telangiectasia patients to oxidative stress
Cancer Lett
 
1990
54
43
50
[62]
Gatei
M.
Shkedy
D.
Khanna
K.K.
Uziel
T.
Shiloh
Y.
Pandita
T.K.
et al
Ataxia–telangiectasia: chronic activation of damage-responsive functions is reduced by alpha-lipoic acid
Oncogene
 
2001
20
289
294
[63]
Shackelford
R.E.
Innes
C.L.
Sieber
S.O.
Heinloth
A.N.
Leadon
S.A.
Paules
R.S.
The ataxia–telangiectasia gene product is required for oxidative stress-induced G1 and G2 checkpoint function in human fibroblasts
J Biol Chem
 
2001
276
21951
21959
[64]
Barlow
C.
Dennery
P.A.
Shigenaga
M.K.
Smith
M.A.
Morrow
J.D.
Roberts
L.J.
2nd
et al
Loss of the ataxia–telangiectasia gene product causes oxidative damage in target organs
Proc Natl Acad Sci U S A
 
1999
96
9915
9919
[65]
Quick
K.L.
Dugan
L.L.
Superoxide stress identifies neurons at risk in a model of ataxia–telangiectasia
Ann Neurol
 
2001
49
627
635
[66]
Reichenbach
J.
Schubert
R.
Schwan
C.
Muller
K.
Bohles
H.J.
Zielen
S.
Anti-oxidative capacity in patients with ataxia–telangiectasia
Clin Exp Immunol
 
1999
117
535
539
[67]
Tibbetts
R.
Brumbaugh
K.
Williams
J.
Sarkaria
J.
Cliby
W.
Shieh
S.
et al
A role for ATR in the DNA damage-induced phosphorylation of p53
Genes Dev
 
1999
13
152
157
[68]
Hammond
E.
Denko
N.
Dorie
M.
Abraham
R.
Giaccia
A.
Hypoxia links ATR and p53 through replication arrest
Mol Cell Biol
 
2002
22
1834
1843
[69]
Heffernan
T.
Simpson
D.
Frank
A.
Heiloth
A.
Pauls
R.
Cordeiro-Stone
M.
et al
An ATR and ChK1 dependent S checkpoint inhibits replicon initiation following UVC-induced DNA damage
Mol Cell Biol
 
2002
22
8552
8561
[70]
Brown
E.
Baltimore
D.
ATR disruption leads to chromosomal fragmentation and early embryonic lethality
Genes Dev
 
2000
14
397
402
[71]
O'Driscoll
M.
Ruiz-Perez
V.
Woods
C.
Jeggo
P.
Goodship
J.
A splicing mutation affecting expression of ataxia–telangiectasia and Rad3-related protein (ATR) results in Seckel syndrome
Nat Genet
 
2003
33
497
501
[72]
Sorensen
C.
Syljuasen
R.
Falck
J.
Schroeder
T.
Ronnstrand
L.
Zhou
B.
et al
ChK1 regulates the S-phase checkpoint by coupling the physiological turnover of ionising radiation-induced accelerated proteolysis of Cdc25A
Cancer Cell
 
2003
3
247
258
[73]
Takai
H.
Tominaga
K.
Motoyama
N.
Miamishima
Y.
Nagahama
H.
Tsukiyama
T.
et al
Aberrant cell cycle checkpoint function and early embryonic death in ChK1(− / −) mice
Genes Dev
 
2000
14
1439
1447
[74]
Zhao
H.
Watkins
J.
Piwnica-Worms
H.
Disruption of the checkpoint kinase 1/cell division cycle 25A pathway abrogates ionising radiation-induced S and G2 checkpoint
Proc. Natl. Acad. Sci
 
2002
99
14795
14800
[75]
Gatei
M.
Sloper
K.
Sorensen
C.
Syljuasen
R.
Falk
J.
Hobson
J.
et al
ATM and DBS1 dependent phosphorylation of ChK1 on S317 in response to IR
J Biol Chem
 
2003
278
14806
14811
[76]
Lukas
C.
Bartkova
J.
Latella
L.
Falck
J.
Mainland
N.
Schroeder
T.
et al
DNA damage activated kinase ChK2 is independent of proliferation or differentiation yet correlates with tissue biology
Cancer Res
 
2001
61
4990
4993
[77]
Xu
X.
Tsvetkov
L.M.
Stern
D.F.
Chk2 activation and phosphorylation-dependent oligomerization
Mol Cell Biol
 
2002
22
4419
4432
[78]
Vahteristo
P.
Tamminen
A.
Karvinen
P.
Eerola
H.
Eklund
C.
Aaltonen
L.A.
et al
p53, CHK2, and CHK1 genes in Finnish families with Li-Fraumeni syndrome: further evidence of CHK2 in inherited cancer predisposition
Cancer Res
 
2001
61
5718
5722
[79]
Schutte
M.
Seal
S.
Barfoot
R.
Meijers-Heijboer
H.
Wasielewski
M.
Evans
D.G.
et al
Variants in CHEK2 other than 1100delC do not make a major contribution to breast cancer susceptibility
Am J Hum Genet
 
2003
72
1023
1028
[80]
Hirao
A.
Cheung
A.
Duncan
G.
Girard
P.M.
Elia
A.J.
Wakeham
A.
et al
Chk2 is a tumor suppressor that regulates apoptosis in both an ataxia–telangiectasia mutated (ATM)-dependent and an ATM-independent manner
Mol Cell Biol
 
2002
22
6521
6532
[81]
Gordon
D.
Reidy
M.A.
Benditt
E.P.
Schwartz
S.M.
Cell proliferation in human coronary arteries
Proc Natl Acad Sci U S A
 
1990
87
4600
4604
[82]
Lutgens
E.
de Muinck
E.D.
Kitslaar
P.J.
Tordoir
J.H.
Wellens
H.J.
Daemen
M.J.
Biphasic pattern of cell turnover characterizes the progression from fatty streaks to ruptured human atherosclerotic plaques
Cardiovasc Res
 
1999
41
473
479
[83]
Bennett
M.R.
Evan
G.I.
Schwartz
S.M.
Apoptosis of human vascular smooth muscle cells derived from normal vessels and coronary atherosclerotic plaques
J Clin Invest
 
1995
95
2266
2274
[84]
Burney
S.
Niles
J.C.
Dedon
P.C.
Tannenbaum
S.R.
DNA damage in deoxynucleosides and oligonucleotides treated with peroxynitrite
Chem Res Toxicol
 
1999
12
513
520
[85]
Petersen
S.
Saretzki
G.
von Zglinicki
T.
Preferential accumulation of single-stranded regions in telomeres of human fibroblasts
Exp Cell Res
 
1998
239
152
160
[86]
von Zglinicki
T.
Saretzki
G.
Docke
W.
Lotze
C.
Mild hyperoxia shortens telomeres and inhibits proliferation of fibroblasts: a model for senescence?
Exp Cell Res
 
1995
220
186
193
[87]
Chen
Q.
Fischer
A.
Reagan
J.D.
Yan
L.J.
Ames
B.N.
Oxidative DNA damage and senescence of human diploid fibroblast cells
Proc Natl Acad Sci U S A
 
1995
92
4337
4341
[88]
Moon
S.
Thompson
L.
Madamanchi
N.
Ballinger
S.
Papaconstantinou
J.
horaist
C.
et al
Aging, oxidative responses, and proliferative capacity in cultured mouse aortic smooth muscle cells
Am J Physiol, Heart Circ Physiol
 
2001
280
H2779
H2788
[89]
Durand
E.
Scoazec
A.
Lafont
A.
Boddaert
J.
Al Hajzen
A.
Addad
F.
et al
In vivo induction of endothelial apoptosis leads to vessel thrombosis and endothelial denudation: a clue to the understanding of the mechanisms of thrombotic plaque erosion
Circulation
 
2004
109
2503
2506
[90]
Geng
Y.
Libby
P.
Evidence for apoptosis in advanced human atheroma: colocalization with interleukin-1β converting enzyme
Am J Pathol
 
1995
147
251
266
[91]
Bauriedel
G.
Hutter
R.
Welsch
U.
Bach
R.
Sievert
H.
Luderitz
B.
Role of smooth muscle cell death in advanced coronary primary lesions: implications for plaque instability
Cardiovasc Res
 
1999
41
480
488
[92]
Rossi
M.L.
Marziliano
N.
Merlini
P.A.
Bramucci
E.
Canosi
U.
Belli
G.
et al
Different quantitative apoptotic traits in coronary atherosclerotic plaques from patients with stable angina pectoris and acute coronary syndromes
Circulation
 
2004
110
1767
1773
[93]
von der Thusen
J.H.
van Vlijmen
B.J.
Hoeben
R.C.
Kockx
M.M.
Havekes
L.M.
van Berkel
T.J.
et al
Induction of atherosclerotic plaque rupture in apolipoprotein E − / − mice after adenovirus-mediated transfer of p53
Circulation
 
2002
105
2064
2070
[94]
Wassmann
S.
Laufs
U.
Baumer
A.T.
Muller
K.
Ahlbory
K.
Linz
W.
et al
HMG-CoA reductase inhibitors improve endothelial dysfunction in normocholesterolemic hypertension via reduced production of reactive oxygen species
Hypertension
 
2001
37
1450
1457
[95]
Vecchione
C.
Brandes
R.P.
Withdrawal of 3-hydroxy-3-methylglutaryl coenzyme A reductase inhibitors elicits oxidative stress and induces endothelial dysfunction in mice
Circ Res
 
2002
91
173
179
[96]
Wassmann
S.
Laufs
U.
Muller
K.
Konkol
C.
Ahlbory
K.
Baumer
A.T.
et al
Cellular antioxidant effects of atorvastatin in vitro and in vivo
Arterioscler Thromb Vasc Biol
 
2002
22
300
305
[97]
Tsubouchi
H.
Inoguchi
T.
Sonta
T.
Sato
N.
Sekiguchi
N.
Kobayashi
K.
et al
Statin attenuates high glucose-induced and diabetes-induced oxidative stress in vitro and in vivo evaluated by electron spin resonance measurement
Free Radic Biol Med
 
2005
39
444
452
[98]
Takayama
T.
Wada
A.
Tsutamoto
T.
Ohnishi
M.
Fujii
M.
Isono
T.
et al
Contribution of vascular NAD(P)H oxidase to endothelial dysfunction in heart failure and the therapeutic effects of HMG-CoA reductase inhibitor
Circ J
 
2004
68
1067
1075
[99]
Harangi
M.
Seres
I.
Varga
Z.
Emri
G.
Szilvassy
Z.
Paragh
G.
et al
Atorvastatin effect on high-density lipoprotein-associated paraoxonase activity and oxidative DNA damage
Eur J Clin Pharmacol
 
2004
60
685
691

Author notes

Time for primary review 11 days